Article pubs.acs.org/Langmuir
Antibacterial Polyelectrolyte Micelles for Coating Stainless Steel Céline Falentin-Daudré,† Emilie Faure,† Tiziana Svaldo-Lanero,‡ Fabrice Farina,∥ Christine Jérôme,† Cécile Van De Weerdt,§ Joseph Martial,§ Anne-Sophie Duwez,‡ and Christophe Detrembleur*,† †
Center for Education and Research on Macromolecules (CERM) and ‡Nanochemistry and Molecular Systems, Chemistry Department, University of Liège, Sart-Tilman B6a, 4000 Liège, Belgium § Laboratoire de Biologie Moléculaire et de Génie Génétique, Center of Biomedical Integrative Genoproteomics, CHU, B34 Sart-Tilman, B-4000 Liège, Belgium ∥ ArcelorMittal Liège Research, Bd de Colonster B57, 4000 Liège, Belgium S Supporting Information *
ABSTRACT: In this study, we report on the original synthesis and characterization of novel antimicrobial coatings for stainless steel by alternating the deposition of aqueous solutions of positively charged polyelectrolyte micelles doped with silver-based nanoparticles with a polyanion. The micelles are formed by electrostatic interaction between two oppositely charged polymers: a polycation bearing 3,4-dihydroxyphenylalanine units (DOPA, a major component of natural adhesives) and a polyanion (poly(styrene sulfonate), PSS) without using any block copolymer. DOPA units are exploited for their well-known ability to anchor to stainless steel and to form and stabilize biocidal silver nanoparticles (Ag0). The chlorine counteranion of the polycation forms and stabilizes biocidal silver chloride nanoparticles (AgCl). We demonstrate that two layers of micelles (alternated by PSS) doped with silver particles are enough to impart to the surface strong antibacterial activity against gram-negative E. coli. Moreover, micelles that are reservoirs of biocidal Ag+ can be easily reactivated after depletion. This novel water-based approach is convenient, simple, and attractive for industrial applications.
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biocide-loaded coatings22 or by chemically grafting the biocides to the surfaces. In the first case, the AB activity is due to the diffusion of the biocide out of the coating. In the second case, bacteria are killed when in contact with the surface. Many examples can be found in the literature of anchoring biocides to SS such as the electrografting of acrylates postmodified to obtain antibacterial properties,14 the grafting of macromolecules bearing quaternary ammonium groups,15,16 the postquaternization of amino groups grafted onto a surface prefunctionalized with cold plasma,17 the formation of biocidal multilayered polyelectrolyte films,18 and the grafting of peptides onto macromolecules adsorbed or grafted onto stainless steel.19−21 However, these are generally multistep processes and mostly use organic solvents that are toxic and are not desirable for industrial applications. Simpler, water-based AB coating solutions are therefore potentially more desirable for coating substrates. The layer-by-layer (LbL) assembly of oppositely charged polyelectrolyte layers is a simple and environmentally friendly method of modifying surface properties.23 In a previous study, we reported on an all-in-one approach to preparing refillable
INTRODUCTION Because of its resistance to corrosion and chemicals and its mechanical and aesthetic properties, stainless steel (SS) is widely used in daily life such as in the food industry, household appliances, and surgery.1 Despite good cleanability, bacteria absorb easily. On a solid surface, bacteria form colonies and subsequently biofilms that serve as reservoirs for the development of pathogenic infection.2 Therefore, for hygienic reasons, it is necessary to develop new strategies to protect the surface against these microorganisms by preventing them from adhering and, in the case of adhesion, to kill them while removing the biofilm. This can be accomplished by physical or chemical surface modifications. To avoid the colonization of the surface by bacteria, two principal strategies have been developed: the first one is to immobilize antifouling coatings that prevent bacterial adhesion,3−13 and the second one is to develop antibacterial surface films.14−21 Many examples can be found in the scientific literature of antifouling coatings such as self-assembled monolayers,3−5 the formation of a multilayer film with poly(ethylene glycol) (PEG),7 the immobilization of dopamine end-functionalized by PEG,8 the grafting of PEG by a cold plasma technique9 or by silane coupling agents,10 and the grafting of lysozymes or PEG onto poly(ethylene imine) adsorbed onto substrates,11−13 to name a few. Antibacterial surfaces (AB) can be generated by painting the substrates with © 2012 American Chemical Society
Received: January 27, 2012 Revised: March 13, 2012 Published: April 16, 2012 7233
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Scheme 1. Strategy for the Formation of (a) Positively Charged Micelles (PSS/P(mDOPA)-co-P(DMAEMA+)) and (b) Positively Charged Silver-Loaded Micelles (PSS/P(mDOPA)-co-P(DMAEMA+)/Ag0/AgCl)
Scheme 2. AB Multilayer Films from Micellesa
a
First layer, PSS/P(mDOPA)-co-P(DMAEMA+) micelles; second layer, PSS; third layer, silver-loaded PSS/P(mDOPA)-co-P(DMAEMA+)/Ag0/ AgCl micelles.
