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Characterization of bacterial biofilms formed on urinary catheters
American Journal of Infection Control 40 (2012) 854-9
Contents lists available at ScienceDirect
American Journal of Infection Control
American Journal of Infection Control
journal homepage: www.ajicjournal.org
Major article
Characterization of bacterial biofilms formed on urinary catheters Ryad Djeribi PhD a, *, Warda Bouchloukh MSc a, Thierry Jouenne PhD b, Bouzid Menaa PhD a, c a
Biofilms and Biocontamination of Materials Laboratory, Faculty of Science, Badji Mokhtar University, Annaba, Algeria UMR 6522 CNRS, Faculty of Science of Rouen, Mont Saint Aignan, France c Fluorotronics, Inc., Vista, CA b
Key Words: Acinetobacter baumannii SEM
Background: The formation of bacterial biofilms on urinary catheters is a leading cause of urinary tract infections in intensive care units. Cytobacteriological examination of urine from patients is often misleading, due to the formation of these biofilms. Therefore, characterizing these biofilms and identifying the bacterial species residing on the surface of catheters are of major importance. Methods: We studied the formation of biofilms on the inner surface of urinary catheters using microbiological culture techniques, with the direct contact of catheter pieces with blood agar. The bacterial species on the surface were characterized by scanning electron microscopy, and the kinetic profile of biofilm formation on a silicone substrate for an imipenem-resistant Acinetobacter baumannii bacterium was evaluated with a crystal violet staining assay. Results: The bacterial species that constituted these biofilms were identified as a variety of gramnegative bacilli, with a predominance of strains belonging to Pseudomonas aeruginosa. The other isolated strains belonged to A baumannii and Klebsiella ornithinolytica. Kinetic profiling of biofilm formation identified the transient behavior of A baumannii between its biofilm and planktonic state. This strain was highly resistant to all of the antibiotics tested except colistin. Scanning electron microscopy images showed that the identified isolated species formed a dense and interconnected network of cellular multilayers formed from either a single cell or from different species that were surrounded and enveloped by a protective matrix. Conclusions: Microbiological analysis of the intraluminal surface of the catheter is required for true identification of the causative agents of catheter-associated urinary tract infections. This approach, combined with a routine cytobacteriological examination of urine, allows for the complete characterization of biofilm-associated species, and also may help prevent biofilm formation in such devices and help guide optimum antibiotic treatment. Copyright Ó 2012 by the Association for Professionals in Infection Control and Epidemiology, Inc. Published by Elsevier Inc. All rights reserved.
Bacterial biofilms are able to colonize the inner surfaces of indwelling urinary catheters recovered from patients in intensive care units (ICUs) and can cause infectious diseases. Placement of urinary catheters is the primary factor responsible for the development of nosocomial urinary tract infections (UTIs).1,2 Long-term use of urinary catheters can lead to a quasi-permanent bacterial colonization of the urine and, as a reservoir of microorganisms, trigger nosocomial infections.3 In fact, 25 % of patients undergoing * Address correspondence to Ryad Djeribi, BP 12 Annaba RP, 23000 Annaba, Algeria. E-mail address: [email protected] (R. Djeribi). Supported by Grants-in-Aid from the Algerian Ministry of Education and Scientific Research (Direction of Scientific Research and Technological Development). Conflict of interest: None to report.
short-term urinary catheterization (<7 days) acquire bacteria, with an increasing daily risk of developing an infection of w5%. The estimated likelihood of developing an infection after long-term catheterization (30 days) is 100%.4 When inserted for long periods, urinary catheters may readily acquire biofilms, composed of gram-positive or gram-negative bacteria, on their inner and/or outer surfaces.5,6 The initial event in the mechanism of bacterial adhesion is the deposition of urinary components on the surface of the catheter, leading to the formation of a protein film. This protein film enhances the adhesion of microorganisms, leading to biofilm formation.2,7 This biofilm poses a public health problem for patients who need these medical devices. Treatment with antibiotics becomes difficult, and in some cases ineffective, because of the continuous swarming of planktonic bacteria resulting from biofilm formation.8,9
0196-6553/$36.00 - Copyright Ó 2012 by the Association for Professionals in Infection Control and Epidemiology, Inc. Published by Elsevier Inc. All rights reserved. doi:10.1016/j.ajic.2011.10.009
R. Djeribi et al. / American Journal of Infection Control 40 (2012) 854-9
Furthermore, the formation of biofilms can cause unrealistic and misleading results from cytobacteriological examination of urine (CBEU) in a patient undergoing urinary catheterization. Detecting biofilms before the appearance of a UTI may be difficult. Thus, there is a need to characterize these biofilms and identify the bacterial species on surfaces. Detection and proper identification requires total detachment of adherent bacteria before cultivation of the bacterial suspension.10 Most identification and analysis of bacteria starts with ultrasound application to release the cells from the urinary catheter surface. However, this mechanical process might not be sufficiently efficient for obtaining all of the important information about the bacterial species, and may lead to cellular damage. Indeed, many previous studies11-13 have applied the ultrasound method to detach bacterial biomass from surfaces to culture the bacterial suspension; however, this method appears to limit the complete characterization of biofilms formed on catheter surfaces because of the lethal action of the ultrasound on some of the species initially present in the biofilms.14 Thus, new, reliable techniques are needed to better understand and control biofilms on indwelling medical devices. In the present study, we aimed to detect, identify, and characterize biofilms formed on urinary catheter surfaces directly using microbiological culture techniques, and then analyze these biofilms by scanning electron microscopy (SEM). Rather than using the ultrasound method, we performing culture testing by cutting discs of urinary catheter pieces and depositing them directly onto blood agar. This process is direct, efficient, simple, and reliable and allows identification of all bacteria present. From these cultures, we identified the presence of both monobacterial and polybacterial assemblages. In addition, SEM analysis of the contaminated catheter pieces enabled us to identify the isolated species and their organization. Along with the culture tests, we assessed the kinetic profile of biofilm formation using crystal violet staining assays on a silicone substrate for imipenem-resistant Acinetobacter baumannii. We identified the transient behavior of the bacterium between the biofilm (sessile) state and planktonic (free) state, demonstrating that bacterial species can pass from one state to another depending on different environments. This behavior can modify the results of any potential CBEU. Moreover, by directly testing the contaminated catheter disks, we were able to test the response of A baumannii to a series of 9 antibiotic agents. The results reported in this article are significant and demonstrate the efficacy of our novel approach for identifying and characterizing bacterial biofilms in urinary catheters used in ICU patients, which can be the source of serious catheter-associated UTIs. Our findings also underscore the importance of ensuring direct contact between contaminated urinary catheter samples and the agar surface to fully characterize all biofilm-associated species and help guide the appropriate antibiotic therapy for potential UTIs.
