Medical Biofilms - American Chemical Society

Chapter 4

Medical Biofilms Kedar Diwakar Mandakhalikar*

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National Heart Centre Singapore, 5 Hospital Drive, Level 9, Singapore 169609 *E-mail: [email protected].

Biofilm formation by bacteria plays a very important role in healthcare-associated infections. It enhances their pathogenesis and severity and causes remarkable human morbidity and mortality. Numerous types of medical devices and surgical implants used in patients provide surfaces for bacterial attachment and compromise host immune responses. Most biofilm infections are associated with medical devices or implants. Biofilm infections are also seen intracellularly or in wounds. It is important to understand the complex processes of bacterial biofilm formation and the interplay between host immunity, biotic or abiotic surfaces, and bacterial biofilms to decide on a therapeutic strategy for combatting these infections. In this chapter, we describe medical biofilms briefly with respect to their formation, importance, and control.

Bacterial infections are one of the major causes of human morbidity and mortality. Biofilm formation by bacteria enhances the pathogenesis and severity of infections considerably. While the universal presence and significance of biofilms are acknowledged, the role of biofilms in healthcareassociated infections (HCAIs) should not be underestimated. In addition to bacteria, yeasts, fungi, protozoa, and viruses have also been isolated from medical biofilms (1–3); however, the scope of this chapter will be limited to bacterial biofilms. We describe the significance of biofilms in the clinical context by answering the following questions: 1. 2. 3. 4.

What are biofilms? Where do medical biofilms form? Why are medical biofilms important? How can biofilms be prevented or treated?

© 2019 American Chemical Society

Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

What Are Biofilms? Bacterial biofilms are multicellular colonies of bacteria embedded in a self-generated extracellular matrix (ECM). Well-coordinated, cooperative association between cells is a distinctive feature of bacterial biofilms that makes them similar to multicellular organisms in certain processes (4, 5). Examples of ex vivo bacterial biofilms are shown in the scanning electron micrographs in Figure 1 (6).

Figure 1. Single-species bacterial biofilms (A and B), and multiple-species bacterial biofilms (C and D). White arrows indicate bacterial cells embedded in the matrix. Adapted with permission from reference (6). Kedar Diwakar Mandakhalikar Copyright 2018. It has been extensively shown that biofilm formation is the predominant mode of bacterial life (7). Either in nature or as pathogens, bacteria prefer to exist as communities attached to a surface, as opposed to the free-swimming planktonic lifestyle. It has also been suggested that the planktonic mode is only an evolutionary step for easy dispersal, whereas bacteria are naturally inclined to attach to a surface (8, 9). Formation of biofilm is a defensive as well as survival mechanism adopted by bacteria. Overall, biofilm formation can be briefly described with the sequence of events shown in Figure 2: reversible attachment to a surface, irreversible attachment, colonization, maturation, and dispersal. The formation begins with the nonspecific reversible attachment of planktonic bacterial cells to a conditioned surface followed by specific irreversible attachment (11). Bacteria can attach to biotic as well as abiotic surfaces with the help of many factors such as bacterial motility and electrostatic 84 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

interactions between the bacterial cells and surface. Nonspecific forces such as Van der Waals interactions bring about the initial bacterial attachment to abiotic surfaces where bacteria behave like colloidal microparticles (12). Various factors such as the pH of the medium as well as the hydrophobicity of the surface play an important role in the irreversible attachment of bacteria to a surface. Medical implants are known to be coated within nanoseconds with proteins from either blood or interstitial fluid. These proteins, along with polysaccharides and extracellular DNA, form a conditioning film on the surface that strongly aids the irreversible bacterial attachment by changing the physicochemical properties of the surface (13, 14).

Figure 2. Schematic representation of biofilm formation. (i) Reversible attachment of planktonic bacteria to surfaces. (ii) Irreversible attachment to surfaces. (iii) Formation of the external matrix. (iv) Biofilms acquire a three-dimensional structure. (v) Biofilm detachment. Reproduced with permission from reference (10). Sara M. Soto Copyright 2014. Once bacteria attach to a surface, they proliferate and grow gradually over time to develop into a mature biofilm. The ECM is secreted by the bacteria, and it covers the cells individually as well as the whole colony. The ECM or the extracellular polymeric substance, previously also called glycocalyx, is mainly composed of polysaccharides (15). This chemically complex matrix stores high amounts of moisture and nutrients and can capture other microorganisms as well as minerals and waste products (16, 17). Biofilm dispersal is the step in which parts of biofilm or specialized planktonic bacterial cells slough off from the surface of mature biofilm and attach to other available sites, disseminating the biofilm within the host. These cells are different from the bacteria that break out from the biofilm due to disturbance or adverse conditions. The biofilm dispersal is initiated by multiple signals such as alterations in nutrients, temperature, and oxygen concentration. It can also be impacted by other microbes releasing quorum-sensing (QS) molecules (e.g., lactones and peptides) (9, 18). The phenomenon of QS plays a critical role in biofilm formation and maintenance. It is a way of communicating between bacterial cells in the biofilm. This cell-to-cell signaling modulates the gene expression in the biofilm bacteria via various chemical molecules called autoinducers (19). The autoinducers are constitutively produced, and their concentration is known to rise with increasing density of the biofilm. These changes in gene expression bring about modulation in various physiological processes such as motility and the release of virulence factors (20). This leads to bacterial cells being in different physiological stages within a biofilm. Different bacteria also differ in the signal molecules that they release. For example, Gram-positive bacteria secrete oligopeptides, whereas Gram-negative bacteria use lactones (7).

85 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

Encrustation as a Complication of Biofilm Certain bacteria such as Proteus mirabilis secrete the enzyme urease that converts urea into ammonia and carbon dioxide. Ammonia increases the pH of the medium, causing the precipitation of mineral salts in crystal form, which is termed encrustation. The crystals of the salts provide additional surfaces for the bacteria to attach to, which makes eradication of crystalline biofilms even more difficult. Encrustation as a complication of biofilm formation is a major concern, especially in urinary tract infections (Figure 3A) (21, 22). These crystalline biofilms are implicated in urinary stone formation. When they are formed on the outer surface of the catheter, they can cause damage to the bladder and urethral epithelia. The catheter lumen can become blocked due to the crystalline biofilm preventing the flow of urine through the catheter, thus making the catheter unusable (Figure 3B, 3C) (23).

Figure 3. Encrustation in ex vivo urinary catheters. (A) Deposition of mineral salts near the eye and the balloon of Foley catheter. (B) Partial and (C) complete blockage of urinary catheter lumen due to encrustation.

