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Biofilm and Urogenital Infections

Written By

Peter Tenke, Bela Koves, Karoly Nagy, Shinya Uehara, Hiromi Kumon, Scott J. Hultgren, Chia Hung and Werner Mendling

Submitted: 14 December 2010 Published: 06 September 2011

DOI: 10.5772/24858

From the Edited Volume

Clinical Management of Complicated Urinary Tract Infection

Edited by Ahmad Nikibakhsh

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1. Introduction

Bacterial adherence and the growth of bacteria on solid surfaces as biofilm are both naturally occurring phenomena. Biofilms can be defined as an accumulation of microorganisms and their extracellular products forming structured communities attached to a surface. Biofilms are able to build up under natural circumstances, for instance on the urothelium or prostate stones and they can also colonize the surfaces of implanted medical devices. Biofilm infections have a major role on temporary and permanent implants or devices placed in the human body. In the process of endourological development a great variety of foreign bodies have been invented besides urethral catheters like ureter, prostatic stents, percutan nephrostomy, penile, testicular implants and artificial urinary sphincters. Many biofilms are quite harmful but others can have a positive impact, namely lining healthy intestine and female genito-urinary tract. Biofilms have significant implications for clinical pharmacology, particularly related to antibiotic resistance, drug adsorption onto and off of devices, and minimum inhibitory concentrations of drugs required for effective therapy.

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2. Biofilm formation and growth

A biofilm is an aggregate of microorganisms in which cells adhere to each other and/or to a surface. These adherent cells are frequently embedded within a self-produced matrix of extracellular polymeric substance (EPS). Formation of a biofilm begins with the attachment of free-floating microorganisms to a surface. The first step of biofilm formation is always the deposition of a conditioning film produced by the host to the foreign body. It is followed by the attachment of microorganisms. The microbial adhesion and anchorage to the surface are made by exopolymer production. After this process their growth, multiplication and dissemination can be observed [1,2,3,4,5].

After insertion of the device into the body the material surface enters into contact with body fluids around the implant. In case of the urinary tract Tamm-Horsfall glycoprotein, various ions, polysaccharides and other components diffuse toward the implant surface from the urine within minutes [6]. Macromolecular components (serum albumin, fibrinogen, collagen, fibronectin) from these body fluids adsorb extremely fast onto the material surfaces to form a conditioning film, prior to the arrival of the first organisms [7]. The creation of a conditioning film alters the surface characteristics of implants. The role of the conditioning film is vital as many pathogens do not have mechanisms allowing them to adhere directly or strongly onto bare implant surfaces [8].

The next step in the development of a biofilm is the approach and attachment of microorganisms. The ability of microorganisms to adhere to surfaces is influenced by electrostatic and hydrophobic interactions, ionic strength, osmolality and urinary pH [9,10].

In order for bacteria to react to a surface or an interface like an air-water interface, these cells must be able to ’sense’ their proximity to these surfaces. The planktonic ’free-floating’ bacterial cells release both protons and signaling molecules as they move through the bulk fluid. These protons and signaling molecules must diffuse radially away from the floating cell, if not adjacent to any surface or interface. But a significantly higher concentration of either protons or signaling molecules can develop on the side of the bacterial cell close to any surface. This allows the cell to sense that it is near a surface because diffusion is limited on this side [4]. After the planktonic bacterial cell has sensed the surface, it may commit to the active process of adhesion and biofilm formation.

There is no single process or theory, which can completely describe microbial adhesion. The initial adhesion is reversible and involves hydrophobic and electrostatic forces. It is followed by irreversible attachment provided by bacterial polysaccharides which anchor the organisms to the surface. Subsequently, colonization takes by species factors, such as slow migration and spreading, rolling, packing and adhesion of the progress. A developed biofilm consists of groups of microorganisms, sometimes in mushroom-like forms, separated by interstitial spaces that are filled with the surrounding fluid [11]. The growth rates of organisms on a surface as well as the strategies used by microorganisms to spread over a surface are important for colonization. These strategies are species specific which can influence the distribution of a biofilm on a surface [12].

The final stage of microbial colonization of a surface is the formation of a biofilm structure. At this point, the microorganisms have created a microenvironment protective against many antimicrobial agents and host immune defense mechanisms. Biofilm has been described as having a heterogeneous structure with a rough surface [13]. The microcolony is actually the basic structural unit of the biofilm, similar to the tissue which is the basic unit of growth of more complex organisms. Depending on the species involved, the microcolony may be composed of 10-25% cells and 75-90% exopolysaccharide (EPS) matrix. The biofilm contains ‘water channels’ which allow transporting of essential nutrients and oxygen for the growth of the cells [14]. Microorganisms within the biofilm also secrete chemical signals that mediate population density-dependent gene expression, which has an important role in biofilm development [15]. In summary, the biofilm is usually built up of three layers [16]:

  1. the linking film which attaches to the surface of tissue or biomaterials

  2. the base film of compact microorganisms

  3. the surface film as an outer layer, where planktonic organisms can be released free-floating and spreading over the surface.