855
Table 1 Bacterial strains identified from catheter samples Patient characteristics
Urinary catheter
Sex
UC- 1 UC- 2
Male Male
18 68
Brain abscess; meningitis Hemorrhagic stroke
UC- 3
Male
52
Prostatitis
Age, years
Pathology
Bacterial isolate(s) P aeruginosa P aeruginosa K ornithinolytica A baumannii P aeruginosa K ornithinolytica
a solution of sodium hypochlorite,15,16 followed by successive washes of the inner surface with jets of sterile distilled water using a syringe with the tip of the needle at the upper end of the catheter. After this treatment, the catheter was carefully and aseptically cut into 3- to 4-mm thick discs. Three or 4 discs were placed on the surface of blood agar plates,17 and the inoculated plates were then incubated at 37 C for 48-72 hours. Isolation and identification of bacterial strains After 48 hours of incubation, bacterial colonies observed on both the cultured discs and their neighborhood were carefully collected as individual samples. Each colony was resuspended in sterile physiological water. Dilutions were prepared from each suspension and grown on standard media tryptic soy agar, blood agar, and cystine lactose electrolyte- deficient medium agar, the latter of which is widely used to study the bacteria responsible for UTIs.18 The inoculated plates were then incubated aerobically at 37 C for 24 hours. The purity of the bacterial colonies was checked, and the bacterial species found on the culture plates were identified by macroscopic, microscopic, and biochemical tests, including an oxidase test and a catalase test using API 20 E and API 20 NE (bioMérieux, Marcy L’Etoile, France). The bacterial strains identified by the bacteriological analysis of urinary catheters from ICU patients are listed in Table 1. Quantification of biofilm formation and kinetics of adhesion on silicone The kinetics of the formation of the biofilm on silicone was assayed using the standard crystal violet staining method.19 In brief, bacteria were grown in Luria-Bertani broth. After incubation, unattached cells were removed by rinsing the support thoroughly with water, and attached cells were stained with 1% crystal violet for 20 min. The dye bound to the adherent cells was then solubilized with ethanol-acetone (75:15, v/v) and the optical density of the solution was measured at 570 nm. Antibiotic resistance
MATERIALS AND METHODS
Urinary catheters used by hospitalized ICU patients for more than 1 week were carefully collected under aseptic conditions. The catheters were placed individually in sterile glass bottles and transported immediately to the laboratory for analysis.
An antibiotic resistance test of the isolated bacteria strains against selected antibiotics was performed using the standard protocol for dissemination of antimicrobial agents on MuellerHinton agar. This test was used to determine the resistance patterns of the isolated strains to 9 commonly used antibiotics: penicillin G, ticarcillin, cefixime, imipenem, gentamicin, amikacin, tobramycin, colistin, and trimethoprim/sulfamethoxazole.
Treatment of medical devices
Biofilm observation by SEM
There is currently no standardized method for the detection and analysis of biofilms on urinary catheter surfaces. The treatment of these catheters consisted of disinfection of the outer surface with
The bacterial biofilms colonizing the intraluminal surfaces of the urinary catheters were examined by SEM analysis. Sterile silicone Foley catheters were cut into 2-mm-thick discs, which were
Sample preparation
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R. Djeribi et al. / American Journal of Infection Control 40 (2012) 854-9
Fig 1. Bacterial cultures obtained from the direct vicinity of the disks incubated on blood agar.
aseptically introduced into tubes containing nutrient broth that had been inoculated with various bacterial cultures obtained from Mueller-Hinton agar. After homogenization, the cultures were incubated at 37 C. After 1 week, the tubes were cleaned of bacterial cultures, and the discs were washed extensively with sterile distilled water before SEM analysis. Samples were fixed in a 2% glutaraldehyde and 0.1 M cacodylate buffer (pH 7.4) for 30 min and then rinsed three times for 10 minutes each rinse in 0.2 M cacodylate buffer (pH 7.4). Samples were then dehydrated by passing them through the following ethanol series: 30%, 50%, and 80% ethanol, each for 10 minutes, followed by 100% ethanol twice for 10 minutes each time. Support samples were then dried at 37 C for 24 hours. After being coated with gold-palladium (via sputter coating), samples were examined under a Cambridge S200 scanning electron microscope (Cambridge Instruments, LEO Electron Microscopy, Inc., Thornwood, NY). RESULTS Identification of biofilm-associated organisms In this study, we collected urinary catheters from patients catheterized for longer than 1 week. Only 3 catheters were selected for analysis, and only one of these was fully analyzed due to the diversity of infection and the presence of multiple potential pathogens typically found in this patient community. Discs were cut from these urinary catheters and cultured on blood agar. Dense bacterial growth was observed around the discs (Fig 1). Macroscopic and microscopic examination of cultures from the inner surface of the catheters revealed that the bacterial masses were composed of both single species (monobacterial biofilms) and a mixture of several species (polybacterial biofilms). The isolated and identified bacterial species that constituted the biomass were mainly varieties of gram-negative bacilli, with a predominance of strains identified as P aeruginosa and other strains identified as A baumannii and Klebsiella ornithinolytica. Kinetics of biofilm formation The kinetic profile of biofilm formation on silicone was assayed for imipenem-resistant A baumannii over a 1-week period using the crystal violet staining method. The results demonstrate the transient behavior of the bacterium between the biofilm (sessile state) and the planktonic state (free bacteria) (Fig 2). These results also show that biofilm formation on the media occurred constantly and continuously despite the cellular release observed during incubation. Based on the observed curve, the kinetic profile is characterized by 2 specific periods during the first 96 hours of incubation, with the first period corresponding to significant biofilm formation and the second period corresponding to biofilm dispersion, with release of bacterial cells into the culture medium.
Fig 2. Biofilm of A baumannii formed on silicone material. (A) Kinetics of biofilm formation of A baumannii. (B) Growth of A baumannii (planktonic state).
In addition, the kinetic profile of biofilm formation on silicon for A baumannii suggests the importance of strong hydrophobicity of the bacterial strain at urine pH and the biofilms’ adhesive strength on such a hydrophobic surface. Our findings demonstrate the transient behavior of the bacterium between the biofilm formed in the inner surface of the catheter and the planktonic state. This behavior indicates that bacterial species are able to pass from one state to another depending on the environment. Thus, this transient behavior may have a significant affect on the results of any potential CBEU. Antibiotic resistance The results of the susceptibility and resistance tests performed using the biofilm bacteria and commonly used antibiotics showed
R. Djeribi et al. / American Journal of Infection Control 40 (2012) 854-9
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Fig 3. SEM images of the biofilms formed in the urinary catheter. (A-C) A baumannii biofilms. (Original magnifications: A, 1,000; B, 3,000; C, 10,000.) (D and E) Biofilms composed of P aeuroginosa and A baumannii. (Original magnification, 3,000.) (F) Biofilms composed of P aeuroginosa, A baumannii, and K ornithinolytica. (Original magnification, 3,000.)
