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J Clin Microbiol. 2007 May; 45(5): 1551–1555.
Published online 2007 February 21. doi:  10.1128/JCM.02424-06
PMCID: PMC1865886

Surveillance Cultures and Duration of Carriage of Multidrug-Resistant Acinetobacter baumannii[down-pointing small open triangle]


Isolating carriers of multidrug-resistant (MDR) Acinetobacter baumannii is the main measure to prevent its spread. Identification of carriers accompanied by contact precautions is essential. We aimed to determine the appropriate surveillance sampling sites and the duration of carriage of MDR A. baumannii. We studied prospectively two groups of patients from whom MDR A. baumannii was previously isolated: (i) those with recent clinical isolation (≤10 days) and (ii) those with remote clinical isolation (≥6 months). Screening for carriage was conducted from six sites: nostrils, pharynx, skin, rectum, wounds, and endotracheal aspirates. Strains recovered concurrently from different sites were genotyped using pulsed-field gel electrophoresis. Twelve of 22 with recent clinical isolation of MDR A. baumannii had ≥1 positive screening culture, resulting in a sensitivity of 55% when six body sites were sampled. Sensitivities of single sites ranged from 13.5% to 29%. Among 30 patients with remote clinical isolation, screening cultures were positive in 5 (17%), with a mean duration of 17.5 months from the last clinical culture. Remote carriers had positive screening cultures from the skin and pharynx but not from nose, rectum, wounds, or endotracheal aspirates. Eleven strains from five patients were genotyped. In all but one case, isolates from different sites in a given patient were clonal. Current methodology is suboptimal to detect MDR A. baumannii carriage. The sensitivity of surveillance cultures is low, even when six different body sites are sampled. The proportion of individuals with previous MDR A. baumannii isolation who remain carriers for prolonged periods is substantial. These data should be considered when designing measures to limit the spread of MDR A. baumannii.

In the past decade, multidrug-resistant (MDR) Acinetobacter baumannii has emerged as a major nosocomial pathogen in many parts of the world, resulting in devastating outcomes in terms of morbidity, mortality, and costs (1, 2, 11, 14, 18, 26). Therapeutic options are often scarce, and in many instances there is not a single drug to administer against this pathogen (1, 11, 13, 14, 17-20). The International Network for the Study and Prevention of Emerging Antimicrobial Resistance defined the emergence of carbapenem resistance in Acinetobacter baumannii infections as a “global sentinel event” warranting prompt epidemiological and microbiological interventions (30).

The most effective measures reported thus far to reduce the burden of MDR A. baumannii infections in hospitals are strict contact precautions, cohorting, application of a routine surveillance program in order to identify silent carriers, intense cleansing of the carrier's environment, appropriate management of infections, and attempts at decolonization (of questionable value) (2, 14, 23). These interventions, though cost-effective in the long run, pose a vast burden on hospital resources and personnel (2, 7, 9, 10, 13, 17, 23, 25, 26, 28).

Acinetobacter species are widely distributed in the environment and may be common commensals in humans (14). Rates of skin colonization as high as 25 to 40% for healthy ambulatory volunteers and up to 75% for hospitalized patients are reported (1-3, 26). The bacteria colonize the pharynx intermittently in 7% of the general population and in addition can be isolated from sputum, urine, stool, and vaginal discharges (2, 14, 26). In contrast, MDR A. baumannii is primarily a nosocomial pathogen acquired in the health care setting, and its carriage and natural history have not been well studied (14).

This study had two principal aims: (i) to examine the sensitivity of culturing various body site samples to detect carriage of MDR A. baumannii among patients with recent MDR A. baumannii clinical isolation, assuming that this group of patients continues to carry the organism at the time of surveillance; and (ii) to evaluate the long-term carriage of MDR A. baumannii among patients with a remote MDR A. baumannii clinical isolation who are readmitted.



