Review
Emerging rules for effective antimicrobial coatings Mario Salwiczek1,2, Yue Qu2,3, James Gardiner1, Richard A. Strugnell4, Trevor Lithgow2, Keith M. McLean1, and Helmut Thissen1 1
CSIRO Materials Science and Engineering, Bayview Avenue, Clayton, Victoria 3168, Australia Department of Microbiology, Monash University, Melbourne, Victoria 3800, Australia 3 Department of Biochemistry and Molecular Biology, Monash University, Melbourne, Victoria 3800, Australia 4 Department of Microbiology and Immunology, University of Melbourne, Parkville, Victoria 3010, Australia 2
In order to colonize abiotic surfaces, bacteria and fungi undergo a profound change in their biology to form biofilms: communities of microbes embedded into a matrix of secreted macromolecules. Despite strict hygiene standards, biofilm-related infections associated with implantable devices remain a common complication in the clinic. Here, the application of highly dosed antibiotics is problematic in that the biofilm (i) provides a protective environment for microbes to evade antibiotics and/or (ii) can provide selective pressure for the evolution of antibiotic-resistant microbes. However, recent research suggests that effective prevention of biofilm formation may be achieved by multifunctional surface coatings that provide both non-adhesive and antimicrobial properties imparted by antimicrobial peptides. Such coatings are the subject of this review. Biofilms and implant infections An aging population and advances in materials technology have brought about, over the past 50 years, an increase in the usage of biomaterials and medical devices such as catheters, cardiac pacemakers, hip implants, and contact lenses, which can restore function to diseased or damaged human tissue. However, the application of such devices involves some challenges – in particular, implant-associated infections resulting from the presence of biofilms. Biofilm formation typically results from peri-operative procedures (where organisms enter the wound or adhere to the implant during surgery) and post-operative procedures (where organisms infect the patient during hospitalization) [1,2]. Infectious diseases are responsible for tens of millions of deaths representing approximately 20% of all fatalities world-wide [3], and it is estimated that 80% of human infections are associated with biofilm formation.
Corresponding author: Salwiczek, M. ([email protected]). Keywords: anti-infective coatings; antimicrobial peptides; biofilms; implant infections; low-fouling polymers. 0167-7799/$ – see front matter . Crown Copyright ß 2013 Published by Elsevier Ltd. All rights reserved. http:// dx.doi.org/10.1016/j.tibtech.2013.09.008
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Furthermore, many of the causative organisms exhibit growing antimicrobial resistance [4]. A range of organisms has been specifically implicated in device/biomaterial-related infections, including many species of bacteria and major fungal pathogens affecting human health (Table 1) [1,2,5–18]. Device-related infection may result in substantial clinical complications, including death, as well as economic consequences such as increased healthcare costs generated by prolonged hospital stays or revision surgery. In the United States, recent estimates of direct costs for healthcare-associated infections were estimated to range from US$28 billion to $45 billion in 1 year with upward of 60% of these being related to medical devices [19]. Several medical interventions are currently used to treat device-related infections, including long-term antimicrobial strategies and combinations of antibiotics and surgical revision. Unfortunately, these interventions carry the risk of re-infection, often at a higher rate, and the development of antibiotic resistance. The application of non-adhesive and antimicrobial coatings has been researched and tested clinically as an alternative approach but has yet to find widespread application. Here, we review coating strategies combining low-fouling polymer coatings with antimicrobial peptides and the continued development required for the prevention of microbial biofilms on medical devices. Low-fouling coatings Effective control of biointerfacial interactions is the key to developing improved biomedical materials and devices, including infection-resistant medical implants. Many of these applications require surfaces that prevent non-specific interactions with the biological environment, in particular the adsorption of proteins and other biomolecules. Such ‘low-fouling’ coatings also reduce the ability of planktonic microbes to adhere, thereby interfering with the earliest stages of biofilm formation. Various chemical approaches that are suitable for establishing such coatings have been described. These include self-assembled monolayers (SAMs), various polymer-based approaches [20–22], and, very recently, liquid-infused nanostructured surfaces that present a dynamic surface structure [23]. Although SAMs are easily applied, their
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Box 1. Biofilm formation The stages of biofilm formation (Figure I) by Staphylococcus epidermidis, a well-known biofilm-producer, are used here as a model. Stage 1: attachment and monolayer formation. Free-floating cells attach within seconds after encountering an abiotic surface [69]. Nonspecific interactions between bacteria and the surface are mediated by physiochemical forces, such as van der Waals forces, hydrophobic interactions, and polar and ionic interactions [70,71]. In addition, specific interactions can be mediated by a preformed ‘conditioning film’ of biomolecules derived from plasma components adsorbed to the device surface [72,73]. This conditioning film is thought to provide specific binding sites for bacterial surface proteins (adhesins), and the interactions often result in tight attachment of bacteria [74,75]. By the end of this stage, a confluent layer of S. epidermidis cells, referred to as an ‘adherent monolayer’, covers the device surface. Stage 2: formation of microcolonies. Within the monolayer, bacteria multiply locally and then assemble to form a mound-shaped cellular aggregate — a ‘microcolony’. In the case of S. epidermidis, microcolony formation depends on secreted and surface-adsorbed bio-macromolecules, including polysaccharide intercellular adhesin (PIA), surface proteins, teichoic acid, and extracellular DNA [76]. Stage 3: maturation and structuring. After the formation of microcolonies, S. epidermidis cells undergo further adaptation and development into a mature biofilm consisting of bacterial macrocolonies,
Stage 1
Stage 2
which eventually converge, being encased by an extracellular polymeric substance (EPS) that is highly penetrated by channels [77]. PIA is still the key extracellular component of macrocolonies. Its expression during the maturation stage is regulated by quorum-sensing systems and other global gene regulators such as SarA, RsbU, and SigB [78–82]. Bacterial surfactant peptides and shear force (e.g., by flowing body fluids) also play a vital part in the biofilm shaping and maturation. All of these factors determine the density of the biofilm matrix, the overall cell density and the strength of surface attachment [83–85]. Stage 4: detachment and return to the planktonic growth model. During this last stage, bacteria return to the planktonic mode, causing a risk of spreading the infection. Low-level sloughing as well as active dispersion of bacteria generated from the biofilm occur synchronously [86]. Once a biofilm structure is formed, cells on the surface exit from the biofilm and re-enter the planktonic state in response to certain environmental cues and self-stress signals [77,87]. During attachment and the initial formation of colonies, microbes are relatively drug-sensitive and susceptible to immune cell response. Bacteria in biofilms are reportedly 100 to 1000 times more resistant than their planktonic counterparts. Drug resistance is provided through adaptive changes, genetic changes, production of a subpopulation of dormant cells, and physical protection from antibiotics by the matrix.