Where Do Medical Biofilms Form? It has been reported that nearly 80% of bacterial infections in humans are linked to biofilm formation (24). Bacterial biofilms are increasingly being recognized as a vital cause of highly recalcitrant, chronic infections, especially in patients with medical devices (25). In the literature, about one-quarter to one-half of all the HCAIs are attributed to medical devices (26, 27). The most frequently reported HCAIs involving biofilms are ventilator-associated pneumonia (VAP), lower respiratory tract infections, catheter-associated urinary tract infections (CAUTI), and surgical-site infections. In addition, dental biofilms are implicated in a significant percentage of chronic oral diseases. The different bacterial genera isolated from medical-device biofilms vary according to the physiological system involved and, in addition to pathogenic bacteria, can also be normal microflora (28). The pathogens regularly associated with HCAIs include but are not limited to: 1. Staphylococcus aureus, Staphylococcus epidermidis, and Enterococcus faecalis (Gram-positive) 2. Escherichia coli, Klebsiella pneumoniae, Acinetobacter baumannii, Proteus mirabilis, and Pseudomonas aeruginosa (Gram-negative) 3. Candida species (yeast) (29) Multidrug-resistant bacteria are being extensively identified in long-term care facilities and acute care hospitals, which further intensifies the seriousness of the problem (30–32). Additionally, bacteria previously not known to be biofilm-forming have also been reported in various cases of infectious diseases (e.g., melioidosis caused by Burkholderia pseudomallei (33) and chronic osteomyelitis caused by a biofilm-producing strain of Morganella morganii (34)). 86 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

Medical Biofilms Associated with Medical Devices The formation of biofilms on medical devices and the resulting infections were first validated in 1972 (35). Since then, biofilms have been extensively shown to develop on the surfaces of medical devices. It has been previously shown in a rabbit model that a substantially lower bacterial titer is required for infection at a surgical site in the presence of a foreign body than in the absence of one (36). Medical devices can get contaminated through contact with skin, contaminated water, or other sources, initiating biofilm formation and further infection. Medically implanted invasive devices that come in contact with mucous membranes such as urinary catheters can easily act as a channel for the migration and entry of bacteria. Additionally, due to the biofilm dispersal capability, these devices pose a considerable risk of systemic (bloodstream) bacterial spread within the host and also of chronic or relapsing infection (37). Even though the risk of infection differs, a surgically implanted invasive device is always considered a foreign body irrespective of the factors such as its location in human body, the type of device, and the indwelling time. Infection of an implanted device is influenced by complex mechanisms of the host immune response to both the implant as well as to the invading bacteria. Host immune responses typically clear out contaminations or minor infections by opportunistic bacteria in the absence of a foreign body. However, fibrous encapsulation as an immune response to the medical device (foreign body) creates an area of low immune response, thereby allowing bacterial colonization and proliferation (38). This area of low immunity is called a locus minoris resistentiae, which is unguarded against less virulent, opportunistic bacteria (39). With an estimated 500,000 types of medical devices available globally implanted in various anatomic sites of the human body, every type of human tissue comes in contact with these biomaterials, increasing the risk of biofilm formation and infection (40). Bacterial biofilms have been frequently associated with a wide range of polymeric medical devices, such as catheters and cardiac pacemakers (9). It has been reported that most, if not all, medical or prosthetic devices may result in biofilm infections. Some such implants that have been studied are endotracheal tubes (41), urinary catheters (42), central venous catheters (43), peritoneal dialysis catheters (44), artificial voice prostheses (45), joint prostheses and orthopedic fixation devices (46, 47), cardiac pacemakers (48), cerebrospinal fluid shunts (49), vascular prostheses (50), prosthetic heart valves (29), contact lenses (51), biliary tract stents (52), prosthetic devices for erectile dysfunction (53), dentures (54), and breast implants (55, 56). It has been seen that, initially, a single bacterial species colonizes the medical implant (Figure 1A, 1B), but as the biofilm grows, multiple bacterial species quickly form consortia in the biofilm (Figure 1C, 1D) (57). The phenotype of a biofilm also depends upon the bacteria involved and changes with the variety of bacteria for different biofilms (58, 59). Bacterial colonization and biofilm formation on central venous catheters have been shown to occur as early as 24 h after catheter insertion, irrespective of the clinical status of the patient or the bacteria involved (60). VAP is commonly observed after two to three days in patients who are on mechanical ventilation with breathing tubes in place. Biofilm formation on the endotracheal tube has been reported even within 24 h of its insertion (61). Mortality due to VAP is naturally notably higher than mortality due to other biofilm-related infections such as urinary tract infections or wound infections (62). In addition to mortality, VAP leads to prolonged hospital stays and very high healthcare costs, thus greatly affecting both the patient and the healthcare resources (63). Bacteria commonly associated


with VAP are A. baumannii, Pseudo. aeruginosa, K. pneumoniae, Staph. aureus, and Candida spp. (64, 65) Urinary catheters are among the most common medical devices used in clinics. Among other factors, the duration of catheterization is very important, and the risk of CAUTI is between 3 and 7% daily (66). The bacteria that colonize the periurethral region may contaminate the catheter, migrating into the bladder via the mucoid layer between the urethra and the catheter. Contamination within the urine drainage bag can also be a source of bacteria that migrate to the bladder and cause infection (67). The most prevalent pathogens that cause CAUTI are uropathogenic E. coli, Enterococcus spp., K. pneumoniae, Prot. mirabilis, Pseudo. aeruginosa, Candida spp., group B Streptococcus, and others (68, 69). The use of central venous catheters for venous access is a common clinical practice to instill blood products or other fluids and for hemodialysis. Catheter-related biofilms can lead to lifethreatening infections such as endocarditis or sepsis (70). Biofilms can form on the inner lumen as well as on the outer surface of the catheter. Catheter-associated bloodstream infections occur within the first week of catheterization mainly due to extraluminal biofilms. On the other hand, mainly intraluminal biofilm formation is seen in patients with long-term catheterization (more than 30 days) leading to a higher risk of systemic infection (71, 72). Coagulase-negative staphylococcus, Staph. aureus, and Candida spp., as well as Gram-negative bacilli such as Enterobacter spp., Klebsiella spp., Stenotrophomonas, Pseudomonas, and Acinetobacter species have been known to be the major pathogens in catheter‐related infections (73, 74). In a recent preliminary clinical study, Staphylococcus spp. was reported to be the most frequently detected genus in catheter-related infections (75). However, biofilms may or may not be associated with a foreign body. Medical Biofilms Not Associated with Foreign Body Some examples of biofilms that are not related to indwelling medical devices are chronic airway infections in individuals with cystic fibrosis, native valve endocarditis, chronic otitis media (inflammatory diseases of middle ear), chronic sinusitis, and diabetic wound infections (76, 77). The introduction of microbes to a surgical site can occur through contaminated medical instruments or transfer from the skin of either the patient or a healthcare professional. Dental Biofilms Normal oral microflora containing more than 700 bacterial species, including those in biofilm form, are beneficial to the host, as they prevent colonization by pathogenic bacteria. When this microbial homeostasis is disturbed due to factors such as bad oral hygiene, diet changes, or medical intervention, opportunistic pathogens can establish biofilms that may lead to oral disease (78). Biofilms that lead to diseases in teeth and their supporting tissues are complex, may contain up to 100 bacterial species, and have been shown to vary significantly with respect to the bacterial species involved depending upon the availability of nutrients and oxygen, the location in the oral cavity, and the individual (79, 80). For example, Gram‐positive bacteria, in the presence of abundant carbohydrates, cause demineralization of teeth and dental caries. On the other hand, anaerobic Gram‐negative bacteria in subgingival biofilms can cause gingivitis and chronic periodontitis (81). The three major bacterial genera involved in dental biofilms are the Streptococcus mitis group, Actinomyces spp., and Fusobacterium spp. (82) Other bacterial genera shown to be important in 88 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