Tel-Aviv Sourasky Medical Center (TASMC) is a 1,200-bed tertiary-care teaching hospital comprising 45 wards, with 84,000 admissions and over 87,500 clinical cultures processed annually. Hospital computerized databases record patients with previously isolated MDR A. baumannii presenting to the emergency rooms, and “flagging” of a patient in the system results in the immediate application of contact precautions. Israeli hospitals over the past decade have been considered an environment where MDR A. baumannii is hyperendemic, with a rate of detection of 1 per 140 hospital admissions of medical or surgical patients (1, 23).

Acinetobacter isolates were identified to the species level by use of the Vitek 2 system (bioMérieux, Hazelwood, MO). In order to differentiate the genomic patterns of the Acinetobacter species identified, PCR of the 16S-23S rRNA intergenic spacer (internal transcribed spacer) was carried out for the study isolates according to an established protocol (8). Antimicrobial susceptibility testing was performed using the Vitek 2 AST GN09 card, and susceptibilities to imipenem and meropenem were confirmed by disc diffusion or Etest (AB Biodisk, Solna, Sweden). An A. baumannii isolate was defined as MDR if it was resistant to at least three classes of antibiotics (including penicillins, cephalosporins, monobactams, β-lactamase inhibitor combinations, aminoglycosides, and fluoroquinolones), while susceptibilities to amikacin, ampicillin-sulbactam, imipenem, meropenem, and minocycline were allowed. All isolates were processed according to Clinical and Laboratory Standards Institute (CLSI) criteria (12).

Infection control practices during the study period.

Throughout the study period, contact precautions for the duration of the hospital stay were advised for patients from whom MDR A. baumannii was isolated. Upon admission of a previously identified carrier, or within 24 h of initial culture of MDR A. baumannii from a hospitalized patient, ward personnel received a daily e-mail list of patients who needed to be under contact precautions and were called by infection control practitioners to confirm adherence to contact precautions. In most cases, patients were in multipatient rooms, their beds were marked with signs bearing the words “contact precautions,” and gowns and gloves for patient contact as well as alcohol-based hand disinfectant were present nearby. Periodic surveys revealed that the material required for contact precautions was present over 90% of the time; however, compliance with use was not systematically recorded.

Study design.

Prospective surveillance was conducted from 1 June 2006 to 31 August 2006. Hospitalized adults (>18 years of age) who had positive clinical cultures of MDR A. baumannii isolated between 1 December 2002 and 31 August 2006 were considered for inclusion. Two groups of patients were included. The first group consisted of patients from whom MDR A. baumannii was isolated from a clinical specimen in the preceding 10 days. We considered these patients with a recent positive clinical isolation to be carriers by definition. This group was studied to determine and compare the sensitivities of surveillance from six different body locations. The second group consisted of patients with a positive clinical isolation during previous admissions ≥6 months prior to study long-term carriage.

Each patient was approached by infection control personnel, who after obtaining consent cultured patient specimens from four different surveillance sites: the nose (nostrils, bilaterally), the pharynx, the skin (the swab was premoistened in the transport media; the same swab was then used to culture the axillae, the antecubital fossae, and the groin bilaterally, in that order), and the rectum. Two additional sites were sampled for subsets of patients: wounds, if a draining ulcer was present; and endotracheal aspirates, if the patient was intubated.

Swabs (CE0373; MEUS, Piove di Sacco, Italy) were inoculated within 1 h in enriched brain heart infusion broth and incubated for 24 h at 35°C. Samples were then streaked on selective MacConkey agar plates (Novamed Ltd.; Jerusalem, Israel) containing 2 μg/ml amphotericin B and 8 μg/ml ceftazidime. Quality control of the selective plates was performed on a regular basis, using Escherichia coli ATCC strain 25922 as a susceptible strain and a ceftazidime-resistant E. coli clinical isolate as a resistant strain. In a preliminary study, the enrichment method showed sensitivity superior to that obtained by the direct plating of swabs onto selective media when performed in parallel (unpublished data). Representative colonies of each morphotype were picked in duplicate and transferred to Enterotubes (Enterotest; Hy Laboratories Ltd., Rehovot, Israel). Oxidase-negative nonfermentors were further identified by use of the Vitek 2 system to the species level, and the antimicrobial susceptibility profiles were determined. Isolates were stored at −70°C for further workup.