Stage 3
Stage 4
Rate of establishment dictated by properes of surface
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Figure I. The four stages of biofilm formation.
versatility with respect to substrates and stability is limited. By contrast, polymer coatings can be applied to almost any substrate material and provide a much broader range of architectures and properties. Among polymer based coating approaches, two techniques stand out in their ability to yield low-fouling surfaces: the ‘grafting-to’ technique (Figure 1A), in which polymers carrying suitable functional groups are synthesized in solution and then tethered to surfaces by reacting with complementary functional groups on the surface, and the ‘grafting-from’ technique (Figure 1B), in which polymer chains are grown from surface-immobilized initiators or chain transfer agents. Multiple factors, including the density and molecular weight of graft polymer chains, have been shown to determine the effectiveness of the coating [24]. An important feature of these graft polymers is that functional groups or biologically active signals can be introduced along the graft polymer chain or at its terminal end to further modulate the biological response [25]. Particularly effective low-fouling polymers include polyacrylamide (PAM) [26], polysaccharides such as dextrane [27], zwitterionic polymers such as poly(N-sulfobetaine methacrylamide) (PSBMA) [28,29], and poly(N-hydroxypropyl methacrylamide) (PHPMA) [30] (Figure 1C). However, polyethylene glycol (PEG)-based polymers and their
low-fouling properties have received most of the attention to date. PEG polymers have been described with linear [31] and star-shaped architectures [32] as well as ‘bottle brush’type polymers with pendant PEG chains such as those based on poly(ethylene glycol) methacrylate (PEGMA) [33]. Impeding biomolecule adsorption disrupts a broad range of processes that require the interaction of proteins or other biomolecules with substrate materials, including cell attachment, platelet adhesion, and blood clot formation, as well as the foreign-body reaction and microbial colonization/biofilm formation [34,35]. However, even ultra-low fouling surfaces might eventually form substrates for the formation of biofilms – for example, due to degradation or inhomogeneity that might also be the result of damage during handling. Therefore, robust antimicrobial coatings would require more than one mechanism of defense. A wide range of molecules that inhibit or disperse biofilms has been identified [36,37]. Among those that can be embedded or immobilized on surfaces, or combined with polymer coatings, silver [38], ammonium, and guanidinium salts, as well as peptides and proteins [39], have attracted much attention. In this context, cationic antimicrobial peptides (AMPs) have shown particularly promising results. 83
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Table 1. Incidence and causative agents of infections associated with commonly used medical devices and implants [1,2,9–18] Implants or devices
Major causative organisms a
Urinary catheter Central venous catheter Peritoneal catheter Mechanical heart valve Ventricular assist device Coronary stents Cardiac pacemakers Vascular grafts
Escherichia coli, Candida spp., CoNS, Enterococcus faecalis, Proteus mirabilis CoNS, Staphylococcus aureus, Enterococcus spp., Candida spp., Klebsiella pneumoniae S. aureus, Pseudomonas aeruginosa, Candida spp. CoNS, S. aureus, Streptococcus spp., Enterococcus spp. CoNS, S. aureus, Candida spp., P. aeruginosa S. aureus, CoNS, P. aeruginosa, Candida spp. S. aureus, CoNS, Streptococcus spp., Candida spp. S. aureus, Staphylococcus epidermidis, Pseudomonas spp., Enterococcus spp., Enterobacter spp. P. aeruginosa, Serratia marcescens, S. aureus S. epidermidis CoNS; S. aureus, Propionibacterium spp., Streptococcus spp., Corynebacterium spp. S. aureus, CoNS, Streptococcus spp., Enterobacteriaceae Streptococcus spp., Actinomyces spp., Porphyromonas spp., Prevotella spp. CoNS; Staphylococcus spp.; Enterobacter spp., Pseudomonas spp. CoNS; Streptococcus spp., P. mirabilis, Enterococcus spp., Actinomyces spp. S. aureus, P. aeruginosa, Haemophilus influenzae, Streptococcus spp. S. aureus, S. epidermidis, CoNS, Peptostreptococcus spp., Bacteroides fragilis, E. coli, Enterococcus spp., P. aeruginosa, Serratia spp. S. aureus, CoNS, Streptococcus, Propionibacterium spp.