forming dental biofilms include Veillonella, Granulicatella, Neisseria, Haemophilus, Corynebacterium, Rothia, Prevotella, Capnocytophaga, and Porphyromonas (83). Biofilms and Wounds A wound, either surgical or as a result of other trauma, is an injury to live tissue mainly due to a cut or breach in the skin. The wound provides a surface for the attachment of bacteria. The woundhealing mechanism also provides an extremely favorable environment that is rich in nutrition for the growth and pathogenicity of bacteria. It is well-established that at least half of all chronic wounds contain biofilm. Some studies have shown that more than 90% of chronic wounds contain bacterial and fungal biofilms, whereas only 6% of acute wounds show the presence of biofilms (59, 84, 85). All wounds are highly likely to get contaminated by bacteria. Once bacteria colonize, it is very difficult to dislodge them due to the formation of biofilm. In addition, wound dressing has been hypothesized to act as a bioreactor, providing an additional reservoir for both planktonic and biofilm bacteria. Bacterial biofilm formation can hamper the wound-healing process remarkably for a variety of reasons, including diversion of resources, extra damage due to the host’s immune reaction to bacteria, and a chronic low-grade inflammatory response (85). It has also been reported that the bacterial isolates from wounds have a higher likelihood of biofilm formation than the normal skin microflora (86). In addition, it has been shown that biofilms in chronic wounds are multispecies bacterial infections, making it more difficult to treat them with antimicrobials. It has been shown that polymicrobial biofilms delay wound healing substantially as compared to single-species biofilms (87). Other Clinically Relevant Biofilms In some cases, persistent chronic infections, in spite of a strong host immune reaction, were explained with the discovery of intracellular bacterial communities (IBCs) (88). The IBCs were first reported for uropathogenic E. coli in bladder infections and have also been found in K. pneumoniae (89, 90). The IBCs are biofilm-like pods that contain bacterial cells enveloped in a polysacchariderich matrix with a protective shell of uroplakin. It has been shown that dry-surface biofilms are a persistent source of pathogens such as drugresistant Staph. aureus in hospitals, and polymicrobial biofilms have been retrieved from the majority of the surfaces tested from hospital wards (91–93). The initial contamination of the medical device most likely occurs because of a small number of microorganisms, which are often transferred to the medical device via the patient’s or healthcare workers skin, contaminated water, dry surface biofilms, or other external environmental sources (30, 32). Therefore, it is important to highlight here that storage tanks, hydrotherapy pools, dialysis water systems, humidifiers, and air-conditioning systems in hospitals can have biofilms growing in them and may become the source of contamination and infection in immunocompromised individuals (94). Similarly, it has been suggested that dental-unit water systems expose patients to opportunistic bacteria because of the biofilms inside the water lines (57, 95). The importance of water distribution systems in hospitals cannot be overstated and should not be overlooked (96).


Why Are Medical Biofilms Important? Various aspects of increased bacterial virulence in humans are directly associated with the ability of bacteria to form biofilms, as this leads to highly enhanced bacterial survival and growth. Biofilm formation confers definite selective growth advantages to the bacteria, especially during scarcity of nutrients, compared to planktonic bacteria. Formation of polymicrobial synergistic consortia is a key feature of biofilms. For better utilization of limited resources, polymicrobial biofilms have been shown to maintain commensalism among different species of mixed biofilm communities (97). Bacteria not only survive in biofilms, but also continuously regenerate and disseminate during the dispersal phase. Furthermore, biofilm acts as a reservoir for drug-resistant bacteria, compounding the clinical problems. The ECM plays an enormous role in protecting the bacterial cells in a biofilm. In addition to providing architectural stability, it also makes the structure impenetrable by antimicrobials as well as the host immune cells. It has been shown that up to 5000 times more antimicrobial agent is required to have an inhibitory response against bacteria in biofilm than against planktonic bacteria (98). The ECM has also been reported to dilute as well as neutralize the antimicrobial agents by creating a diffusion barrier (9). However, even though reduced penetration of the ECM by antimicrobial agents is well established in the literature, the ECM provides more resistance against some antimicrobials than others and does not work against all agents (99). The ECM can also protect the biofilm bacteria against exposure to ultraviolet light, acids, phagocytosis, metal toxicity, and dehydration (9, 100). On top of the structural benefits provided by the ECM, biofilm bacteria also undergo different physiological changes than planktonic cells. In addition to a modified phenotype, subpopulations of bacteria in a biofilm differ considerably from planktonic bacteria with respect to gene expression in response to nutrient and oxygen availability, as well as cell-to-cell interaction (24, 101). Bacteria are known to activate certain pathways, such as the rpoS-mediated general stress response, and adjust the QS pathways to protect against antimicrobial compounds (102). Biofilm bacteria have also been shown to alter the cell-membrane transport systems and bacterial surface-associated molecules that can bind to and neutralize antimicrobial agents (103). Transfer of genetic material in the form of plasmids between bacterial cells occurs more frequently in biofilms, thereby rapidly spreading the antimicrobial resistance (104). Since polymicrobial biofilms are very common in HCAIs, as opposed to single-species biofilms, various resistance mechanisms of different bacterial species can act synergistically. This can be one of the reasons for the high multidrug resistance to antimicrobial agents regularly reported in medical biofilms (105). Furthermore, the microenvironment within biofilms is modified to suit the bacterial cells; antimicrobials may not be able to function due to factors such as altered nutrient levels or reduced oxygen. Some bacterial cells in a biofilm display slower metabolic and growth rates, making them resistant to antimicrobials that target the metabolic pathways and require a certain degree of physiological activity. Even though these metabolically inactive, nondividing, “persister cells” do not have the genetic mutations for drug resistance, they are still highly multidrug resistant (106–108). It has been reported that sublethal concentrations of antimicrobials activate the formation of persister cells, and since the concentration of antimicrobials required to control biofilm bacteria is much higher, it is natural that persisters are observed in biofilms (109). Biofilm formation aids in the rise and persistence of multidrug-resistant bacterial strains that pose a huge infection risk to the human population. The biofilm mode of bacteria is associated 90 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

with several chronic infections such as cystic fibrosis, where Pseudo. aeruginosa forms biofilms in the patient’s lungs. Colonization in such cases has been reported to be a lifelong problem in spite of using strong antimicrobials (110, 111). As many biofilm infections are associated with a medical implant, the formation of biofilm renders that medical device inoperative. Some cases, such as with cardiovascular implants, require an elaborate surgical procedure to replace the infected medical device. Additionally, dispersal of a device-associated biofilm may lead to a systemic infection, especially in immunocompromised individuals (29). These complications extend the hospital stay of the patients as well as excessively burden the healthcare system. It can be inferred from these examples that, in addition to high patient morbidity and mortality, a vast financial burden on healthcare services can be attributed to biofilm infections (72).

How Can Biofilms Be Prevented or Treated? Treatment The age and composition of biofilms have a big influence on the outcomes of antibiofilm therapy. Biofilms grow and mature very rapidly. There is only a short period where the cells are physiologically active and the ECM is still penetrable for antimicrobial agents. Therefore, to determine an accurate treatment schedule, it is very important to quickly identify the genus of the infecting bacteria as well as its antimicrobial susceptibility. Viable but nonculturable bacteria and small-colony variants pose a diagnostic challenge through traditional microbiological methods and may lead to false negatives (112, 113). Furthermore, considering the speed limitations of the traditional microbiological methods, molecular tools such as polymerase chain reactions are recommended for a rapid diagnosis (114). With less time pressure, the molecular diagnosis can be confirmed using the routine microbiological testing. In absence of a “gold standard” method for extraction of biofilm, we previously developed a method using sonication and vortexing for extracting bacterial biofilm from samples collected from in vitro or in vivo experiments as well as from human patients (115). It has been recommended that for a precise diagnosis, a combination of traditional and molecular methods should be utilized. It was reported that biofilms develop resistance to antimicrobial therapy only 2 to 4 days after formation, which is likely due to active cells and weakly developed ECM in the early stages. Newly formed biofilms were shown to be more susceptible than mature biofilms to antimicrobial treatment as well as to the host immune response. The genera of the bacteria involved is an important factor in deciding the duration of therapy against biofilm formation. Furthermore, polymicrobial biofilms are extremely difficult to treat unless a very specific approach is considered. Each cluster of bacteria within a biofilm forms a microcolony with differing characteristics. Exploring the vulnerabilities of each bacterial genus involved in a multispecies biofilm can achieve this effect to some extent. Such a therapy requires a combination of multiple antimicrobial agents (116). These factors justify the recommendations for an early and aggressive treatment by a combination of antimicrobial agents for an appropriate duration (117, 118). Several innovative approaches have also been reported, such as QS molecule inhibitors, as QS plays a critical role in biofilm development (119). Biofilm infections are a very difficult and complex problem for microbiologists and clinicians. Antibiotic treatment alone seems to be an incomplete solution to this problem. Rather than treating the infecting biofilms individually, a systematic multidisciplinary approach to antibiofilm therapy has been suggested for better clinical results. This multidisciplinary approach includes but may not be limited to the following steps (76, 120–122): 91 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

1. 2. 3. 4.