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3. Antimicrobial susceptibility of bacteria in biofilm

Infections caused as a result of biofilm formation are characterized by particularly strong antibiotic and immune resistance patterns. Bacteria within the biofilms differ in behaviour and in phenotypic form from the planktonic bacteria. Antimicrobial agents are effective against planktonic bacteria and appear to clear mucosal surfaces of adherent bacterial microcolonies but frequently fail to eradicate bacterial biofilms on urological devices. The use of antibiotics is currently one of the possibilities of the prevention of biofilm formation. However, even in the presence of antibiotics bacteria can adhere, colonize and survive on implanted medical devices as has been shown for urinary catheters and ureteral stent surfaces in vitro and in vivo [17,18,19]. The problem in conventional clinical microbiology is how to treat patients in the best way when choosing antibiotics is based on bacterial cultures derived from planktonic bacterial cells which differ very much from bacteria in the biofilm mode. This can stand behind the clinical failure rate of treating chronic bacterial infection.

The failure of antimicrobial agents to treat biofilms has been associated with a variety of mechanisms (4) [18,19,20,21,22,23,24]. One mechanism of biofilm resistance to antimicrobial agents is the failure of an agent to penetrate the full depth of the biofilm (extrinsic resistance). The extracellular matrix for instance may block the penetration at the very beginning.

  • One mechanism is the failure of an agent to penetrate the full depth of the biofilm (extrinsic resistance). The extracellular matrix may block the penetration at the very beginning.

  • The organisms growing at a slower rate within the biofilm are more resistant to the effects of antimicrobial agents, which require active growth.

  • Bacteria within biofilm are phenotypically so different from their planktonic counterparts that antimicrobial agents developed against the latter often fail to eradicate organisms in the biofilm. Bacteria within a biofilm activate many genes which alter the cell envelope, the molecular targets and the susceptibility to antimicrobial agents (intrinsic resistance). Current opinion is that phenotypic changes caused by a genetic switch, when approximately 65-80 proteins change, play a more important role in the protection from antimicrobial agents than the external resistance provided by the exopolysaccharide matrix.

  • Bacteria within a biofilm can sense the external environment, communicate with each other and transfer genetic information and plasmids within biofilm.

  • Bacteria in a biofilm can usually survive the presence of antimicrobial agents at a concentration 1000-1500 times higher than the concentration that kills planktonic cells of the same species.

According to in vitro and in vivo studies aminoglycosides and beta-lactam antibiotics can prevent the formation of ‘young’ biofilms, while fluoroquinolones are effective in case of both ‘young’ and ‘older’ biofilms because of their good penetrative qualities. They are present in biofilms even one or two weeks after the end of the antibiotic treatment [25-28].

Most researchers believe that antibiotics can only slow down the progress of biofilm formation by eliminating unprotected planktonic bacteria and reducing the metabolic activity of bacteria on the biofilm surface [23, 29-30]. However, during an acute febrile phase of a biofilm infection.

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4. Indwelling urethral catheters

Due to the urinary catheter the development of bacteriuria and biofilm formation is inevitable. Urinary catheters are readily targets of biofilm development on their inner and outer surfaces once they are inserted. The long-term use of them leads to infection in most of the cases. The surface of a catheter (depending on its material) provides sufficient circumstances for bacteria to adhere and spread along in two ways. One route is when organisms ascend the catheter extraluminally by direct inoculation at the time of the catheter insertion or migrate in the mucous sheath that surrounds the external aspect of the catheter. Extraluminal organisms are primarily endogenous, originating from the gastrointestinal tract. These organisms colonize the patient’s perineum and ascend the urethra after catheter insertion [13,31,32,33,34] Approximately 70% of bacteriuria in catheterized women is believed to occur through the extraluminal entry.

Bacteria can ascend the catheter also by an intraluminal route, which occurs when organisms gain access to the internal lumen of the catheter. These organisms are usually introduced from exogenous sources, for instance with cross transmission from the hands of health care personnel [13,32,33,35]. Adhesion of microorganisms to catheter materials depends on the hydrophobicity of the organism and catheter surface.

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5. The biofilms and the encrustation and blockage of catheters

An additional problem in use of medical biomaterials in the urinary tract environment is the development of encrustation and consecutive obstruction. When the drained urinary tract becomes infected by urease producing bacteria such as Proteus mirabilis, the bacterial urease generates ammonia from urea and elevates the pH of the urine. Under these alkaline conditions, crystals of calcium phosphate (hydroxyapatite) and magnesium ammonium phosphate (struvite) are formed and trapped in the organic matrix surrounding the cells [20, 21, 36,37]. Progression of these encrustations eventually blocks the catheter lumen.

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6. Ureteral stents

In vitro and in vivo studies confirmed the difficulty in detecting biofilm formation by using conventional laboratory procedures [38, 39]. Reid at al found that 90% of indwelling silicone double J stents were colonized by adherent bacteria, however the incidence of urinary infection detected clinically was only 27% [38]. The difficulty in detecting biofilm formation by using conventional laboratory procedures was confirmed in a large study where 237 ureteral stents were tested. It was shown that 68% of stents were actually colonized but only 30% of patients were found to have bacteriuria [39]. Therefore, a negative urine culture does not rule out the possibility of stent colonization. The study testified correlation between the length of the indwelling time and the development of infection.

