Substrate independent silver nanoparticle based antibacterial coatings

Biomaterials 35 (2014) 4601e4609

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Biomaterials journal homepage: www.elsevier.com/locate/biomaterials

Substrate independent silver nanoparticle based antibacterial coatings Shima Taheri a, Alex Cavallaro a, Susan N. Christo c, Louise E. Smith d, Peter Majewski a, Mary Barton b, John D. Hayball b, c, Krasimir Vasilev a, d, * a

School of Engineering, University of South Australia, Mawson Lakes, SA 5095, Australia School of Pharmacy and Medical Sciences, University of South Australia, SA 5000, Australia c Sansom Institute, University of South Australia, Adelaide, SA 5000, Australia d Mawson Institute, University of South Australia, Mawson Lakes, SA 5095, Australia b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 20 December 2013 Accepted 20 February 2014 Available online 12 March 2014

Infections arising from bacterial adhesion and colonization on medical device surfaces are a significant healthcare problem. Silver based antibacterial coatings have attracted a great deal of attention as a potential solution. This paper reports on the development of a silver nanoparticles based antibacterial surface that can be applied to any type of material surface. The silver nanoparticles were surface engineered with a monolayer of 2-mercaptosuccinic acid, which facilitates the immobilization of the nanoparticles to the solid surface, and also reduces the rate of oxidation of the nanoparticles, extending the lifetime of the coatings. The coatings had excellent antibacterial efficacy against three clinically significant pathogenic bacteria i.e. Staphylococcus epidermidis, Staphylococcus aureus and Pseudomonas aeruginosa. Studies with primary human fibroblast cells showed that the coatings had no cytotoxicity in vitro. Innate immune studies in cultures of primary macrophages demonstrated that the coatings do not significantly alter the level of expression of pro-inflammatory cytokines or the adhesion and viability of these cells. Collectively, these coatings have an optimal combination of properties that make them attractive for deposition on medical device surfaces such as wound dressings, catheters and implants. Ó 2014 Elsevier Ltd. All rights reserved.

Keywords: Antibacterial surfaces Silver nanoparticles Infections Innate inflammatory response Cytotoxicity Medical devices

1. Introduction Today, medical devices are an important part of medical practice, improving patient wellbeing and saving millions of lives each year. Despite all these benefits, infections associated with medical devices constitute a significant healthcare burden [1,2]. These types of complications account for more than half of all hospital acquired infections (HAI) and are the most complex and costly to treat [3,4]. Hospital acquired infections annually affect more than two million patients in the USA and cause more than 100,000 deaths [5,6]. Infection rates in the case of biomedical implants can be as high as 4% for some devices and can cost in excess of $50,000 per patient to rectify the problem if revision of the implanted site is required [7e 10]. Catheters are another type of medical device that are often the cause of infections. For example, central venous catheters cause an estimated 80,000 catheter-related bloodstream infections in the

* Corresponding author. University of South Australia, Mawson Institute and School of Engineering, Mawson Lakes, SA 5095, Australia. Tel.: þ61 8 83025697; fax: þ61 8 83025689. E-mail address: [email protected] (K. Vasilev). http://dx.doi.org/10.1016/j.biomaterials.2014.02.033 0142-9612/Ó 2014 Elsevier Ltd. All rights reserved.

USA, resulting in 28,000 deaths per year [11,12]. Likewise, urinary catheters are estimated to be associated with 80% of all urinary tract infections [13,14]. Medical device-associated infections are mainly caused by bacterial attachment to, and colonization of the device surface [15e19]. For this reason, it is well accepted that preventing bacterial adhesion to the device surface through the application of an antibacterial coating is a potential solution [20,21]. Amongst the various strategies for generating antibacterial surfaces are those that are based on silver compounds and silver nanoparticles [14,22e24]. Silver (Ag) is precious metal that has been known to humans for more than seven millennia and has been appreciated not only because of its beauty but also because of its antibacterial properties [25e29]. The historical use of silver and silver formulations was mainly based on empirical observations that liquids did not spoil and wounds healed better. Over time, the medical use of silver became more sophisticated and by the end of the 19th century a number commercial products were available [28]. Silver was almost replaced by other antibacterial compounds after the penicillin revolution of the 1930s. However, the development of antibiotic resistance led to the resurgence of silver in medicine in the 1970s [28,29]. Currently, there is