Epidemiologic data collection.

Epidemiologic data were collected via patient interviews and chart reviews. Parameters assessed included demographics (age and sex), microbiological parameters of previous clinical isolations, medical diagnosis at the current admission, long-term care facility residency, functional status, level of consciousness, comorbidities (including calculation of the Charlson comorbidity index [22]), severity of illness (according to the McCabe score [4]), use of chronic invasive devices, recent invasive procedures, recent use of antibiotics, chronic medications, tobacco or alcohol use, recent immunosuppressive treatment (glucocorticoids or oncologic chemotherapy), malignant diseases, renal function, nutritional status, and time intervals from most recent hospitalizations and/or intensive care unit stays.


Patients with ≥2 positive cultures from different body sites were genotyped and classified into genetic clusters by use of pulsed-field gel electrophoresis (PFGE). Since the initial isolates from previous admissions of the remote carriers were not available, we could not compare the PFGE patterns of previous and current isolations. MDR A. baumannii was cultured on MacConkey agar and afterwards in brain heart infusion broth for 18 h. Agarose discs of genomic DNA were prepared as previously described (21, 24). DNA was then cleaved using 20 U of the restriction enzyme ApaI endonuclease (New England Biolabs, Beverly, MA) for 3 h at 25°C (15, 21, 24). Electrophoresis was performed in a 1% agarose gel (BMA Products) prepared and run in 0.5× Tris-borate-EDTA buffer on a CHEF-DR III apparatus (Bio-Rad Laboratories, Richmond, CA). The initial switch time was 5 seconds, the final switch time was 35 seconds, and the run time was 23 h at 6 V/cm. Gels were stained with ethidium bromide, destained in distilled water, and photographed by using a Bio-Rad GelDoc 2000 camera. DNA patterns were analyzed visually and by using Diversity software (Bio-Rad). PFGE DNA patterns were compared and interpreted according to an established protocol (6, 24, 27).

Statistical analysis.

Continuous variables were compared between groups by use of an unpaired t test and a paired t test within each group. Categorical variables were compared by use of the Pearson χ2 test. For small samples, analysis of variance and the Fisher exact test were used to analyze continuous and categorical variables, respectively. Statistical analyses were conducted using SPSS (version 13.0; SPSS Inc., Chicago, IL) and Stata (version 9.0; Stata Corp., College Station, TX). P values of ≤0.05 were considered significant.


Surveillance sensitivity study.

Twenty-two patients with a recent clinical isolation of MDR A. baumannii (≤10 days) were considered “carriers” by definition. These patients were surveyed. Sixteen of the patients were men (73%), the mean age was 68 years (range, 29 to 88), and the mean number of days since the last positive MDR A. baumannii isolation was 6 (range, 3 to 10). The index clinical cultures of MDR A. baumannii were obtained from endotracheal aspirates (seven patients), from wounds (six patients), from urine specimens (three patients), from intravascular catheter tips (three patients), and from blood specimens (three patients).

Of the 22 patients studied, 12 had at least one positive surveillance isolation (range, 1 to 4). Therefore, the overall sensitivity of the multisite surveillance approach was 55%. The surveillance site and clinical site cultures are depicted in Table Table1.1. No statistical difference between the various sites in terms of yield was found (P = 0.36). The clinical syndrome and the number of days since the previous MDR A. baumannii isolation were not significantly associated with surveillance culture results.

Sensitivities of surveillance cultures from different body sites among patients with recent clinical culture of MDR A. baumannii (≤10 days)

The clinical culture location most correlated with positive surveillance isolation was the endotracheal aspirate, followed by a wound: five of seven patients (71%) with a positive clinical culture from the endotracheal aspirate and four of six patients (67%) with a positive clinical culture from a wound had a positive surveillance isolation at any site. Other sites of positive clinical culture—urine, intravascular catheter tip, and blood—were less predictive of positive surveillance isolation, with a correlation of one of three patients (33%) for each site.