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7. Penile prostheses

The prosthesis-associated chronic pain due to subclinical infection is more common than clinically apparent infection (3). Staphylococcus species, especially Staphylococcus epidermidis are the most common pathogens found in penile prostheses infection (35-56%) [40], while Gram-negative enteric bacteria are liable for 20 % of infections [41,42]. S. epidermidis was cultured in 40% of penile prosthesis removed for malfunction with no clinical evidence of infection [43]. Staphylococcal species were also found to enhance biofilm formation. These cases can be ‘silent’ for many years before becoming clinically evident [44] in contrast to Gram-negative bacterial infection (Pseudomonas aeruginosa, E.coli, Serratia marcescens, and Proteus mirabilis) being responsible for 20% of infections, which usually become manifest in a month after implantation [43].

To reduce the risk of device associated infections many modifications have been developed such as antibiotic and hydrophilic coated devices.Hydrophilic penile prosthesis coating was has been shown to decrease bacterial adherence in vitro and in animal models [45]. Antibiotic prophylaxis is desirable for the above-mentioned facts. Since the most common pathogen is the Staphylococcus epidermidis, first-generation cephalosporins, broad-spectrum penicillin should be used [46]. In cases of chronic pain, long-term administration of quinolones eased 60% of symptoms. Lack of success involves the necessity of implant removal.

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8. Artificial urinary sphincters (AUS)

Around 3% of the AUS become infected and symptoms are mainly associated with the control pump device. Avoiding the risk factors as infected urine, prolonged urinary retention and large bladder residual can reduce this high occurrence [43,46]. Since the parts of the sphincter device form one continuous surface, the AUS is suggested to be removed entirely as the first step to eliminate the infection. The reimplantation must be preceded by the complete treatment of the infected area. This is not always achievable as many of these patients are paraplegic or have a neurogenic bladder with recurrent UTIs [43,46].

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9. Infected urinary calculi

In case of urease-producing bacteriuria the infection can be conjoined with the formation of struvite and calcium phosphate calculi as described above. The infected calculi grow rapidly and provide safe environment for the bacteria adhered to the biofilm [47]. The complete removal of all stone fragments during stone operation (PCNL, URS, ±combined with ESWL), prolonged administration of antibiotics (8-10 weeks for destroying urease-producing bacteria) and metaphylaxis are the features of the most effective treatment strategy.

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10. Chronic bacterial prostatitis

Although the diagnosis and classification of chronic prostatitis have been standardized, the differentiation of chronic non-bacterial from bacterial inflammation is still challenging. Being out of the sweeping effect of streaming urine the prostatic ducts and acini provides safe circumstances to planktonic bacteria to multiply rapidly and induce a host response with infiltration of acute inflammatory cells into the ducts. The ducts become engorged with infiltrate composed of dead and living bacteria as well as living and dying acute inflammatory cells, desquamated epithelial cells and cellular debris. At this point it is relatively easy to eradicate all the offending organisms which are in a ‘planktonic state’ with appropriate antibiotic therapy. If the bacteria persist from either clinically acute or more likely, subacute inflammation, they can form sporadic bacterial microcolonies or biofilms adherent to the epithelium of the ductal system (2b) [48,49]. These bacteria also produce an exopolysaccharide slime or glycocalyx that envelops these adherent microcolonies. The bacteria persisting in the prostate gland within these focal biofilms can provoke persistent immunological stimulation and subsequent chronic inflammation [48]. The diagnosis of chronic bacterial prostatitis can be difficult as colonized bacteria will not get into the prostatic secretion or urine sample. Antimicrobial therapy eradicates the planktonic bacteria but not the adherent bacterial biofilms deep within the prostate gland. Another cause of unsuccessful treatment may be the fact that the bacteria within biofilms differ significantly from their planktonic counterparts in metabolic rate, molecular targets and expression of antimicrobial binding proteins [3,19]. There is a need in development of diagnostic tools which would be able to recognize small adherent bacterial biofilms which exist deep within the prostate gland in chronic bacterial prostatitis. New treatment regimens should be carried out in order to be able to deliver much higher antibiotic concentrations to the biofilm within the prostatic duct.

11. Intracellular bacterial biofilm-like pods in the recurrent cystitis

Entry of E. coli into the urinary tract is not well understood, although sexual intercourse is the most clearly defined predisposing factor. Presumably, a small number of E. coli from the vaginal or enteric flora are introduced into the bladder during an average incident, and it seems plausible that in most cases the innate defenses in the bladder would be able to prevent infection. However, sometimes Uropathogenic E.Coli (UPEC) clearly possess mechanisms to overcome these defenses and establish a foothold in the bladder. UPEC pathogenesis initiates with bacterial binding of superficial bladder epithelial cells. Initial colonization events activate inflammatory and apoptotic cascades in the epithelium, which is normally inert and only turns over every 6 to 12 months. Bladder epithelial cells respond to invading bacteria in part by recognizing bacterial lipopolysaccharide (LPS) via the Toll-like receptor pathway, which results in strong neutrophil influx into the bladder. In addition, interactions mediated by adhesin FimH at the tips of type 1 pili with the bladder epithelium stimulate exfoliation of superficial epithelial cells, causing many of the pathogens to be shed into the urine. Genetic programs are activated that lead to differentiation and proliferation of the underlying transitional cells in an effort to renew the exfoliated superficial epithelium. Despite the robust inflammatory response and epithelial exfoliation, UPEC are able to maintain high titers in the bladder for several days.