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substantial research in silver coated medical devices [21,23,24,30e33]. This is mainly driven by the fact that silver is active against both Gram-positive and Gram-negative bacteria and resistance has not been yet been convincingly demonstrated for clinically-relevant pathogens. For more on the extensive efforts on development of silver based antibacterial coatings, we refer the reader to several instructive recent reviews on the topic [21,23,24,31e33]. When an antibacterial coating is being developed, several factors need to be taken into consideration in order to ensure efficacy against bacteria and successful wound healing. Among them are the capacity to kill bacteria, the need to eliminate the coating’s cytotoxicity toward eukaryotic cells and the ability to avoid any undesired inflammatory response that may occur [34]. However, most reports are confined to presenting efficacy against one or more bacterial strains but fall short of reporting biocompatibility with mammalian tissues and cells and even more rarely deal with possible adverse innate immune inflammatory consequences. In the case of silver, it is generally accepted that mammalian cells are capable of tolerating higher concentrations compared to bacteria. However, it is evident that the concentration of silver which is demonstrably cytotoxic varies greatly for different mammalian cells [27]. In addition, the studies were not always conducted with the same silver compound and when nanoparticles were investigated, they had various sizes and surface functionalities. Nanocrystalline silver has been reported to reduce inflammation [35e 37]. However, silver nitrate has been reported to increase inflammatory responses, and silver nanoparticles may cause the necrosis of immune cells which would also result in exacerbated inflammation [35]. It has also been reported that metallic silver surfaces induced cell death and a pro-inflammatory response in cultured J774 macrophages [38]. These contradictions clearly highlight the need for the full characterization of any new coating that contains silver. Without a doubt, the ultimate tests are in vivo studies. However, these types of studies are expensive and animal welfare and ethical processes require in vitro assessment prior to the use of an in vivo model. Here, silver nanoparticle-based antibacterial surfaces were generated, such that deposition onto any type of substrate material was possible, for the purpose of coating a wide variety of medical devices. To achieve this goal, silver nanoparticles with appropriate surface functionality were synthesized in solution and adsorbed to model substrata pre-modified with a functional plasma polymer film. Plasma polymers were used because they have been demonstrated to grow independently from the substrate material except in the very early stages of deposition [39e 41]. This approach also provides a ready control over the amount of immobilized silver nanoparticles. The efficacy of the coatings against both Gram-positive and Gram-negative bacteria was determined. The cytotoxicity was evaluated using primary human dermal fibroblast cells. Finally, evaluation of the inflammatory response potential was assessed by culture of primary macrophages and analysis of the cell viability and levels of expression of pro-inflammatory cytokines. 2. Experimental 2.1. Materials 2.1.1. Substrate preparation All chemicals were used as received. Allylamine (AA) (98%), silver nitrate (99.99%), sodium borohydride, 2-mercaptosuccinic acid (97%), nitric acid (70%) were supplied by Aldrich (Australia). Hydrochloric acid (36%) by Ajax Finechem Pty. Ltd (Australia), Safranin O, acetic acid glacial and sodium hydroxide pellet were purchased from Chem Supply(Australia). All solution preparation and glassware cleaning procedures were performed using ultrapure (Milli-Q) water (resistivity 18.2 U). All glassware and magnet stirrer were soaked in aqua regia solution (3:1 conc.HCl: conc.HNO3) and then rinsed thoroughly with the Milli-Q water before nanoparticle synthesis.