antimicrobial films using the LbL deposition of polyelectrolytes.18 The first layer of synthetic glue was used as an anchoring layer of a biocidal multilayered polyelectrolyte film onto SS. This glue was a copolymer of a cationic methacrylate bearing an ammonium group and a methacrylamide bearing 3,4-dihydroxyphenylalanine (P(mDOPA)-co-P(DMAEMA+), 1, Scheme 1). Indeed, 3,4-dihydroxyphenylalanine (DOPA) is a major component of natural glue proteins secreted by mussels,24 and the catechol group of DOPA is thought to be responsible for adhesion to inorganic surfaces such as glasses,
metals (gold and stainless steel),8,25 and metal oxides (TiO2, Al2O3, Fe2O3, and SiO2),8,25b,c,26 although the actual adhesion mechanism is not yet fully understood.27 Polyelectrolyte (P(mDOPA)-co-P(DMAEMA+) (1) was therefore used as the anchoring layer for the antibacterial coating made by the LbL deposition of a polycation doped with AB silver-based nanoparticles (P(DOPA)-co-P(DMAEMA+)/AgCl/Ag0) with a polyanion (poly(styrene sulfonate), 2, PSS). Although this surface modification provided high antibacterial activity against gram-negative E. coli bacteria, it required the LbL deposition of 7234
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same technique, except that the P(mDOPA)-co-P(DMAEMA+)/PSS/ Ag0/AgCl hybrid micelle solution substitutes for the P(mDOPA)-coP(DMAEMA+)/PSS one after the first two layers. Characterization Methods. Dynamic Light Scattering (DLS). A Delsa Nano C particle analyzer (Beckman Coulter) equipped with a laser diode source (wavelength 658 nm, power 30 mW) was used to measure the hydrodynamic diameter of the aqueous micelle solutions. Scattering data were collected for at least 70 individual measurements at a constant scattering angle and averaged for each sample. The obtained scattering data were fitted using volume-weighted cumulative analysis to estimate the diffusion coefficient of the micelles in solution.31a,b The hydrodynamic diameter of the samples (DH) was obtained using the Stokes−Einstein relationship. Zeta Potential Measurement. The coated glass surfaces were characterized by using a Delsa Nano C particle analyzer (Beckman Coulter) equipped with a laser diode source (wavelength 658 nm, power 30 mW) at 25 °C. The polyacrylamide-coated quartz cell with a 24 mm × 50 mm × 5 mm groove was covered with the glass slide sample of 15 mm × 35 mm × 1 mm size. The space between the sample and the cell was then filled with a 7.5 mM NaCl solution. Quartz Crystal Microbalance. Film growth is followed in real time using a Q-Sense E4 quartz crystal microbalance with a dissipation technique (QCM-D). The stainless steel-coated AT-cut resonator (fundamental frequency 5 MHz) was used as received. First, deionized water was introduced into the cell, and the circulation was maintained until obtaining a stable baseline. LbL deposition was then carried out by switching the liquid exposed to the crystal from deionized water to the micelle solution (flow rate 250 μL/min, temperature 25.09 °C ± 0.02 °C). The micelle solution was allowed to adsorb onto the substrate for 10 min before being rinsed with deionized water to get a uniform positive coating on the SS sensor. After the micelle solution was used, the sensor was rinsed with deionized water and then a polystyrene sulfonate solution was injected. This cyclic procedure was repeated several times. Transmission Electron Microscopy. TEM images were recorded using a Phillips CM100. To prepare the TEM samples, a drop of the dilute aqueous solution was deposited onto a copper grid precoated with a thin film of Formvar and carbon. Two minutes after deposition, the excess aqueous solution was blotted away with a strip of filter paper. Atomic Force Microscopy (AFM). Acoustic intermittent contact mode AFM measurements were performed in air at room temperature using a PicoPlus 5500 microscope (Agilent Technologies, Inc.) and silicon cantilevers (PPP-NCH, nanosensors, nominal spring constant 42 N/m). Gwyddion SPM data analysis software has been used to analyze the data. Spectroscopic Ellipsometry (SE). SE measurements were made using a SOPRA GES 5 working in the UV−visible range from 300 to 900 nm. The angle of incidence was 75°. To extract information from the SE spectra, an optical model of the presumed surface structure has to be built. The interpretation of the SE data is then carried out by fitting the calculated responses Ψ(λ) and Δ(λ) of the optical model to the experimental data by using simulation. The optical model consists of a stratified structure of layers with flat and parallel interfaces, with each layer being described by its thickness and optical constants. Data obtained for the uncoated substrate (stainless steel) are expressed in terms of pseudo-optical constants. The polymeric layers are described using a Cauchy dispersion relation in order to model the refractive index of the film. Antimicrobial Assessment. The antibacterial activity of the multilayered polyelectrolyte films on stainless steel against the gramnegative bacteria Escherichia coli was assessed by a viable cell-counting method.29a A freeze-dried ampule of E. coli (DH5α) was opened, and the culture was picked out with a micropipet and placed in 2 mL of nutrient broth, which was then incubated (incubator shaker model G25, New Brunswick, Scientific Co. Inc., Edison, NJ, USA) at 37 °C overnight. (The composition of 1 L of nutrient broth (Luria−Bertani) was 10 g of bactotryptone and 5 g of yeast extract in sodium chloride.) Then, 200 μL of the culture was placed in 100 mL of nutrient broth,