Removal of the infected medical devices Selection of ECM-penetrating combination of antibiotics specific to the bacteria involved Systemic or topical antimicrobials in high doses for appropriate duration Use of novel technology such as QS inhibitors and amyloid-like fibers inhibitors

Prevention Complete removal of the biofilm associated with the wound is said to be unlikely, as the biofilm can spread below the surface of the wound and reform very rapidly (123, 124). Since biofilm infections begin with contamination by a small number of bacteria, it is highly recommended to adhere to various clinical guidelines regarding hygiene protocols (e.g., National Institute for Health and Care Excellence guidelines and Comprehensive Unit-Based Safety Program) (125, 126). Additionally, several important guidelines and reports have been published to improve understanding of the mechanisms of biofilm-associated infections (127, 128). Antimicrobial coating and surface alterations of medical devices have shown promising but inconsistent results with respect to clinical outcomes in the prevention of biofilm formation on medical devices (37, 129). The aim is to prevent initial attachment onto biomaterials with the use of antimicrobial agents as well as changing the physicochemical properties of the medical device surfaces (130). One major limitation of this approach is the quick release of the antimicrobial agent or deposition of the conditioning film on the device surface neutralizing the effect of coating. To overcome this challenge, we have recently reported a novel antifouling coating on a urinary catheter with controllable and sustained silver release in a proof-of-concept in vivo study (130). There are many more novel antimicrobial technologies and coatings being developed and studied, including bio-reduced graphene oxide (131) and copper-based composite coating (132). The majority of bacterial studies use the planktonic bacteria in liquid cultures as a model and have helped us better understand the physiology of a pure culture. However, these studies do not correlate well with the natural growth of bacteria as biofilm, and there is a need to emphasize the biofilm mode of microbial life (5). More studies on this aspect will also help identify molecules as targets for novel therapeutic approaches.

Conclusion Biofilm infections are a serious medical problem. Medical devices, surgical implants, and wounds provide surfaces for primary bacterial attachment and compromise host immune reactions. Bacteria form biofilms and use several other mechanisms to survive by evading antimicrobial agents and host immunity, leading to high morbidity and mortality in patients, prolonged hospitalization, severe financial losses, and excessive strain on critical healthcare resources. Therefore, prevention of bacterial colonization on biomaterials and in wounds is vital. An ideal biomaterial should have similar properties to a healthy biological surface or a membrane that would minimize bacterial attachment and thus limit biofilm formation and infection. A multipronged, early, and aggressive therapeutic approach, along with a quick and accurate diagnosis, is necessary to control biofilm infections. It can only be done with a better understanding of the complex processes of biofilm formation, bacterial survival, communication, proliferation, and antimicrobial resistance. Further understanding the interactions of bacteria with biotic and abiotic surfaces and the role of host immunity in this interplay can help in establishing potential antibiofilm therapeutic strategies. 92 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

References 1. 2. 3. 4. 5. 6.

7. 8. 9. 10. 11. 12.

13. 14. 15. 16. 17. 18. 19. 20. 21. 22.

Jabra-Rizk, M. A.; Falkler, W. A.; Meiller, T. F. Fungal biofilms and drug resistance. Emerg. Infect. Dis. 2004, 10 (1), 14–19. Patterson, T. F. Advances and challenges in management of invasive mycoses. Lancet 2005, 366 (9490), 1013–1025. Laffey, S. F.; Butler, G. Phenotype switching affects biofilm formation by Candida parapsilosis. Microbiology 2005, 151 (Pt. 4), 1073–1081. Davey, M. E.; O’Toole, G. A. Microbial biofilms: from ecology to molecular genetics. Microbiol. Mol. Biol. Rev. 2000, 64 (4), 847–867. Parsek, M. R.; Fuqua, C. Biofilms 2003: emerging themes and challenges in studies of surface-associated microbial life. J. Bacteriol. 2004, 186 (14), 4427–4440. Mandakhalikar, K. D. Prevention of infection in urology patients through the use of novel devices. Ph.D. Thesis, National University of Singapore, Singapore, 2018; http://scholarbank.nus.edu.sg/handle/10635/ 149509. Lindsay, D.; von Holy, A. Bacterial biofilms within the clinical setting: what healthcare professionals should know. Journal of Hospital Infection 2006, 64 (4), 313–325. Costerton, B. Microbial ecology comes of age and joins the general ecology community. Proc. Natl. Acad. Sci. U. S. A. 2004, 101 (49), 16983–1684. Hall-Stoodley, L.; Costerton, J. W.; Stoodley, P. Bacterial biofilms: from the natural environment to infectious diseases. Nat. Rev. Microbiol. 2004, 2 (2), 95–108. Soto, S. M. Importance of biofilms in urinary tract infections: new therapeutic approaches. J. Adv. Biol. 2014, 2014, 13. Dunne, W. M., Jr. Bacterial adhesion: seen any good biofilms lately? Clin. Microbiol. Rev. 2002, 15 (2), 155–166. Ribeiro, M.; Monteiro, F. J.; Ferraz, M. P. Infection of orthopedic implants with emphasis on bacterial adhesion process and techniques used in studying bacterial-material interactions. Biomatter 2012, 2 (4), 176–194. Franz, S.; Rammelt, S.; Scharnweber, D.; Simon, J. C. Immune responses to implants - a review of the implications for the design of immunomodulatory biomaterials. Biomaterials 2011, 32 (28), 6692–6709. Geng, J.; Henry, N. Short time-scale bacterial adhesion dynamics. Adv. Exp. Med. Biol. 2011, 715, 315–331. Branda, S. S.; Vik, S.; Friedman, L.; Kolter, R. Biofilms: the matrix revisited. Trends Microbiol. 2005, 13 (1), 20–26. Costerton, J. W.; Irvin, R. T.; Cheng, K. J. The bacterial glycocalyx in nature and disease. Annu. Rev. Microbiol. 1981, 35, 299–324. Costerton, J. W.; Cheng, K. J.; Geesey, G. G.; Ladd, T. I.; Nickel, J. C.; Dasgupta, M.; Marrie, T. J. Bacterial biofilms in nature and disease. Annu. Rev. Microbiol. 1987, 41, 435–464. Kaplan, J. B. Biofilm dispersal: Mechanisms, clinical implications, and potential therapeutic uses. J. Dent. Res. 2010, 89 (3), 205–218. Parsek, M. R.; Greenberg, E. P. Sociomicrobiology: the connections between quorum sensing and biofilms. Trends Microbiol. 2005, 13 (1), 27–33. Mangwani, N.; Dash, H. R.; Chauhan, A.; Das, S. Bacterial quorum sensing: functional features and potential applications in biotechnology. J. Mol. Microbiol. Biotechnol. 2012, 22 (4), 215–227. Stickler, D.; Young, R.; Jones, G.; Sabbuba, N.; Morris, N. Why are Foley catheters so vulnerable to encrustation and blockage by crystalline bacterial biofilm? Urol. Res. 2003, 31 (5), 306–311. Stickler, D.; Ganderton, L.; King, J.; Nettleton, J.; Winters, C. Proteus mirabilis biofilms and the encrustation of urethral catheters. Urol. Res. 1993, 21 (6), 407–411.