Contact lenses Intraocular lenses Fracture fixation devices Hip/knee implants Dental implants Penile implants Pelvic organ prolapse mesh Cochlear implants Sutures Breast implants
Infection incidence over life of device 33% 2–10% 3–5% 1–4% 13–80% 50% reduction of bacterial attachment to the background coating, whereas combined coatings with Tet-26 and Tet-20 showed 90% and 80% reduction in biofilm formation, respectively. A 7-day in vivo implant infection model using S. aureus showed an 85% reduction of surface-attached bacteria compared to a control sample presenting the low-fouling polymer only. Structure–activity relationship (SAR) studies revealed that low-graft-density brushes allow for higher peptide surface density. However, these coatings were also more adhesive compared to higher-graft-density brushes that present lower peptide densities [61]. Kim et al. used an engineered arginine and tryptophane rich peptide immobilized via a H2N-PEG-maleimide tethered to terminal epoxide groups of allylglycidyl ether (AGE) polymer brushes on polydimethylsiloxane (PDMS) slides [62]. The peptide-modified surfaces were shown to inhibit growth of three bacterial strains (E. coli, S. aureus, and P. aeruginosa), and the bactericidal properties against E. coli were retained even after up to 3 days of immersion in water. Two complementary biofilm assays (crystal violet staining after 24 h and live/dead staining after 5 days) indicated that their background coating significantly reduces biofilm formation. Additional tethering of the peptides completely prevented bacterial attachment. Moreover, the surfaces were judged to be ‘non-toxic’ as measured by haemolytic activity and muscle cell viability. One advantage of the bottle brush architecture is that the amount of loaded peptide is more easily controlled by the ratio of monomers used for co-polymerization. Furthermore, the peptides do not shield the low-fouling background as an icing on top of it. Importantly, however, the study on switchable polymers provides indirect evidence that the castle architecture, although showing appreciable low-fouling properties, might not provide for bactericidal activity.
Multilayer architecture The LBL assembly of polyelectrolytes on surfaces has been studied for various applications [63] owing to its simplicity and low cost. LBL assembly is typically carried out by incubation of the substrate in a solution of one polyelectrolyte followed by a washing step and subsequent incubation with an oppositely charge polyelectrolyte. Such multilayered films have been demonstrated to effectively mask the underlying surface chemistry and properties. The properties of the resulting coating can be tailored by the assembly conditions, which include the identity and charge of the substrate, the nature and molecular weight of the polyelectrolytes as well as the pH and ionic strength of the coating buffer. Importantly, multilayer architecture allows the coating of complex geometries and is extremely versatile [64]. Native polyelectrolytes, such as charged polysaccharides and AMPs, are also suitable for creating biocompatible multilayer coatings. Wang et al. described the application of PEG diacrylate-co-polyacrylic acid (PEGDA/PAA) to generate negatively charged microgels for deposition on silicon wafers that were previously primed with poly(L-lysine) [65] and subsequently loaded with bovine cathelicidin-derived peptide L5. Non-loaded microgels were shown to prevent Staphylococcus epidermidis adhesion for at least 6 h, and anti-adhesiveness was shown to correlate with surface density. After 10 h of exposure, however, anti-adhesiveness was lost. Owing to additional ionic interactions between the positively charged side chains of the peptide and the carboxyl groups, greater peptide loadings could be achieved with gels containing acrylic acid. Microgels that did not incorporate PAA were quickly depleted of peptide and bacterial colonization levels rose significantly after 6 h. High-density L5-loaded PEG-co-PAA microgels almost completely prevented colonization for at least 10 h. The authors concluded that the slow release of L5 provides for high local concentrations of the peptide, keeping systemic concentrations low while ensuring extended activity. Vreuls et al. designed ‘easy-cleaning’ antibacterial coatings based on native polyelectrolyte multilayers on stainless steel [66]. Synthetic polyelectrolyte co-polymers of 87
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(A)
(B)
(C)
(D)
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Figure 3. Possible coating architectures to combine low-fouling with antibacterial properties: (A) icing; (B) bottle brush; (C) multilayer; and (D) castle.
