Characterized by suppuration and frank pus formation, tissue destruction during anaerobic infections may be extensive and gangrenous. Gas may form in infected tissues and septic thrombophlebitis may become manifest. Members of the B. fragilis (Table 4) group are the most commonly recovered and the most resistant of all anaerobic pathogens to antimicrobial therapy. B. fragilis is the most frequently encountered and most pathogenic of the group, but others including B. thetaiotaomicron, are also commonly seen and are typically more resistant to the antibiotics generally used for the treatment of anaerobic infections than is B. fragilis. The pigmented Porphyromonas and Prevotella spp. are recovered infrequently in pure culture and are typically more fastidious in their in-vitro growth requirements than are members of the B. fragilis group. The Prevotella spp. are important contributors to infection, particularly following insult to the oropharyngeal or female genital tract mucosal surfaces. One recently described organism, Sutterella wadsworthensis, recovered from mixed intra-abdominal infections, has demonstrated significant resistance to a number of antibiotics with recognized activity against anaerobes including metronidazole, clindamycin, cefotetan, ceftizoxime, piperacillin, piperacillin/tazobactam, and trova- floxacin . This organism is also apparently more virulent than the related species B. ureolyticus and Campylobacter gracilis.
Of the non-spore-forming anaerobes, Fusobacterium necrophorum is without question the most virulent. Despite typical susceptibility to any of a number of antimicrobial agents, the organism often produces overwhelming bacteraemia with metastatic infections. Lemierre's syndrome (necrobacillosis or postanginal anaerobic septicemia) is an acute oropharyngeal infection with secondary septic thrombophlebitis of the internal jugular vein, frequently leading to metastatic infections that are caused by this pathogen. The disease characteristically occurs in young healthy adults. One recent report suggests an association between the syndrome and Epstein–Barr virus-induced acute mononucleosis .
Another recently described Gram-negative anaerobic bacillus, Bilophila wadsworthia, is seen in approximately half of all cases of gangrenous or perforated appendicitis. This virulent species is nutritionally fastidious and may require up to 1 week to grow in culture. Almost always β-lactamase-positive, the organism has also been reported in cases of liver abscess, pericarditis, purulent arthritis, empyema, bacteraemia and soft tissue infection [23,24].
Extremely virulent, Cl. perfringens is the most commonly isolated of the clostridial species. Cl. tertium and Cl. septicum are of importance because of their association with lower gastrointestinal malignancy, particularly the cecum, and their role in the enterocolitis that sometimes strikes neutropenic individuals. Among the anaerobic cocci , Peptostreptococcus magnus appears to be the most pathogenic .
ANTIMICROBIAL SUSCEPTIBILITY TESTING OF ANAEROBES
To manage individual patients, to determine the activity of new compounds, and to monitor resistance trends locally, nationally, or internationally, antibiotic susceptibility testing of anaerobic bacteria is of significant value. When used for guiding therapy in individual patients, these tests are usually performed on isolates recovered in the following situations: serious infections, in pure culture or from usually sterile body sites, infections requiring extended therapy (e.g. osteomyelitis and endocarditis), patients failing empiric treatment and particularly virulent species. Examples of serious infections from which isolates should be tested for antimicrobial susceptibility include brain abscess, joint infection, infection of a vascular graft or prosthetic device, and recurrent bacteraemia. Organisms recognized as being commonly resistant to antibiotics and/or especially virulent include members of the B. fragilis group, Prevotella and Porphyromonas spp., certain fusobacteria, Cl. ramosum, Cl. septicum, Cl. perfringens, Bilophila wadsworthia, and Campylobacter gracilis. Appropriate methods for susceptibility testing include agar dilution on supplemented Brucella blood agar, broth (macro) dilution, and broth microdilution as described by the National Committee for Clinical Laboratory Standards (NCCLS) . The E-test represents another commonly used option .