23. Percival, S. L.; Sabbuba, N. A.; Kite, P.; Stickler, D. J. The effect of EDTA instillations on the rate of development of encrustation and biofilms in Foley catheters. Urol. Res. 2009, 37 (4), 205–209. 24. Attinger, C.; Wolcott, R. Clinically addressing biofilm in chronic wounds. Adv. Wound Care (New Rochelle) 2012, 1 (3), 127–132. 25. Donlan, R. M. Biofilms: microbial life on surfaces. Emerg. Infect. Dis. 2002, 8 (9), 881–890. 26. Paredes, J.; Alonso-Arce, M.; Schmidt, C.; Valderas, D.; Sedano, B.; Legarda, J.; Arizti, F.; Gomez, E.; Aguinaga, A.; Del Pozo, J. L.; Arana, S. Smart central venous port for early detection of bacterial biofilm related infections. Biomed. Microdevices 2014, 16 (3), 365–374. 27. Magill, S. S.; Edwards, J. R.; Bamberg, W.; Beldavs, Z. G.; Dumyati, G.; Kainer, M. A.; Lynfield, R.; Maloney, M.; McAllister-Hollod, L.; Nadle, J.; Ray, S. M.; Thompson, D. L.; Wilson, L. E.; Fridkin, S. K. Emerging Infections Program Healthcare-Associated Infections and Antimicrobial Use Prevalence Survey Team, Multistate point-prevalence survey of health care-associated infections. N. Engl. J. Med. 2014, 370 (13), 1198–1208. 28. Vandecandelaere, I.; Coenye, T. Microbial composition and antibiotic resistance of biofilms recovered from endotracheal tubes of mechanically ventilated patients. Adv. Exp. Med. Biol. 2015, 830, 137–155. 29. Donlan, R. M. Biofilms and device-associated infections. Emerg. Infect. Dis. 2001, 7 (2), 277–281. 30. von Eiff, C.; Heilmann, C.; Peters, G. New aspects in the molecular basis of polymer-associated infections due to staphylococci. Eur. J. Clin. Microbiol. Infect. Dis. 1999, 18 (12), 843–846. 31. Niveditha, S.; Pramodhini, S.; Umadevi, S.; Kumar, S.; Stephen, S. The isolation and the biofilm formation of uropathogens in the patients with catheter associated urinary tract infections (UTIs). J. Clin. Diagn. Res. 2012, 6 (9), 1478–1482. 32. von Eiff, C.; Jansen, B.; Kohnen, W.; Becker, K. Infections associated with medical devices: pathogenesis, management and prophylaxis. Drugs 2005, 65 (2), 179–214. 33. Nuryastuti, T.; Umaroh, N.; Asdie, R. H.; Sari, I. P.; Musthafa, A. Pan-drug-resistant and biofilmproducing strain of Burkholderia pseudomallei: first report of melioidosis from a diabetic patient in Yogyakarta, Indonesia. Int. Med. Case. Rep. J. 2018, 11, 319–323. 34. De, A.; Raj, H. J.; Maiti, P. K. Biofilm in osteomyelitis caused by a rare pathogen, Morganella morganii: a case report. J. Clin. Diagn. Res. 2016, 10 (6), DD06–DD08. 35. Johanson, W. G., Jr.; Pierce, A. K.; Sanford, J. P.; Thomas, G. D. Nosocomial respiratory infections with gram-negative bacilli. The significance of colonization of the respiratory tract. Ann. Intern. Med. 1972, 77 (5), 701–706. 36. Southwood, R. T.; Rice, J. L.; McDonald, P. J.; Hakendorf, P. H.; Rozenbilds, M. A. Infection in experimental hip arthroplasties. J. Bone Joint Surg. Br. 1985, 67 (2), 229–231. 37. Percival, S. L.; Suleman, L.; Vuotto, C.; Donelli, G. Healthcare-associated infections, medical devices and biofilms: risk, tolerance and control. J. Med. Microbiol. 2015, 64 (Pt. 4), 323–334. 38. Schierholz, J. M.; Beuth, J. Implant infections: a haven for opportunistic bacteria. J. Hosp. Infect. 2001, 49 (2), 87–93. 39. Arciola, C. R.; Campoccia, D.; Montanaro, L. Implant infections: adhesion, biofilm formation and immune evasion. Nat. Rev. Microbiol. 2018, 16 (7), 397–409. 40. Feldman, M. D.; Petersen, A. J.; Karliner, L. S.; Tice, J. A. Who is responsible for evaluating the safety and effectiveness of medical devices? The role of independent technology assessment. J. Gen. Intern. Med. 2008, 23 (Suppl. 1), 57–63. 41. Danin, P. E.; Girou, E.; Legrand, P.; Louis, B.; Fodil, R.; Christov, C.; Devaquet, J.; Isabey, D.; Brochard, L. Description and microbiology of endotracheal tube biofilm in mechanically ventilated subjects. Respir. Care 2015, 60 (1), 21–29. 42. Trautner, B. W.; Darouiche, R. O. Role of biofilm in catheter-associated urinary tract infection. Am. J. Infect. Control 2004, 32 (3), 177–183.


