Correspondence Cardiff School of Biosciences, Main Building, Cardiff University, Museum Avenue, Cardiff CF10 3TL, UK

Email stickler@cardiff.ac.uk

Introduction

The biofilm mode of growth is a basic survival strategy deployed by bacteria in a wide range of environmental, industrial and clinical aquatic settings.¹ Bacterial cells have a strong preference for life on surfaces rather than in planktonic suspension.² These cells have an array of adhesins in their cell walls that allow them to colonize many types of substrate, and, on contact with a surface, the cells secrete exopolysaccharides that secure their attachment. The bacteria then multiply to form microcolonies of cells that subsequently spread over the surface, forming populations embedded in a gel-like polysaccharide matrix (Figure 1). The cells in these biofilm communities are protected from environmental stresses, and this protection has particular advantages for the bacteria in biofilms that develop in vivo. Microorganisms that are apparently fully sensitive to antibiotics and antiseptics in conventional laboratory testing methods become fully resistant in the biofilm mode in vivo.

Figure 1 Conceptualization of biofilm development and dynamic behaviors.

The figure was compiled from laboratory and natural observations of pure-culture (both Gram-positive and Gram-negative organisms) and mixed-culture biofilms. Image courtesy of P Dirckx, Center for Biofilm Engineering, USA. Permission obtained from Nature Publishing Group © Hall-Stoodley L et al. (2004) Nat Rev Microbiol 2: 95–108.

Full figure and legend (69K)Figures & Tables indexDownload Power Point slide (274K)

Catheter biofilms

Prolonged urinary tract infections can facilitate the development of catheter biofilms. While indwelling (Foley) catheters are effective in relieving urinary retention and managing urinary incontinence, external bacteria have easy access to the bladder, and catheterization can often result in bacteriuria. The risk of urinary tract infection is related to the length of time the catheter is in place. Most patients catheterized for a week or less should escape infection, but for the many elderly and disabled patients who are catheterized for several months or years, bacteriuria is inevitable.^{3, 4}

Urinary tract infections in catheterized patients can occur in several ways. Organisms that colonize the periurethral skin can migrate into the bladder through the mucoid film that forms between the epithelial surface of the urethra and the catheter. In addition, contamination of the urine in the drainage bag can allow organisms to access the bladder through the drainage tube and the catheter lumen.^{5, 6} The initial bacteria that cause the urinary tract infections are usually Staphylococcus epidermidis, Escherichia coli or Enterococcus faecalis.^{6, 7} As time goes by, other species appear in the residual bladder urine, including Pseudomonas aeruginosa, Proteus mirabilis, Providencia stuartii, Morganella morganii and Klebsiella pneumoniae.^{3, 8} The bacteria that present in the latter stages of urinary tract infection are difficult to eradicate with antibiotics while the catheter is in place.^{8, 9} As the infections are usually asymptomatic, and because of the danger of promoting antibiotic resistance, catheter-associated bacteriuria is generally not treated.^{10, 11, 12} In patients with long-term indwelling catheters, catheter changes are commonly scheduled at 10–12-week intervals; contaminated urine can, therefore, be flowing through individual catheters for periods of 3 months at a time. Thus, catheters provide attractive sites for bacterial colonization: the biofilm bacteria thrive in their matrix gel and the gentle flow of warm nutritious urine. Enormous populations develop, and become visible to the naked eye as thick coatings. Biofilms containing 5 10⁹ viable cells per centimeter can be found on long-term indwelling catheters removed from patients.¹³ The biofilm populations, therefore, often outnumber those in the urine.

A variety of bacterial species colonize catheters, and many of these biofilms can induce serious complications.^{10, 13, 14, 15, 16, 17}Table 1 summarizes the bacterial species identified from a set of 106 catheter biofilms;¹⁷ 14 species were commonly found. Isolated cases of single-species biofilms were observed, but most biofilms contained mixed bacterial communities containing up to five species. The most common species present in the mixed-population biofilms were E. faecalis, P. aeruginosa, E. coli, and P. mirabilis. In patients who develop bacteriuria during short-term catheterization, bacterial colonization of the catheter does occur.¹⁶ The biofilms formed are generally sparse, and because the catheter is removed within a few days, they cause few problems. By contrast, long-term catheters become colonized by extensive biofilms, which can have profound effects on the health of the patient. By far the most troublesome biofilms are those that become crystalline in nature.^{18, 19} These biofilms can form on the outer surface of the catheter around the balloon and catheter tip, and can cause trauma to the bladder and urethral epithelia. On deflation of the retention balloon, crystalline debris from the biofilm can be shed into the bladder and initiate stone formation. The main complication, however, is blockage in the flow of urine through the catheter that results from the build up of the crystalline material on the lumenal surfaces (Figure 2). As a consequence, urine often leaks along the outside of the catheter and patients become incontinent, resulting in the increased need for nursing assistance. In addition, blockage of the catheter can lead to retention of urine in the bladder and vesicoureteric reflux of infected urine; if the blockage is not detected and if the catheter is not changed, patients can suffer episodes of pyelonephritis and septicemia.^{15, 20}

Figure 2 Examples of crystalline biofilms on blocked catheters taken from patients.