Historically it has been argued that antimicrobial susceptibility testing of anaerobes is not necessary because: (i) they remain highly susceptible to antibiotics with recognized anaerobic activity; (ii) their slow-growing nature results in delays that limit the test's clinical utility; (iii) standardized methods for such testing are not available. These arguments are now invalid. Unfortunately, the antimicrobial susceptibility patterns of anaerobes are no longer predictable and growing resistance has been clearly documented [27,28]. In addition, standardized techniques are now commonly used in clinical laboratories following the guidelines of the NCCLS. Furthermore, Snydman et al. have shown that results of in-vitro susceptibility testing for cefoxitin are predictive of patients’ response to therapy . Although surgical intervention and drainage, when appropriate, remain the mainstays of therapy for most anaerobic infections, antibiotics clearly play a major role in treating many of these patients successfully. Patients’ overall health and the microenvironments in which infections occur are also contributing factors in the determination of how patients will respond to therapeutic interventions. Furthermore, most anaerobic infections are polymicrobial in nature. Therefore, to expect precise correlation between susceptibility test results and clinical outcome with anaerobic infections is unrealistic. The fact, however, that most such testing is conducted on isolates from chronic or refractory infections underscores its clinical relevance, even if 48 h of incubation are required for accurate testing. In the most serious infections caused by anaerobic bacteria, a consensus group of infectious disease physicians have concluded that patient clinical response does, in fact, correlate with in-vitro antimicrobial susceptibility testing results , clearly supporting the use of these tests in selected cases.
MECHANISMS OF ANTIMICROBIAL RESISTANCE
Among anaerobes, the mechanisms resulting in antimicrobial resistance to various classes of antibiotics are described below and are outlined in Table 5. Multiple-resistant organisms are sometimes isolated.
Production of β-lactamases by anaerobes is common. Up to 97–100 % of B. fragilis group isolates in the USA [31,32] and 76 % of such isolates in the UK  have been shown to produce these enzymes. Among Bacteroides spp., other than those in the B. fragilis group, approximately 65 % produce β-lactamases . Most of the β-lactamases produced by Bacteroides spp. are chromosomally-mediated and produced constitutively. Some contain serine at their active site whereas others, known as metallo-enzymes, require a Zn2+ at their active site for effective β-lactam hydrolysis . Organisms producing these enzymes are particularly disturbing because, with the exception of the monobactams, metallo-β-lactamases hydrolyse all β-lactam agents, including carbapenems such as imipenem and meropenem, and are not inhibited by available β-lactamase inhibitors. Although usually chromosomal, plasmid-mediated metallo-β-lactamases also occur.
β-lactamase production has also been described among a number of other species of anaerobes including Cl. clostridioforme , Cl. butyricum , Cl. ramosum , Fusobacterium nucleatum [34,39], Prevotella spp. and Porphyromonas spp. . Although β-lactamase production by Clostridium spp. remains uncommon, when produced such enzymes are generally inducible, with the exception of TEM-1 β-lactamase in Cl. ramosum.
Members of the B. fragilis group produce group 2e cephalosporinases, which are inhibited by the β-lactamase inhibitors sulbactam, clavulanic acid and tazobactam, thereby explaining their general susceptibility to the commercially available β-lactam/β-lactamase inhibitor antibiotic combinations. Many of the enzymes produced by Bacteroides spp. have been characterized molecularly as class A serine cephalosporinases, which are smaller in size than the inducible group 1 (class C) common to many aerobic Gram-negative species. However, a strain of B. intermedius has been reported that produces a β-lactamase refractory to inhibition by clavulanic acid. Strains of B. vulgatus have been described that produce a class A cefoxitin-hydrolysing enzyme capable of degrading the compound slowly in overnight culture. This degradation may be accompanied by decreased permeability, thus explaining their clinical resistance to cefoxitin therapy.
Rather than cephalosporinases, clostridia and fusobacteria produce at least three types of penicillinase. Enzymes from Cl. butyricum and fusobacteria are inhibited by clavulanic acid whereas the penicillinase from Cl. clostridioforme is refractory to inhibition by all three commercially available β-lactamase inhibitors. This penicillinase is similar in nature to the class D, group 2d cloxacillin-hydrolysing enzymes.