that all of the isolated bacteria (P aeruginosa, A baumannii, and K ornithinolytica) are resistant to penicillin G, cefixime, and trimethoprim/sulfamethoxazole and sensitive to colistin. However, the resistance profiles obtained for the A baumannii strain revealed resistance to the full range of antibiotics tested except colistin. A baumannii is known to be resistant to imipenem, which is widely used to treat infections caused by this bacterium. Of note, an A baumannii strain was isolated from a mixed biofilm formed on the inner surface of a catheter recovered from a 68-year-old patient admitted to the ICU for a cerebrovascular accident. The patient had a history of persistent urinary infection with no apparent symptoms. Bacteriological examinations of urine samples obtained from the catheter pocket revealed the presence of P aeruginosa. In addition, analysis of the urinary catheter surface revealed the
presence of 4 bacterial strainsdA baumannii, Klebsiella spp, and 2 strains of P aeruginosadwith significantly different antibiograms. Biofilm characterization by SEM SEM analysis of the biofilms formed on the inner surface of the catheter samples revealed a dense network of cellular multilayers, formed either from a single cell (Fig 3A-C) or from different species (Fig 3D-F), surrounded and enveloped by a protective matrix. SEM images indicate that the identified isolated species readily developed compatible and strong associations with other species and formed agglomerates on the inner catheter surfaces. The enveloping matrix is composed of extracellular polymeric substances, mainly polysaccharides, which constitute most of the carbon
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organic mass of the biofilm (Fig 3C). This matrix confers resistance to antimicrobial agents and constitutes a reservoir of viable microorganisms.6,20 The biofilms are characterized by a multitude of intercellular pores and interconnected channels, which allow not only the exchange of information and the flow of water and ions, but also the transport of nutrients and the efflux of waste products.
DISCUSSION All of the previous studies on the bacterial biomass responsible for nosocomial infections have reported that gram-negative bacilli are predominant in UTIs.21 In addition, other reports indicate that the strains of P aeruginosa and A baumannii identifed are among the most nonfermented bacterial species frequently isolated from clinical specimens of hospitalized patients.17,22 However, the pathogenicity of these bacterial species is low, and their virulence is expressed only in patients with impaired immune defenses and those requiring invasive devices. Numerous techniques have been developed to detect and characterize biofilm-associated cells on medical devices. Among these is the roll-plate technique, in which infected pieces of catheter are rolled over the surface of a nonselective medium. Characterization and quantification of the biofilm depend on the number of organisms recovered by contact with the agar surface.1 Another method is sonication plus vortexing to detach adherent microorganisms. The developed colonies are allowed to recover after dilution and plating of the resulting suspension on culture media.2 The foregoing methods have limited recovery efficiency, however. The microbiological culture method used in this study involves a longer period of contact between contaminated catheter pieces and the blood agar surface. This method allows the recovery and characterization of most of the bacteria found in polybacterial biofilms with the ability to colonize the inner surfaces of urinary catheters. Thus, unlike methods based on detaching the bacterial biomass with ultrasound, our method allows the identification and complete characterization of all bacterial species present. Ultrasound can damage some cells or even kill certain species initially present in biofilm. Thus, our proposed technique appears better suited to the decontamination of materials compared with the characterization of bacterial biomass fixed on the supports.23 Identification and complete characterization of biofilm is vitally important. The CBEU of urine obtained from contaminated catheters may not necessarily reflect actual UTI status. A sample may be sterile urine with microorganisms located in the luminal catheter biofilm, which could contaminate all urine samples obtained for analysis.24 SEM analysis is important as well. In one patient, SEM images of the inner surface of an indwelling urinary catheter that had been in place for 1 month revealed the presence of biofilms that included Staphylococcus epidermidis within a glycocalyx, despite the fact that CBEU analysis during ablation showed that the urine was free of bacteria.25 According to our findings, CBEU might not reveal all of the bacterial species present in the planktonic state when urine is collected during the initial period (0-24 hours), which coincides with the period of maximum bacterial adhesion on the catheter surface. Therefore, the kinetic profile of biofilm formation and characterization of the transient behavior of A baumannii are significant, because they provide information on the planktonic and biofilm states, allowing identification of the ideal time frame for CBEU. As shown in Figure 2, patients are more likely to develop nosocomial UTIs in the absence of a systematic review and analysis of urinary catheters. Thus, CBEU should be performed at least every 48 hours after placement of the first catheter, to identify the
bacterial cells released from the biofilm matrix formed on the catheter surfaces. In hospitals and other medical institutions, A baumannii has emerged as a significant and problematic human pathogen. This bacterium has a capacity to quickly acquire antimicrobial resistance,26 but also readily adheres to both biological and abiotic surfaces.27,28 Moreover, it has developed a varied pattern of resistance, especially to carbapenems (eg, imipenem), the drugs of choice for treating serious A baumannii infections. Thus, treatment options for these infections appear to be very limited. In this study, we analyzed an imipenem-resistant strain of A baumannii to examine the behavior of such bacteria in immunodeficient patients. The resistance profile revealed a remarkable resistance to all of the antibiotic agents tested, along with a significant sensitivity to colistin. Thus, colistin appears to be an effective antibiotic for treating A baumannii infections in either monotherapy or combination therapy. Our findings may provide useful insight into the nature and effective treatment of biofilmassociated infections caused by this strain. SEM analysis suggests a heterogeneous mosaic architecture of biofilms containing microcolonies of bacterial cells encased in an extracellular polymeric substances matrix and separated from other microcolonies by interstitial voids. This microstructure is similar to that commonly observed in other sessile bacteria, including P aeruginosa. This structural organization shows a dense network of cellular multilayers formed from different species that are surrounded and enveloped by a protective matrix, which constitutes a reservoir of viable microorganisms. This organization allows microorganisms to grow under adverse conditions13,17 and is typical of biofilms found on medical implants and devices (eg, intrauterine devices, central venous catheters). In a study by Morris et al,29 SEM analysis revealed that most of the catheters removed after 1 week of insertion were colonized by bacterial biofilms, particularly on the catheters’ luminal surfaces. These observations also demonstrate that different species can be readily integrated and develop close associations in biofilms. In this study, we investigated and described the different aspects of association of A baumannii in pure