43. Gominet, M.; Compain, F.; Beloin, C.; Lebeaux, D. Central venous catheters and biofilms: where do we stand in 2017? APMIS 2017, 125 (4), 365–375. 44. Martins, M.; Rodrigues, A.; Pedrosa, J. M.; Carvalho, M. J.; Cabrita, A.; Oliveira, R. Update on the challenging role of biofilms in peritoneal dialysis. Biofouling 2013, 29 (8), 1015–1027. 45. Leonhard, M.; Schneider-Stickler, B. Voice prostheses, microbial colonization and biofilm formation. Adv. Exp. Med. Biol. 2015, 830, 123–136. 46. Gbejuade, H. O.; Lovering, A. M.; Webb, J. C. The role of microbial biofilms in prosthetic joint infections. Acta. Orthop. 2015, 86 (2), 147–158. 47. Zoubos, A. B.; Galanakos, S. P.; Soucacos, P. N. Orthopedics and biofilm--what do we know? A review. Med. Sci. Monit. 2012, 18 (6), RA89–RA96. 48. Okuda, K. I.; Nagahori, R.; Yamada, S.; Sugimoto, S.; Sato, C.; Sato, M.; Iwase, T.; Hashimoto, K.; Mizunoe, Y. The composition and structure of biofilms developed by propionibacterium acnes isolated from cardiac pacemaker devices. Front. Microbiol. 2018, 9, 182. 49. Benachinmardi, K. K.; Ravikumar, R.; Indiradevi, B. Role of biofilm in cerebrospinal fluid shunt infections: a study at tertiary neurocare center from South India. J. Neurosci. Rural Pract. 2017, 8 (3), 335–341. 50. Herten, M.; Bisdas, T.; Knaack, D.; Becker, K.; Osada, N.; Torsello, G. B.; Idelevich, E. A. Rapid in vitro quantification of S. aureus biofilms on vascular graft surfaces. Front. Microbiol. 2017, 8, 2333. 51. El-Ganiny, A. M.; Shaker, G. H.; Aboelazm, A. A.; El-Dash, H. A. Prevention of bacterial biofilm formation on soft contact lenses using natural compounds. J. Ophthalmic. Inflamm. Infect. 2017, 7 (1), 11. 52. Vaishnavi, C.; Samanta, J.; Kochhar, R. Characterization of biofilms in biliary stents and potential factors involved in occlusion. World J. Gastroenterol. 2018, 24 (1), 112–123. 53. Faller, M.; Kohler, T. The status of biofilms in penile implants. Microorganisms 2017, 5 (2), 19–25. 54. Øilo, M.; Bakken, V. Biofilm and dental biomaterials. Materials 2015, 8 (6), 2887–2900. 55. Bachour, Y. Response to “Bacterial Adhesion and Biofilm Formation on Textured Breast Implant Shell Materials”. Aesthetic Plast. Surg. 2018. 56. James, G. A.; Boegli, L.; Hancock, J.; Bowersock, L.; Parker, A.; Kinney, B. M. Bacterial adhesion and biofilm formation on textured breast implant shell materials. Aesthetic Plast. Surg. 2019, 43 (2), 490–497. 57. Donlan, R. M.; Costerton, J. W. Biofilms: survival mechanisms of clinically relevant microorganisms. Clin. Microbiol. Rev. 2002, 15 (2), 167–193. 58. James, G. A.; Ge Zhao, A.; Usui, M.; Underwood, R. A.; Nguyen, H.; Beyenal, H.; deLancey Pulcini, E.; Agostinho Hunt, A.; Bernstein, H. C.; Fleckman, P.; Olerud, J.; Williamson, K. S.; Franklin, M. J.; Stewart, P. S. Microsensor and transcriptomic signatures of oxygen depletion in biofilms associated with chronic wounds. Wound Repair Regen. 2016, 24 (2), 373–383. 59. James, G. A.; Swogger, E.; Wolcott, R.; Pulcini, E.; Secor, P.; Sestrich, J.; Costerton, J. W.; Stewart, P. S. Biofilms in chronic wounds. Wound Repair Regen. 2008, 16 (1), 37–44. 60. Anaissie, E.; Samonis, G.; Kontoyiannis, D.; Costerton, J.; Sabharwal, U.; Bodey, G.; Raad, I. Role of catheter colonization and infrequent hematogenous seeding in catheter-related infections. Eur. J. Clin. Microbiol. Infect. Dis. 1995, 14 (2), 134–137. 61. Bauer, T. T.; Torres, A.; Ferrer, R.; Heyer, C. M.; Schultze-Werninghaus, G.; Rasche, K. Biofilm formation in endotracheal tubes. Association between pneumonia and the persistence of pathogens. Monaldi. Arch. Chest Dis. 2002, 57 (1), 84–87. 62. Craven, D. E.; Hjalmarson, K. I. Ventilator-associated tracheobronchitis and pneumonia: thinking outside the box. Clin. Infect. Dis. 2010, 51 (Suppl. 1), S59–S66. 63. Palmer, L. B. Ventilator-associated infection: the role for inhaled antibiotics. Curr. Opin. Pulm. Med. 2015, 21 (3), 239–249.


64. Bonell, A.; Azarrafiy, R.; Huong, V. T. L.; Viet, T. L.; Phu, V. D.; Dat, V. Q.; Wertheim, H.; van Doorn, H. R.; Lewycka, S.; Nadjm, B. A systematic review and meta-analysis of ventilator-associated pneumonia in adults in Asia: an analysis of national income level on incidence and etiology. Clin. Infect. Dis. 2019, 68 (3), 511–518. 65. Huang, Y.; Jiao, Y.; Zhang, J.; Xu, J.; Cheng, Q.; Li, Y.; Liang, S.; Li, H.; Gong, J.; Zhu, Y.; Song, L.; Rong, Z.; Liu, B.; Jie, Z.; Sun, S.; Li, P.; Wang, G.; Qu, J. Infection assembly of Shanghai Respiratory Society, Microbial etiology and prognostic factors of ventilator-associated pneumonia: a multicenter retrospective study in Shanghai. Clin. Infect. Dis. 2018, 67 (Suppl. 2), S146–S152. 66. Maki, D. G.; Tambyah, P. A. Engineering out the risk for infection with urinary catheters. Emerg. Infect. Dis. 2001, 7 (2), 342–347. 67. Stickler, D. J. Bacterial biofilms in patients with indwelling urinary catheters. Nat. Clin. Pract. Urol. 2008, 5 (11), 598–608. 68. Flores-Mireles, A. L.; Walker, J. N.; Caparon, M.; Hultgren, S. J. Urinary tract infections: epidemiology, mechanisms of infection and treatment options. Nat. Rev. Microbiol. 2015, 13 (5), 269–284. 69. Emori, T. G.; Gaynes, R. P. An overview of nosocomial infections, including the role of the microbiology laboratory. Clin. Microbiol. Rev. 1993, 6 (4), 428–442. 70. Mermel, L. A.; Allon, M.; Bouza, E.; Craven, D. E.; Flynn, P.; O’Grady, N. P.; Raad, I. I.; Rijnders, B. J.; Sherertz, R. J.; Warren, D. K. Clinical practice guidelines for the diagnosis and management of intravascular catheter-related infection: 2009 update by the Infectious Diseases Society of America. Clin. Infect. Dis. 2009, 49 (1), 1–45. 71. Percival, S. L.; Kite, P. Intravascular catheters and biofilm control. J. Vasc. Access 2007, 8 (2), 69–80. 72. Donlan, R. M. Biofilms on central venous catheters: is eradication possible? Curr. Top. Microbiol. Immunol. 2008, 322, 133–161. 73. Bouza, E.; Eworo, A.; Fernandez Cruz, A.; Reigadas, E.; Rodriguez-Creixems, M.; Munoz, P. Catheterrelated bloodstream infections caused by Gram-negative bacteria. J. Hosp. Infect. 2013, 85 (4), 316–320. 74. Hanna, H.; Afif, C.; Alakech, B.; Boktour, M.; Tarrand, J.; Hachem, R.; Raad, I. Central venous catheter-related bacteremia due to gram-negative bacilli: significance of catheter removal in preventing relapse. Infect. Control Hosp. Epidemiol. 2004, 25 (8), 646–649. 75. Okuda, K. I.; Yoshii, Y.; Yamada, S.; Chiba, A.; Hironaka, I.; Hori, S.; Yanaga, K.; Mizunoe, Y. Detection of bacterial DNA from central venous catheter removed from patients by next generation sequencing: a preliminary clinical study. Ann. Clin. Microbiol. Antimicrob. 2018, 17 (1), 44. 76. Wu, H.; Moser, C.; Wang, H. Z.; Hoiby, N.; Song, Z. J. Strategies for combating bacterial biofilm infections. Int. J. Oral Sci. 2015, 7 (1), 1–7. 77. Thornton, R. B.; Rigby, P. J.; Wiertsema, S. P.; Filion, P.; Langlands, J.; Coates, H. L.; Vijayasekaran, S.; Keil, A. D.; Richmond, P. C. Multi-species bacterial biofilm and intracellular infection in otitis media. BMC Pediatr. 2011, 11, 94. 78. Marsh, P. D. Microbiology of dental plaque biofilms and their role in oral health and caries. Dent. Clin. North Am. 2010, 54 (3), 441–454. 79. Takenaka, S.; Ohsumi, T.; Noiri, Y. Evidence-based strategy for dental biofilms: current evidence of mouthwashes on dental biofilm and gingivitis. Jpn. Dent. Sci. Rev. 2019, 55 (1), 33–40. 80. Do, T.; Devine, D.; Marsh, P. D. Oral biofilms: molecular analysis, challenges, and future prospects in dental diagnostics. Clin. Cosmet. Investig. Dent. 2013, 5, 11–19. 81. Larsen, T.; Fiehn, N. E. Dental biofilm infections - an update. APMIS 2017, 125 (4), 376–384. 82. Benitez-Paez, A.; Belda-Ferre, P.; Simon-Soro, A.; Mira, A. Microbiota diversity and gene expression dynamics in human oral biofilms. BMC Genomics 2014, 15, 311. 83. Marsh, P. D.; Head, D. A.; Devine, D. A. Ecological approaches to oral biofilms: control without killing. Caries Res. 2015, 49 (Suppl. 1), 46–54.