2.1.2. Microorganisms Staphylococcus epidermidis strain ATCC 35984, Staphylococcus aureus ATCC 4330 (Methicillin-resistant Staphylococcus aureus (MRSA)), Pseudomonas aeruginosa ATCC 27853 were cultured in tryptone soya broth (TSB, Oxoid, UK) in incubator at 37  C and 60% humidity. 2.1.3. Fibroblasts cell cultures Human dermal fibroblasts (HDFs) were harvested and grown as described previously [42] from split thickness skin grafts (STSG’s) obtained from scavenged tissue specimens following routine breast reductions and abdominoplasties. All patients gave informed consent for skin to be used for research through a protocol approved by the Ethical Committee at the Queen Elisabeth Hospital and the University of South Australia Human Ethics Committee. Briefly fibroblasts were grown in fibroblast culture medium (FCM) consisting of Dulbecco’s Modified Eagle Medium (DMEM) high glucose (Gibco, Life Technologies, Australia), 10% v/v fetal calf serum (FCS, Ausgenex, Australia), 0.625 mg/mL amphotericin B (SigmaeAldrich, Australia), 100 IU/mL penicillin and 100 mg/mL streptomycin (Gibco, Life Technologies, Australia) in an incubator at 37  C, 5% CO2 in a humidified atmosphere. The medium was changed every 3e4 days until the cells were 80% confluent. HDFs between passages 3 and 9 were used. Cells were stained with the Phallotoxin “Alexa Fluor 488Ò phalloidin” (a filamentous actin probe from Life) and DAPI- dilactate were purchased from Life Technologies, Invitrogen. 2.1.4. Immune response study Roswell Park Memorial Institute (RPMI; Sigma Aldrich) medium or DMEM (Sigma Aldrich) was supplemented with 10% fetal calf serum, penicillin (100 U/ml), gentamycin (100 mg/ml), b-mercaptoethanol (2 mM), L-glutamine (2 mM), and HEPES (10 mM) to produce complete RPMI (cRPMI) or complete DMEM (cDMEM), respectively. The L-929 conditioned media was prepared by culturing L-929 cells in culture flasks to >95% confluency and cultured in cDMEM until the media was exhausted. The conditioned media containing the required macrophage colony stimulating factor was removed, sterilely filtered and aliquoted for storage at 20  C until use. 2.2. Silver nanoparticle-based antibacterial coating preparation 2.2.1. Synthesis of silver nanoparticles capped with mercaptosuccinic acid (AgNPs@MSA) Mercaptosuccinic acid (MSA) modified silver nanoparticles (AgNPs) were synthesized by mixing 12 mL of 2 mM silver nitrate (AgNO3) with 5 mL of 2 mM MSA under ice-cold condition and vigorous stirring followed by drop-wise addition of 0.5 mL 0.05 M sodium borohydride (NaBH4). The color of the solution changed from colorless to dark red-brownish within a few seconds indicating formation of silver nanoparticles. The functionalized AgNPs colloidal solutions were sealed and stored in darkness. These nanoparticles have been shown to be stable up to twelve months (data not shown). 2.2.2. Deposition of allylamine plasma polymerized (AApp) thin film Plasma polymerization was carried out in a custom-built reactor described previously [43,44] using a 13.56 MHz plasma generator and a matching network (Advanced Energy, USA). The 13 mm Thermanox cover slips substrates (Thermo Fisher Scientific, USA) were cleaned with ethanol and Milli-Q water before placing in the reactor chamber. Samples were air cleaned by exposure to air plasma for 2 min at pressure of 2.5  102 mbar using a power of 20 W. The deposition of allylamine (AA) was carried out at a pressure of 2  103 mbar and an input radio frequency (RF) power of 10 W for 5 min at a monomer flow rate of 10 sccm. These conditions result in nitrogen-rich films with a thickness of about 24 nm. Allylamine plasma polymer (AApp) coated substrates are kept sealed overnight at room temperature any further treatment. The nanoparticles were deposited on silicon wafers for SEM and AFM analysis, and on Thermanox coverslips for bacterial and cell culture studies. 2.2.3. Immobilization of AgNPs@MSA onto the AApp surface AApp coated substrates were immersed in the solution of AgNPs@MSA for defined time intervals ranging from 1 h to 24 h at 23  C temperature. The samples were rinsed thoroughly with Milli-Q water, dried with nitrogen and kept sealed. 2.3. Characterization of nanoparticles 2.3.1. X-ray photoelectron spectroscopy (XPS) XPS spectra were recorded on a SPECS SAGE spectrometer with an Mg Ka radiation source (hn 1253.6 eV) operating at 10 kV and 10 mA. The hemispherical analyzer was a Phoibos 150, with an MCD-9 detector. The elements present were identified from a survey spectrum recorded over the energy range 0e1000 eV at pass energy of 100 eV and a resolution of 0.5 eV. The areas under the photoelectron peaks in the spectrum were used to calculate the percentage atomic concentrations. High resolution (0.1 eV) spectra were then recorded for pertinent photoelectron peaks at pass energy of 20 eV to identify the chemical states of each element. All binding energies (BEs) were referenced to the C 1s neutral carbon peak at 285 eV to compensate for the effect of surface charging. The analysis area was circular and 5 mm in diameter. Processing and component fitting of high resolution spectra were performed with CasaXPS software (Casa Software Ltd).

S. Taheri et al. / Biomaterials 35 (2014) 4601e4609 2.3.2. Particle size distribution and particle charge measurement The particle size distribution (PSD) of MSA coated AgNPs suspended in water (MilliQ, 18.2 MU) was analyzed by dynamic light scattering (DLS) using a Nicomp 380. Additional measurements were carried out using a Malvern Zetasizer Nano ZS. The zeta potential of the nanoparticles was measured with a Zeta Nanosizer (Malvern Instruments, U.K.) at 25  C. 2.3.3. Imaging For SEM imaging a FEI Quanta 450 FEG Environmental SEM was used. Images were recorded using primary beam energy of 10 kV. Secondary Electron emission was collected signal using the Everhart-Thornley Detector. For SEM characterization the MSA functionalized silver nanoparticles were deposited onto 13 mm  13 mm cut silicon wafers pre-coated with allylamine plasma film. Transmission Electron Microscope (TEM) Images were acquired using a Philips CM200 Transmission electron microscope equipped with a tungsten filament. An accelerating voltage of 200 kV was used, resulting in a final point resolution of w0.4 nm. Images were acquired with a Gatan 832 SC1000 Orius CCD camera, and image analysis conduction using the Olympus analysis program. EDS spectrums were acquired with an EDAX Si(Li) detector with an active area of 10 mm2. Samples were prepared by depositing a drop of diluted AgNPs solution on a copper grid and allowing it to dry overnight (23  C). Atomic force microscopy (AFM) provided topographical images using an NTMDT NTEGRA SPM in non-contact mode. Silicon nitride non-contact tips coated with gold on the reflective side (NT-MDT, NSG03) were used and had resonance frequencies between 65 and 100 kHz. The maximum range of the scanner was 100 mm, and the scan rate of 0.5 Hz. The scanner was calibrated in the x, y, and z directions using 1.5 mm grids with a height of 22 nm. 2.3.4. UVevisible spectroscopy and ICP measurement UVevisible spectra were recorded on a Cary 5 UVevis spectrometer (Varian Australia Pty Ltd) from 250 nm to 700 nm. Milli-Q water was used as blank and 1 ml of sample was diluted with Milli-Q water at the ratio of 1:4, before UVevis analysis were performed. Amount of silver present in colloidal was analyzed by inductively coupled plasma (ICP) spectroscopy Perkin Elmer ICP-OES Optima 7300 DV to analyze Ag (mgl1). The reading wavelength for Ag was 328.068 nm and the collected data was processed with MSF (Multicomponent Spectral Fitting).

