Eur J Clin Microbiol Infect Dis (2013) 32:573–580 DOI 10.1007/s10096-012-1777-5
ARTICLE
Microbiological profile and antimicrobial susceptibility pattern of infected root canals associated with periapical abscesses E. L. R. Sousa & B. P. F. A. Gomes & R. C. Jacinto & A. A. Zaia & C. C. R. Ferraz
Received: 14 July 2012 / Accepted: 4 November 2012 / Published online: 6 December 2012 # Springer-Verlag Berlin Heidelberg 2012
Abstract The aim of this investigation was to identify microorganisms from root canals with periapical abscesses and assess the susceptibility of specific anaerobic bacteria to selected antimicrobials and their β-lactamase production. Sixty root canals were microbiologically investigated. The susceptibility of Anaerococcus prevotii, Fusobacterium necrophorum, F. nucleatum, Parvimonas micra, and Prevotella intermedia/nigrescens to antimicrobials was evaluated with the Etest, whereas β-lactamase production was assessed with nitrocefin. A total of 287 different bacterial strains were recovered, including 201 strict anaerobes. The most frequently strict isolated anaerobes were A. prevotii, P. micra, and F. necrophorum. The selected bacteria were susceptible to all the tested antibiotics, except A. prevotii and Fusobacterium species to azithromycin and erythromycin, as well as A. prevotii and F. necrophorum to metronidazole. None of the microorganisms produced β-lactamase. Gram-positive anaerobic bacteria predominated in the root canals with periapical abscesses. All microorganisms tested were susceptible to benzylpenicillin, amoxicillin, amoxicillin + clavulanate, cefaclor, and clindamycin, producing no β-lactamase. E. L. R. Sousa : R. C. Jacinto Department of Conservative Dentistry, Endodontics Division, Pelotas School of Dentistry, Federal University of Pelotas, Pelotas, RS, Brazil E. L. R. Sousa : B. P. F. A. Gomes : R. C. Jacinto : A. A. Zaia : C. C. R. Ferraz (*) Department of Restorative Dentistry, Endodontics Division, Piracicaba Dental School, State University of Campinas (UNICAMP), Av. Limeira 901., Bairro Areiao, Piracicaba, SP 13414-903, Brazil e-mail:
[email protected]
Introduction Periapical abscesses are odontogenic infections that lead to pain and/or swelling, and have the potential to progress through the cortical bone, spreading over the sinuses and other facial spaces of the head and neck, with eventual lethal consequences. Bacteria from infected root canals gain access to periradicular tissues and initiate an extraradicular infection with purulent inflammation [1]. Therefore, early recognition of the etiology and clinical aspects of periapical abscesses is essential to the establishment of appropriate therapies. Most periapical abscesses do not require antibiotic therapy or bacteriologic investigation. Patients who need antibiotics as an adjuvant to root canal therapy (medically compromised individuals, for example) usually respond well to empirical treatment that is not based on the results of culture and susceptibility testing [2]. However, because both the choice and prescription of the antibiotic are immediate, it is important for dentists to have a current view of the endodontic microbial infection [3]. Systemic antibiotics act as a coadjuvant to the conventional surgical methods and should be used with restraint because of the possibility of allergic reactions, toxicity, side effects, and development of resistant strains of microbes [3]. The increasing resistance of anaerobic bacteria to some widely used antibiotics and the introduction of new antibiotics for the treatment and prophylaxis of anaerobic infections ensure the need for monitoring susceptibility patterns periodically by using susceptibility tests. Thus, the periodical monitoring of possible changes in the type and antibiotic resistance of microorganisms responsible for periapical abscesses is recommended [4, 5]. The Etest (AB
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Biodisk, Solna, Sweden), an agar diffusion susceptibility test, holds the promise of being accurate and flexible enough to be performed in most clinical laboratories [6]. The aim of this study was to identify the microbiota of root canals with periapical abscesses and to determine the susceptibility of Anaerococcus prevotii, Fusobacterium necrophorum, F. nucleatum, Parvimonas micra, and Prevotella intermedia/nigrescens to benzylpenicillin, amoxicillin, amoxicillin + clavulanate potassium, metronidazole, cefaclor, clindamycin, erythromycin, and azithromycin by investigating their production of β-lactamase.