and mixed biofilms. The pure culture biofilm shown in Figure 3A is composed of isolated coccobacilli cells, grouped in pairs or with multiple channels of various lengths embedded in a simile matrix. However, the communities shown in Figure 3B have different multilayers formed by A baumannii cells clustered together and embedded in a dense biofilm. This dense biofilm is crossed with a multitude of pores and visible intercellular channels to ensure bacterial exchanges. Figure 3C shows a mature biofilm with a large amount of glycocalyx obscuring the shape of the organism. The adhesive matrix that developed appears as a network around a group of cells that ensures protection for this bacterium. Taken together, the results shown in Figure 3D and E suggest a strong and close association between the A baumannii and P aeruginosa strains analyzed in this study. In these biofilms, the colonies are intermixed, and the communities are embedded in heterogeneous abundant extracellular polymeric substances. The structural complexity, characterized by towers and mushroom-like shapes, gives volume to the biofilm. These analyses not only are descriptive, but also show that these species are isolated together from specimens of hospitalized patients, and that the interactions between these bacteria have the potential to pose a serious barrier to the effective treatment of nosocomial infections. CONCLUSION The results reported in this article are novel and significant. They demonstrate the efficacy of our approach for identifying and
R. Djeribi et al. / American Journal of Infection Control 40 (2012) 854-9
characterizing bacterial biofilms formed on urinary catheters in daily use in ICU patients, which can be the source of serious catheter-associated UTIs. The use of indwelling urinary catheters in daily urological practice remains widespread. The most common infection associated with these catheters is nosocomial infection due to the formation of bacterial biofilms on the catheter surface. This biofilm formation is accompanied by the risk of spreading planktonic microorganisms, which can cause serious nosocomial infections related the difficulty in detecting the biofilm even before the appearance of UTI. The classical methods of bacteriological diagnosis of a UTI, such as CBEU, often yield misleading or incorrect results when evaluating the presence of a biofilm on the surface of a medical device. The present study demonstrates that the proper identification of infectious agents responsible for UTI often requires microbiological analysis of the inner catheter surfaces in addition to routine bacteriological examination. This method allows for identification and complete characterization of the bacterial species forming the biofilm, which can aid in the detection and prevention of biofilm formation and guide appropriate antibiotic therapy for such devicerelated infections. References 1. Trautner BW, Darouiche RO. Role of biofilm in catheter-associated urinary tract infection. Am J Infect Control 2004;32:177-83. 2. Hatt JK, Rather PN. Role of bacterial biofilms in urinary tract infections. Curr Top Microbiol Immunol 2008;322:163-92. 3. Jacobsen SM, Stickler DJ, Mobley HLT, Shirtliff ME. Complicated catheterassociated urinary tract infections due to Escherichia coli and Proteus mirabilis. Clin Microbiol Rev 2008;21:26-59. 4. Stickler DJ. Bacterial biofilms and the encrustation of urethral catheters. Biofouling 1996;94:293-305. 5. Ohkawa M. Bacterial adherence to Foley urinary catheters. Int Urogynecol J 1991;2:236-41. 6. Donlan RM. Biofilms and device-associated infections. Emerg Infect Dis 2001;7: 277-81. 7. Tenke P, Riedl CR, Jones GL, Williams GJ, Stickler D, Nagy E. Bacterial biofilm formation on urologic devices and heparin coating as preventive strategy. Int J Antimicrob Agents 2004;23:67-74. 8. Costerton JW, Stewart PS, Greenberg EP. Bacterial biofilms: a common cause of persistent infections. Science 1999;284:1318-22. 9. Fux AC, Stoodley P, Hall-Stoodley L, Costerton JW. Bacterial biofilms: a diagnostic and therapeutic challenge. Expert Rev Anti-Infect Ther 2003;4:667-83. 10. Farsi HMA, Mosli HA, Al-Zemaity MF, Bahnassy AA, Alvarez M. Bacteriuria and colonization of double-pigtail ureteral stents: long-term experience with 237 patients. J Endourol 1995;9:469-72.
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11. Gorman SP, Adair CG, Mawhinney WM. Incidence and nature of peritoneal catheter biofilm determined by electron and confocal laser scanning microscopy. Epidemiol Infect 1994;112:551-9. 12. Monsen T, Lovgren E, Widerstrom M, Wallinder L. In vitro effect of ultrasound on bacteria and suggested protocol for sonication and diagnosis of prosthetic infections. J Clin Microbiol 2009;47:2496-501. 13. Pruthi V, Al-Janabi A, Pereira BMJ. Characterization of biofilm formed on intrauterine devices. Indian J Med Microbiol 2003;21:161-5. 14. Brigitte C, Cerf O. Biofilms and their consequences, with particular reference to hygiene in the food industry. J Appl Bacteriol 1993;75:499-511. 15. Donlan RM, Murga R, Bell M, Toscano CM, Carr JH, Novicki TJ, et al. Protocol for detection of biofilms on needleless connectors attached to central venous catheters. J Clin Microbiol 2001;39:750-3. 16. Murga R, Miller JM, Donlan RM. Biofilm formation by gram-negative bacteria on central venous catheter connectors: effect of conditioning films in a laboratory model. J Clin Microbiol 2001;39:2294-7. 17. Storti A, Pizzolitto AC, Pizzolitto EL. Detection of mixed microbial biofilms on central venous catheters removed from intensive care unit patients. Brazil J Microbiol 2005;36:275-80. 18. Taiwo SS, Aderounmu AOA. Catheter-associated urinary tract infection: aetiologic agents and antimicrobial susceptibility pattern in Ladoke Akintola University Teaching Hospital, Osogbo, Nigeria. Afr J Biomed Res 2006;9: 141-8. 19. Tomaras AP, Dorsey CW, Edelmann RE, Actis LA. Attachment to and biofilm formation on abiotic surfaces by Acinetobacter baumannii: involvement of a novel chaperone-usher pili assembly system. Microbiology 2003;149: 3473-84. 20. Cormio L, La Forgia P, La Forgia D, Siitonen A, Ruutu M. Is it possible to prevent bacterial adhesion onto ureteric stents? Urol Res 1997;25:213-6. 21. Hauer T, Lacour M, Gastmeier P, Schulgen G, Schumacher R, Ruden H, et al. Nosocomial infections in intensive care units: a nationwide prevalence study. Anaesthesist 1996;45:1184-91. 22. Karlowsky JA, Jones ME, Draghi DC, Thornsberry C, Sahm DF, Volturo GA. Prevalence and antimicrobial susceptibilities of bacteria isolated from blood cultures of hospitalized patients in the United States in 2002. Ann Clin Microbiol Antimicrob 2004;3:1-8. 23. Merritt K, Hitchins VM, Brown SA. Safety and cleaning of medical materials and devices. New York [NY]: Wiley; 2000. p. 131-6. 24. Nickel JC, Gristina P, Costerton JW. Electron microscopic study of an infected Foley catheter. Can J Surg 1985;28:50-2. 25. Characklis WG, Marshall KC. Biofilms: a basis for an interdisciplinary approach. In: J. Wiley, editor. Biofilms. New York [NY]: Wiley; 1990. p. 3-15. 26. Gordona NC, Warehama DW. Multidrug-resistant Acinetobacter baumannii: mechanisms of virulence and resistance. Int J Antimicrob Agents 2010;35: 219-26. 27. Lee HW, Kah YM, Kim J, Lee JC, Seol SY, Cho DT, et al. Capacity of multidrugresistant clinical isolates of Acinetobacter baumannii to form biofilm and adhere to epithelial cell surfaces. Clin Microbiol Infect 2008;14:49-54. 28. Moura Costa GF, Bronharo TMC, Cardoso CL, Carrara-Marrone FE, Garcia LB. Preliminary evaluation of adherence on abiotic and cellular surfaces of Acinetobacter baumannii strains isolated from catheter tips. Brazil J Infect Dis 2006; 10:346-51. 29. Morris NS, Stickler DJ, McLean RJC. The development of bacterial biofilms on indwelling urethral catheters. World J Urol 1999;17:345-50.