N-methacrolyl-3,4-dihydroxyphenylalanine and 2-methacryloxyethyl trimethlyammonium chloride (DMAEMA+) or polyethylene imine (PEI) served as the basal anchoring layer, onto which different multilayers were built. The authors evaluated the natural anti-adhesion molecules mucin (porcine and bovine), a glycoprotein containing large amounts of sialic acids, or heparin and PAA as the negatively charged polyelectrolytes. Different AMPs (nisin, tritrpticin, and lipopeptide 4K-C16) as well as polyallylamine hydrochloride (PAH) were used as positively charged polyelectrolytes to create low-fouling or antimicrobial as well as combined coatings. The 3,4-dihydroxyphenylalanine-based polymer expectedly outperformed PEI as an anchoring layer, and porcine stomach mucin showed the most favorable anti-adhesion properties. Of the antimicrobial coatings, those combining mixed layers of nisin and 4K-C16 (1:10) on a base of PAA showed optimal activity against both E. coli and Bacillus subtilis (>5 log reduction of both), whereas the individual peptides, combinations with PAH, and coatings presenting either nisin or 4K-C16 as the outer layer showed unfavorable specificities or generally low activities. Different combined coatings were created using either mucin or PAA combined with either nisin or PAH. Antibacterial activity and the amount of adsorbed protein were assessed after three bacterial challenges with intermittent rinsing steps. The results indicate that the antibacterial activity does not depend on the anionic polyelectrolyte used and that it is irrelevant whether the peptide (nisin) or the anti-adhesive polymer (PAA/ mucin) make up the outermost layer. However, the coating incorporating mucin again showed the best low-fouling properties. The authors also demonstrated that PAA–nisin coatings were stable towards immersion and wiping in water. Wiping with 1% household detergent, however, had a deleterious effect on activity. The LBL architecture is technically straight forward and does not require chemical modification of AMPs to achieve highly active coatings. It is highly flexible regarding the structures that can be used. However, by design, the coatings are not stable and are therefore most suitable 88
for short term applications in which release of the antimicrobial agents into the body might be desirable. Concluding remarks and future perspectives Low-fouling and bactericidal coatings have both shown promising results in the reduction of biofilm formation and related adverse events. However, more effective and durable solutions based on coatings that provide multiple complementary lines of defense are needed. These include low-fouling surface coatings that display antimicrobial compounds such as AMPs. There is no doubt that AMPs exhibit many advantageous biological properties, and although there are currently too few examples of combined low-fouling antimicrobial coatings based on AMPs to clearly indicate which chemistries and coating architectures are preferable, the results to date suggest that AMPs are excellent candidates for surface applications. It has been observed on several occasions that AMPs have an antagonizing effect on the low-fouling properties of covalently immobilized coatings. The available data suggest that it is necessary to maintain high-polymer densities with low peptide loadings, and thus low surface-charge density, to keep the balance between sufficient bactericidal activity and maintaining low-fouling properties. In cases in which peptides are released over time and coatings are not covalently immobilized, ‘self-cleaning’ may occur by layer detachment. But as the layers essentially dissipate over time, such coatings would be most appropriate for short-term applications. At this stage, comparing the different approaches remains difficult because the biological readouts vary among the studies, which use different time points, methods, and organisms, and because most studies focus on the early stages of biofilm establishment. A standardized testing procedure including assessment of the later stages of biofilm formation should provide the necessary insights to enable the direct comparison of different approaches and, ultimately, to refine the coating architectures reviewed here.
Review An important clinical challenge that also has to be addressed in line with developing implant coatings is tissue integration. Short-term applications such as wound dressings or urinary catheters might not require a permanent coating, and the release of antimicrobials into the surrounding tissue may even be desirable. Here, multilayer coatings may be most suitable, especially owing to their low production costs. Some implants, for example, heart valves or joint replacements, need to reside within the body for a lifetime. Here, permanent coatings such as immobilized bottle brush and icing coatings are more suitable because they do not dissipate over time, at least not by release. In order to function correctly, they need to be fully integrated into the surrounding tissue. However, low-fouling coatings unselectively reduce cellular adhesion. The answer to who is winning this ‘race for the surface’ [67] lies within multifunctional surface coatings that not only include low-fouling and antimicrobial components but, in addition, give tissue cells an incentive – that is, selective peptide adhesion signals – to colonize the surface first. The recent public interest in implant regulatory affairs [68] underlines the need for appropriate in vivo testing to guarantee the adequate performance and safety of medical devices. The key issues to be addressed here are the stability of the coating, toxicity of eventually released components, and tissue integration (where this is desirable). In this regard, using known AMPs as the antimicrobial component is sensible because the risks involving their use are a known variable. Despite the remaining challenges, we anticipate substantial progress in the prevention of bacterial colonization and biofilm formation with clinically relevant and biocompatible surfaces. Acknowledgments The authors acknowledge the Australian Research Council (ARC) for funding: Y.Q. is an ARC Super Science Fellow and T.L. is an ARC Federation Fellow.