Duration of carriage study.

During the 3-month study period, 30 of 36 patients who had a previous remote clinical isolation (≥6 months prior) of MDR A. baumannii and were readmitted to TASMC agreed to participate in the study. One hundred forty samples were obtained from these patients. For five patients (17%), at least one surveillance isolation yielded MDR A. baumannii. The mean duration from the first MDR A. baumannii isolation from a clinical culture was 20 months (range, 8 to 42), and that from the last isolation was 16 months (range, 1 to 39). The durations from the last clinical isolation were similar between those found to be long-term carriers and those with negative surveillance cultures (Table (Table2).2). Risk factors for prolonged carriage were a bedridden functional state, disorientation at admission, and status after coronary bypass surgery, as depicted in Table Table2.2. Other risk factors, including recent use of immunosuppressants, did not reach significance. The source of the previous positive isolation and the current admission diagnosis did not affect the duration of carriage.

Several epidemiological parameters and their association with MDR A. baumannii prolonged carriage in univariate analysis

Among the cohort of remote clinical MDR A. baumannii isolations, 7 of 140 sites sampled yielded the organism, i.e., three patients had 1 site positive and two patients had 2 sites positive for MDR A. baumannii. The skin was the source for four of the isolates, and the pharynx for three. No MDR A. baumannii was isolated from the nostrils or rectum (30 samples obtained from each site). The clinical sites that were surveyed, wounds (17 samples) and endotracheal aspirates (3 samples), also had no positive isolations.


Among the 52 patients studied (the two cohorts combined), 7 of the 17 patients with positive surveillance isolations had ≥2 positive sites, yielding a total of 16 strains. Eleven isolates from five patients were genotyped (Fig. (Fig.1);1); three distinct clones were detected, all known nosocomial clones that are commonly associated with MDR A. baumannii infections at TASMC (1). For each of the five patients for whose samples genotyping was performed, the same clone was found at two sites; for one patient, an additional clone at a third site was also found (patient 517) (Fig. (Fig.11).

FIG. 1.
Distribution of PFGE clones among patients with ≥2 positive isolates. Lanes: 1, lambda ladder molecular size marker; 2 and 3, clone H, patient 11, with isolates recovered from pharynx and skin, respectively; 4 and 5, clone D, patient 502, with ...


In this surveillance study, we focused on two questions which are cardinal in limiting the spread of MDR A. baumannii: which body site should be cultured in order to detect carriage of MDR A. baumannii, and what is the duration of carriage? We found that culturing a single body site has very low sensitivity, not higher than 30%, and that even when multiple sites are sampled the sensitivity of detecting carriers of MDR A. baumannii reaches only 55%. Sampling multiple body sites is time-consuming and costly, and it is not suitable for routine use by clinicians and clinical laboratories. The low sensitivity of single-site surveillance for MDR A. baumannii is in contrast to the much higher sensitivities of single-site screening for other MDR pathogens, e.g., rectal cultures for the detection of carriers of vancomycin-resistant enterococci and nasal cultures to detect carriers of methicillin-resistant Staphylococcus aureus (16, 29). We also found that MDR A. baumannii may be carried for long durations, up to 42 months, and that prolonged carriage affects at least 17% of patients with previous clinical isolations of MDR A. baumannii. This proportion of long-term carriers is likely an underestimate due to the limited sensitivity of surveillance in the detection of carriers (55%) and implies that prolonged carriage of MDR A. baumannii may affect 30% of patients with remote clinical isolations.