(A) This image shows a catheter that had been indwelling suprapubically for 6 months. It was removed surgically. Crystalline material completely covered the eyehole and balloon of the hydrogel-coated latex catheter. Image kindly supplied by Professor Roger Feneley. (B) A cross-section of a silicone catheter that had been indwelling for 8 weeks. The image shows that the central lumen is occluded by crystalline biofilm. Permission obtained from Elsevier Ltd © Stickler DJ (1999) Eur Urol Update Series 5: 1–8. (C) A longitudinal section of a silver–hydrogel-coated latex catheter that blocked after 11 days in situ.

Full figure and legend (40K)Figures & Tables indexDownload Power Point slide (244K)

Table 1 The incidence of bacterial species isolated from 106 catheter biofilms.¹⁷

Full tableFigures & Tables indexDownload Power Point slide (252K)

About half the patients who undergo long-term catheterization will suffer the complication of catheter encrustation and blockage by bacterial biofilms at some time.^{21, 22, 23} The welfare of many elderly and disabled patients is thus put at risk by the development of these biofilms, and considerable demands are made on the resources of the health-care service to manage the complications. An insight into the scale of the problem was given by a prospective study of 467 patients in community care in the UK. Over a 6-month period, 506 emergency referrals were recorded for these patients, mostly to deal with catheter blockage.²³

Crystalline biofilms

The crystalline deposits on catheters have a similar composition to infection-induced kidney and bladder stones. Struvite (magnesium ammonium phosphate) and a poorly crystalline form of apatite (a hydroxylated calcium phosphate, in which a variable proportion of the phosphate groups are replaced by carbonate) are the principle crystalline components.^{24, 25} Scanning electron microscopy has shown that large numbers of bacilli are associated with the crystals (Figure 3).²⁶ Culture techniques have confirmed the persistence of a range of bacteria. Notably, species capable of producing the enzyme urease are predominantly associated with crystallization.²⁷ Urease is, in fact, the driving force of crystallization: it hydrolyzes urea, leading to the formation of ammonium and carbonate ions and an increase in urinary pH. As the urine becomes alkaline, magnesium and calcium phosphate crystals are precipitated. Aggregates of this crystalline material accumulate in the urine and in the biofilm that develops on the catheter surfaces. The continued accumulation of crystalline bacterial biofilm blocks the flow of urine through the catheter.^{14, 28}

Figure 3 Scanning electron micrographs of encrustation on the surface of a silver–hydrogel-coated latex catheter that had been indwelling for 14 days.

(A) This image shows the crystalline forms present in the biofilm. (B) In this higher-magnification image, the bacterial colonization of these crystals is illustrated.

Full figure and legend (36K)Figures & Tables indexDownload Power Point slide (240K)

Several species commonly found in catheter biofilms produce urease (Table 1). In laboratory tests, urease can be detected in P. aeruginosa, K. pneumoniae and M. morganii, Proteus species, including P. mirabilis, some Providencia species and some strains of Staphylococcus aureus and coagulase-negative staphylococci. Of these species, P. mirabilis is most commonly isolated from the urine of patients suffering from recurrent catheter encrustation and blockage.^{29, 30} Furthermore, P. mirabilis is also the species most commonly recovered from patients' encrusted catheters.²⁷ The urease produced by P. mirabilis is a potent enzyme, and is able to hydrolyze urea several times faster than urease produced by other species.³¹ Experimental work in laboratory models of the catheterized bladder has demonstrated that species such as M. morganii, K. pneumoniae, and P. aeruginosa fail to produce alkaline urine and do not produce appreciable encrustation on catheters.³² In this laboratory work, the only species capable of producing alkaline urine and causing extensive encrustation were P. mirabilis, Proteus vulgaris and Providencia rettgeri. As these latter two species are only found in about 5–10% of catheter biofilms,¹⁷ strong epidemiological and experimental evidence indicates that P. mirabilis is mainly responsible for the formation of crystalline biofilms on catheters.

Proteus mirabilis

P. mirabilis is not usually a pioneer colonizer of the catheterized urinary tract, and is not commonly found in patients undergoing short-term catheterization.⁷ The longer the catheter is in place, however, the more likely it is to be found in the urine. In patients undergoing long-term catheterization, P. mirabilis has been isolated from around 40% of urine samples.³³ Sabbuba et al.³⁴ have developed a technique for genotyping P. mirabilis to help understand the epidemiology and pathogenesis of P. mirabilis catheter-associated urinary tract infections. Pulsed-field gel electrophoresis of restriction enzyme digests of DNA from P. mirabilis produced highly discriminatory genotypic profiles. The application of this technique established the remarkable stability of P. mirabilis strains in the catheterized urinary tract. The same genotype persisted in a patient's urinary tract despite many catheter changes, courses of antibiotic treatment, and even periods when the patient was not catheterized. Further investigation revealed that P. mirabilis was also present in the bladder stones that frequently form in these patients. Genotyping of pairs of P. mirabilis isolates from the encrusted catheters and bladder stones from the same patient demonstrated that, in each case, the strain of P. mirabilis was identical.³⁵ Genotyping also showed that the majority of patients were infected with genetically distinct strains. P. mirabilis is an enteric organism, and subsequent analysis showed that bacteria from fecal and catheter biofilm isolates from the same patients were identical.³⁶ These findings indicate that most long-term catheterized patients who suffer from catheter encrustation probably acquire P. mirabilis from their own fecal flora. These strains will eventually cause chronic colonization of the urine, catheters and bladder stones.