A bacterial mechanism of FimH-mediated invasion into the superficial cells apparently allows evasion of these innate defenses. Initially, bacteria replicate rapidly inside superficial cells as disorganized clusters. Subsequently, bacteria in the clusters divide without much growth in cell size, resulting in coccoid-shaped bacteria, presumably due to changes in genetic programs. Furthermore, the bacterial clusters became highly compact and organized into biofilm-like structures, termed intracellular bacterial communities (IBCs), inside bladder superficial cells [50]. The IBCs push the bladder superficial cell membranes outward to give a “pod” like appearance by scanning electron microscopy. Bacteria in the IBCs are held together by exopolymeric matrices, reminiscent of biofilm structures [51]. At some point during this IBC developmental process, bacteria on the edges of IBCs become elongated again, become motile and start to move away from IBCs. Bacteria can exit out of infected bladder cells, probably due to compromised membrane integrity. UPEC undergo such IBC cascade to increase in numbers, resulting in high bacterial titers in the bladder. In addition, bacteria in these intracellular niches can create a chronic quiescent reservoir in the bladder, which can persist undetected for several months without bacteria shedding in the urine [52,53,54]. Bacteria in IBCs are completely resistant to 3- and 10-day courses of antibiotics [55].

12. Biofilm and pyelonephritis

Once bacteria reach the kidney either by ascending infection or vesicoureteral reflux they are able to adhere to the urothelium and papillae. Nickel et al showed that bacteria could adhere in thin biofilms to the urothelium before invading the renal tissue with resultant pyelonephritis [47]. These bacterial biofilms are more easily eradicated by antimicrobial agents, in contrast to the biofilms on catheter surfaces [51], which may be ascribed to the effective synergistic actions of antimicrobial agents and host defenses against the biofilms on urothelium [56].

13. Biofilm in bacterial vaginosis

Bacterial vaginosis (BV) is the most common vaginal disorder in adult women [57]. Although it is a non-fatal disease, BV presents an increased risk for other more severe clinical outcomes, such as preterm birth and HIV infections [58,59]. As defined by Amsel clinical criteria, BV exhibits at least 3 of the following 4 clinical symptoms:

  1. elevation of vaginal fluid pH to above 4.5;

  2. detectable “fishy odor” of vaginal fluid upon addition of 10% potassium hydroxide;

  3. presence of clue cells, vaginal epithelial cells covered with bacteria, in vaginal fluid; and

  4. milky vaginal discharge.

The vaginal flora of healthy women consists predominantly of Gram-positive lactobacilli, especially Lactobacillus cripatus and Lactobacillus jensenii [60-62]. Productions of antimicrobial proteins as well as the maintenance of acidic pH and hydrogen peroxide (H2O2) in the vaginal fluid by these bacteria contribute critically to the establishment of a healthy ecosystem in the vagina [61-63]. On the other hand, the vaginal microbiota of women with BV showed a loss of Lactobacillus species and an increase in microbial diversity dominated by Gardnerella vaginalis and to a lesser extend, many other bacterial organisms, including Porphyromonas, Mobiluncus, and Prevotella species [64-66].

The attempts to demonstrate G. vaginalis as the causative pathogen of BV have failed [67], many studies have demonstrated unequivocally that G. vaginalis is present in the majority of BV vaginal cultures in high numbers [64, 68, 69]. One additional complication with BV is the high recurrent rate of infection, despite of efficient resolution of infection by antibiotic treatments [70]. The recurrence nature of this disease prompted the speculation that bacterial biofilms are involved in BV. G. vaginalis poses the intrinsic ability to form biofilm in vitro [71-73]. Similar to other bacterial biofilm phenotypes, G. vaginalis biofilm is more resistant to antibiotic treatments compared to it planktonic counterparts [73].

Swidsinski and co-workers demonstrated the presence of bacterial biofilms on the vaginal epithelium of biopsies from women with BV [68]. These biofilm showed characteristics of dense surface bacterial biofilm and were comprised predominantly of G. vaginalis. Although G. vaginalis was also detected in biopsies from healthy women, they were present in very small numbers and infrequent. The sensitivity and specificity of FISH technique also allowed the researchers to identify the presence of Gram-positive (Streptococcus spp., Enterococcus spp. and Staphylococcus spp.) and Gram-negative (Escherichia coli and Proteus spp.) bacteria embedded within the G. vaginalis biofilms. Furthermore, in a subsequent publication, Swidsinski and colleagues reported the resurgence of dense bacterial biofilms at 1-week post-cessation of metronidazole treatment [74]. These biofilms were comprised principally of G. vaginalis and Atopobium vaginae. These clinical data strongly support the presence and involvement of bacterial biofilm in BV. It is interesting to note, however, that Saunders et al. [72] demonstrated that incubation of preformed G. vaginalis biofilm with certain strains of L. reuteri or L. iners resulted in the disruption of biofilm and decreased viability of G. vaginalis.