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2.5.2. Zone of inhibition (KirbyeBauer test) Bacteria suspensions of S. epidermidis and S. aureus and P. aeruginosa in TSB were prepared using the same procedure described above. 150 ml of 1  106 cfu ml1 of bacteria suspension was poured onto the surface of an agar plate and spread uniformly by a sterile swab. Substrates coated with AApp and AgNPs@MSA were applied face down on the agar plate and were incubated overnight at 37  C. The test repeated at least three times for every specimen. 2.6. Immune response 2.6.1. In vitro activation of bone marrow derived macrophages Bone marrow derived macrophage (BMDM) cells were generated by flushing bone marrow cells from the femurs and tibia of 8-week old C57Bl/6 mice in cRPMI supplemented with 20% L-929 conditioned media. On day 4 of culture, nonadherent cells were removed, and media replenished before being seeded onto AgNPs@MSA coated coverslips. On day 7, the in vitro generated BMDMs were stimulated with lipopolysaccharide (LPS) (100 ng/ml; Sigma Aldrich) or left unstimulated for 4 h, and supernatant collected for the analysis of tumor necrosis factoralpha (TNF-a), and interleukin (IL-6). BMDMs were then stimulated with ATP (5 mM; Sigma Aldrich) for 1 h, and supernatant collected for IL-1b analysis using standard ELISA protocols. Transmission images were collected on day 7 using an Olympus IX51 Fluorescence Microscope and the CellSens program. To determine cell viability, adherent BMDMs were gently scraped off the coverslips and stained with the nuclear counterstain DAPI (1 mg/ml) for 10 min and analyzed on the FACSCantoII immediately. Viable cells were gated as side scatter (SSC) and DAPI low. Data was analyzed using the FlowJo (Treestar) software. 2.7. Cell viability and morphology 2.7.1. Cell viability Samples were placed in 24 well tissue culture plates. Human dermal fibroblasts were seeded at a density of 1  104 cells per well in fibroblast cell media. After 2 days cell viability was measured using the resazurin assay. A stock solution of 110 mg/ml resazurin was prepared in phosphate buffered saline and filter sterilized using a 0.2 mm filter. This was then diluted 1:10 in fibroblast culture medium. 500 ml of the resazurin solution was added to each sample. After 4 h 200 ml of the reduced solution was removed from each well and placed into a well of a 96 well plate and the florescent intensity read using a plate reader (lex ¼ 544 nm and lem ¼ 590 nm).

In order to evaluate the amount of silver released from the silver nanoparticle coated surface when exposed to an aqueous media samples were immersed in Milli-Q water for time intervals of 1, 3, 7 and 30 days. The samples were then dried with nitrogen and the amount of silver which remained on the surface was determined by XPS and compared to the control. All studies were conducted in triplicates.

2.7.2. Cell morphology Finally the samples were fixed in 4% paraformaldehyde in PBS for immunocytochemistry staining of the cytoskeleton to assess cell morphology. Fixed cells were incubated at room temperature with 0.1% (v/v) Triton X-100 for 5 min to permeablize the membranes. Phallotoxin Alexa e 488 and DAPI to final concentrations of 0.165 mM and 0.1 mg/ml respectively in 200 mL of PBS was added to each sample and samples incubated further in the dark. Excess dye was washed from the samples and samples were imaged with a Nikon A1-R confocal laser scanning microscope running NIS elements AR software.

2.5. Determination of antibacterial activity in vitro

2.8. Statistical analysis method

In order to evaluate the antibacterial characteristic of a surface or coating, the surfaces were exposed to bacterial culture. The measured reduction in bacterial growth after a set period of time defined the antibacterial nature of the surface. S. aureus ATCC 4330, S. epidermidis ATTC 35984, P. aeruginosa ATCC 27853 were used for this test as the common microorganisms involved in hospital-acquired infections.

All statistical analysis was performed on GraphPad Prism 5 and a one-way ANOVA test was performed on data. The post test performed on immune results was Dunnett’s post test (AAp surfaces were the chosen control in which data was compared against) and for antibacterial and fibroblast cell results a Turkey post test (compare all pairs of columns).