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63 Gribova, V. et al. (2012) Polyelectrolyte multilayer assemblies on materials surfaces: from cell adhesion to tissue engineering. Chem. Mater. 24, 854–869 64 Caruso, F. (2001) Nanoengineering of particle surfaces. Adv. Mater. 13, 11–22 65 Wang, Q. et al. (2012) Self-assembled poly(ethylene glycol)-co-acrylic acid microgels to inhibit bacterial colonization of synthetic surfaces. ACS Appl. Mater. Interfaces 4, 2498–2506 66 Vreuls, C. et al. (2010) Biomolecules in multilayer film for antimicrobial and easy-cleaning stainless steel surface applications. Biofouling 26, 645–656 67 Gristina, A. (1987) Biomaterial-centered infection: microbial adhesion versus tissue integration. Science 237, 1588–1595 68 Storz-Pfennig, P. et al. (2013) Trials are needed before new devices are used in routine practice in Europe. Br. Med. J. 346, f1646 69 Peters, G. et al. (1982) Adherence and growth of coagulase-negative staphylococci on surfaces of intravenous catheters. J. Infect. Dis. 146, 479–482 70 Donlan, R.M. (2001) Biofilm formation: a clinically relevant microbiological process. Clin. Infect. Dis. 33, 1387–1392 71 von Eiff, C. et al. (2002) Pathogenesis of infections due to coagulasenegative staphylococci. Lancet Infect. Dis. 2, 677–685 72 Peters, G. (1988) Pathogenesis of S. epidermidis foreign body infections. Br. J. Clin. Pract. Suppl. 57, 62–65 73 Gottenbos, B. et al. (2002) Pathogenesis and prevention of biomaterial centered infections. J. Mater. Sci. Mater. Med. 13, 717–722 74 Herrmann, M. et al. (1988) Fibronectin, fibrinogen, and laminin act as mediators of adherence of clinical staphylococcal isolates to foreign material. J. Infect. Dis. 158, 693–701 75 Espersen, F. et al. (1990) Attachment of staphylococci to silicone catheters in vitro. APMIS 98, 471–478 76 Otto, M. (2009) Staphylococcus epidermidis – the ‘accidental’ pathogen. Nat. Rev. Microbiol. 7, 555–567 77 Sauer, K. et al. (2002) Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J. Bacteriol. 184, 1140–1154 ´ . et al. (2005) SarA is an assential positive regulator of 78 Tormo, M.A Staphylococcus epidermidis biofilm development. J. Bacteriol. 187, 2348–2356 79 Xu, L. et al. (2006) Role of the luxS quorum-sensing system in biofilm formation and virulence of Staphylococcus epidermidis. Infect. Immun. 74, 488–496 80 Tao, J-H. et al. (2006) Depression of biofilm formation and antibiotic resistance by sarA disruption in Staphylococcus epidermidis. World J. Gastroenterol. 12, 4009–4013 81 Goller, C.C. and Romeo, T. (2008) Environmental influences on biofilm development. In Bacterial Biofilms (Romeo, T., ed.), pp. 37–66, Springer 82 Pintens, V. et al. (2008) The role of sB in persistence of Staphylococcus epidermidis foreign body infection. Microbiology 154, 2827–2836 83 Rupp, C.J. et al. (2005) Viscoelasticity of Staphylococcus aureus biofilms in response to fluid shear allows resistance to detachment and facilitates rolling migration. Appl. Environ. Microbiol. 71, 2175–2178 84 Otto, M. (2008) Staphylococcal biofilms. In Bacterial Biofilms (Romeo, T., ed.), pp. 207–228, Springer 85 Lee, J-H. et al. (2008) Microfluidic devices for studying growth and detachment of Staphylococcus epidermidis biofilms. Biomed. Microdevices 10, 489–498 86 Stoodley, P. et al. (2005) Molecular and imaging techniques for bacterial biofilms in joint arthroplasty infections. Clin. Orthop. Relat. Res. 437, 31–40 87 Bester, E. et al. (2005) Planktonic-cell yield of a pseudomonad biofilm. Appl. Environ. Microbiol. 71, 7792–7798
Emerging rules for effective antimicrobial coatings Mario Salwiczek1,2, Yue Qu2,3, James Gardiner1, Richard A. Strugnell4, Trevor Lithgow2, Keith M. McLean1, and Helmut Thissen1 1
CSIRO Materials Science and Engineering, Bayview Avenue, Clayton, Victoria 3168, Australia Department of Microbiology, Monash University, Melbourne, Victoria 3800, Australia 3 Department of Biochemistry and Molecular Biology, Monash University, Melbourne, Victoria 3800, Australia 4 Department of Microbiology and Immunology, University of Melbourne, Parkville, Victoria 3010, Australia 2
In order to colonize abiotic surfaces, bacteria and fungi undergo a profound change in their biology to form biofilms: communities of microbes embedded into a matrix of secreted macromolecules. Despite strict hygiene standards, biofilm-related infections associated with implantable devices remain a common complication in the clinic. Here, the application of highly dosed antibiotics is problematic in that the biofilm (i) provides a protective environment for microbes to evade antibiotics and/or (ii) can provide selective pressure for the evolution of antibiotic-resistant microbes. However, recent research suggests that effective prevention of biofilm formation may be achieved by multifunctional surface coatings that provide both non-adhesive and antimicrobial properties imparted by antimicrobial peptides. Such coatings are the subject of this review. Biofilms and implant infections An aging population and advances in materials technology have brought about, over the past 50 years, an increase in the usage of biomaterials and medical devices such as catheters, cardiac pacemakers, hip implants, and contact lenses, which can restore function to diseased or damaged human tissue. However, the application of such devices involves some challenges – in particular, implant-associated infections resulting from the presence of biofilms. Biofilm formation typically results from peri-operative procedures (where organisms enter the wound or adhere to the implant during surgery) and post-operative procedures (where organisms infect the patient during hospitalization) [1,2]. Infectious diseases are responsible for tens of millions of deaths representing approximately 20% of all fatalities world-wide [3], and it is estimated that 80% of human infections are associated with biofilm formation.