In addition to enzyme production, permeability changes lead to β-lactam resistance among anaerobes. Pore-forming molecules associated with decreased permeability and β-lactam resistance have been described in B. fragilis, Fusobacterium nucleatum and Porphyromonas endodontalis . Decreased permeability has also been associated with increased β-lactamase production, resulting in even higher levels of resistance to β-lactam antibiotics . Cefoxitin resistance has been correlated specifically with a decrease in outer membrane permeability and the loss of a 49–50 kDa outer membrane protein .
Binding to penicillin binding proteins (PBP) is the critical factor in determining whether a β-lactam antibiotic will effectively inhibit cell wall synthesis in a bacterium. Essential for bacterial growth, PBP function is the terminal stage of cell wall synthesis. When a β-lactam effectively competes for the active site of an essential PBP, bacterial cell death results. Three to five PBP exist in strains of Bacteroides spp.: a PBP1 complex comprised of from one to three different enzymes, PBP2 and PBP3. Most β-lactams bind well to PBP2 as well as the PBP1 complex of Bacteroides spp. By comparison, monobactams such as aztreonam have very poor affinity for these Bacteroides PBP, accounting for their lack of activity against them . Although resistance to cephalosporins is usually attributable to β-lactamase production, resistance in B. fragilis due to decreased affinity for PBP3 has also been incriminated and decreased susceptibility to cefoxitin has been attributed to decreased affinity for PBP2 as well as the PBP1 complex.
Rates of resistance to clindamycin vary geographically and, therefore, surveillance for resistance is critical to assess the utility of clindamycin as a therapeutic option at any given institution . Although all three major mechanisms of antimicrobial resistance (enzyme inactivation, target site modification, and permeability alterations) result in resistance to clindamycin among various species of anaerobes, macrolide/lincosamide/streptogramin B (MLS) is the mechanism most frequently encountered in Bacteroides spp. Based on DNA and protein sequence analysis, it apparently occurs by a mechanism similar to that for clindamycin resistance in staphylococci.
Among Bacteroides spp., resistance is apparently mediated by methylation of 23S rRNA at one of two adenine residues which prevents effective binding of clindamycin to the ribosomes, rendering them refractory to the drug's inhibitory properties. This mechanism may be either inducible or expressed constitutively. Three closely related MLS genes, cloned from different Bacteroides spp., lie either on a conjugal element or on a transposon:ermFS is encoded on Tn 4551, ermFU is encoded on a B. vulgatus conjugal element, and ermF is encoded on Tn 4351. The resulting proteins have sequence identities highly similar to those encoded by the MLS resistance genes of Gram-positive organisms. However, not all Bacteroides clindamycin resistance DNA sequences cross-hybridize with the ermF gene, indicating that another mechanism of resistance is also present in some Bacteroides spp.
Conjugal transfer of clindamycin resistance has been shown to be plasmid-mediated. Many of these plasmids are self-transmissible and range in size from 14.6 kb to approximately 82 kb. Chromosomally-encoded clindamycin resistance is linked to tetracycline resistance and the gene has been shown to lie within the tetracycline resistance transfer element .
Resistance to tetracycline is almost universal among Bacteroides spp.: the rate exceeds 80–90 % at most medical centres. Due to this high rate of resistance, tetracycline should not be used therapeutically for anaerobic infections, as it was through the 1960s, without evidence generated from susceptibility testing that a patient's infecting pathogens are susceptible. Protection or modification of the target site is the only documented mechanism of resistance to tetracycline in Bacteroides spp. The chromosomal tetQ gene encodes a protein that renders the ribosomal protein synthesis apparatus refractory to the inhibitory effects of tetracyclines. DNA cross-hybridization studies indicate that a tetQ or tetQ-like gene is present in most tetracycline-resistant Bacteroides isolates. Identification of tetracycline-resistant isolates that do not contain tetQ DNA sequences indicates that another mechanism (e.g. efflux of tetracycline) or another class of ribosomal protection proteins also contributes to tetracycline resistance.
Tetracycline resistance genes have also been identified in Clostridium spp. The tetA (P) and tetB (P) genes together form an operon encoding two unrelated proteins that result in tetracycline resistance mediated by two separate mechanisms . Two additional genes that cause tetracycline resistance have been identified in Bacteroides species. The tetX gene product inactivates tetracycline by oxidation of the molecule, but is active only under aerobic conditions and has, therefore, not been shown to be operational in Bacteroides spp. A gene encoding a protein that is able to actively efflux tetracycline has also been identified, but has not been shown to confer resistance to tetracycline in these species.