The formation of proteus mirabilis biofilms

Physical factors

In addition to the biological factors, powerful physical forces can initiate the development of crystalline biofilms. Scanning electron microscopy has revealed the rough, irregular nature of catheter surfaces.⁴⁸ Latex-based catheters have particularly uneven surfaces.⁴⁹ The manufacturing techniques used to produce the eyeholes tear through the latex and produce surfaces that must seem like rocky landscapes of craters and crevices to bacteria (Figure 4). The roughness of the lumenal surfaces is exacerbated by the common occurrence of embedded diatom skeletons.⁵⁰ These skeletons come from the diatomaceous earth—a naturally occurring, soft, chalk-like sedimentary rock, which is easily crumbled into a fine powder—that is used to prevent the latex sticking to the metallic formers on which the catheters are produced. All-silicone catheters have smoother surfaces than latex catheters, but irregularities are still common around the eyeholes and where extrusion manufacturing techniques have produced striations on the lumenal surfaces.⁴⁹

Figure 4 Scanning electron micrographs of the surfaces of unused hydrogel-coated latex catheters.

(A) This image illustrates the rough surface produced by cutting the eyeholes. Permission obtained from Springer © Stickler DJ (2003) Urol Res 31: 306–311. (B) This higher-magnification image also shows the rough surface of the eye holes. (C) This high-magnification micrograph shows the craters and crevices produced in the latex around the eyelet. The silica skeletons of diatoms can be seen on the irregular lumenal surfaces. (D) This image shows the presence of the diatom skeletons on the luminal surface of the catheter.

Full figure and legend (48K)Figures & Tables indexDownload Power Point slide (252K)

Eyeholes are particularly vulnerable to bacterial colonization. In experiments where catheters were removed from bladder models at various intervals after urine inoculation with P. mirabilis, scanning electron microscopy revealed that, within 2 h, bacterial cells were trapped in the crevices in the uneven surfaces of the eyelets.⁴⁹ Microcolonies of cells developed in the surface depressions, and then, with the rise in urinary pH, crystals started to form in the biofilm (Figure 5). Extensive crystalline biofilm developed and spread down the catheter lumen. The silica skeletons of the diatoms embedded in the latex were also attractive sites for bacterial colonization.⁵⁰ Blockage with extensive crystalline biofilm generally occurred at the eyehole or in the balloon region of the lumen.

Figure 5 Electron micrographs illustrating the colonization of a hydrogel-coated latex catheter by Proteus mirabilis in a laboratory model of the bladder.

(A) This image shows bacteria trapped in crevices in the surface of the eyeholes 2 h after incubation in the model. (B) Microcolonies of P. mirabilis develop at the eyehole 4 h after incubation. (C) Bacteria attach to a diatom skeleton embedded in the lumenal surface of the catheter 6 h after incubation in the model. (D) Biofilm develops at the eyehole 6 h after incubation in the model. Aggregates typical of apatite can be seen forming in the biofilm as the urine becomes alkaline. Permission obtained from Springer © Stickler DJ (2003) Urol Res 31: 306–311

Full figure and legend (45K)Figures & Tables indexDownload Power Point slide (249K)

Chemical factors

The chemical environment also has a vital role in the development of crystalline biofilms. Brisset et al.⁵¹ reported in 1996 that hydrophobic cells were more likely to colonize hydrophobic than hydrophilic surfaces, and that colonization was increased in alkaline urine. Experiments in parallel-plate flow cells have also shown that when urine cultures flow over smooth, flat, polymer films, the pH of the urine can be a major factor in determining the extent to which bacteria adhere. For example, some polymers with strongly electron donating, hydrophilic surfaces will resist colonization by cells until the pH of the urine rises. In the alkaline urine, however, macroscopic aggregates of cells and crystals will form, settle on the polymer surface, and initiate crystalline biofilm formation.⁴⁶

The chemical environment can also affect the rate at which biofilms develop. A prospective study by Mathur et al.⁵² in patients infected with P. mirabilis revealed that the time taken for catheters to block varied from 2 to 98 days. This variation can be explained by the concept of the nucleation pH of urine (pHn). The pHn of a urine sample is the pH at which the urine becomes turbid, due to microcrystals of apatite and struvite coming out of the solution. The urine becomes turbid as the pH increases. Choong et al.⁵³ found that, for patients whose catheters were blocked by crystalline biofilms, the mean pHn of their urine was 7.58, while the mean pH of the voided urine was 7.85; these results clearly indicate that catheters become encrusted if the pH of the urine is greater than the pHn.