84. Kirketerp-Moller, K.; Jensen, P. O.; Fazli, M.; Madsen, K. G.; Pedersen, J.; Moser, C.; Tolker-Nielsen, T.; Hoiby, N.; Givskov, M.; Bjarnsholt, T. Distribution, organization, and ecology of bacteria in chronic wounds. J. Clin. Microbiol. 2008, 46 (8), 2717–2722. 85. Gurjala, A. N.; Geringer, M. R.; Seth, A. K.; Hong, S. J.; Smeltzer, M. S.; Galiano, R. D.; Leung, K. P.; Mustoe, T. A. Development of a novel, highly quantitative in vivo model for the study of biofilmimpaired cutaneous wound healing. Wound Repair Regen. 2011, 19 (3), 400–410. 86. Westgate, S. J.; Percival, S. L.; Knottenbelt, D. C.; Clegg, P. D.; Cochrane, C. A. Chronic equine wounds: what is the role of infection and biofilms? Wounds 2010, 22 (6), 138–145. 87. Metcalf, D. G.; Bowler, P. G. Biofilm delays wound healing: a review of the evidence. Burns Trauma 2013, 1 (1), 5–12. 88. Anderson, G. G.; Palermo, J. J.; Schilling, J. D.; Roth, R.; Heuser, J.; Hultgren, S. J. Intracellular bacterial biofilm-like pods in urinary tract infections. Science 2003, 301 (5629), 105–107. 89. Mulvey, M. A.; Lopez-Boado, Y. S.; Wilson, C. L.; Roth, R.; Parks, W. C.; Heuser, J.; Hultgren, S. J. Induction and evasion of host defenses by type 1-piliated uropathogenic Escherichia coli. Science 1998, 282 (5393), 1494–1497. 90. Rosen, D. A.; Pinkner, J. S.; Jones, J. M.; Walker, J. N.; Clegg, S.; Hultgren, S. J. Utilization of an intracellular bacterial community pathway in Klebsiella pneumoniae urinary tract infection and the effects of FimK on type 1 pilus expression. Infect. Immun. 2008, 76 (7), 3337–3345. 91. Otter, J. A.; Vickery, K.; Walker, J. T.; deLancey Pulcini, E.; Stoodley, P.; Goldenberg, S. D.; Salkeld, J. A.; Chewins, J.; Yezli, S.; Edgeworth, J. D. Surface-attached cells, biofilms and biocide susceptibility: implications for hospital cleaning and disinfection. J. Hosp. Infect. 2015, 89 (1), 16–27. 92. Ledwoch, K.; Dancer, S. J.; Otter, J. A.; Kerr, K.; Roposte, D.; Rushton, L.; Weiser, R.; Mahenthiralingam, E.; Muir, D. D.; Maillard, J. Y. Beware biofilm! Dry biofilms containing bacterial pathogens on multiple healthcare surfaces; a multi-centre study. J. Hosp. Infect. 2018, 100 (3), e47–e56. 93. Chowdhury, D.; Tahir, S.; Legge, M.; Hu, H.; Prvan, T.; Johani, K.; Whiteley, G. S.; Glasbey, T. O.; Deva, A. K.; Vickery, K. Transfer of dry surface biofilm in the healthcare environment: the role of healthcare workers’ hands as vehicles. J. Hosp. Infect. 2018, 100 (3), e85–e90. 94. Exner, M.; Kramer, A.; Lajoie, L.; Gebel, J.; Engelhart, S.; Hartemann, P. Prevention and control of health care-associated waterborne infections in health care facilities. Am. J. Infect. Control 2005, 33 (5) (Suppl. 1), S26–S40. 95. Ji, X. Y.; Fei, C. N.; Zhang, Y.; Zhang, W.; Liu, J.; Dong, J. Evaluation of bacterial contamination of dental unit waterlines and use of a newly designed measurement device to assess retraction of a dental chair unit. Int. Dent. J. 2016, 66 (4), 208–214. 96. Anaissie, E. J.; Penzak, S. R.; Dignani, M. C. The hospital water supply as a source of nosocomial infections: a plea for action. Arch. Intern. Med. 2002, 162 (13), 1483–1492. 97. Jefferson, K. K. What drives bacteria to produce a biofilm? FEMS Microbiol. Lett. 2004, 236 (2), 163–173. 98. Khoury, A. E.; Lam, K.; Ellis, B.; Costerton, J. W. Prevention and control of bacterial infections associated with medical devices. ASAIO J. 1992, 38 (3), M174–M178. 99. Singh, R.; Ray, P.; Das, A.; Sharma, M. Penetration of antibiotics through Staphylococcus aureus and Staphylococcus epidermidis biofilms. J. Antimicrob. Chemother. 2010, 65 (9), 1955–1958. 100. Hall-Stoodley, L.; Stoodley, P. Biofilm formation and dispersal and the transmission of human pathogens. Trends Microbiol. 2005, 13 (1), 7–10. 101. Cho, K. H.; Caparon, M. G. Patterns of virulence gene expression differ between biofilm and tissue communities of Streptococcus pyogenes. Mol. Microbiol. 2005, 57 (6), 1545–1556. 102. Rani, S. A.; Pitts, B.; Beyenal, H.; Veluchamy, R. A.; Lewandowski, Z.; Davison, W. M.; BuckinghamMeyer, K.; Stewart, P. S. Spatial patterns of DNA replication, protein synthesis, and oxygen


103. 104. 105. 106. 107. 108.

109. 110.

111. 112. 113.

114. 115. 116.

117.

118.

119.

120.