A combination of several factors may explain the low sensitivity of surveillance cultures. First, it was assumed that all patients from whom MDR A. baumannii was isolated from clinical cultures within the last 10 days were carriers, and this may not have been true in all cases, i.e., certain patients may have had MDR A. baumannii at the infection site only. This is likely the explanation in very few cases, since both surveillance sites and clinically relevant sites (draining wounds and endotracheal aspirates from intubated patients) were sampled. Second, patients may have received an appropriate antimicrobial therapy against MDR A. baumannii, which may have eradicated the pathogen prior to our sampling. However, among our cohort, only two had received such a regimen, as one patient received cefepime (for which the in vitro results have a questionable significance if an extended-spectrum-β-lactamase-producing organism is present), and the second patient received, for only 1 day prior to culture, colistin. Therefore, we believe that this factor did not considerably bias our results. Third, the sampling and microbiological methods used may not be sensitive enough to detect MDR A. baumannii bacteria, particularly if they are present at the sampled body sites in low concentrations. Although an enrichment method was used after its superiority to direct culturing onto selective media was confirmed, this method may still not be sensitive enough. Regarding the surveillance skin samples, for example, we can only postulate that another method which samples a skin surface area larger than that sampled by swabbing would increase the yield of the surveillance cultures. Fourth, MDR A. baumannii may occupy different body sites in different patients.

One might expect that the pharynx would be the best site to sample for carriage of MDR A. baumannii for a number of reasons: Acinetobacter spp. were reported to be common commensals in the human pharynx, pneumonia is the most common clinical syndrome of A. baumannii infections, and the bacterium is known for its ability to rapidly colonize tracheotomies (2). In fact, we found that the pharynx had a low sensitivity (23%) as a surveillance site. The skin was also suggested to be appropriate site for the surveillance of MDR A. baumannii (14). In our cohort of patients with remote clinical isolations, four of five long-term carriers were identified by skin sampling. However, the skin had the lowest sensitivity as a surveillance site for patients with recent clinical isolation (13.5%), though as we previously mentioned, the appropriate methodology for obtaining skin samples is yet not well defined.

Isolating MDR A. baumannii from hospitalized patients depends on external ecological variables and risk factors related to the patients themselves (5). Several previous reports have discussed the risk factors associated with the development of MDR A. baumannii infections in hospitalized patients (2, 14, 26). As far as we know, this is the first report that investigates the risk factors associated with prolonged MDR A. baumannii carriage. Two of the risk factors identified, a bedridden state and a disoriented state, were both reported in the past as being associated with MDR A. baumannii infections in hospitalized patients (2, 26). Small sample sizes limit the generalizability of these results and do not allow for meaningful multivariate analysis. In addition, due to the low sensitivity of surveillance and the possible resultant misclassification of MDR A. baumannii carriers as noncarriers, the identification of these risk factors should be interpreted cautiously.

Another study limitation was that the previous MDR A. baumannii strains of the remote carriers were not available for genotyping, and therefore we can only assume that remote carriers remained carriers of the same clone. However, when we genotyped isolates that were colonizing concurrently different body sites, we saw that a given clone can be isolated from multiple sites and cause different clinical syndromes. In addition, the clones identified were all familiar, having been previously associated with MDR A. baumannii outbreaks at TASMC (1), and therefore it is reasonable to assume that the long-term carriers acquired the strains in the nosocomial setting, continued to harbor them for several years, and probably spread them in their local outpatient environment as well.

In conclusion, our study demonstrates that the current methodology to detect MDR A. baumannii carriage is suboptimal and that persistent carriage of MDR A. baumannii occurs in a substantial proportion of patients. Improved methods of surveillance are necessary, and long-term contact precautions for MDR A. baumannii carriers should be considered.


We thank Keren Strauss and Rina Moskovitch for their assistance in conducting the study.

This study did not receive any financial support.

We have no commercial association or other conflict of interest regarding any of the products used in the study.


[down-pointing small open triangle]Published ahead of print on 21 February 2007.