Further analysis⁵⁴ of the data from the study by Mathur et al.⁵² showed that, in patients infected with P. mirabilis, the pHn of the urine was the most important factor in predicting the rate of catheter encrustation. The higher the mean pHn value, the slower the rate of encrustation, and the longer catheters took to block. As Mathur et al.⁵² had shown that the pHn of any patient's urine varied from week to week, the authors of the later analysis suggested that manipulation of pHn might be possible; raising the pHn above urinary pH values would thus prevent catheter encrustation.⁵⁴ A study in healthy, noncatheterized volunteers demonstrated that dilution of the urine by increasing fluid intake, and increasing the urinary concentration of citrate (a chelating agent that can keep divalent metal ions, such as Ca²⁺ and Mg²⁺, in solution) elevated the pHn to values that were rarely exceeded by the urinary pH of patients infected with P. mirabilis.⁵⁵ Subsequent experiments in a laboratory model of P. mirabilis infection confirmed that when the models were supplied with dilute, citrate-containing urine with a pHn >8.3, crystalline biofilms did not form (Figure 6).⁵⁶

Figure 6 Electron micrographs of catheters removed from bladder models with varying citrate concentrations and nucleation pHs.

(A) This micrograph shows a section of Proteus mirabilis crystalline biofilm on an all-silicone catheter that was removed 24 h after incubation in a bladder model supplied with urine containing 0.41 mg/ml citrate, which had a nucleation pH of 7.4. Permission obtained from The Society for General Microbiology © Stickler DJ and MorganSD (2006) J Med Microbiol 55: 489–494. (B) Micrograph of the surface of a silicone catheter removed 24 h after incubation from a model supplied with urine containing citrate at 1.5 mg/ml, which had a nucleation pH of 8.3. The biofilm is sparse and is composed of microcolonies of cells with no signs of crystalline material. Permission obtained from The Society for General Microbiology © Stickler DJ and Morgan SD (2006) J Med Microbiol55: 489–494

Full figure and legend (27K)Figures & Tables indexDownload Power Point slide (231K)

The advice for patients to increase their fluid intake by drinking steadily throughout the day⁵⁷ clearly has a sound basis in physiology and physical chemistry. The dilution of urine resulting from an increased fluid intake will elevate the pHn and slow the rate of catheter encrustation. If the citrate content of urine can also be elevated by encouraging patients to take, for example, lemon-based drinks, the rate of crystal formation should reduce further. These observations should encourage a clinical trial to examine the effect of increasing patient's fluid intake with citrate-containing drinks on the encrustation and blockage of catheters.

Recurrent crystalline biofilms

In many patients who suffer recurrent catheter encrustation, the usual management involves simply replacing the blocked catheter with a new one. Fresh catheters are thus placed directly into urine cultures of P. mirabilis at alkaline pHs, which contain microcrystals of calcium and magnesium phosphates. Examination of the early stages of crystalline biofilm formation under these circumstances in a laboratory model has revealed a common sequence in the development of crystalline biofilm on all-silicone, silicone-coated latex, hydrogel-coated latex, and silver–hydrogel-coated latex catheters.⁵⁰ After only 1 h in the model, the catheter surfaces were covered by a microcrystalline layer. X-ray microanalysis confirmed that this material was composed largely of calcium and phosphate. Bacterial colonization of this foundation layer followed, with microcolonies of cells developing on the microcrystals (Figure 7). By 18 h, the eyelets and lumenal surfaces of all these catheters were comprehensively covered by densely populated, crystalline P. mirabilis biofilm. Examination of catheters removed from patients has confirmed that a microcrystalline foundation layer forms on catheters in vivo (Figure 8).⁵⁰

Figure 7 An early stage in the formation of a crystalline biofilm on a hydrogel-coated latex catheter.

The catheter had been placed in a bladder model containing a culture of Proteus mirabilis in urine at pH 8.5. Cells can be seen colonizing a foundation layer composed of aggregates of apatite microcrystals.

Full figure and legend (31K)Figures & Tables indexDownload Power Point slide (234K)

Figure 8 A crystalline biofilm developing around the eyehole of a silver–hydrogel-coated latex catheter.

The catheter had been removed from a patient just 5 days after it had been inserted. Bacilli and cocci can be seen colonizing a microcrystalline foundation layer that had formed on the catheter surface. Permission obtained from Woodhead Publishing (Cambridge, UK) © Stickler DJ (In Press) The challenge of the special problems for bladder catheters produced by infection with Proteus mirabilis In Biomaterials and Tissue Engineering in Urology, (Eds Atala A and Denstedt J).

Full figure and legend (35K)Figures & Tables indexDownload Power Point slide (240K)

Future catheter design

The formation of P. mirabilis crystalline biofilms has important implications for the development of encrustation-resistant catheters. Attempts to inhibit bacterial attachment and biofilm development by immobilizing an antibacterial in the catheter are unlikely to prevent encrustation in patients infected with P. mirabilis. In the case of silver-coated catheters, for example, the deposition of the crystalline foundation layer allows cells to attach and grow, protected from contact with the underlying silver. The obvious lesson for preventing catheter encrustation is to stop the pH of the urine rising above the pH at which crystals form. If antimicrobials are to be incorporated into catheters to prevent pH rising, they must diffuse out from the catheter surface and reduce the viable cell populations of P. mirabilis in the urine. Silver does not elute from silver–hydrogel catheters in sufficient quantities to inhibit the activity of P. mirabilis.⁵⁰ Chakravarti et al.⁵⁸ demonstrated that antibacterial concentrations of silver ions could be generated by passing an electric current through silver electrodes attached to catheters. Biofilm development was inhibited, but the effect was temporary as the silver electrodes disintegrated after about 150 h.