Contents lists available at ScienceDirect
American Journal of Infection Control
American Journal of Infection Control
journal homepage: www.ajicjournal.org
Major article
Characterization of bacterial biofilms formed on urinary catheters Ryad Djeribi PhD a, *, Warda Bouchloukh MSc a, Thierry Jouenne PhD b, Bouzid Menaa PhD a, c a
Biofilms and Biocontamination of Materials Laboratory, Faculty of Science, Badji Mokhtar University, Annaba, Algeria UMR 6522 CNRS, Faculty of Science of Rouen, Mont Saint Aignan, France c Fluorotronics, Inc., Vista, CA b
Key Words: Acinetobacter baumannii SEM
Background: The formation of bacterial biofilms on urinary catheters is a leading cause of urinary tract infections in intensive care units. Cytobacteriological examination of urine from patients is often misleading, due to the formation of these biofilms. Therefore, characterizing these biofilms and identifying the bacterial species residing on the surface of catheters are of major importance. Methods: We studied the formation of biofilms on the inner surface of urinary catheters using microbiological culture techniques, with the direct contact of catheter pieces with blood agar. The bacterial species on the surface were characterized by scanning electron microscopy, and the kinetic profile of biofilm formation on a silicone substrate for an imipenem-resistant Acinetobacter baumannii bacterium was evaluated with a crystal violet staining assay. Results: The bacterial species that constituted these biofilms were identified as a variety of gramnegative bacilli, with a predominance of strains belonging to Pseudomonas aeruginosa. The other isolated strains belonged to A baumannii and Klebsiella ornithinolytica. Kinetic profiling of biofilm formation identified the transient behavior of A baumannii between its biofilm and planktonic state. This strain was highly resistant to all of the antibiotics tested except colistin. Scanning electron microscopy images showed that the identified isolated species formed a dense and interconnected network of cellular multilayers formed from either a single cell or from different species that were surrounded and enveloped by a protective matrix. Conclusions: Microbiological analysis of the intraluminal surface of the catheter is required for true identification of the causative agents of catheter-associated urinary tract infections. This approach, combined with a routine cytobacteriological examination of urine, allows for the complete characterization of biofilm-associated species, and also may help prevent biofilm formation in such devices and help guide optimum antibiotic treatment. Copyright Ó 2012 by the Association for Professionals in Infection Control and Epidemiology, Inc. Published by Elsevier Inc. All rights reserved.
Bacterial biofilms are able to colonize the inner surfaces of indwelling urinary catheters recovered from patients in intensive care units (ICUs) and can cause infectious diseases. Placement of urinary catheters is the primary factor responsible for the development of nosocomial urinary tract infections (UTIs).1,2 Long-term use of urinary catheters can lead to a quasi-permanent bacterial colonization of the urine and, as a reservoir of microorganisms, trigger nosocomial infections.3 In fact, 25 % of patients undergoing * Address correspondence to Ryad Djeribi, BP 12 Annaba RP, 23000 Annaba, Algeria. E-mail address: [email protected] (R. Djeribi). Supported by Grants-in-Aid from the Algerian Ministry of Education and Scientific Research (Direction of Scientific Research and Technological Development). Conflict of interest: None to report.
short-term urinary catheterization (<7 days) acquire bacteria, with an increasing daily risk of developing an infection of w5%. The estimated likelihood of developing an infection after long-term catheterization (30 days) is 100%.4 When inserted for long periods, urinary catheters may readily acquire biofilms, composed of gram-positive or gram-negative bacteria, on their inner and/or outer surfaces.5,6 The initial event in the mechanism of bacterial adhesion is the deposition of urinary components on the surface of the catheter, leading to the formation of a protein film. This protein film enhances the adhesion of microorganisms, leading to biofilm formation.2,7 This biofilm poses a public health problem for patients who need these medical devices. Treatment with antibiotics becomes difficult, and in some cases ineffective, because of the continuous swarming of planktonic bacteria resulting from biofilm formation.8,9
0196-6553/$36.00 - Copyright Ó 2012 by the Association for Professionals in Infection Control and Epidemiology, Inc. Published by Elsevier Inc. All rights reserved. doi:10.1016/j.ajic.2011.10.009
R. Djeribi et al. / American Journal of Infection Control 40 (2012) 854-9
Furthermore, the formation of biofilms can cause unrealistic and misleading results from cytobacteriological examination of urine (CBEU) in a patient undergoing urinary catheterization. Detecting biofilms before the appearance of a UTI may be difficult. Thus, there is a need to characterize these biofilms and identify the bacterial species on surfaces. Detection and proper identification requires total detachment of adherent bacteria before cultivation of the bacterial suspension.10 Most identification and analysis of bacteria starts with ultrasound application to release the cells from the urinary catheter surface. However, this mechanical process might not be sufficiently efficient for obtaining all of the important information about the bacterial species, and may lead to cellular damage. Indeed, many previous studies11-13 have applied the ultrasound method to detach bacterial biomass from surfaces to culture the bacterial suspension; however, this method appears to limit the complete characterization of biofilms formed on catheter surfaces because of the lethal action of the ultrasound on some of the species initially present in the biofilms.14 Thus, new, reliable techniques are needed to better understand and control biofilms on indwelling medical devices. In the present study, we aimed to detect, identify, and characterize biofilms formed on urinary catheter surfaces directly using microbiological culture techniques, and then analyze these biofilms by scanning electron microscopy (SEM). Rather than using the ultrasound method, we performing culture testing by cutting discs of urinary catheter pieces and depositing them directly onto blood agar. This process is direct, efficient, simple, and reliable and allows identification of all bacteria present. From these cultures, we identified the presence of both monobacterial and polybacterial assemblages. In addition, SEM analysis of the contaminated catheter pieces enabled us to identify the isolated species and their organization. Along with the culture tests, we assessed the kinetic profile of biofilm formation using crystal violet staining assays on a silicone substrate for imipenem-resistant Acinetobacter baumannii. We identified the transient behavior of the bacterium between the biofilm (sessile) state and planktonic (free) state, demonstrating that bacterial species can pass from one state to another depending on different environments. This behavior can modify the results of any potential CBEU. Moreover, by directly testing the contaminated catheter disks, we were able to test the response of A baumannii to a series of 9 antibiotic agents. The results reported in this article are significant and demonstrate the efficacy of our novel approach for identifying and characterizing bacterial biofilms in urinary catheters used in ICU patients, which can be the source of serious catheter-associated UTIs. Our findings also underscore the importance of ensuring direct contact between contaminated urinary catheter samples and the agar surface to fully characterize all biofilm-associated species and help guide the appropriate antibiotic therapy for potential UTIs.