Corresponding author: Salwiczek, M. ([email protected]). Keywords: anti-infective coatings; antimicrobial peptides; biofilms; implant infections; low-fouling polymers. 0167-7799/$ – see front matter . Crown Copyright ß 2013 Published by Elsevier Ltd. All rights reserved. http:// dx.doi.org/10.1016/j.tibtech.2013.09.008
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Furthermore, many of the causative organisms exhibit growing antimicrobial resistance [4]. A range of organisms has been specifically implicated in device/biomaterial-related infections, including many species of bacteria and major fungal pathogens affecting human health (Table 1) [1,2,5–18]. Device-related infection may result in substantial clinical complications, including death, as well as economic consequences such as increased healthcare costs generated by prolonged hospital stays or revision surgery. In the United States, recent estimates of direct costs for healthcare-associated infections were estimated to range from US$28 billion to $45 billion in 1 year with upward of 60% of these being related to medical devices [19]. Several medical interventions are currently used to treat device-related infections, including long-term antimicrobial strategies and combinations of antibiotics and surgical revision. Unfortunately, these interventions carry the risk of re-infection, often at a higher rate, and the development of antibiotic resistance. The application of non-adhesive and antimicrobial coatings has been researched and tested clinically as an alternative approach but has yet to find widespread application. Here, we review coating strategies combining low-fouling polymer coatings with antimicrobial peptides and the continued development required for the prevention of microbial biofilms on medical devices. Low-fouling coatings Effective control of biointerfacial interactions is the key to developing improved biomedical materials and devices, including infection-resistant medical implants. Many of these applications require surfaces that prevent non-specific interactions with the biological environment, in particular the adsorption of proteins and other biomolecules. Such ‘low-fouling’ coatings also reduce the ability of planktonic microbes to adhere, thereby interfering with the earliest stages of biofilm formation. Various chemical approaches that are suitable for establishing such coatings have been described. These include self-assembled monolayers (SAMs), various polymer-based approaches [20–22], and, very recently, liquid-infused nanostructured surfaces that present a dynamic surface structure [23]. Although SAMs are easily applied, their
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Box 1. Biofilm formation The stages of biofilm formation (Figure I) by Staphylococcus epidermidis, a well-known biofilm-producer, are used here as a model. Stage 1: attachment and monolayer formation. Free-floating cells attach within seconds after encountering an abiotic surface [69]. Nonspecific interactions between bacteria and the surface are mediated by physiochemical forces, such as van der Waals forces, hydrophobic interactions, and polar and ionic interactions [70,71]. In addition, specific interactions can be mediated by a preformed ‘conditioning film’ of biomolecules derived from plasma components adsorbed to the device surface [72,73]. This conditioning film is thought to provide specific binding sites for bacterial surface proteins (adhesins), and the interactions often result in tight attachment of bacteria [74,75]. By the end of this stage, a confluent layer of S. epidermidis cells, referred to as an ‘adherent monolayer’, covers the device surface. Stage 2: formation of microcolonies. Within the monolayer, bacteria multiply locally and then assemble to form a mound-shaped cellular aggregate — a ‘microcolony’. In the case of S. epidermidis, microcolony formation depends on secreted and surface-adsorbed bio-macromolecules, including polysaccharide intercellular adhesin (PIA), surface proteins, teichoic acid, and extracellular DNA [76]. Stage 3: maturation and structuring. After the formation of microcolonies, S. epidermidis cells undergo further adaptation and development into a mature biofilm consisting of bacterial macrocolonies,
Stage 1
Stage 2
which eventually converge, being encased by an extracellular polymeric substance (EPS) that is highly penetrated by channels [77]. PIA is still the key extracellular component of macrocolonies. Its expression during the maturation stage is regulated by quorum-sensing systems and other global gene regulators such as SarA, RsbU, and SigB [78–82]. Bacterial surfactant peptides and shear force (e.g., by flowing body fluids) also play a vital part in the biofilm shaping and maturation. All of these factors determine the density of the biofilm matrix, the overall cell density and the strength of surface attachment [83–85]. Stage 4: detachment and return to the planktonic growth model. During this last stage, bacteria return to the planktonic mode, causing a risk of spreading the infection. Low-level sloughing as well as active dispersion of bacteria generated from the biofilm occur synchronously [86]. Once a biofilm structure is formed, cells on the surface exit from the biofilm and re-enter the planktonic state in response to certain environmental cues and self-stress signals [77,87]. During attachment and the initial formation of colonies, microbes are relatively drug-sensitive and susceptible to immune cell response. Bacteria in biofilms are reportedly 100 to 1000 times more resistant than their planktonic counterparts. Drug resistance is provided through adaptive changes, genetic changes, production of a subpopulation of dormant cells, and physical protection from antibiotics by the matrix.
Stage 3
Stage 4
Rate of establishment dictated by properes of surface
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Figure I. The four stages of biofilm formation.
versatility with respect to substrates and stability is limited. By contrast, polymer coatings can be applied to almost any substrate material and provide a much broader range of architectures and properties. Among polymer based coating approaches, two techniques stand out in their ability to yield low-fouling surfaces: the ‘grafting-to’ technique (Figure 1A), in which polymers carrying suitable functional groups are synthesized in solution and then tethered to surfaces by reacting with complementary functional groups on the surface, and the ‘grafting-from’ technique (Figure 1B), in which polymer chains are grown from surface-immobilized initiators or chain transfer agents. Multiple factors, including the density and molecular weight of graft polymer chains, have been shown to determine the effectiveness of the coating [24]. An important feature of these graft polymers is that functional groups or biologically active signals can be introduced along the graft polymer chain or at its terminal end to further modulate the biological response [25]. Particularly effective low-fouling polymers include polyacrylamide (PAM) [26], polysaccharides such as dextrane [27], zwitterionic polymers such as poly(N-sulfobetaine methacrylamide) (PSBMA) [28,29], and poly(N-hydroxypropyl methacrylamide) (PHPMA) [30] (Figure 1C). However, polyethylene glycol (PEG)-based polymers and their