In light of the possibility that catheters releasing effective concentrations of antibacterial agents into the urine for the lifetime of catheter could prevent biofilm encrustation, Bibby et al.⁵⁹ had the idea that the retention balloon could be used as a reservoir for antibacterial agents. The authors suggested that the membrane of the balloon might control the release of the active agent over long periods. An investigation of the activity of a wide range of antibacterial agents against strains of P. mirabilis from catheters showed that the biocide triclosan was particularly active against these organisms.^{60, 61} Experiments in laboratory models demonstrated that triclosan diffused through the balloons of all-silicone and latex-based catheters. The bacterial population of the urine was reduced, the rise in urinary pH was prevented, and crystalline biofilm formation on the catheters was inhibited.^{62, 63} Loading the retention balloons of silicone catheters with 10 ml of triclosan (10 mg/ml in 5% w/v polyethylene glycol) resulted in the daily diffusion of around 115 g of the agent into the urine.⁶⁴ Assuming this diffusion rate is maintained, the antibacterial activity should persist in the urine for well over the current 12-week maximum life span of each long-term catheter. As with any intervention with an antibacterial agent, the possibility of resistance to triclosan developing is a concern. In clinical trials of this strategy, monitoring the urinary flora of the catheterized patients for signs of the emergence of less susceptible strains or the selection of intrinsically resistant species will be important.⁶⁵

Williams and Stickler⁶¹ explored the possibility of delivering other agents directly to the bladder through silicone catheter balloons. The authors reported that, of 18 biocides and antibiotics investigated, only triclosan and nalidixic acid were capable of diffusing through the balloon in sufficient concentrations to significantly inhibit the encrustation process. Polyurethane balloons, however, were permeable to gentamicin and the fluoroquinolones,⁶¹ and might be considered as an alternative to silicone or latex in catheter manufacture.

In an important editorial in 1988, Kunin⁶⁶ posed the question "can we build a better catheter?" Kunin expressed disappointment about the substantial technological advances that were common in many other medical fields, while draining urine from a debilitated bladder without resulting in infections was still not possible. Kunin commented that catheter manufacturers had been reluctant to invest in research and development, and that catheters used in 1988 were essentially the same as those introduced in the 1930s. Little has changed in the 20 years since Kunin's call for action. Currently, catheters are not costly, but the eventual price we have to pay for their use is enormous.¹⁰

Current management of crystalline catheter biofilms

Bacteria in biofilms are notoriously difficult to eradicate with bactericidal drugs.⁶⁷ This universal resistance of biofilm cells to antibacterial agents⁴¹ is of course clinically important, and has to be considered when instigating antibiotic therapy for symptomatic infections. Replacing the old, biofilm-laden catheters before antibiotic treatment is a sensible option.¹¹ Treatment should then be based on the susceptibility of organisms that are isolated from urine aspirated from the new catheter, as samples collected from the old catheter can contain different species and greater numbers of organisms.^{68, 69} In a prospective, randomized trial, Raz et al.⁷⁰ found that routine replacement of long-term catheters prior to instigation of antibiotic therapy led to shorter times to return to afebrile status, and a significantly lower rate of symptomatic clinical relapse 28 days after therapy.

For patients with a history of catheter encrustation and blockage, Norberg et al.⁷¹ suggested that monitoring the times that patients' catheters take to block would be sensible. If a characteristic pattern of blockage could then be identified for a patient, a schedule of catheter replacement could be planned so that catheters would be changed before the predicted time of blockage. A difficulty here, however, is the great variability in the times catheters take to block.⁵² Norberg et al.⁷¹ suggested that, to obtain a reasonable estimate of catheter life span for an individual patient, determining the median life span for between three and five consecutive catheters is necessary. Such a strategy might help to reduce the incidences of the clinical crises induced by blockage in some patients who are prone to recurrent blockage. An alternative approach is to use a simple sensor located in the drainage system that signals the early stages of catheter encrustation and the need to change the catheter. Such a sensor has been shown to have the added advantage of giving early warning of catheter encrustation in patients who have recently developed P. mirabilis infections and have not previously been prone to the complication.⁷²

Noncrystalline biofilms

Many species, in addition to P. mirabilis, form extensive biofilms on urinary catheters. While these biofilms do not generate crystalline formations, they are certainly of clinical interest. P. aeruginosa and K. pneumoniae, for example, produce copious amounts of exopolysaccharide and form mucoid biofilms that can occlude the catheter lumen (Figure 9).^{13, 64} Experiments in laboratory models have shown that, while these biofilms are not as effective as crystalline material at blocking catheters, they can markedly impede the flow of urine.^{17, 64} In some patients, catheters become blocked by mucoid material rather than by encrustation. It would be interesting to investigate these catheters, and determine whether the mucus is in fact mucoid biofilm.