2.4. Silver nanoparticle dissolution

3. Results 2.5.1. Surface immobilized antibacterial activity Bacteria were inoculated onto Blood Agar plates and incubated overnight at 37  C. Individual bacterial colonies were isolated and incubated overnight at 37  C in 10 ml Tryptic Soya Broth (TSB). 1 ml of solution was diluted in 9 ml of fresh TSB and incubated for 2 h at 37  C. 1  106 cfu ml1 of bacteria was prepared using fresh TSB. Immobilized AgNPs@MSA on AApp surfaces were placed in 24 well plates. Each well was filled with 400 ml of diluted culture of bacteria (1  106 cfu ml1) in TSB. Two set of AApp cover slips covered with 400 ml TSB were used as positive and negative control. The sample box was incubated in a container with moist foam for 4 h at 37  C, the broth in each well was replaced with 400 ml fresh TSB and incubated in similar fashion for 24 h at 37  C. After second incubation period, the TSB was removed and wells were rinsed twice with 400 ml of sterile saline (0.9% sterile solution of sodium chloride in water) to clean any loose biofilm formed on the surface. The biofilm formed on the other side of each cover slip was removed with a swab immersed in ethanol (EtOH, 70%). 400 ml of SafranineO (0.1%, Ajax Chemical) solution was added to each well to stain the biofilm adhered to the surface. After 30 min Safranin solution was removed and wells were rinsed twice with 400 ml of Milli-Q water to remove excess Safranin and allowed to dry at ambient temperature. 400 ml of acetic acid (AcOH, 33%) was introduced into each well to lyse the bacterial cells and release the trapped dye. 100 ml of the content of each well was transferred into a 96 well plate and the optical density (OD) was measured by an ELISA plate reader at 490 nm.

The experimental strategy for generation antibacterial surfaces is presented in Fig. 1. First, silver nanoparticles were synthesized using MSA as a capping and stabilizing agent. In the next step, a substrata modified with a plasma polymer layer deposited from vapor of allylamine was immersed in the solution of silver nanoparticles. The positive surface charge of a substrate modified with an AApp film [46] facilitates the electrostatic binding of MSA functionalized silver nanoparticles. MSA has two carboxylic acid groups which are dissociated in water and acquire a negative charge. As it will be demonstrated later in the paper, this immobilization strategy allows for control over the amount of silver nanoparticles immobilized onto the surface. 3.1. Silver nanoparticles: synthesis and characterization The synthesis of silver nanoparticles was carried out in ice cold conditions in order to obtain particles with relatively small size (ca

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Fig. 1. Schematic representation of the strategy for the generation of antibacterial coatings.

11 nm). The selection of MSA as a capping and stabilizing agent had two purposes: Firstly, it was expected that the thiol group of MSA would attach to the surface of the nanoparticles, thus leaving carboxyl acid functionalities exposed which can then be used for immobilization reactions. Secondly, it was expected that the covalent coupling of the thiol group of the MSA to the silver [45] would slow down the rate of oxidation of the silver nanoparticles. Oxidation of the nanoparticles will inevitably happen in the presence of oxygen dissolved in the medium, and thus slowing this will extend the lifetime of the antibacterial surfaces. The insert in Fig. 2a shows the resultant solution of silver nanoparticles. The solution is clear without any signs of precipitation and when in a closed vial is stable for longer than a year. The plasmon resonance adsorption band appears at about 400 nm (Fig. 2a). A TEM image of the nanoparticles is shown in Fig. 2b and the nanoparticles size distribution is presented in Fig. 2c. The particles have spherical shapes and the average size of the nanoparticles is 12.27 nm. The size distribution is relatively narrow with only few larger (still below 30 nm) and smaller particles. The TEM analysis correlated well with DLS measurement which showed average size of 10.3 nm and polydispersity index (PDI) of 0.14. The surface potential of the nanoparticles was determined to be 26.9 mV which is consistent with our expectations. The silver content of the nanoparticle solution was 165 ppm as determined by ICP. 3.2. Preparation of antibacterial surfaces These nanoparticles were further used for surface immobilization which was achieved by immersion of a substrate modified with

an allylamine plasma polymer film into the solution of nanoparticles for a predetermined time period (Fig. 1). Analysis on the chemical composition of the surface of the different model substrata used for nanoparticles immobilization (Thermanox and glass cover slips, silicon wafer) by XPS showed that there was no difference in the silver content which demonstrated the applicability of the technology to different type of surface. The SEM and AFM images presented in Fig. 3a and b show that the nanoparticles are uniformly distributed with no aggregations or uncoated areas being detectable. This is an indication of the homogenous distribution of binding sites (amine groups) on the auxiliary plasma polymer layer and the stability of the colloidal solution of nanoparticles synthesized for this work. The visual appearance of the Thermanox coverslips before and after binding of silver nanoparticles is shown in Fig. 3c. The samples after nanoparticle immobilization (bottom image) had a light brownish color which was homogeneous throughout the surface. Control over the amount of silver nanoparticles that are bound to the surface may be important when different applications are concerned. To demonstrate such control, we chose to vary the time of immobilization as a convenient and flexible approach. Fig. 4 shows the atomic surface concentration of silver determined by XPS on samples where silver nanoparticles were deposited for time intervals between 1 h and 24 h. The concentration of silver on the sample surface increased from 5.2  0.1% (1 h) to 14.6  0.2% (24 h) which demonstrates that the time of immobilization is an efficient method to control the amount of bound silver nanoparticles. Next, we examined the retention of silver on the sample surface of the samples upon immersion in aqueous medium. The rational for these studies was the following. In the presence of oxygen, silver

Fig. 2. a) UVevis spectrum of the synthesized MSA functionalized silver nanoparticles. The insert shows the appearance of the solution; b) a representative TEM image, and c) nanoparticles size distribution (12.27  10.27 nm).