low-fouling properties have received most of the attention to date. PEG polymers have been described with linear [31] and star-shaped architectures [32] as well as ‘bottle brush’type polymers with pendant PEG chains such as those based on poly(ethylene glycol) methacrylate (PEGMA) [33]. Impeding biomolecule adsorption disrupts a broad range of processes that require the interaction of proteins or other biomolecules with substrate materials, including cell attachment, platelet adhesion, and blood clot formation, as well as the foreign-body reaction and microbial colonization/biofilm formation [34,35]. However, even ultra-low fouling surfaces might eventually form substrates for the formation of biofilms – for example, due to degradation or inhomogeneity that might also be the result of damage during handling. Therefore, robust antimicrobial coatings would require more than one mechanism of defense. A wide range of molecules that inhibit or disperse biofilms has been identified [36,37]. Among those that can be embedded or immobilized on surfaces, or combined with polymer coatings, silver [38], ammonium, and guanidinium salts, as well as peptides and proteins [39], have attracted much attention. In this context, cationic antimicrobial peptides (AMPs) have shown particularly promising results. 83
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Table 1. Incidence and causative agents of infections associated with commonly used medical devices and implants [1,2,9–18] Implants or devices
Major causative organisms a
Urinary catheter Central venous catheter Peritoneal catheter Mechanical heart valve Ventricular assist device Coronary stents Cardiac pacemakers Vascular grafts
Escherichia coli, Candida spp., CoNS, Enterococcus faecalis, Proteus mirabilis CoNS, Staphylococcus aureus, Enterococcus spp., Candida spp., Klebsiella pneumoniae S. aureus, Pseudomonas aeruginosa, Candida spp. CoNS, S. aureus, Streptococcus spp., Enterococcus spp. CoNS, S. aureus, Candida spp., P. aeruginosa S. aureus, CoNS, P. aeruginosa, Candida spp. S. aureus, CoNS, Streptococcus spp., Candida spp. S. aureus, Staphylococcus epidermidis, Pseudomonas spp., Enterococcus spp., Enterobacter spp. P. aeruginosa, Serratia marcescens, S. aureus S. epidermidis CoNS; S. aureus, Propionibacterium spp., Streptococcus spp., Corynebacterium spp. S. aureus, CoNS, Streptococcus spp., Enterobacteriaceae Streptococcus spp., Actinomyces spp., Porphyromonas spp., Prevotella spp. CoNS; Staphylococcus spp.; Enterobacter spp., Pseudomonas spp. CoNS; Streptococcus spp., P. mirabilis, Enterococcus spp., Actinomyces spp. S. aureus, P. aeruginosa, Haemophilus influenzae, Streptococcus spp. S. aureus, S. epidermidis, CoNS, Peptostreptococcus spp., Bacteroides fragilis, E. coli, Enterococcus spp., P. aeruginosa, Serratia spp. S. aureus, CoNS, Streptococcus, Propionibacterium spp.
Contact lenses Intraocular lenses Fracture fixation devices Hip/knee implants Dental implants Penile implants Pelvic organ prolapse mesh Cochlear implants Sutures Breast implants
Infection incidence over life of device 33% 2–10% 3–5% 1–4% 13–80% 50% reduction of bacterial attachment to the background coating, whereas combined coatings with Tet-26 and Tet-20 showed 90% and 80% reduction in biofilm formation, respectively. A 7-day in vivo implant infection model using S. aureus showed an 85% reduction of surface-attached bacteria compared to a control sample presenting the low-fouling polymer only. Structure–activity relationship (SAR) studies revealed that low-graft-density brushes allow for higher peptide surface density. However, these coatings were also more adhesive compared to higher-graft-density brushes that present lower peptide densities [61]. Kim et al. used an engineered arginine and tryptophane rich peptide immobilized via a H2N-PEG-maleimide tethered to terminal epoxide groups of allylglycidyl ether (AGE) polymer brushes on polydimethylsiloxane (PDMS) slides [62]. The peptide-modified surfaces were shown to inhibit growth of three bacterial strains (E. coli, S. aureus, and P. aeruginosa), and the bactericidal properties against E. coli were retained even after up to 3 days of immersion in water. Two complementary biofilm assays (crystal violet staining after 24 h and live/dead staining after 5 days) indicated that their background coating significantly reduces biofilm formation. Additional tethering of the peptides completely prevented bacterial attachment. Moreover, the surfaces were judged to be ‘non-toxic’ as measured by haemolytic activity and muscle cell viability. One advantage of the bottle brush architecture is that the amount of loaded peptide is more easily controlled by the ratio of monomers used for co-polymerization. Furthermore, the peptides do not shield the low-fouling background as an icing on top of it. Importantly, however, the study on switchable polymers provides indirect evidence that the castle architecture, although showing appreciable low-fouling properties, might not provide for bactericidal activity.
Multilayer architecture The LBL assembly of polyelectrolytes on surfaces has been studied for various applications [63] owing to its simplicity and low cost. LBL assembly is typically carried out by incubation of the substrate in a solution of one polyelectrolyte followed by a washing step and subsequent incubation with an oppositely charge polyelectrolyte. Such multilayered films have been demonstrated to effectively mask the underlying surface chemistry and properties. The properties of the resulting coating can be tailored by the assembly conditions, which include the identity and charge of the substrate, the nature and molecular weight of the polyelectrolytes as well as the pH and ionic strength of the coating buffer. Importantly, multilayer architecture allows the coating of complex geometries and is extremely versatile [64]. Native polyelectrolytes, such as charged polysaccharides and AMPs, are also suitable for creating biocompatible multilayer coatings. Wang et al. described the application of PEG diacrylate-co-polyacrylic acid (PEGDA/PAA) to generate negatively charged microgels for deposition on silicon wafers that were previously primed with poly(L-lysine) [65] and subsequently loaded with bovine cathelicidin-derived peptide L5. Non-loaded microgels were shown to prevent Staphylococcus epidermidis adhesion for at least 6 h, and anti-adhesiveness was shown to correlate with surface density. After 10 h of exposure, however, anti-adhesiveness was lost. Owing to additional ionic interactions between the positively charged side chains of the peptide and the carboxyl groups, greater peptide loadings could be achieved with gels containing acrylic acid. Microgels that did not incorporate PAA were quickly depleted of peptide and bacterial colonization levels rose significantly after 6 h. High-density L5-loaded PEG-co-PAA microgels almost completely prevented colonization for at least 10 h. The authors concluded that the slow release of L5 provides for high local concentrations of the peptide, keeping systemic concentrations low while ensuring extended activity. Vreuls et al. designed ‘easy-cleaning’ antibacterial coatings based on native polyelectrolyte multilayers on stainless steel [66]. Synthetic polyelectrolyte co-polymers of 87
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(A)
(B)
(C)
(D)
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Figure 3. Possible coating architectures to combine low-fouling with antibacterial properties: (A) icing; (B) bottle brush; (C) multilayer; and (D) castle.