14. Prevention of biofilm formation in the urinary tract

The harsh and potentially fatal consequences of microbial biofilm infections generated efforts to prevent their formation, particularly on indwelling medical devices using chemical and mechanical approaches. Catheters coated with hydrogel, silver salts, and antimicrobials have been evaluated; however, they provide minimal reduction in infection incidence (75).

Antibiotic (minocycline, rifampicin, nitrofurantoin) impregnated catheters lowered the rate of asymptomatic bacteriuria compared to catheters without impregnation at less than one week but difference was not statistically significant at greater than one week, and the authors concluded that the data were too few to draw conclusions about long-term catheterization. [76]

Silver alloy catheters significantly reduced the incidence of asymptomatic bacteriuria at less than one week of catheterization [76]. Beyond one week the estimated effect was smaller but the risk of asymptomatic bacteriuria was still less in the silver alloy group. There are no available clinical trials with appropriate setting about the effect of silver alloy coated catheters on bacteriuria or biofilm formation in case of long-term catheterisation.

De Ridder et al found that fewer patients using hydrophilic-coated catheter (64%) for CIC experienced UTIs compared to the uncoated catheter group (82%)[77]. However, in a randomised controlled study the authors did not find significant difference between hydrophilic-coated and uncoated indwelling urethral catheters in place for 6 weeks with respect to symptomatic urinary tract infection and microbiological analysis of urine culture [78].

Heparin coated ureteral stents did not show any organic (biofilms) or anorganic (crystals) deposits after being in situ for up to 6 weeks whereas significant biofilms were demonstrated in 33% of uncoated stents [79].

15. Use of low-energy surface acoustic waves (SAW)

Biofilm formation can be prevented- or delayed- by applying low intensity nanowaves along the surfaces of an indwelling catheter. This approach opens new options for pharmacological prevention of urinary tract infections (80,81).The concept of using low-energy SAW is based on the hypothesis that these acoustic waves are able to disrupt the formation of biofilms if transmitted directly to indwelling medical devices by inhibiting the adhesion of planktonic bacteria to their surface. Hazan et al. demonstrated the the effectiveness of Low-Energy Surface Acoustic Waves in the prevention of biofilm formation in an animal model in vivo. They found that SAW treatment reduced biofilm formation in vitro, leaving catheters virtually clean of adherent microorganisms, irrespective of the types of bacteria that were examined. In the animal model SAW treated catheters showed strong inhibition of bacterial biofilm compared to controls [82].

In a double blind sham controlled randomized study related to short term catheterization, applying SAW releasing device to catheters prevented biofilm formation in all of the catheters whereas biofilm was present in 63% of the control group [83].

A workgroup of the authors of the present article performed a prospective parallel group comparative study on the efficacy of the SAW treatment in case of long-term catheterisation (8 weeks). SAW treatment lowered the rate of significant bacteriuria (33% vs. 81%) and the rate of biofilm formation was also significantly lower in the SAW group compared to the controls[84].

16. Conclusion

The number of biomaterial devices used in urology has been increasing permanently. Biofilm infections have a major impact on implants or devices placed in the human body. The mechanism and the different bacterial and host factors taking part in the formation of biofilms have been extensively researched in the last decades, such ideal method has not been developed yet. Antimicrobial agents are effective against planktonic bacteria and appear to clear mucosal surfaces of adherent bacterial microcolonies but frequently fail to eradicate bacterial biofilms on urological devices. Several different approaches to disease prevention are being investigated and some promising results have been obtained.