The tetQ resistance gene is both inducible and transferable. The frequency of transfer of resistance to tetracycline is, however, very low unless an organism is exposed to tetracycline. Gene transfer occurs by conjugation mediated by the tetracycline resistance transfer element itself. Transfer is controlled by a two-component regulatory system. The two regulatory genes, rteA and rteB, are located in the tetQ operon downstream from the tetQ gene. Their expression is enhanced greatly by the presence of tetracycline, explaining the more efficient transfer of genetic material following tetracycline exposure. The rteA gene encodes the cytoplasmic membrane component of the system, RteA. Encoded by the rteB gene, the regulatory response protein, RteB, appears to elicit its effect through an interaction with a σ54-like protein . RteB plays an essential role in the mobilization and transfer of the tetracycline transfer element. A third gene, rteC, is also involved in the self-transfer of tetracycline resistance. Although its specific role remains undefined, RteC, its gene product, follows RteB in the regulation cascade.
In addition to regulating the movement of the tetracycline resistance transfer element, RteA and RteB are also involved in the regulation of transfer of unlinked chromosomal elements called non-replicating Bacteroides units (NBU). Although most NBU do not result in any type of phenotypic expression, a cefoxitin-hydrolysing β-lactamase gene, cfxA, has been shown to reside on a NBU. Transfer of the cefoxitin-hydrolysing β-lactamase is facilitated by tetracycline pretreatment. Similar to the conjugal transposon Tn 916 in Enterococcus faecalis, these tetracycline resistance transfer elements, with estimated sizes ranging from 70 to 80 kbp, reside in the chromosome and frequently contain other resistance genes (e.g. ermF).
Metronidazole, the first 5-nitroimidazole to be used clinically, was introduced in 1960. It was not until 1978, however, that a metronidazole-resistant B. fragilis isolate was recovered from a patient following long-term therapy . Metronidazole-resistant isolates from patients not treated with the compound have also been reported. One theory put forward for clinical failure with metronidazole is that the drug is inactivated by other organisms present in polymicrobial infections. Enterococcus faecalis has been reported to inactivate metronidazole and protect B. fragilis from its antimicrobial effects in mixed cultures. Despite these reports, metronidazole resistance among anaerobic Gram-negative bacilli remains extraordinarily low, typically < 1 %, and the compound continues to be viewed as a primary drug for treatment of infections caused by Bacteroides and other anaerobic Gram-negative species.
The 5-nitroimidazoles, including metronidazole, tinidazole and ornidazole, must undergo reduction to form an active antibacterial agent. This reduction is rapidly reversible in the presence of oxygen and is stable only under anaerobic conditions. Resistance to metronidazole is attributed to a combination of reduced nitroreductase and decreased drug uptake by the organism. Both of these occur simultaneously. Because entry of 5-nitroimidazoles into the cell depends on the rate of reduction of the nitro group, any decrease in the reducing environment within the bacterium will result in reduced nitroreductase activity and a concomitant reduction in drug uptake. Decreased pyruvate : ferredoxin oxireductase activity in combination with a compensatory increase in the lactate dehydrogenase activity results in a decrease in the reducing power of the bacterium. These changes in enzyme activity are uncommon and difficult for the organism to accomplish. Metronidazole-resistant phenotypes have, however, been reported, albeit rarely. Although the exact mechanism of this metronidazole resistance has not been identified, efflux and reduced permeability to the drug have been excluded.
Two genes, designated nimA and nimB, capable of conferring moderate- to high-level resistance have been described. DNA sequencing of these two genes shows approximately 73 % similarity, and they presumably represent two unique genes that confer resistance by the same mechanism. These genes have been localized to the chromosome as well as to a variety of different non-self-transmissible plasmids. These plasmids can, however, be mobilized by other conjugal elements or acquired by transformation. Of note, the transcriptional start genome for both nimA and nimB has been sequenced and is located on insertion sequence (IS) elements integrated 12–14 bp upstream from the protein-coding regions. IS 1118, the element for the nimA gene, is very closely related to the IS element providing the transcriptional start information for the nimB gene and is almost identical to IS 1186, an IS element that provides the transcriptional initiation signals for the metallo-β-lactamase gene described in Bacteroides spp.