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Fig. 3. Representative a) SEM and b) AFM images (3D view) of MSA functionalized silver nanoparticles immobilized to AApp modified silicon wafer. c) Thermanox coverslip coated with AApp only (top) and after immobilization of silver nanoparticles.

oxidizes to silver oxide which is then dissolved in an aqueous medium. This well-known process is the reason for the release of silver ions which are the actual species that kill bacteria [47]. However, this gradual process will also result of exhaustion of the silver present in the coating. To provide a continuous protection from bacteria, the silver nanoparticles should remain on the surface for a reasonable time period. Fig. 5 shows the rate of reduction in the concentration of silver after immersion of the samples in water for 30 days. We considered this time is sufficient for applications since most of the wound dressings or catheters would be changed within a month and most often in much shorter timeframe. In the case of medical implants, it can be assumed that after one month the wound has sufficiently healed and that the adaptive immune system is strong enough to handle the presence of invading pathogens. The data shows that the coatings developed in this study retained more than 20% of the initial concentration of silver even after 30 days of immersion in an aqueous solution. In the case of silver nanoparticles of this size, the slow oxidation and dissolution can be attributed to the capping action of MSA, which is covalently bound to the surface of the nanoparticles and provides a tight selfassembled monolayer (SAM). 3.3. Assessment of antibacterial efficacy The antibacterial efficacy of the coatings was examined against three clinically significant pathogenic bacteria i.e. S. epidermidis,

Fig. 4. Silver atomic percentage (At%) as a function of the time of immobilization of AgNPs onto AApp films obtained from quantification of the XPS survey spectra.

S. aureus (MRSA) and P. aeruginosa. The examined surfaces contained 14% silver as it was shown by XPS analysis. The quantification of the biofilm development was performed using safranin-O staining method. The application of this technology for determining the biofilm formation is feasible when the value of optical density at 492 nm (OD492 nm) is more than 0.05. Fig. 6 shows the level of reduction in biofilm formation of the coatings containing silver nanoparticles relative to AApp control. The reductions after biofilm safranin-staining were 95.74% for S. epidermidis, 80.45% for S. aureus and 95.77% for P. aeruginosa. An example of the visual appearance of the stained biofilm (red (in the web version)) on silver nanoparticles modified substrate (right) and AApp control (left) is shown in Fig. 6a. Further a disc diffusion assay was conducted. In this experiment, the substrate is placed face down on agar plate where bacteria are grown. The formation of an area around the sample where bacteria cannot grow is termed “zone of inhibition” and is evidence of leaching of the antibacterial agents. It has been shown that the antibacterial action of silver is due to their oxidation and release of silver ions [47e49]. This is clearly evident in Fig. 6b, for S. epidermidis, a clear zone of inhibition of approximately 5 mm is formed around a sample containing MSA modified silver nanoparticles. Such zone is absent in the case of AApp only coverslip. The zone of inhibition in cultured S. aureus was 3 mm. For P. aeruginosa, there was no clear zone of inhibition like S. epidermidis or S. aureus however there was no visible colony growth under the coverslips and there is an area around 2 mm with

Fig. 5. Concentration of silver on the surface of the coating after immersion in Milli-Q water for up to 30 days. The concentration of silver was obtained from quantification of the XPS spectra.

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Fig. 6. (a) An illustration of a biofilm staining of surface coated with AgNPs@MSA (left) and AApp film (right) showing the ability of bacteria to grow on the surface without AgNPs; (b) Zone of inhibition study for S. epidermidis (106 cfu ml1); (c, d, e) relative level of surface bound bacteria in 24 h (Filter 492 nm) of S. epidermidis (c) S. aureus (d) and P. aeruginosa (e), n ¼ 3 data mean  SEM, *** means p < 0.001.

a distinct yellowish color (in the web version). These data show that the coatings developed in this work are active against both gram positive and gram negative bacteria. This is consistent with published literature and often pointed as an advantage of silver to other antibacterial agents which may be active only against only gram positive or gram negative bacteria. 3.4. Cytotoxicity to primary fibroblast cells The potential toxicity of these antibacterial surfaces to primary human dermal fibroblast cells was then examined. Surfaces containing the highest concentration of silver nanoparticles were selected (14% of silver as determined by XPS). The viability of fibroblast cells was studied using the Resazurin assay (Fig. 7). Cells were also cultured on AApp coated glass coverslips in order to examine the effect of the plasma polymer itself. All data was compared to the results obtained on the uncoated glass coverslips and is shown in Fig. 7a. The viability of the cells appears slightly higher on the AApp and silver nanoparticle modified samples. AApp coatings have previously been shown to be non-cytotoxic. The viability of cells on the silver nanoparticle modified surfaces appears even higher; however this was not statistically significant, p ¼ 0.2818. Confocal microscopy images of the cells grown on three types of surfaces after 24 h of incubation are shown in Fig. 7b. Cytotoxicity results suggest that the antibacterial coating obtained by immobilization of MSA-modified silver nanoparticles may not be cytotoxic to mammalian cells. To some extent this result is not surprising. A recent review [24] on the topic summarized data from published studies, which indicated that mammalian cells may be able to tolerate higher amounts of silver than bacteria. It should be stressed that unlike usual practice, the data in our study was obtained from a culture of primary cells and did not utilize immortalized cell lines.