concentration within bacterial biofilms reveal diverse physiological states. J. Bacteriol. 2007, 189 (11), 4223–4233. Neu, T. R. Significance of bacterial surface-active compounds in interaction of bacteria with interfaces. Microbiol. Rev. 1996, 60 (1), 151–166. Hausner, M.; Wuertz, S. High rates of conjugation in bacterial biofilms as determined by quantitative in situ analysis. Appl. Environ. Microbiol. 1999, 65 (8), 3710–3713. Dodds, M. G.; Grobe, K. J.; Stewart, P. S. Modeling biofilm antimicrobial resistance. Biotechnol. Bioeng. 2000, 68 (4), 456–465. Lewis, K. Multidrug tolerance of biofilms and persister cells. Curr. Top. Microbiol. Immunol. 2008, 322, 107–131. Lewis, K. Persister cells. Annu. Rev. Microbiol. 2010, 64, 357–372. Percival, S. L.; Hill, K. E.; Malic, S.; Thomas, D. W.; Williams, D. W. Antimicrobial tolerance and the significance of persister cells in recalcitrant chronic wound biofilms. Wound Repair Regen. 2011, 19 (1), 1–9. Brauner, A.; Fridman, O.; Gefen, O.; Balaban, N. Q. Distinguishing between resistance, tolerance and persistence to antibiotic treatment. Nat. Rev. Microbiol. 2016, 14 (5), 320–330. Anderson, G. G.; Moreau-Marquis, S.; Stanton, B. A.; O’Toole, G. A. In vitro analysis of tobramycintreated Pseudomonas aeruginosa biofilms on cystic fibrosis-derived airway epithelial cells. Infect. Immun. 2008, 76 (4), 1423–1433. Hoiby, N.; Ciofu, O.; Bjarnsholt, T. Pseudomonas aeruginosa biofilms in cystic fibrosis. Future Microbiol. 2010, 5 (11), 1663–1674. Lee, S.; Bae, S. Molecular viability testing of viable but non-culturable bacteria induced by antibiotic exposure. Microb. Biotechnol. 2018, 11 (6), 1008–1016. Trombetta, R. P.; Dunman, P. M.; Schwarz, E. M.; Kates, S. L.; Awad, H. A. A high-throughput screening approach to repurpose FDA-approved drugs for bactericidal applications against Staphylococcus aureus small-colony variants. mSphere 2018, 3 (5), e00422-18. Wolcott, R. D.; Dowd, S. E. A rapid molecular method for characterising bacterial bioburden in chronic wounds. J. Wound Care 2008, 17 (12), 513–516. Mandakhalikar, K. D.; Rahmat, J. N.; Chiong, E.; Neoh, K. G.; Shen, L.; Tambyah, P. A. Extraction and quantification of biofilm bacteria: method optimized for urinary catheters. Sci. Rep. 2018, 8 (1), 8069. Herrmann, G.; Yang, L.; Wu, H.; Song, Z.; Wang, H.; Hoiby, N.; Ulrich, M.; Molin, S.; Riethmuller, J.; Doring, G. Colistin-tobramycin combinations are superior to monotherapy concerning the killing of biofilm Pseudomonas aeruginosa. J. Infect. Dis. 2010, 202 (10), 1585–1592. Wolcott, R. D.; Rumbaugh, K. P.; James, G.; Schultz, G.; Phillips, P.; Yang, Q.; Watters, C.; Stewart, P. S.; Dowd, S. E. Biofilm maturity studies indicate sharp debridement opens a time-dependent therapeutic window. J. Wound Care 2010, 19 (8), 320–328. Hengzhuang, W.; Wu, H.; Ciofu, O.; Song, Z.; Hoiby, N. Pharmacokinetics/pharmacodynamics of colistin and imipenem on mucoid and nonmucoid Pseudomonas aeruginosa biofilms. Antimicrob. Agents Chemother. 2011, 55 (9), 4469–4474. Hentzer, M.; Wu, H.; Andersen, J. B.; Riedel, K.; Rasmussen, T. B.; Bagge, N.; Kumar, N.; Schembri, M. A.; Song, Z.; Kristoffersen, P.; Manefield, M.; Costerton, J. W.; Molin, S.; Eberl, L.; Steinberg, P.; Kjelleberg, S.; Hoiby, N.; Givskov, M. Attenuation of Pseudomonas aeruginosa virulence by quorum sensing inhibitors. EMBO J. 2003, 22 (15), 3803–3815. Romero, D.; Sanabria-Valentin, E.; Vlamakis, H.; Kolter, R. Biofilm inhibitors that target amyloid proteins. Chem. Biol. 2013, 20 (1), 102–110.


121. Cegelski, L.; Pinkner, J. S.; Hammer, N. D.; Cusumano, C. K.; Hung, C. S.; Chorell, E.; Aberg, V.; Walker, J. N.; Seed, P. C.; Almqvist, F.; Chapman, M. R.; Hultgren, S. J. Small-molecule inhibitors target Escherichia coli amyloid biogenesis and biofilm formation. Nat. Chem. Biol. 2009, 5 (12), 913–919. 122. Sintim, H. O.; Smith, J. A.; Wang, J.; Nakayama, S.; Yan, L. Paradigm shift in discovering nextgeneration anti-infective agents: targeting quorum sensing, c-di-GMP signaling and biofilm formation in bacteria with small molecules. Future Med Chem 2010, 2 (6), 1005–1035. 123. Schaber, J. A.; Triffo, W. J.; Suh, S. J.; Oliver, J. W.; Hastert, M. C.; Griswold, J. A.; Auer, M.; Hamood, A. N.; Rumbaugh, K. P. Pseudomonas aeruginosa forms biofilms in acute infection independent of cell-tocell signaling. Infect. Immun. 2007, 75 (8), 3715–3721. 124. Davis, S. C.; Ricotti, C.; Cazzaniga, A.; Welsh, E.; Eaglstein, W. H.; Mertz, P. M. Microscopic and physiologic evidence for biofilm-associated wound colonization in vivo. Wound Repair Regen. 2008, 16 (1), 23–29. 125. Timmel, J.; Kent, P. S.; Holzmueller, C. G.; Paine, L.; Schulick, R. D.; Pronovost, P. J. Impact of the Comprehensive Unit-based Safety Program (CUSP) on safety culture in a surgical inpatient unit. Jt. Comm. J. Qual. Patient Saf. 2010, 36 (6), 252–260. 126. Thornhill, M. H.; Dayer, M.; Lockhart, P. B.; McGurk, M.; Shanson, D.; Prendergast, B.; Chambers, J. B. A change in the NICE guidelines on antibiotic prophylaxis. Br. Dent. J. 2016, 221 (3), 112–114. 127. Hoiby, N.; Bjarnsholt, T.; Moser, C.; Bassi, G. L.; Coenye, T.; Donelli, G.; Hall-Stoodley, L.; Hola, V.; Imbert, C.; Kirketerp-Moller, K.; Lebeaux, D.; Oliver, A.; Ullmann, A. J.; Williams, C. ESCMID Study Group for Biofilms and Consulting External Expert Werner Zimmerli, ESCMID guideline for the diagnosis and treatment of biofilm infections 2014. Clin. Microbiol. Infect. 2015, 21 (Suppl. 1), S1–S25. 128. Scudeller, L.; Rodriguez-Bano, J.; Zinkernagel, A.; Tacconelli, E.; Akova, M.; Friedrich, A. W.; Sanguinetti, M.; Paul, M.; Poljak, M. ESCMID white paper: a guide on ESCMID guidance documents. Clin. Microbiol. Infect. 2019, 25 (2), 155–162. 129. Mandakhalikar, K. D.; Chua, R. R.; Tambyah, P. A. New technologies for prevention of catheter associated urinary tract infection. Current Treatment Options in Infectious Diseases 2016, 8 (1), 24–41. 130. Mandakhalikar, K. D.; Wang, R.; Rahmat, J. N.; Chiong, E.; Neoh, K. G.; Tambyah, P. A. Restriction of in vivo infection by antifouling coating on urinary catheter with controllable and sustained silver release: a proof of concept study. BMC Infect. Dis. 2018, 18 (1), 370. 131. Choudhary, P.; Das, S. K. Bio-reduced graphene oxide as a nanoscale antimicrobial coating for medical devices. ACS Omega 2019, 4 (1), 387–397. 132. Montero, D. A.; Arellano, C.; Pardo, M.; Vera, R.; Galvez, R.; Cifuentes, M.; Berasain, M. A.; Gomez, M.; Ramirez, C.; Vidal, R. M. Antimicrobial properties of a novel copper-based composite coating with potential for use in healthcare facilities. Antimicrob. Resist. Infect. Control 2019, 8, 3.


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