Resistance to chloramphenicol among anaerobes, as reported in most antimicrobial susceptibility surveys, remains uncommon probably due to its relatively infrequent use in areas where such surveillance studies have been conducted. Two different classes of chloramphenicol resistance genes have, however, been reported in Bacteroides spp. Both result in drug inactivation, either through acetylation or by nitro-reduction at the p-nitro group of the benzene ring. The chloramphenicol acetyltransferase gene is transferable and plasmid-mediated.
Most currently available quinolones and fluoroquinolones have limited activity against many anaerobic bacterial species. Recently-developed agents including trovafloxacin, however, demonstrate far superior activity than previously released drugs . Whether the typically low rate of susceptibility to most quinolones is due to poor drug penetration, low affinity for the target topoisomerases II and IV, or some other mechanism has not yet been clearly ascertained. Concomitant reduced cell permeability has, however, been described in clinical isolates of B. fragilis to cefoxitin and norfloxacin.
Another reservation to the use of quinolone class antibiotics for treatment of anaerobic infections is the finding that they are generally bacteriostatic rather than bactericidal under anaerobic conditions. As a group, fluoroquinolones have significantly more activity against oral anaerobes than members of the B. fragilis group. As a result, they may eventually prove useful for treatment of mixed aerobic and anaerobic pulmonary infections such as aspiration pneumonia.
Resistance to aminoglycosides is uniform among anaerobic bacteria. This universal resistance is not, however, due to decreased target sensitivity. Both gentamicin and streptomycin bind to and inhibit protein synthesis occurring on Cl. perfringens and B. fragilis ribosomes in a cell-free system . Drug inactivation was not detectable in cell extracts of these same two species  and, therefore, does not appear to account for this resistance. Instead, it appears as if aminoglycoside resistance is the result of the drugs’ failure to reach their target sites. Aminoglycoside uptake is a two-step process involving both energy-independent and energy-dependent phases. Either an oxygen- or a nitrogen-dependent electron transport system has the capability to provide the energy necessary for the energy-driven phase of aminoglycoside uptake. Strict anaerobes lack these electron transport systems and are therefore impermeable to aminoglycosides.
Anaerobic bacteria are common causes of clinical infection. Their recovery from patient specimens is compromised, however, by their innate sensitivity to molecular oxygen. As a result, these important pathogens are often underappreciated as significant contributors to infectious processes. Because infections caused by anaerobes are frequently rapidly destructive, resulting in significant morbidity and on occasion mortality, hospitals should take measures to assure that optimal specimen transport systems (e.g. PRAS transport media) are in place so that these organisms can be recovered efficiently in culture.
In addition, clinical microbiology laboratories should assess their methods of anaerobic culture to optimize recovery of these pathogens further and to assure that their presence in patient specimens is reported rapidly and accurately. Although surgical intervention remains critical to the effective management of many types of infectious processes involving anaerobes, antimicrobial therapy is often an equally important therapeutic modality.
Because of evolving resistance to all major classes of antimicrobial agents used for treatment of clinical infection, antibiotic susceptibility testing of anaerobes is important to direct therapy. Many clinical microbiology laboratories have to date either been unable or unwilling to conduct such testing. All hospital laboratories should develop a strategy to offer susceptibility testing to clinicians when clinically relevant anaerobes are recovered from infections. In addition, a mechanism should be developed whereby cumulative antimicrobial susceptibility results are collected such that changing patterns of resistance can be monitored and direction can be afforded to physicians in terms of drugs useful for empirical treatment of anaerobic and mixed aerobic/anaerobic infections. Only by instituting such measures will optimal patient care be guaranteed in cases of serious anaerobic infection.
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Keywords:© 2001 Lippincott Williams & Wilkins, Inc.
Anaerobes; gangrene; susceptibility; antibiotics; Bacteroides; resistance