of their role in mediating early innate immune inflammatory responses. Primary bone marrow-derived macrophages (BMDM) were incubated on surfaces containing the highest concentration of silver (14% At% of silver as determined by XPS) and stimulated for the secretion of pro-inflammatory cytokines (Fig. 8a). There were no statistically significant differences in TNF-a, IL-1b or IL-6 secretion between BMDM incubated on AApp films and silver nanoparticle-modified surfaces. Cytokine expression was not detected in unstimulated cultures (data not shown). This was consistent with optical microscopy imaging (Fig. 8b), revealing no differences in cellular attachment nor morphology. Furthermore,

3.5. Effect on innate immune cell function in vitro Finally, the silver nanoparticle-based antibacterial coatings were assessed for their effect on innate immune cell function in vitro, by measuring cytokine secretion from bone marrowderived primary macrophages. These cells were selected because

Fig. 7. (a) Relative fibroblast viability from Resazurin assay compared to a glass only control n ¼ 3 data mean  SEM and n.s: means statistically not significant; (b) confocal images of fibroblast cells viability study on glass (left), AApp (middle) and AgNPs@MSA þ AApp (right) substrates (Scale ¼ 100 mm).

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Fig. 8. Activation response of Bone Marrow Derived Macrophages on AgNP derivatized surfaces. (a) Bone marrow derived macrophages (BMDM) were incubated on AgNP derivatized surfaces then stimulated with LPS and ATP for the detection of (i) TNF-a, (ii) IL-6 and (iii) IL-1b secretion (mean  SEM). (b) Transmission images were taken of adherent BMDM on AgNP surfaces. (c) Viability of adherent BMDM viability was assessed using flow cytometry, gating on side scatter (SSC) lo and DAPI lo cells (i), and quantified as mean percentage  SEM (ii). Results represent two independent experiments, with a one way ANOVA statistical analysis being performed using a Dunnett’s post test (AAp comparison group). ns, p > 0.05; *p  0.05.

there were no statistical differences observed on the viability of the cells, as detected using flow cytometric analyses, to assess the side scatter (SSC) low and DAPI low populations (Fig. 8c). 4. Discussion The surface of medical devices such as implants and wound dressings are an ideal place for colonization by microorganisms [17].

Therefore, tuning the properties of medical device surfaces in a manner that protects it from bacterial adhesion and colonization will reduce infection rates and consequently improve health outcomes. An engaging solution that captured researchers and medical professionals is the placements of antibacterial coating onto device surface [1,50,51], However, achieving prolonged antibacterial activity, low or no toxicity to mammalian cells and a controlled innate immune inflammatory response potential remains a critical issue [1,50,51].

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This study describes a substrate independent approach to create antibacterial coatings that have high antibacterial activity, no cytotoxic effects on primary human fibroblast cells and do not significantly alter the inflammatory response potential of cultured BMDM. The substrate independence of the technology was achieved by using an intermediate plasma polymer film. This type of coatings are well-known to grow independently of the substrate material [39,40]. This was specifically demonstrated for allylamine based plasma polymer films that were chosen for this study [39,40]. The allylamine based plasma polymer film plays another important role. This type of films contain a significant amount of amine groups which are protonated at pH ¼ 7 and give the surface a positive charge [46]. This charge was used to electrostatically immobilize silver nanoparticles which were engineered with a SAM of MSA. We suggest that the SAM of MSA has also a role in reducing the rate of oxidation of the nanoparticles, which will inevitably happen when exposed to an oxygen containing environment. This role is indicated in published studies where a SAM of MSA was demonstrated to reduce the rate of dissolution of gold nanoparticles in potassium cyanide (KCN) solution when compared to non-modified nanoparticles [52]. The immobilization strategy utilized here also allowed for a control over the amount of surface bound silver nanoparticles. Such control may be required for some applications and in particular, when possible cytotoxicity towards mammalian cells is relevant. Mammalian cells have been shown to have a wide range of tolerance to silver nanoparticles ranging from 10 to 100 mg/L [24]. Although these numbers need to be interpreted with care because the studies have not been conducted with particles of the same size and surface functionality, they do point to potential differential capacity of mammalian cells to tolerate silver, which should be considered in the context of different medical applications. The major contribution of this work is that it assesses the same coatings for antibacterial efficacy, cytotoxicity to primary fibroblast cells and their effect on inflammatory cells. The coatings were highly effective against pathogenic bacteria species i.e. S. epidermidis, S. aureus, P. aeruginosa, which are responsible for significant number of infections associated with medical devices. In addition to inhibiting biofilm growth on the surface, the diffusion assay showed an inhibition zone formed around the coated sample which suggests that the surrounding of the device could also be protected. The lack of cytotoxicity to primary human fibroblast may be attributed to two interpretations: 1) higher tolerance of these cells to silver and 2) the slow oxidation and release of silver ions from the immobilized nanoparticles due to the SAM of MSA. At present stage, these interpretations are only hypothetical and further studies would be required to confirm them. Silver nanoparticles have been shown to induce a number of pro-inflammatory responses, markedly increased expression of TNF-a [53,54] and they have also been shown to induce macrophage cell death [38]. Other studies report that nanocrystalline silver suppressed expression of pro-inflammatory cytokines and reduced inflammation [35]. Overall however, the published literature highlights the current controversies here, which is not surprising because studies to date were conducted using different silver sources. Our results point to the fact that silver nanoparticlebased antibacterial coatings developed in this study do not affect BMDM function or viability, which may be advantageous in the context of controlling innate immune effector responses. To this end, we demonstrate that the expression of TNF-a, IL-1b and IL-6 was not significantly altered by the presence of silver nanoparticles. These cytokines are important messengers and their enhanced or suppressed expression may lead to a cascade of physiological events that can alter the natural healing process. The coatings also did not affect the number and morphology of adhered