Materials and methods Patient selection and clinical features The present study was approved by the Piracicaba Dental School Research Ethics Committee at the State University of Campinas, São Paulo, Brazil, and informed consent was obtained from all subjects. Patients with systemic disease and those who had used antibiotics in the last 3 months were excluded from the study. Impossibility of tooth isolation, poor access to the apical region for sampling (as determined by previous radiographic examination), or the presence of marginal periodontitis were also exclusion factors. Thus, from the 100 patients initially examined, only 60 root canals were included. Patients aged 11 to 63 years, comprising 28 males and 32 females, participated in the study. Among them, 56 presented pulp necrosis determined by vitality tests and four had previous root canal treatment. The diagnosis of periapical abscesses was based on the presence of spontaneous pain, percussion pain, palpation pain, and fistula, associated or not with localized or diffuse swelling. Also, their symptoms matched the criteria for acute periapical abscesses described by Torabinejad and Walton [7]. Radiographic examination was performed to evaluate the root canal treatment status and presence of periapical pathology. Patient sampling The method followed for microbiological procedures has been previously described elsewhere [2, 8, 9]. The microbiological investigation was performed under strict aseptic conditions. A standardized routine of root canal therapy was instituted, and, in each case, a single root canal was sampled. In multi-rooted teeth, only the largest canal was sampled so as to preserve the identity of a single endodontic/microbiologic ecosystem. Prior to root canal sampling, restorations and carious lesions were completely removed and the tooth was immediately isolated with a rubber dam. The tooth and rubber
Eur J Clin Microbiol Infect Dis (2013) 32:573–580
dam were disinfected with 5.25 % sodium hypochlorite for 30 s by using a cotton-tipped applicator. Sodium hypochlorite was neutralized with 5 % sodium thiosulphate [10] and sterile saline solution was used for final washing. Access cavity was prepared with sterile burs (Gates-Glidden, Dentsply-Maillefer, Ballaigues, Switzerland) without the use of water spray to preserve the contents of the radicular pulp tissue. The coronal necrotic pulp tissue was carefully removed, and subsequent enlargement of the coronal third of the root canal was performed to prevent contamination of the paper point by the residues of the coronal pulp. When the tooth had an atresic root canal interfering with the penetration of the paper point, patency was accomplished through minimal instrumentation with a #10 file (C+ files, Dentsply-Maillefer, Ballaigues, Switzerland) without any irrigant. Preexisting root fillings were mechanically removed before microbial sampling. In the case of dry root canal, 0.5 mL of sterile saline solution was injected before sampling. Microbiologic samples were then collected in the beginning of the first endodontic intervention by introducing three sterile paper points (Tanari-Tariman Ltda, Manaus, AM, Brazil) into the full length of the canal, which was determined through standard initial radiographs. The paper points were held in place for 60 s under a continuous flow of nitrogen to preserve anaerobic bacteria [11]. Teeth in which it was not possible to reach the full length of the canal were excluded from the microbiologic sampling. The paper points were immediately placed in an Eppendorf tube containing 1.0 mL of VMGA III transport medium [10, 12] for microbiological investigation within 15 min. Inside the anaerobic workstation (Don Whitley Scientific, Bradford, England), the Eppendorf tube was vortexed for 1 min. The VMGA III medium was diluted to 1/ 10, 1/100, 1/1,000, and 1/10,000 by using prereduced suspension medium (Fastidious Anaerobe Broth; FAB; Lab M, Bury, England). Fifty microliters of each dilution were inoculated onto plates containing 5 % defibrinated sheep blood (Fastidious Anaerobe Agar; FAA, Lab M) and incubated in the anaerobic workstation at 37 °C in an atmosphere of 10 % H2, 10 % CO2, and 80 % N2 for 2, 5, and 14 days to differentiate strict anaerobes from the facultative ones. Selection for Gram-positive anaerobes and actinomycetes involved the use of 5 % defibrinated sheep blood– FAA + nalidixic acid (0.001 % wt/vol; Lab M) agar plates at 37 °C anaerobically for 2, 5, and 14 days. Selection for Gram-negative anaerobes employed 5 % defibrinated sheep blood–FAA + nalidixic acid + vancomycin (0.00025 % wt/ vol; Lab M) agar plates at 37 °C anaerobically for 2, 5, and 14 days. Finally, selection for Clostridia and other anaerobes involved the use of 5 % defibrinated sheep blood– FAA + neomycin (0.0075 % wt/vol; Lab M) agar plates at 37 °C anaerobically for 2, 5, and 14 days.