Figure 9 Examples of mucoid, noncrystalline biofilms formed on all-silicone catheters after 4 days of incubation in a laboratory model of the bladder.

(A) This image shows a biofilm in a model infected with Pseudomonas aeruginosa. (B) This image shows a biofilm in a model infected with Klebsiella pneumoniae.

Full figure and legend (9K)Figures & Tables indexDownload Power Point slide (212K)

Conclusions

Infection with bacteria that produce urease, particularly P. mirabilis, exposes the many faults of the currently available Foley catheters, and can result in potentially disastrous consequences for patient care and substantial financial implications for health-care authorities. Catheters that are available today have roughly engineered surfaces, thick walls and narrow central channels that are extremely vulnerable to blockage by crystalline P. mirabilis biofilms. Plenty of scope exists for improving the design and manufacture of catheters, and many interesting ideas are presented in the literature.³⁸ The complications caused by these biofilms undermine patients' quality of life and threaten the health of so many people; these complications are no longer acceptable. Catheter manufacturers should take up the challenge for reducing the incidence of catheter biofilms.

Acknowledgments

The author would like to thank Dr Rob Broomfield, Dr Sheridan Morgan and Dr Rob Young, who are members of Dr Stickler's research team, for providing the previously unpublished images shown in this Review. Charles P Vega, University of California, Irvine, CA, is the author of and is solely responsible for the content of the learning objectives, questions and answers of the Medscape-accredited continuing medical education activity associated with this article.

