Biofilms: survival mechanisms of clinically relevant microorganisms

Abstract

Though biofilms were first described by Antonie van Leeuwenhoek, the theory describing the biofilm process was not developed until 1978. We now understand that biofilms are universal, occurring in aquatic and industrial water systems as well as a large number of environments and medical devices relevant for public health. Using tools such as the scanning electron microscope and, more recently, the confocal laser scanning microscope, biofilm researchers now understand that biofilms are not unstructured, homogeneous deposits of cells and accumulated slime, but complex communities of surface-associated cells enclosed in a polymer matrix containing open water channels. Further studies have shown that the biofilm phenotype can be described in terms of the genes expressed by biofilm-associated cells. Microorganisms growing in a biofilm are highly resistant to antimicrobial agents by one or more mechanisms. Biofilm-associated microorganisms have been shown to be associated with several human diseases, such as native valve endocarditis and cystic fibrosis, and to colonize a wide variety of medical devices. Though epidemiologic evidence points to biofilms as a source of several infectious diseases, the exact mechanisms by which biofilm-associated microorganisms elicit disease are poorly understood. Detachment of cells or cell aggregates, production of endotoxin, increased resistance to the host immune system, and provision of a niche for the generation of resistant organisms are all biofilm processes which could initiate the disease process. Effective strategies to prevent or control biofilms on medical devices must take into consideration the unique and tenacious nature of biofilms. Current intervention strategies are designed to prevent initial device colonization, minimize microbial cell attachment to the device, penetrate the biofilm matrix and kill the associated cells, or remove the device from the patient. In the future, treatments may be based on inhibition of genes involved in cell attachment and biofilm formation.

Figures

FIG. 1.

Scanning electron micrograph of a biofilm on a metal surface from an industrial water system.

FIG. 2.

Confocal laser scanning micrograph of a biofilm, showing cell clusters and water channels. Reproduced with the permission of Paul Stoodley.

FIG. 3.

Mixed-species heterotrophic biofilm grown on stainless steel in a potable-water biofilm reactor containing Pseudomonas aeruginosa, Klebsiella pneumoniae, and Flavobacterium spp. This image of a biofilm was obtained, after staining with 4′,6′-diamidino-2-phenylindole, with a Zeiss Axioskop 2 epifluorescence microscope and the Zeiss deconvolution system.

FIG. 4.

Biofilm structure cartoon. Copyright Center for Biofilm Engineering, Montana State University, Bozeman, Mont. Reprinted with permission.

FIG. 5.

Transmission electron micrograph of a prostatic duct in an area of focal chronic inflammation from a patient with an E. coli chronic prostatitis. Arrows point to bacterial microcolonies amid inflammatory cells and debris. These bacteria were cultured from both expressed prostatic secretions and tissue biopsies obtained 4 weeks after antibiotics were discontinued. Bar, 1 μm. Reprinted from reference 151 with permission of Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc.

FIG. 6.

(A) Cut section of a urinary catheter collected from a patient, revealing a worm-like structure occluding the lumen; (B) low-power scanning electron micrograph of a freeze-fractured cross-section of a blocked catheter; (C) crystalline formations on the outer surface of a freeze-dried preparation of material blocking the catheter; (D) fixed and critical-point-dried specimen showing that, below their crystalline coats, the catheter casts are composed of a mass of cocci and bacilli. Reprinted from reference 190 with permission of the publisher (W. B. Saunders).

FIG. 7.

Effect of acetohydroxamic acid, a urease inhibitor, on the encrustation of silicone catheters by Proteus mirabilis biofilms. Each value is the mean calculated from three replicated experiments. ∗∗, significant difference (P < 0.01) from the control values (analysis of variance). The mean values for the log of the number of viable cells per milliliter of urine at 24 h were 8.02 (control), 8.16 (0.01 mg of acetohydroxamic acid per ml), 8.20 (0.5 mg/ml), and 8.09 (1.0 mg/ml). Reprinted from reference 141 with permission of Springer-Verlag Co.

FIG. 8.

(A) Effect of a low-strength electric field with a low current density followed by biocide application (arrows) on P. aeruginosa colonization (mean, n = 2). At 24 h, glutaraldehyde (5 ppm) (open and solid squares) or kathon (1 ppm) (open and solid triangles) was applied to both electrified and control devices. (B) Effect of biocides on an established (24-h) P. aeruginosa biofilm in the presence and absence of a low-strength electric field with a low current density. Glutaraldehyde (5 ppm) (open and solid squares) or quaternary ammonium compound (10 ppm) (open and solid diamonds) was supplied to both electrified and control devices for 24 h (mean, n = 2). The electrified devices are represented by solid symbols. Reprinted from reference 17 with permission of the American Society for Microbiology.