References

  1. 1. Mardis HK, Kroeger RM 1988 Ureteral stents. Urol ClinNorth Am 15 471 479
  2. 2. Biering-Sorensen F. 2002 Urinary tract infection in individuals with spinal cord lesion. Current Opinion in Urology 12 45 49
  3. 3. Choong S. Whitfield H. 2000Biofilms and their role in infections in urology Brit J Urology 86 935 941
  4. 4. Costerton JW 1999 Introduction to biofilm.Int J Antimicrob Agents 11 217 221
  5. 5. Habash M. Reid G. 1999 Microbial Biofilms: Their development and significance for medical device-related infections.J Clin Pharmacology 39 887 898
  6. 6. Fletcher M. (ed 1996 Bacterial Adhesion: Molecular and Ecological Diversity.New York: Wiley-Liss
  7. 7. Busscher H. J. Stokoos I. Schakenraad J. M. 1991Two-dimensional spatial arrangement of fibronectin adsorbed to biomaterials with different wettabilities. Cells Mater 1: 49 57
  8. 8. Busscher HJ, Weerkamp AH 1987 Specific and non-specific interactions in bacterial adhesion to solid substrataFEMS Microbiol Rev 46 165 173
  9. 9. Van Loosdrecht MCM, Lyklema J, Norde W, Schraa G, Zehnder AJB (1987) The role of bacterial cell hydrophobicity in adhesion. Appl Environ Microbiol 53 1893 1897
  10. 10. Van Loosdrecht MCM, Lyklema J, Norde W, Schraa G, Zehnder AJB (1987) Electrophoretic mobility and hydrophobicity as a measure to predict the initial steps of bacterial adhesion. Appl Environ Microbiol 53 1989 1901
  11. 11. Denstedt J. D. Wollin T. A. Reid G. 1998 Biomaterials used in urology: Current issues of biocompatibility, infection and encrustation.J of Endourology 12 493 500
  12. 12. Lawrence JR, Caldwell DE 1987Behaviour of bacterial stream populations within the hydrodynamic boundary layers of surface microenvironments. Microbial Ecol 14 15 27
  13. 13. Reid G. Habash M. B. 1998Urogenital microflora and urinary tract infections. In, Tannock GW (ed.): Medical Importance of the Normal Microflora. London: Chapman & Hall 423 440
  14. 14. Densted J. D. Reid G. Sofer M. 2000Advances in ureteral stent tecnology. World J Urol 18 237 242
  15. 15. Costerton J. Lewandowski Z. Caldwell D. Korber D. Lappin-Scott H. 1995Microbial biofilms. Annu. Rev. Microbiol 49 711 745
  16. 16. Busscher G. J. Bos R. van der Mei H. C. 1995Initial microbial adhesion is a determinant for strength of biofilm adhesion. FEMS Microbiol Lett 128 229 234
  17. 17. Caldwell DE.Cultivation and study of biofilm communities. In Lappin Scott HM, Costerton JW eds Microbial Biofilms Cambridge: Cambridge University Press, 1195 4 69
  18. 18. Brown M. R. W. Collier P. J. Gilbert P. 1990 Influence of growth rate on susceptibility to antimicrobial agents: modification of the cell envelope and batch and continuous culture studies.Antimicrob Agents Chemother 34 1623 1628
  19. 19. Brown M. W. Allison D. G. Gilbert P. 1988Resistance of bacterial biofilms to antibiotics: a growth-related effect J Antimicrob Chemother 22 777 783
  20. 20. Goto T. Nakame Y. Nishida M. 1999 Bacterial biofilms and catheters in experimental urinary tract infection.Int. J of Antimicrob. Agents 11 227 231
  21. 21. Choong S. Wood S. Whitfield H. F. 2001 Catheter-associated urinary tract infection and encrustation.Int J of Antimicrobial Agents 17 305 310
  22. 22. Nickel J. C. Wright J. B. Ruseska I. Marrie T. J. Whitfield C. Costerton J. W. 1985Antibiotic resistance of Pseudomonas aeruginosa colonising a urinary catheter in vitro. Eur J Clin Microbiol 4 213 218
  23. 23. Goto T. Nakame Y. Nishida M. Oh Y. 1999 In vitro bactericidal activities of beta-lactamases, amikacin and fluoroquinolones against Pseudomonas aeruginosa biofilm in artificial urine. Urology 53 1058 1062
  24. 24. Tsukamoto T. Matsukawa M. Sano M. et al. 1999 Biofilm in complicated urinary tract infection.Int. J.of Antimicrob. Agents 11 233 236
  25. 25. Kumon H. 1996Pathogenesis and management of bacterial biofilms in the urinary tract. J Infect Chemother 2 18 28
  26. 26. Reid G. Habash M. 2001 Oral fluoroquinolone therapy results in drug adsorption on ureteral stents and prevention of biofilm formation.Int J of Antimicrob Agents 17 317 332
  27. 27. Reid G. Potter P. Dalenay G. Hsieh J. Nicoshia S. Hayes K. 2000 Ofloxacin for treatment of urinary tract infections and biofilms in spinal cord injury. IntJ Antimicrob. Agents 4 305 307
  28. 28. Shigeta M. Komatsuzawa H. Sugai M. Suginaka H. Usui T. 1997 Effect of the growth rate of Pseudomonas aeruginosa biofilms on the susceptibility to antimicrobial agents Chemotherapy 43 137 141
  29. 29. Reid G. 1999Biofilms in infectious diseases and on medical devices. Int. J. of Antimicrob.l Agents 11 223 226
  30. 30. Nickel J. C. Downey J. 1992Movement of pseudomonas aeruginosa along catheter surfaces. Urology39 93 98
  31. 31. Warren J. Bakke A. Desgranchamps F. Johnson J. R. Kumon H. Shah J. Tambyah P. 2000Catheter-Associated Bacteriuria and the Role of Biomaterial in Prevention. Nosocomial and Health Care Associated Infections In Urology 153 177