References

1. Costerton JW et al. (1987) Bacterial biofilms in nature and disease. Annu Rev Microbiol 41: 435–464 | Article | PubMed | ISI | ChemPort |
2. Donlan RM (2002) Biofilms: microbial life on surfaces. Emerg Infect Dis 8: 881–890 | PubMed | ISI |
3. Warren JW (1991) The catheter and urinary tract infection. Med Clin North Am 75: 481–493 | PubMed | ChemPort |
4. Kunin CM (1997) Care of the urinary catheter. In Urinary Tract Infections: Detection, Prevention and Management, edn 5, 226–278 (Ed. Kunin CM) Baltimore: Williams & Wilkins
5. Stamm WE (1991) Catheter-associated urinary tract infections: epidemiology, pathogenesis, and prevention. Am J Med 91 (Suppl 3B): 65S–71S | Article |
6. Tambyah PA et al. (1999) A prospective study of pathogenesis of catheter-associated urinary tract infections. Mayo Clin Proc 74: 131–136 | PubMed | ChemPort |
7. Matsukawa M et al. (2005) Bacterial colonization on intraluminal surface of urethral catheter. Urology 65: 440–444 | Article | PubMed |
8. Clayton CL et al. (1982) Some observations on urinary tract infections in patients undergoing long-term bladder catheterization. J Hosp Infect 3: 39–47 | Article | PubMed | ChemPort |
9. Warren JW et al. (1982) Cephalexin for susceptible bacteriuria in afebrile, long-term catheterized patients. JAMA 248: 454–458 | Article | PubMed | ChemPort |
10. Saint S and Chenoweth CE (2003) Biofilms and catheter-associated urinary tract infections. Infect Dis Clin North Am 17: 411–432 | Article | PubMed |
11. Trautner BW and Darouiche RO (2004) Role of biofilm in catheter-associated urinary tract infection. Am J Infect Control 32: 177–183 | Article | PubMed |
12. Tenke P et al. (2008) European and Asian guidelines on management and prevention of catheter-associated urinary tract infections. Int J Antimicrob Agents 31 (Suppl 1): S68–S78 | Article |
13. Ganderton L et al. (1992) Scanning electron microscopy of bacterial biofilms on indwelling bladder catheters. Eur J Clin Microbiol Infect Dis 11: 789–796 | Article | PubMed | ChemPort |
14. Morris NS et al. (1999) The development of bacterial biofilms on indwelling urethral catheters. World J Urol 17: 345–350 | Article | PubMed | ChemPort |
15. Liedl B (2001) Catheter-associated urinary tract infections. Curr Opin Urol 11: 75–79 | Article | PubMed | ChemPort |
16. Ohkawa M et al. (1990) Bacterial and crystal adherence to the surfaces of indwelling urethral catheters. J Urol 143: 717–721 | PubMed | ChemPort |
17. Macleod SM and Stickler DJ (2007) Species interactions in mixed-community crystalline biofilms on urinary catheters. J Med Microbiol 56: 1549–1557 | Article | PubMed |
18. Getliffe KA and Mulhall AB (1991) The encrustation of indwelling catheters. Br J Urol 67: 337–341 | PubMed | ChemPort |
19. Stickler DJ and Zimakoff J (1994) Complications of urinary tract infections associated with devices used for long-term bladder management. J Hosp Infect 28: 177–194 | Article | PubMed | ChemPort |
20. Kunin CM (1987) Care of the urinary catheter. In Detection, Prevention and Management of Urinary Tract Infections, edn 4, 245–298 (Ed. Kunin CM) Philadelphia: Lea & Febiger
21. Cools HJ and Van der Meer JW (1986) Restriction of long-term indwelling urethral catheterisation in the elderly. Br J Urol 58: 683–688 | PubMed | ChemPort |
22. Getliffe KA (1994) The characteristics and management of patients with recurrent blockage of long-term urinary catheters. J Adv Nurs 20: 140–149 | Article | PubMed | ChemPort |
23. Kohler-Ockmore J and Feneley RC (1996) Long-term catheterization of the bladder: prevalence and morbidity. Br J Urol 77: 347–351 | PubMed | ChemPort |
24. Hedelin H et al. (1984) The composition of catheter encrustations, including the effects of allopurinol treatment. Br J Urol 56: 250–254 | PubMed | ChemPort |
25. Cox AJ and Hukins DW (1989) Morphology of mineral deposits on encrusted urinary catheters investigated by scanning electron microscopy. J Urol 142: 1347–1350 | PubMed | ChemPort |
26. Cox AJ et al. (1989) Infection of catheterised patients: bacterial colonisation of encrusted Foley catheters shown by scanning electron microscopy. Urol Res 17: 349–352 | Article | PubMed | ChemPort |
27. Stickler D et al. (1993) Proteus mirabilis biofilms and the encrustation of urethral catheters. Urol Res 21: 407–411 | Article | PubMed | ISI | ChemPort |
28. McLean RJ et al. (1996) Biofilm mediated calculus formation in the urinary tract. Cells Mater 6: 165–174
29. Mobley HL and Warren JW (1987) Urease-positive bacteriuria and obstruction of long-term urinary catheters. J Clin Microbiol 25: 2216–2217 | PubMed | ISI | ChemPort |
30. Kunin CM (1989) Blockage of urinary catheters: role of microorganisms and constituents of the urine on formation of encrustations. J Clin Epidemiol 42: 835–842 | Article | PubMed | ChemPort |
31. Jones BD and Mobley HL (1987) Genetic and biochemical diversity of ureases of Proteus, Providencia, and Morganella species isolated from urinary tract infection. Infect Immun 55: 2198–2203 | PubMed | ISI | ChemPort |
32. Stickler D et al. (1998) Studies on the formation of crystalline bacterial biofilms on urethral catheters. Eur J Clin Microbiol Infect Dis 17: 649–652 | Article | PubMed | ChemPort |
33. Mobley HT (1996) Virulence of Proteus mirabilis. In Urinary tract infections: molecular pathogenesis and clinical management, 245–270 (Eds Mobley HL and Warren JW) Washington DC: ASM Press
34. Sabbuba NA et al. (2003) Molecular epidemiology of Proteus mirabilis infections of the catheterized urinary tract. J Clin Microbiol 41: 4961–4965 | Article | PubMed | ChemPort |
35. Sabbuba NA et al. (2004) Genotyping demonstrates that the strains of Proteus mirabilis from bladder stones and catheter encrustations of patients undergoing long-term bladder catheterization are identical. J Urol 171: 1925–1928 | Article | PubMed | ChemPort |
36. Mathur S et al. (2005) Genotyping of urinary and fecal Proteus mirabilis isolates from individuals with long-term urinary catheters. Eur J Clin Microbiol Infect Dis 24: 643–644 | Article | PubMed | ChemPort |