macrophages. However, understanding the direct connection between silver nanoparticles based antibacterial coatings and immune responses requires in vivo assessment which will be a subject of further studies. 5. Conclusion In summary, we have developed silver nanoparticle-based antibacterial coatings which have excellent antibacterial efficacy and a good mammalian cell biocompatibility profile. Stable colloidal silver nanoparticles were synthesized using sodium borohydride as a reducing agent for AgNO3 and mercaptosuccinic acid as a capping and stabilizing agent. Mean nanoparticle diameter of 11 nm was determined by electron microscopy and DLS analysis. Antibacterial coatings that can be applied to any type of material surface were prepared by the electrostatic immobilization of the carboxyl acid groups functionalized nanoparticles to an amine group rich interlayer prepared by plasma deposition from a vapor of allylamine. Ready control over the amount of silver nanoparticles on the coatings was achieved by immobilization for different time intervals. Excellent antibacterial efficacy was demonstrated against three clinically significant pathogens i.e. S. epidermidis, S. aureus (MRSA) and P. aeruginosa. Studies with primary human dermal fibroblast cells demonstrated no toxicity to mammalian cells. Finally, the effect of the coatings on the innate immune response was studied in culture of primary macrophages. The coatings did not significantly alter the level of expression of pro-inflammatory cytokines and the adhesion and viability of the cells. This is an indication that the coatings might not adversely alter natural innate immune inflammatory processes. Collectively, the coating developed in this work possess and optimal combination of properties which make them interesting for further in vivo testing towards application on medical devices. Acknowledgments KV thanks to The Australian Research Council for support through fellowship FT100100292. The authors would like to thank the group of Professor Michael Roberts at the School of Pharmacy, University of South Australia for help with the sourcing and collection of the skin. References [1] Vasilev K, Cook J, Griesser HJ. Antibacterial surfaces for biomedical devices. Expert Rev Med Devices 2009;6:553e67. [2] Paitoonpong L, Wong CKB, Perl TM. Healthcare-associated infections. In: Melson KE, Williams CM, editors. Infectious disease epidemiology theory and practice. Jones&Bartlet Learning; 2013. pp. 369e466. [3] Weston D. Types of healthcare associated infection. Infection prevention and control. John Wiley & Sons Ltd; 2008. pp. 85e109. [4] Campoccia D, Montanaro L, Arciola CR. The significance of infection related to orthopedic devices and issues of antibiotic resistance. Biomaterials 2006;27: 2331e9. [5] Anderson DJ, Pyatt DG, Weber DJ, Rutala WA. Statewide costs of health careassociated infections: estimates for acute care hospitals in North Carolina. Am J Infect Control 2013;41:764e8. [6] Klevens RM, Edwards JR, Richards Jr CL, Horan TC, Gaynes RP, Pollock DA, et al. Estimating health care-associated infections and deaths in U.S. Hospitals, 2002. Public Health Rep 2007;122:160e6. [7] Graves N, Halton K, Paterson D, Whitby M. Economic rationale for infection control in Australian hospitals. Healthc Infect 2009;14:81e8. [8] Gottenbos B, Busscher HJ, van der Mei HC, Nieuwenhuis P. Pathogenesis and prevention of biomaterial centered infections. J Mater Sci-Mater Med 2002;13:717e22. [9] Halton K, Graves N. Economic evaluation and catheter-related bloodstream infections. Emerg Infect Dis 2007;13:815e23. [10] Darouiche RO. Current conceptsetreatment of infections associated with surgical implants. N Engl J Med 2004;350:1422e9.

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