1. Abbo, A., S. Navon-Venezia, O. Hammer-Muntz, T. Krichali, Y. Siegman-Igra, and Y. Carmeli. 2005. Multidrug-resistant Acinetobacter baumannii. Emerg. Infect. Dis. 11:22-29. [PubMed]
2. Allen, D. M., and B. J. Hartman. 2004. Acinetobacter species, p. 2631-2635. In G. L. Mandell, J. E. Bennett, and R. Dolin (ed.), Mandell, Douglas, and Bennett's principles and practice of infectious diseases, 6th ed. Churchill Livingstone, Philadelphia, PA.
3. Berlau, J., H. Aucken, H. Malnick, and T. Pitt. 1999. Distribution of Acinetobacter species on skin of healthy humans. Eur. J. Clin. Microbiol. Infect. Dis. 18:179-183. [PubMed]
4. Bion, J. F., S. A. Edlin, G. Ramsay, S. McCabe, and I. M. Ledingham. 1985. Validation of a prognostic score in critically ill patients undergoing transport. Br. Med. J. (Clin. Res. Ed.) 291:432-434.
5. Bonten, M. J., S. Slaughter, A. W. Ambergen, M. K. Hayden, J. van Voorhis, C. Nathan, and R. A. Weinstein. 1998. The role of “colonization pressure” in the spread of vancomycin-resistant enterococci: an important infection control variable. Arch. Intern. Med. 158:1127-1132. [PubMed]
6. Bou, G., G. Cervero, M. A. Dominguez, C. Quereda, and J. Martinez- Beltran. 2000. PCR-based DNA fingerprinting (REP-PCR, AP-PCR) and pulsed-field gel electrophoresis characterization of a nosocomial outbreak caused by imipenem- and meropenem-resistant Acinetobacter baumannii. Clin. Microbiol. Infect. 6:635-643. [PubMed]
7. The Brooklyn Antibiotic Resistance Task Force. 2002. The cost of antibiotic resistance: effect of resistance among Staphylococcus aureus, Klebsiella pneumoniae, Acinetobacter baumannii, and Pseudomonas aeruginosa on length of hospital stay. Infect. Control Hosp. Epidemiol. 23:106-108. [PubMed]
8. Chang, H. C., Y. F. Wei, L. Dijkshoorn, M. Vaneechoutte, C. T. Tang, and T. C. Chang. 2005. Species-level identification of isolates of the Acinetobacter calcoaceticus-Acinetobacter baumannii complex by sequence analysis of the 16S-23S rRNA gene spacer region. J. Clin. Microbiol. 43:1632-1639. [PMC free article] [PubMed]
9. Chen, H. P., T. L. Chen, C. H. Lai, C. P. Fung, W. W. Wong, K. W. Yu, and C. Y. Liu. 2005. Predictors of mortality in Acinetobacter baumannii bacteremia. J. Microbiol. Immunol. Infect. 38:127-136. [PubMed]
10. Choi, J. Y., Y. S. Park, C. O. Kim, Y. S. Park, H. J. Yoon, S. Y. Shin, Y. A. Kim, Y. G. Song, D. Yong, K. Lee, and J. M. Kim. 2005. Mortality risk factors of Acinetobacter baumannii bacteraemia. Intern. Med. J. 35:599-603. [PubMed]
11. Cisneros, J. M., and J. Rodriguez-Bano. 2002. Nosocomial bacteremia due to Acinetobacter baumannii: epidemiology, clinical features and treatment. Clin. Microbiol. Infect. 8:687-693. [PubMed]
12. Clinical and Laboratory Standards Institute. 2006. Performance standards for antimicrobial susceptibility testing; 16th informational supplement. Approved standard M100-S16. Clinical and Laboratory Standards Institute, Wayne, PA.
13. Coelho, J., N. Woodford, J. Turton, and D. M. Livermore. 2004. Multiresistant acinetobacter in the UK: how big a threat? J. Hosp. Infect. 58:167-169. [PubMed]
14. Fournier, P. E., and H. Richet. 2006. The epidemiology and control of Acinetobacter baumannii in health care facilities. Clin. Infect. Dis. 42:692-699. [PubMed]