37. Morris NS et al. (1997) Which indwelling urethral catheters resist encrustation by Proteus mirabilis? biofilms Br J Urol 80: 58–63 | PubMed | ChemPort |
38. Stickler DJ and Sabbuba NA (2007) Antimicrobial catheters. In Disinfection and Decontamination Principles, Applications and Related Issues, 415–458 (Ed. Manivannan G) Boca Raton: CRC Press
39. Capewell AE and Morris SL (1993) Audit of catheter management provided by District Nurses and Continence Advisors. Br J Urol 71: 259–264 | PubMed | ChemPort |
40. Stickler DJ (1996) Biofilms, catheters and urinary tract infections. Eur Urol Update Series 5: 1–8
41. Donlan RM and Costerton JW (2002) Biofilms: survival mechanisms of clinically relevant microorganisms. Clin Microbiol Rev 15: 167–193 | Article | PubMed | ISI | ChemPort |
42. Ong CL et al. (2008) Identification of type 3 fimbriae in uropathogenic Escherichia coli reveals a role in biofilm formation. J Bacteriol 190: 1054–1063 | Article | PubMed | ChemPort |
43. Rocha SP et al. (2007) Fimbriae of uropathogenic Proteus mirabilis. FEMS Immunol Med Microbiol 51: 1–7 | Article | PubMed | ChemPort |
44. Jacobsen SM et al. (2008) Complicated catheter-associated urinary tract infections due to Escherichia coli and Proteus mirabilis. Clin Microbiol Rev 21: 26–59 | Article | PubMed | ChemPort |
45. Downer A et al. (2003) Polymer surface properties and their effect on the adhesion of Proteus mirabilis. Proc Inst Mech Eng [H] 217: 279–289
46. Stickler DJ et al. (2006) Observations on the adherence of Proteus mirabilis onto polymer surfaces. J Appl Microbiol 100: 1028–1033 | Article | PubMed | ChemPort |
47. Jones BV et al. (2005) Role of swarming in the formation of crystalline Proteus mirabilis biofilms on urinary catheters. J Med Microbiol 54: 807–813 | Article | PubMed |
48. Cox AJ (1990) Comparison of catheter surface morphologies. Br J Urol 65: 55–60 | PubMed | ChemPort |
49. Stickler D et al. (2003) Why are Foley catheters so vulnerable to encrustation and blockage by crystalline bacterial biofilm> Urol Res 31: 306–311 | Article | PubMed |
50. Stickler DJ and Morgan SD (2008) Observations on the development of the crystalline bacterial biofilms that encrust and block Foley catheters. J Hosp Infect 69: 350–360 | Article | PubMed | ChemPort |
51. Brisset L et al. (1996) In vivo and in vitro analysis of the ability of urinary catheter to microbial colonization. Pathol Biol (Paris) 44: 397–404 | PubMed | ChemPort |
52. Mathur S et al. (2006) Prospective study of individuals with long-term urinary catheters colonized with Proteus species. BJU Int 97: 121–128 | Article | PubMed |
53. Choong S et al. (2001) Catheter associated urinary tract infection and encrustation. Int J Antimicrob Agents 17: 305–310 | Article | PubMed | ChemPort |
54. Mathur S et al. (2006) Factors affecting crystal precipitation from urine in individuals with long-term urinary catheters colonized with urease-positive bacterial species. Urol Res 34: 173–177 | Article | PubMed |
55. Suller MT et al. (2005) Factors modulating the pH at which calcium and magnesium phosphates precipitate from human urine. Urol Res 33: 254–260 | Article | PubMed | ChemPort |
56. Stickler DJ and Morgan SD (2006) Modulation of crystalline Proteus mirabilis biofilm development on urinary catheters. J Med Microbiol 55: 489–494 | Article | PubMed | ChemPort |
57. Burr RG and Nuseibeh IM (1997) Urinary catheter blockage depends on urine pH, calcium and rate of flow. Spinal Cord 35: 521–525 | Article | PubMed | ChemPort |
58. Chakravarti A et al. (2005) An electrified catheter to resist encrustation by Proteus mirabilis biofilm. J Urol 174: 1129–1132 | Article | PubMed |
59. Bibby JM et al. (1995) Feasibility of preventing encrustation of urinary catheters. Cells Mater 2: 183–195
60. Stickler DJ (2002) Susceptibility of antibiotic-resistant Gram-negative bacteria to biocides: a perspective from the study of catheter biofilms. J Appl Microbiol 92 (Suppl): 163S–170S | Article |
61. Williams GJ and Stickler DJ (2007) Some observations on the diffusion of antimicrobial agents through the retention balloons of Foley catheters. J Urol 178: 697–701 | Article | PubMed | ChemPort |
62. Stickler DJ et al. (2003) Control of encrustation and blockage of Foley catheters. Lancet 361: 1435–1437 | Article | PubMed | ChemPort |
63. Jones GL et al. (2005) A strategy for the control of catheter blockage by crystalline Proteus mirabilis biofilm using the antibacterial agent triclosan. Eur Urol 48: 838–845 | Article | PubMed |
64. Jones GL et al. (2006) Effect of triclosan on the development of bacterial biofilms by urinary tract pathogens on urinary catheters. J Antimicrob Chemother 57: 266–272 | Article | PubMed | ISI | ChemPort |
65. Stickler DJ and Jones GL (2008) Reduced susceptibility of Proteus mirabilis to triclosan. Antimicrob Agents Chemother 52: 991–994 | Article | PubMed | ChemPort |
66. Kunin CM (1988) Can we build a better urinary catheter. N Engl J Med 319: 365–366 | PubMed | ChemPort |
67. Lewis K (2001) Riddle of biofilm resistance. Antimicrob Agents Chemother 45: 999–1007 | Article | PubMed | ISI | ChemPort |
68. Tenney JH and Warren JW (1988) Bacteriuria in women with long-term catheters: paired comparison of indwelling and replacement catheters. J Infect Dis 157: 199–202 | PubMed | ChemPort |
69. Ramsay JW et al. (1989) Biofilms, bacteria and bladder catheters. A clinical study. Br J Urol 64: 395–398 | PubMed | ChemPort |
70. Raz R et al. (2000) Chronic indwelling catheter replacement before antimicrobial therapy for symptomatic urinary tract infection. J Urol 164: 1254–1258 | Article | PubMed | ChemPort |
71. Norberg B et al. (1983) The spontaneous variation of catheter life in long-stay geriatric inpatients with indwelling catheters. Gerontology 29: 332–335 | PubMed | ChemPort |
72. Stickler DJ et al. (2006) A clinical assessment of the performance of a sensor to detect crystalline biofilm formation on indwelling bladder catheters? BJU Int 98: 1244–1249 | Article | PubMed |

Contact the journal about this article

Subject areas under which this article appears: Infections, inflammation and prostatitis | Urinary incontinence, urodynamics and lower urinary tract dysfunction