N-methacrolyl-3,4-dihydroxyphenylalanine and 2-methacryloxyethyl trimethlyammonium chloride (DMAEMA+) or polyethylene imine (PEI) served as the basal anchoring layer, onto which different multilayers were built. The authors evaluated the natural anti-adhesion molecules mucin (porcine and bovine), a glycoprotein containing large amounts of sialic acids, or heparin and PAA as the negatively charged polyelectrolytes. Different AMPs (nisin, tritrpticin, and lipopeptide 4K-C16) as well as polyallylamine hydrochloride (PAH) were used as positively charged polyelectrolytes to create low-fouling or antimicrobial as well as combined coatings. The 3,4-dihydroxyphenylalanine-based polymer expectedly outperformed PEI as an anchoring layer, and porcine stomach mucin showed the most favorable anti-adhesion properties. Of the antimicrobial coatings, those combining mixed layers of nisin and 4K-C16 (1:10) on a base of PAA showed optimal activity against both E. coli and Bacillus subtilis (>5 log reduction of both), whereas the individual peptides, combinations with PAH, and coatings presenting either nisin or 4K-C16 as the outer layer showed unfavorable specificities or generally low activities. Different combined coatings were created using either mucin or PAA combined with either nisin or PAH. Antibacterial activity and the amount of adsorbed protein were assessed after three bacterial challenges with intermittent rinsing steps. The results indicate that the antibacterial activity does not depend on the anionic polyelectrolyte used and that it is irrelevant whether the peptide (nisin) or the anti-adhesive polymer (PAA/ mucin) make up the outermost layer. However, the coating incorporating mucin again showed the best low-fouling properties. The authors also demonstrated that PAA–nisin coatings were stable towards immersion and wiping in water. Wiping with 1% household detergent, however, had a deleterious effect on activity. The LBL architecture is technically straight forward and does not require chemical modification of AMPs to achieve highly active coatings. It is highly flexible regarding the structures that can be used. However, by design, the coatings are not stable and are therefore most suitable 88
for short term applications in which release of the antimicrobial agents into the body might be desirable. Concluding remarks and future perspectives Low-fouling and bactericidal coatings have both shown promising results in the reduction of biofilm formation and related adverse events. However, more effective and durable solutions based on coatings that provide multiple complementary lines of defense are needed. These include low-fouling surface coatings that display antimicrobial compounds such as AMPs. There is no doubt that AMPs exhibit many advantageous biological properties, and although there are currently too few examples of combined low-fouling antimicrobial coatings based on AMPs to clearly indicate which chemistries and coating architectures are preferable, the results to date suggest that AMPs are excellent candidates for surface applications. It has been observed on several occasions that AMPs have an antagonizing effect on the low-fouling properties of covalently immobilized coatings. The available data suggest that it is necessary to maintain high-polymer densities with low peptide loadings, and thus low surface-charge density, to keep the balance between sufficient bactericidal activity and maintaining low-fouling properties. In cases in which peptides are released over time and coatings are not covalently immobilized, ‘self-cleaning’ may occur by layer detachment. But as the layers essentially dissipate over time, such coatings would be most appropriate for short-term applications. At this stage, comparing the different approaches remains difficult because the biological readouts vary among the studies, which use different time points, methods, and organisms, and because most studies focus on the early stages of biofilm establishment. A standardized testing procedure including assessment of the later stages of biofilm formation should provide the necessary insights to enable the direct comparison of different approaches and, ultimately, to refine the coating architectures reviewed here.
Review An important clinical challenge that also has to be addressed in line with developing implant coatings is tissue integration. Short-term applications such as wound dressings or urinary catheters might not require a permanent coating, and the release of antimicrobials into the surrounding tissue may even be desirable. Here, multilayer coatings may be most suitable, especially owing to their low production costs. Some implants, for example, heart valves or joint replacements, need to reside within the body for a lifetime. Here, permanent coatings such as immobilized bottle brush and icing coatings are more suitable because they do not dissipate over time, at least not by release. In order to function correctly, they need to be fully integrated into the surrounding tissue. However, low-fouling coatings unselectively reduce cellular adhesion. The answer to who is winning this ‘race for the surface’ [67] lies within multifunctional surface coatings that not only include low-fouling and antimicrobial components but, in addition, give tissue cells an incentive – that is, selective peptide adhesion signals – to colonize the surface first. The recent public interest in implant regulatory affairs [68] underlines the need for appropriate in vivo testing to guarantee the adequate performance and safety of medical devices. The key issues to be addressed here are the stability of the coating, toxicity of eventually released components, and tissue integration (where this is desirable). In this regard, using known AMPs as the antimicrobial component is sensible because the risks involving their use are a known variable. Despite the remaining challenges, we anticipate substantial progress in the prevention of bacterial colonization and biofilm formation with clinically relevant and biocompatible surfaces. Acknowledgments The authors acknowledge the Australian Research Council (ARC) for funding: Y.Q. is an ARC Super Science Fellow and T.L. is an ARC Federation Fellow.
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