  32. 32. Warren J. 2001Catheter-associated urinary tract infections. Int J Antimicrob Agents 17 299 303
  33. 33. Liedl B. 2001Catheter-associated urinary tract infections. Current Opinion in Urology 11 75 79
  34. 34. Nickel JC 1991Catheter-associated urinary tract infection: new perspectives on old problems. Can J Infect Contrl 6 38 42
  35. 35. Ganderton L. Chawla J. Winters C. Wimpenny J. Stickler D. 1992Scanning electron microscopy of bacterial biofilms on indwelling bladder catheters. Eur J Clin Microbiol Infect Dis 11 789 797
  36. 36. Stickler D. J. Williams T. Jarman C. Howe N. Winters C. 1995The encrustation of urethral catheters. In: Wimpenny J, Handley P, Gilbert P. Lappin-Scott H, eds. The life and death of biofilm. Cardiff: Bioline 119 125
  37. 37. Kunin C. M. Chin Q. F. Chambers S. 1987Formation of encrustations on indwelling catheters in the elderly: a comparison of different types of catheter material in „blockers” and „non-blockers”. J Urol 138 899 902
  38. 38. Reid G. JD Denstedt Kang. Y. S. Lam D. Naus C. 1992Microbial adhesion and biofilm formation on ureteral stents in vitro and in vivo. J Urol 148 1592 1594
  39. 39. Farsi HMA, Mosli HA, Al-Zemaity 1995Bacteriuria and colonisation of double pigtail ureteral stents: long-term experience with 237 patients. J Endourol 9 469 472
  40. 40. Carson CC 1999Management of prosthesis infections in urologic surgery. Urol Clin North Am 26 829 839
  41. 41. Abouassaly, R., D.K. Montague, and K.W. Angermeier, Antibiotic-coated medical devices: with an emphasis on inflatable penile prosthesis.Asian J Androl, 2004 249 257
  42. 42. Carson, C.C., Diagnosis, treatment and prevention of penile prosthesis infection.Int J Impot Res, 2003Suppl 5: S139 S146
  43. 43. Licht MR, Montague DK, Angermeier KW et al 1995Cultures from genitourinary prostheses at re-operation: Questioning the role of Staphylococcus epidermidis in periprosthetic infection. J Urol 154 387 390
  44. 44. Fishman IJ, Scott FB, Selam IN 1987Rescue procedure: an alternative to complete removal for treatment of infected penile prosthesis J Urol 137: 202A
  45. 45. Rajpurkar, A., et al., Antibiotic soaked hydrophilic coated bioflex: a new strategy in the prevention of penile prosthesis infection.J Sex Med, 2004 215 220
  46. 46. Carson CC. 1989Infections in genitourinary prostheses. Ural Clin North Am 16 139 147
  47. 47. Nickel JC, Olson ME, Mclean RJ, Grant SK, Costerton JW 1987An ecological study of infected urinary stone genesis in an animal model. Br J Urol 59 21 3145
  48. 48. Nickel J. C. ME Olson Barabas. A. Benediktsson H. Dasgupta M. K. Costerton J. W. 1990Pathogenesis of chronic bacterial prostatitis in an animal model. B.J of Urol 66 47 54
  49. 49. Nickel J. C. ME Olson Ceri. H. 1993Experimental prostatitis. In Prostatitis (Weidner W, Madson P.O.P., Schiefer H.G, Eds). Springer-Verlag, Berlin
  50. 50. Justice S. S. Hung C. Theriot J. A. Fletcher D. A. Anderson G. G. MJ Footer Hultgren. S. J. Differentiation and developmental pathways of uropathogenic Escherichia coli in urinary tract pathogenesis. Proc Natl Acad Sci U S A. 2004Feb 3;101 5 1333 8
  51. 51. Nickel C. Costerton W. Mc Lean R. J. C. Olson M. 1994Bacterial biofilms: influence on the pathogenesis, diagnosis and treatment of urinary tract infections. Antimicrobial Chemotherapy 33 (Suppl. A): 31 41
  52. 52. Mysorekar IU and Hultgren SJ.Mechanisms of uropathogenic Escherichia coli persistence and eradication from the urinary tract. Proc Natl Acad Sci U S A. 2006Sep 19;103 38 14170 5
  53. 53. Mulvey MA, Schilling JD, Hultgren SJ.Establishment of a persistent Escherichia coli reservoir during the acute phase of a bladder infection. Infect Immun. 2001Jul;69 7 4572 9
  54. 54. Anderson G. G. Palermo J. J. JD Schilling Roth. R. et al. Intracellular bacterial biofilm-like pods in urinary tract infections Science. Washington: Jul 4, 2003 301Iss. 5629; 105
  55. 55. Schilling JD, Lorenz RG, Hultgren SJ.Effect of trimethoprim-sulfamethoxazole on recurrent bacteriuria and bacterial persistence in mice infected with uropathogenic Escherichia coli. Infect Immun. 2002Dec;70 12 7042 9
  56. 56. Nickel JC 1990The bottle of the bladder: the pathogenesis and treatment of uncomplicated cystitis Int Urogynecol J 1 218 222
  57. 57. Sobel, J.D., 2005 What’s new in bacterial vaginosis and trichomoniasis? Infect Dis Clin North Am,387 406
  58. 58. Hillier, S.L., et al., Association between bacterial vaginosis and preterm delivery of a low-birth-weight infant.The Vaginal Infections and Prematurity Study Group. N Engl J Med, 1995 1737 1742
  59. 59. Taha, T.E., et al., Bacterial vaginosis and disturbances of vaginal flora: association with increased acquisition of HIV.AIDS, 1998 1699 1706
  60. 60. Vasquez, A., et al., Vaginal lactobacillus flora of healthy Swedish women.J Clin Microbiol, 2002 2746 2749