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Table 1 Bacterial strains identified from catheter samples Patient characteristics
Urinary catheter
Sex
UC- 1 UC- 2
Male Male
18 68
Brain abscess; meningitis Hemorrhagic stroke
UC- 3
Male
52
Prostatitis
Age, years
Pathology
Bacterial isolate(s) P aeruginosa P aeruginosa K ornithinolytica A baumannii P aeruginosa K ornithinolytica
a solution of sodium hypochlorite,15,16 followed by successive washes of the inner surface with jets of sterile distilled water using a syringe with the tip of the needle at the upper end of the catheter. After this treatment, the catheter was carefully and aseptically cut into 3- to 4-mm thick discs. Three or 4 discs were placed on the surface of blood agar plates,17 and the inoculated plates were then incubated at 37 C for 48-72 hours. Isolation and identification of bacterial strains After 48 hours of incubation, bacterial colonies observed on both the cultured discs and their neighborhood were carefully collected as individual samples. Each colony was resuspended in sterile physiological water. Dilutions were prepared from each suspension and grown on standard media tryptic soy agar, blood agar, and cystine lactose electrolyte- deficient medium agar, the latter of which is widely used to study the bacteria responsible for UTIs.18 The inoculated plates were then incubated aerobically at 37 C for 24 hours. The purity of the bacterial colonies was checked, and the bacterial species found on the culture plates were identified by macroscopic, microscopic, and biochemical tests, including an oxidase test and a catalase test using API 20 E and API 20 NE (bioMérieux, Marcy L’Etoile, France). The bacterial strains identified by the bacteriological analysis of urinary catheters from ICU patients are listed in Table 1. Quantification of biofilm formation and kinetics of adhesion on silicone The kinetics of the formation of the biofilm on silicone was assayed using the standard crystal violet staining method.19 In brief, bacteria were grown in Luria-Bertani broth. After incubation, unattached cells were removed by rinsing the support thoroughly with water, and attached cells were stained with 1% crystal violet for 20 min. The dye bound to the adherent cells was then solubilized with ethanol-acetone (75:15, v/v) and the optical density of the solution was measured at 570 nm. Antibiotic resistance
MATERIALS AND METHODS
Urinary catheters used by hospitalized ICU patients for more than 1 week were carefully collected under aseptic conditions. The catheters were placed individually in sterile glass bottles and transported immediately to the laboratory for analysis.
An antibiotic resistance test of the isolated bacteria strains against selected antibiotics was performed using the standard protocol for dissemination of antimicrobial agents on MuellerHinton agar. This test was used to determine the resistance patterns of the isolated strains to 9 commonly used antibiotics: penicillin G, ticarcillin, cefixime, imipenem, gentamicin, amikacin, tobramycin, colistin, and trimethoprim/sulfamethoxazole.
Treatment of medical devices
Biofilm observation by SEM
There is currently no standardized method for the detection and analysis of biofilms on urinary catheter surfaces. The treatment of these catheters consisted of disinfection of the outer surface with
The bacterial biofilms colonizing the intraluminal surfaces of the urinary catheters were examined by SEM analysis. Sterile silicone Foley catheters were cut into 2-mm-thick discs, which were
Sample preparation
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Fig 1. Bacterial cultures obtained from the direct vicinity of the disks incubated on blood agar.
aseptically introduced into tubes containing nutrient broth that had been inoculated with various bacterial cultures obtained from Mueller-Hinton agar. After homogenization, the cultures were incubated at 37 C. After 1 week, the tubes were cleaned of bacterial cultures, and the discs were washed extensively with sterile distilled water before SEM analysis. Samples were fixed in a 2% glutaraldehyde and 0.1 M cacodylate buffer (pH 7.4) for 30 min and then rinsed three times for 10 minutes each rinse in 0.2 M cacodylate buffer (pH 7.4). Samples were then dehydrated by passing them through the following ethanol series: 30%, 50%, and 80% ethanol, each for 10 minutes, followed by 100% ethanol twice for 10 minutes each time. Support samples were then dried at 37 C for 24 hours. After being coated with gold-palladium (via sputter coating), samples were examined under a Cambridge S200 scanning electron microscope (Cambridge Instruments, LEO Electron Microscopy, Inc., Thornwood, NY). RESULTS Identification of biofilm-associated organisms In this study, we collected urinary catheters from patients catheterized for longer than 1 week. Only 3 catheters were selected for analysis, and only one of these was fully analyzed due to the diversity of infection and the presence of multiple potential pathogens typically found in this patient community. Discs were cut from these urinary catheters and cultured on blood agar. Dense bacterial growth was observed around the discs (Fig 1). Macroscopic and microscopic examination of cultures from the inner surface of the catheters revealed that the bacterial masses were composed of both single species (monobacterial biofilms) and a mixture of several species (polybacterial biofilms). The isolated and identified bacterial species that constituted the biomass were mainly varieties of gram-negative bacilli, with a predominance of strains identified as P aeruginosa and other strains identified as A baumannii and Klebsiella ornithinolytica. Kinetics of biofilm formation The kinetic profile of biofilm formation on silicone was assayed for imipenem-resistant A baumannii over a 1-week period using the crystal violet staining method. The results demonstrate the transient behavior of the bacterium between the biofilm (sessile state) and the planktonic state (free bacteria) (Fig 2). These results also show that biofilm formation on the media occurred constantly and continuously despite the cellular release observed during incubation. Based on the observed curve, the kinetic profile is characterized by 2 specific periods during the first 96 hours of incubation, with the first period corresponding to significant biofilm formation and the second period corresponding to biofilm dispersion, with release of bacterial cells into the culture medium.
Fig 2. Biofilm of A baumannii formed on silicone material. (A) Kinetics of biofilm formation of A baumannii. (B) Growth of A baumannii (planktonic state).
In addition, the kinetic profile of biofilm formation on silicon for A baumannii suggests the importance of strong hydrophobicity of the bacterial strain at urine pH and the biofilms’ adhesive strength on such a hydrophobic surface. Our findings demonstrate the transient behavior of the bacterium between the biofilm formed in the inner surface of the catheter and the planktonic state. This behavior indicates that bacterial species are able to pass from one state to another depending on the environment. Thus, this transient behavior may have a significant affect on the results of any potential CBEU. Antibiotic resistance The results of the susceptibility and resistance tests performed using the biofilm bacteria and commonly used antibiotics showed
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Fig 3. SEM images of the biofilms formed in the urinary catheter. (A-C) A baumannii biofilms. (Original magnifications: A, 1,000; B, 3,000; C, 10,000.) (D and E) Biofilms composed of P aeuroginosa and A baumannii. (Original magnification, 3,000.) (F) Biofilms composed of P aeuroginosa, A baumannii, and K ornithinolytica. (Original magnification, 3,000.)