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Fifty microliters of the dilutions were also inoculated onto plates containing 5 % sheep blood + Brain–Heart Infusion agar (BHI, Lab M), which were aerobically incubated at 37 °C and examined after 18 h and 2 days to differentiate aerobes from facultative anaerobes. Microbial identification After the incubation period, a bacterial morphology analysis was performed with a light stereomicroscope (Lambda Let 2; Atto Instruments Co., Hong Kong, China). The different colony types were subcultured onto fresh prereduced medium (FAA + 5 % sheep blood) to obtain pure cultures. Different colonies were selected on the basis of their appearance on the agar plates. The pure cultures were initially identified in terms of their Gram morphology, gaseous requirements, and ability to produce catalase. Each colony obtained by anaerobic incubation was used to inoculate two plates of fresh FAA + 5 % sheep blood (Lab M). One plate was incubated aerobically and the other anaerobically, both of which were incubated for 2 days. The bacterial growth on each plate was compared to determine the gaseous requirements of the bacterial species. These procedures allowed us to make the primary identification of the strain as Gram-positive or Gram-negative, coccus or bacillus, catalase-positive or catalase-negative, and aerobic or anaerobic. On the basis of these primary results, the appropriate kit for identification was selected. Microbial speciation The following biochemical identification kits were used for the primary speciation of individual isolates: Rapid ID 32A (bioMérieux, Marcy-l’Etoile, France) for strict anaerobic Gram-negative and Gram-positive rods; RapID ANA II system (Innovative Diagnostic Systems Inc., Atlanta, GA, USA) for strict anaerobic Gram-positive cocci; API Staph (bioMérieux) for staphylococci and micrococci (Grampositive cocci; catalase-positive cocci); Rapid ID 32 Strep (bioMérieux) for streptococci (Gram-positive cocci; catalase-negative cocci); and the Rapid NH system (Innovative Diagnostic Systems Inc.) for Eikenella, Haemophilus, Neisseria, and Actinobacillus. Additional tests were performed for black-pigmented Gram-negative anaerobes, including fluorescence testing under long-wave (366-nm) ultraviolet light, hemagglutination of 3 % sheep erythrocytes, and lactose fermentation by application of fluorogenic substrate for 4-methylumbelliferyl-βgalactosidase (MUG) (Sigma Chemical Co., St Louis, MO, USA), as described by Alcoforado et al. [13]. Also, the determination of trypsin-like activity was performed by the application of synthetic fluorogenic peptide (CAAM), as described by Alcoforado et al. [13], Nakamura et al. [14], and Slots [15].
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For additional tests, 20-μL aliquots of MUG and CAAM solutions were placed in plates containing bacterial colonies with characteristic morphologies for the identification of Porphyromonas gingivalis, Porphyromonas endodontalis, Prevotella intermedia/nigrescens, Prevotella melaninogenica, Prevotella corporis, Prevotella denticola, and Prevotella loescheii. Control groups consisted of commercially available species pattern: Porphyromonas gingivalis ATCC 33277, Porphyromonas endodontalis ATCC 35406, Porphyromonas asaccharolytica ATCC 25260, Prevotella intermedia ATCC 25611, Prevotella nigrescens ATCC 33536, Prevotella melaninogenica ATCC 25845, Prevotella loescheii ATCC 15930, Prevotella corporis ATCC 33547, and Prevotella denticola ATCC 33185. Etest The susceptibility of isolated strains of Anaerococcus prevotii (formerly Peptostreptococcus prevotii) and Parvimonas micra (formerly Peptostreptococcus micros), both Gram-positive anaerobic bacteria, as well as Fusobacterium necrophorum, Prevotella intermedia/nigrescens, and Fusobacterium nucleatum (all Gram-negative anaerobic bacteria), was determined by the minimum inhibitory concentration (MIC) using the Etest method (AB Biodisk, Solna, Sweden). Pure colonies of each of the tested microorganisms were suspended in Brucella broth to achieve a density corresponding to 1.0 McFarland turbidity standard (3×108 cfu/mL). A cotton wool swab soaked in the inoculum was used to inoculate the surface of plates containing 5 % defibrinated sheep blood Brucella agar enriched with 5 mg/ml of hemin and 1 mg/ml of vitamin K. The Etest strips containing benzylpenicillin, amoxicillin, and amoxicillin + clavulanate potassium, cefaclor, metronidazole, clindamycin, erythromycin, and azithromycin were applied to each plate in duplicate. After allowing the agar surface to dry for 10 min, the plates were incubated in the anaerobic workstation at 37 °C for 48 h in total. After incubation, an elliptical zone of growth inhibition was seen around the strip. In all species, the tests were read after 24 and 48 h of incubation. At the intersection point of the ellipse with the strip, the MIC was read from the interpretative scale provided (AB Biodisk). MICs less than or equal to the breakpoints recommended by the Clinical and Laboratory Standards Institute (CLSI) [16] were considered to be susceptible; those above the breakpoints were considered to be resistant. The susceptibility breakpoints against anaerobes were determined by using the CLSI 2007 criteria and the values provided by van Winkelhoff et al. [17] for erythromycin and azithromycin, as the breakpoints of these antimicrobials against anaerobes have not been determined by the CLSI [16]. Isolates were considered to be susceptible if the MIC for each antibiotic was as follows: penicillin