  61. 61. Vallor, A.C., et al., Factors associated with acquisition of, or persistent colonization by, vaginal lactobacilli: role of hydrogen peroxide production.J Infect Dis, 2001 1431 1436
  62. 62. Hillier, S.L., et al., Characteristics of three vaginal flora patterns assessed by gram stain among pregnant women.Vaginal Infections and Prematurity Study Group. Am J Obstet Gynecol, 1992 938 944
  63. 63. Aroutcheva A. A. Simoes J. A. Faro S. Antimicrobial protein produced by vaginal Lactobacillus acidophilus that inhibits Gardnerella vaginalis. Infect Dis Obstet Gynecol, 2001 33 39
  64. 64. Fredricks, D.N., T.L. Fiedler, and J.M. Marrazzo, Molecular identification of bacteria associated with bacterial vaginosis. N Engl J Med, 2005. 353(18): 1899 1911
  65. 65. Sobel J. D. 2000Bacterial vaginosis. Annu Rev Med349 356
  66. 66. Nugent, R.P., M.A. Krohn, and S.L. Hillier, Reliability of diagnosing bacterial vaginosis is improved by a standardized method of gram stain interpretation.J Clin Microbiol, 1991 297 301
  67. 67. Srinivasan, S. and D.N. Fredricks, The human vaginal bacterial biota and bacterial vaginosis. Interdiscip Perspect Infect Dis, 2008. 2008: 750479
  68. 68. Swidsinski A. et al. Adherent biofilms in bacterial vaginosis. Obstet Gynecol, 2005Pt 1): 1013 1023
  69. 69. Gardner, H.L. and C.D. Dukes, Haemophilus vaginalis vaginitis: a newly defined specific infection previously classified non-specific vaginitis.Am J Obstet Gynecol, 1955 962 976
  70. 70. Wilson J. Managing recurrent bacterial vaginosis.Sex Transm Infect, 200480(1):8 11
  71. 71. Patterson, J.L., et al., Effect of biofilm phenotype on resistance of Gardnerella vaginalis to hydrogen peroxide and lactic acid.Am J Obstet Gynecol, 2007 170e1-7.
  72. 72. Saunders, S., et al., Effect of Lactobacillus challenge on Gardnerella vaginalis biofilms.Colloids Surf B Biointerfaces, 2007 138 142
  73. 73. Muli F. Struthers J. K. Use of a continuous-culture biofilm system to study the antimicrobial susceptibilities of Gardnerella vaginalis and Lactobacillus acidophilus.Antimicrob Agents Chemother, 1998 1428 1432
  74. 74. Swidsinski, A., et al., An adherent Gardnerella vaginalis biofilm persists on the vaginal epithelium after standard therapy with oral metronidazole.Am J Obstet Gynecol, 2008 97e1-6.
  75. 75. Thibon P. Le Coutour X. Leroyer R. Fabry J. 2000Randomized multi-centre trial of the effects of a catheter coated with hydrogel and silver salts on the incidence of hospital-acquired urinary tract infections. J. Hosp. Infect. 45 117 1124
  76. 76. Schumm K. Lam T. B. Types of urethral catheters for management of short-term voiding problems in hospitalised adults.Cochrane Database Syst Rev, 2008 CD004013
  77. 77. De Ridder, D.J., et al., Intermittent catheterisation with hydrophilic-coated catheters (SpeediCath) reduces the risk of clinical urinary tract infection in spinal cord injured patients: a prospective randomised parallel comparative trial.Eur Urol, 2005 991 995
  78. 78. Sarica, S., et al., Comparison of the use of conventional, hydrophilic and gel-lubricated catheters with regard to urethral micro trauma, urinary system infection, and patient satisfaction in patients with spinal cord injury: a randomized controlled study.Eur J Phys Rehabil Med, 2010 473 479
  79. 79. Riedl, C.R., et al., Heparin coating reduces encrustation of ureteral stents: a preliminary report.Int J Antimicrob Agents, 2002 507 510
  80. 80. Hazan Z, Zumeris J, Jacob H, Raskin H, Kratysh G, Vishnia M, Dror N, Barliya T, Mandel M, Lavie G. 2006. Effective Prevention of Microbial Biofilm Formation on Medical Devices by Low-Energy Surface Acoustic Waves. Antimicrob Agents Chemother. 2006 Dec; Epub 2006 Aug 2812 50 4144 52
  81. 81. Ikinger U., Zillich S., Weber C. 2007. Biofilm Prevention by Surface Acoustic Nanowaves: A New Approach to Urinary Tract Infections? 25th World Congress of Endourology and SWL Cancun, Mexico, October 2007
  82. 82. Hazan, Z., et al., Effective prevention of microbial biofilm formation on medical devices by low-energy surface acoustic waves.Antimicrob Agents Chemother, 2006 4144 4152
  83. 83. Ikinger U. Zillich S. Weber C. Biofilm Prevention by Surface Acoustic Nanowaves: A New Approach to Urinary Tract Infections?Poster presented at: 25th World Congress of Endourology and SWL; 2007Cancun, Mexico.
  84. 84. Nagy K. Koves B. Tenke P. The effectiveness of acoustic energy induced by UroShield device in the prevention of bacteriuria and the reduction of patients’ complaints related to long-term indwelling urinary catheters. Poster accepted to: 26th Annual EAU Congress; 2011March 18 22Vienna, Austria

Written By

Peter Tenke, Bela Koves, Karoly Nagy, Shinya Uehara, Hiromi Kumon, Scott J. Hultgren, Chia Hung and Werner Mendling

Submitted: 14 December 2010 Published: 06 September 2011