that all of the isolated bacteria (P aeruginosa, A baumannii, and K ornithinolytica) are resistant to penicillin G, cefixime, and trimethoprim/sulfamethoxazole and sensitive to colistin. However, the resistance profiles obtained for the A baumannii strain revealed resistance to the full range of antibiotics tested except colistin. A baumannii is known to be resistant to imipenem, which is widely used to treat infections caused by this bacterium. Of note, an A baumannii strain was isolated from a mixed biofilm formed on the inner surface of a catheter recovered from a 68-year-old patient admitted to the ICU for a cerebrovascular accident. The patient had a history of persistent urinary infection with no apparent symptoms. Bacteriological examinations of urine samples obtained from the catheter pocket revealed the presence of P aeruginosa. In addition, analysis of the urinary catheter surface revealed the
presence of 4 bacterial strainsdA baumannii, Klebsiella spp, and 2 strains of P aeruginosadwith significantly different antibiograms. Biofilm characterization by SEM SEM analysis of the biofilms formed on the inner surface of the catheter samples revealed a dense network of cellular multilayers, formed either from a single cell (Fig 3A-C) or from different species (Fig 3D-F), surrounded and enveloped by a protective matrix. SEM images indicate that the identified isolated species readily developed compatible and strong associations with other species and formed agglomerates on the inner catheter surfaces. The enveloping matrix is composed of extracellular polymeric substances, mainly polysaccharides, which constitute most of the carbon
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organic mass of the biofilm (Fig 3C). This matrix confers resistance to antimicrobial agents and constitutes a reservoir of viable microorganisms.6,20 The biofilms are characterized by a multitude of intercellular pores and interconnected channels, which allow not only the exchange of information and the flow of water and ions, but also the transport of nutrients and the efflux of waste products.
DISCUSSION All of the previous studies on the bacterial biomass responsible for nosocomial infections have reported that gram-negative bacilli are predominant in UTIs.21 In addition, other reports indicate that the strains of P aeruginosa and A baumannii identifed are among the most nonfermented bacterial species frequently isolated from clinical specimens of hospitalized patients.17,22 However, the pathogenicity of these bacterial species is low, and their virulence is expressed only in patients with impaired immune defenses and those requiring invasive devices. Numerous techniques have been developed to detect and characterize biofilm-associated cells on medical devices. Among these is the roll-plate technique, in which infected pieces of catheter are rolled over the surface of a nonselective medium. Characterization and quantification of the biofilm depend on the number of organisms recovered by contact with the agar surface.1 Another method is sonication plus vortexing to detach adherent microorganisms. The developed colonies are allowed to recover after dilution and plating of the resulting suspension on culture media.2 The foregoing methods have limited recovery efficiency, however. The microbiological culture method used in this study involves a longer period of contact between contaminated catheter pieces and the blood agar surface. This method allows the recovery and characterization of most of the bacteria found in polybacterial biofilms with the ability to colonize the inner surfaces of urinary catheters. Thus, unlike methods based on detaching the bacterial biomass with ultrasound, our method allows the identification and complete characterization of all bacterial species present. Ultrasound can damage some cells or even kill certain species initially present in biofilm. Thus, our proposed technique appears better suited to the decontamination of materials compared with the characterization of bacterial biomass fixed on the supports.23 Identification and complete characterization of biofilm is vitally important. The CBEU of urine obtained from contaminated catheters may not necessarily reflect actual UTI status. A sample may be sterile urine with microorganisms located in the luminal catheter biofilm, which could contaminate all urine samples obtained for analysis.24 SEM analysis is important as well. In one patient, SEM images of the inner surface of an indwelling urinary catheter that had been in place for 1 month revealed the presence of biofilms that included Staphylococcus epidermidis within a glycocalyx, despite the fact that CBEU analysis during ablation showed that the urine was free of bacteria.25 According to our findings, CBEU might not reveal all of the bacterial species present in the planktonic state when urine is collected during the initial period (0-24 hours), which coincides with the period of maximum bacterial adhesion on the catheter surface. Therefore, the kinetic profile of biofilm formation and characterization of the transient behavior of A baumannii are significant, because they provide information on the planktonic and biofilm states, allowing identification of the ideal time frame for CBEU. As shown in Figure 2, patients are more likely to develop nosocomial UTIs in the absence of a systematic review and analysis of urinary catheters. Thus, CBEU should be performed at least every 48 hours after placement of the first catheter, to identify the
bacterial cells released from the biofilm matrix formed on the catheter surfaces. In hospitals and other medical institutions, A baumannii has emerged as a significant and problematic human pathogen. This bacterium has a capacity to quickly acquire antimicrobial resistance,26 but also readily adheres to both biological and abiotic surfaces.27,28 Moreover, it has developed a varied pattern of resistance, especially to carbapenems (eg, imipenem), the drugs of choice for treating serious A baumannii infections. Thus, treatment options for these infections appear to be very limited. In this study, we analyzed an imipenem-resistant strain of A baumannii to examine the behavior of such bacteria in immunodeficient patients. The resistance profile revealed a remarkable resistance to all of the antibiotic agents tested, along with a significant sensitivity to colistin. Thus, colistin appears to be an effective antibiotic for treating A baumannii infections in either monotherapy or combination therapy. Our findings may provide useful insight into the nature and effective treatment of biofilmassociated infections caused by this strain. SEM analysis suggests a heterogeneous mosaic architecture of biofilms containing microcolonies of bacterial cells encased in an extracellular polymeric substances matrix and separated from other microcolonies by interstitial voids. This microstructure is similar to that commonly observed in other sessile bacteria, including P aeruginosa. This structural organization shows a dense network of cellular multilayers formed from different species that are surrounded and enveloped by a protective matrix, which constitutes a reservoir of viable microorganisms. This organization allows microorganisms to grow under adverse conditions13,17 and is typical of biofilms found on medical implants and devices (eg, intrauterine devices, central venous catheters). In a study by Morris et al,29 SEM analysis revealed that most of the catheters removed after 1 week of insertion were colonized by bacterial biofilms, particularly on the catheters’ luminal surfaces. These observations also demonstrate that different species can be readily integrated and develop close associations in biofilms. In this study, we investigated and described the different aspects of association of A baumannii in