15. Gouby, A., M. J. Carles-Nurit, N. Bouziges, G. Bourg, R. Mesnard, and P. J. Bouvet. 1992. Use of pulsed-field gel electrophoresis for investigation of hospital outbreaks of Acinetobacter baumannii. J. Clin. Microbiol. 30:1588-1591. [PMC free article] [PubMed]
16. Hill, R. L., and M. W. Casewell. 1990. Nasal carriage of MRSA: the role of mupirocin and outlook for resistance. Drugs Exp. Clin. Res. 16:397-402. [PubMed]
17. Jain, R., and L. H. Danziger. 2004. Multidrug-resistant Acinetobacter infections: an emerging challenge to clinicians. Ann. Pharmacother. 38:1449-1459. [PubMed]
18. Jones, R. N. 2001. Resistance patterns among nosocomial pathogens: trends over the past few years. Chest 119:397S-404S. [PubMed]
19. Levin, A. S. 2002. Multiresistant Acinetobacter infections: a role for sulbactam combinations in overcoming an emerging worldwide problem. Clin. Microbiol. Infect. 8:144-153. [PubMed]
20. Levin, A. S. 2003. Treatment of Acinetobacter spp infections. Expert Opin. Pharmacother. 4:1289-1296. [PubMed]
21. Maslow, J., A. Slutsky, and R. Arbeit. 1993. Application of pulsed-field gel electrophoresis to molecular epidemiology, p. 563-572. In D. Persing, T. Smith, F. Tenover, and T. White (ed.), Diagnostic molecular microbiology: principles and applications. American Society for Microbiology, Washington, DC.
22. McGregor, J. C., P. W. Kim, E. N. Perencevich, D. D. Bradham, J. P. Furuno, K. S. Kaye, J. C. Fink, P. Langenberg, M. C. Roghmann, and A. D. Harris. 2005. Utility of the Chronic Disease Score and Charlson Comorbidity Index as comorbidity measures for use in epidemiologic studies of antibiotic-resistant organisms. Am. J. Epidemiol. 161:483-493. [PubMed]
23. Paul, M., M. Weinberger, Y. Siegman-Igra, T. Lazarovitch, I. Ostfeld, I. Boldur, Z. Samra, H. Shula, Y. Carmeli, B. Rubinovitch, and S. Pitlik. 2005. Acinetobacter baumannii: emergence and spread in Israeli hospitals 1997-2002. J. Hosp. Infect. 60:256-260. [PubMed]
24. Tenover, F. C., R. D. Arbeit, R. V. Goering, P. A. Mickelsen, B. E. Murray, D. H. Persing, and B. Swaminathan. 1995. Interpreting chromosomal DNA restriction patterns produced by pulsed-field gel electrophoresis: criteria for bacterial strain typing. J. Clin. Microbiol. 33:2233-2239. [PMC free article] [PubMed]
25. Theaker, C., B. Azadian, and N. Soni. 2003. The impact of Acinetobacter baumannii in the intensive care unit. Anaesthesia 58:271-274. [PubMed]
26. Van Looveren, M., and H. Goossens. 2004. Antimicrobial resistance of Acinetobacter spp. in Europe. Clin. Microbiol. Infect. 10:684-704. [PubMed]
27. Vila, J., M. A. Marcos, and M. T. Jimenez de Anta. 1996. A comparative study of different PCR-based DNA fingerprinting techniques for typing of the Acinetobacter calcoaceticus-A. baumannii complex. J. Med. Microbiol. 44:482-489. [PubMed]
28. Villegas, M. V., and A. I. Hartstein. 2003. Acinetobacter outbreaks, 1977-2000. Infect. Control Hosp. Epidemiol. 24:284-295. [PubMed]
29. Weinstein, J. W., S. Tallapragada, P. Farrel, and L. M. Dembry. 1996. Comparison of rectal and perirectal swabs for detection of colonization with vancomycin-resistant enterococci. J. Clin. Microbiol. 34:210-212. [PMC free article] [PubMed]
30. Wisplinghoff, H., T. Bischoff, S. M. Tallent, H. Seifert, R. P. Wenzel, and M. B. Edmond. 2004. Nosocomial bloodstream infections in US hospitals: analysis of 24,179 cases from a prospective nationwide surveillance study. Clin. Infect. Dis. 39:309-317. [PubMed]

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