pure and mixed biofilms. The pure culture biofilm shown in Figure 3A is composed of isolated coccobacilli cells, grouped in pairs or with multiple channels of various lengths embedded in a simile matrix. However, the communities shown in Figure 3B have different multilayers formed by A baumannii cells clustered together and embedded in a dense biofilm. This dense biofilm is crossed with a multitude of pores and visible intercellular channels to ensure bacterial exchanges. Figure 3C shows a mature biofilm with a large amount of glycocalyx obscuring the shape of the organism. The adhesive matrix that developed appears as a network around a group of cells that ensures protection for this bacterium. Taken together, the results shown in Figure 3D and E suggest a strong and close association between the A baumannii and P aeruginosa strains analyzed in this study. In these biofilms, the colonies are intermixed, and the communities are embedded in heterogeneous abundant extracellular polymeric substances. The structural complexity, characterized by towers and mushroom-like shapes, gives volume to the biofilm. These analyses not only are descriptive, but also show that these species are isolated together from specimens of hospitalized patients, and that the interactions between these bacteria have the potential to pose a serious barrier to the effective treatment of nosocomial infections. CONCLUSION The results reported in this article are novel and significant. They demonstrate the efficacy of our approach for identifying and
R. Djeribi et al. / American Journal of Infection Control 40 (2012) 854-9
characterizing bacterial biofilms formed on urinary catheters in daily use in ICU patients, which can be the source of serious catheter-associated UTIs. The use of indwelling urinary catheters in daily urological practice remains widespread. The most common infection associated with these catheters is nosocomial infection due to the formation of bacterial biofilms on the catheter surface. This biofilm formation is accompanied by the risk of spreading planktonic microorganisms, which can cause serious nosocomial infections related the difficulty in detecting the biofilm even before the appearance of UTI. The classical methods of bacteriological diagnosis of a UTI, such as CBEU, often yield misleading or incorrect results when evaluating the presence of a biofilm on the surface of a medical device. The present study demonstrates that the proper identification of infectious agents responsible for UTI often requires microbiological analysis of the inner catheter surfaces in addition to routine bacteriological examination. This method allows for identification and complete characterization of the bacterial species forming the biofilm, which can aid in the detection and prevention of biofilm formation and guide appropriate antibiotic therapy for such devicerelated infections. References 1. Trautner BW, Darouiche RO. Role of biofilm in catheter-associated urinary tract infection. Am J Infect Control 2004;32:177-83. 2. Hatt JK, Rather PN. Role of bacterial biofilms in urinary tract infections. Curr Top Microbiol Immunol 2008;322:163-92. 3. Jacobsen SM, Stickler DJ, Mobley HLT, Shirtliff ME. Complicated catheterassociated urinary tract infections due to Escherichia coli and Proteus mirabilis. Clin Microbiol Rev 2008;21:26-59. 4. Stickler DJ. Bacterial biofilms and the encrustation of urethral catheters. Biofouling 1996;94:293-305. 5. Ohkawa M. Bacterial adherence to Foley urinary catheters. Int Urogynecol J 1991;2:236-41. 6. Donlan RM. Biofilms and device-associated infections. Emerg Infect Dis 2001;7: 277-81. 7. Tenke P, Riedl CR, Jones GL, Williams GJ, Stickler D, Nagy E. Bacterial biofilm formation on urologic devices and heparin coating as preventive strategy. Int J Antimicrob Agents 2004;23:67-74. 8. Costerton JW, Stewart PS, Greenberg EP. Bacterial biofilms: a common cause of persistent infections. Science 1999;284:1318-22. 9. Fux AC, Stoodley P, Hall-Stoodley L, Costerton JW. Bacterial biofilms: a diagnostic and therapeutic challenge. Expert Rev Anti-Infect Ther 2003;4:667-83. 10. Farsi HMA, Mosli HA, Al-Zemaity MF, Bahnassy AA, Alvarez M. Bacteriuria and colonization of double-pigtail ureteral stents: long-term experience with 237 patients. J Endourol 1995;9:469-72.
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11. Gorman SP, Adair CG, Mawhinney WM. Incidence and nature of peritoneal catheter biofilm determined by electron and confocal laser scanning microscopy. Epidemiol Infect 1994;112:551-9. 12. Monsen T, Lovgren E, Widerstrom M, Wallinder L. In vitro effect of ultrasound on bacteria and suggested protocol for sonication and diagnosis of prosthetic infections. J Clin Microbiol 2009;47:2496-501. 13. Pruthi V, Al-Janabi A, Pereira BMJ. Characterization of biofilm formed on intrauterine devices. Indian J Med Microbiol 2003;21:161-5. 14. Brigitte C, Cerf O. Biofilms and their consequences, with particular reference to hygiene in the food industry. J Appl Bacteriol 1993;75:499-511. 15. Donlan RM, Murga R, Bell M, Toscano CM, Carr JH, Novicki TJ, et al. Protocol for detection of biofilms on needleless connectors attached to central venous catheters. J Clin Microbiol 2001;39:750-3. 16. Murga R, Miller JM, Donlan RM. Biofilm formation by gram-negative bacteria on central venous catheter connectors: effect of conditioning films in a laboratory model. J Clin Microbiol 2001;39:2294-7. 17. Storti A, Pizzolitto AC, Pizzolitto EL. Detection of mixed microbial biofilms on central venous catheters removed from intensive care unit patients. Brazil J Microbiol 2005;36:275-80. 18. Taiwo SS, Aderounmu AOA. Catheter-associated urinary tract infection: aetiologic agents and antimicrobial susceptibility pattern in Ladoke Akintola University Teaching Hospital, Osogbo, Nigeria. Afr J Biomed Res 2006;9: 141-8. 19. Tomaras AP, Dorsey CW, Edelmann RE, Actis LA. Attachment to and biofilm formation on abiotic surfaces by Acinetobacter baumannii: involvement of a novel chaperone-usher pili assembly system. Microbiology 2003;149: 3473-84. 20. Cormio L, La Forgia P, La Forgia D, Siitonen A, Ruutu M. Is it possible to prevent bacterial adhesion onto ureteric stents? Urol Res 1997;25:213-6. 21. Hauer T, Lacour M, Gastmeier P, Schulgen G, Schumacher R, Ruden H, et al. Nosocomial infections in intensive care units: a nationwide prevalence study. Anaesthesist 1996;45:1184-91. 22. Karlowsky JA, Jones ME, Draghi DC, Thornsberry C, Sahm DF, Volturo GA. Prevalence and antimicrobial susceptibilities of bacteria isolated from blood cultures of hospitalized patients in the United States in 2002. Ann Clin Microbiol Antimicrob 2004;3:1-8. 23. Merritt K, Hitchins VM, Brown SA. Safety and cleaning of medical materials and devices. New York [NY]: Wiley; 2000. p. 131-6. 24. Nickel JC, Gristina P, Costerton JW. Electron microscopic study of an infected Foley catheter. Can J Surg 1985;28:50-2. 25. Characklis WG, Marshall KC. Biofilms: a basis for an interdisciplinary approach. In: J. Wiley, editor. Biofilms. New York [NY]: Wiley; 1990. p. 3-15. 26. Gordona NC, Warehama DW. Multidrug-resistant Acinetobacter baumannii: mechanisms of virulence and resistance. Int J Antimicrob Agents 2010;35: 219-26. 27. Lee HW, Kah YM, Kim J, Lee JC, Seol SY, Cho DT, et al. Capacity of multidrugresistant clinical isolates of Acinetobacter baumannii to form biofilm and adhere to epithelial cell surfaces. Clin Microbiol Infect 2008;14:49-54. 28. Moura Costa GF, Bronharo TMC, Cardoso CL, Carrara-Marrone FE, Garcia LB. Preliminary evaluation of adherence on abiotic and cellular surfaces of Acinetobacter baumannii strains isolated from catheter tips. Brazil J Infect Dis 2006; 10:346-51. 29. Morris NS, Stickler DJ, McLean RJC. The development of bacterial biofilms on indwelling urethral catheters. World J Urol 1999;17:345-50.