about 45−60 bilayers to impart high AB activities to the surface, which makes the process difficult to scale up for production lines in industry. In the present work, we report on a convenient, straightforward approach to the fast formation of antibacterial coatings by the deposition of an aqueous solution of novel hybrid polyelectrolyte micelles made of a polycation bearing DOPA units (P(mDOPA)-co-P(DMAEMA+)) doped with silver nanoparticles and a polyanion (polystyrene sulfonate (PSS)). Scheme 1 illustrates the formation of the micelles, and Scheme 2 illustrates the deposition technique used. Mixing oppositely charged polymers in water in suitable proportions led to the spontaneous formation of micelles that are stable in water without using any block copolymers. The AB coating is built in only five dipping steps (Scheme 2). The SS surface is first dipped into a solution of positively charged micelles bearing DOPA groups on their surfaces used as an adhesion promoter. The second layer is formed by dipping the surface into an aqueous solution of the polyanion (poly(styrene sulfonate)). And the third layer is deposited from an aqueous solution of positively charged micelles containing ammonium groups and silver-based nanoparticles for high AB properties over a broad spectrum of bacterial strains that are commonly found in daily life.28 These last two steps are repeated once again.
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EXPERIMENTAL SECTION
Materials. P(mDOPA)-co-P(DMAEMA+) (Mw = 106 g/mol, 20 mol % DOPA) was prepared by copolymerizing 2-methacryloxyethyltrimethylammonium chloride with N-methacryloyl-3,4-dihydroxy-Lphenylalanine methyl ester in water according to our reported procedure.18 Poly(sodium 4-styrene sulfonate) (PSS, Aldrich, Mw = 7000 g/mol) was used as received. All other reagents and solvents were purchased from Aldrich and used as received. Stainless steel (SS, 304 2R) was used as a substrate for LBL assembly and was provided by ArcelorMittal (Arcelor Research Industry Liege, ARIL). Glass surfaces used for the LBL assembly characterization were purchased from VWR (Geschliffen microscope slide). Formation of a P(DOPA)-co-P(DMAEMA+)/PSS Micelle Solution. A stock solution of P(mDOPA)-co-P(DMAEMA+) was freshly prepared in water at a concentration of 0.5 g/L. Then, 10 mL of this stock solution was placed in a vial, and 915 μL of a solution of polystyrene sulfonate (PSS) was added at a concentration of 1 g/L under vigorous stirring. The solution was allowed to react for 1 h under vigorous stirring at room temperature. Formation of a P(DOPA)-co-P(DMAEMA+)/PSS/Ag0/AgCl Hybrid Micelle Solution. A solution of AgNO3 (6.67 × 10−5 mol) in water (1 mL) was slowly added to an aqueous solution of P(mDOPA)-co-P(DMAEMA+) (0.5 g/L, 10 mL, nDOPA = 6.67 × 10−5 mol) in the dark under vigorous stirring at room temperature overnight. The experiment has been carried out in the dark because AgNO3 is photosensitive. Then, a solution of PSS (1 g/L, 915 μL) was slowly added to the suspension under vigorous stirring. The final suspension was stirred for 1 h at room temperature before use. Preparation of the Multilayered Polyelectrolyte Films. Stainless steel samples were cut from as-received 1-mm-thick foils into 2.7 cm × 2 cm pieces for ellipsometry and 2 cm × 2 cm pieces for antimicrobial tests and AFM. Glass samples were cut out from asreceived 1-mm-thick foils, 1.7 cm × 3.5 cm for the zeta potential. Each substrate was first cleaned with acetone and ethanol, dried in a flow of clean nitrogen, and then wiped with optical paper. The substrate was first immersed in an aqueous micelle solution (P(mDOPA)-coP(DMAEMA+)/PSS, as prepared above), rinsed with deionized water, and then dipped into a solution of PSS (1 g/L). The immersion time was 2 min for each step at room temperature. By repeating the above two steps in a cyclic procedure, the LBL multilayered film was fabricated. Silver-loaded multilayered films were also prepared by the 7235
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and the bacterial culture was incubated at 37 °C for 4 h. At this stage, the culture of E. coli contained ca. 108 cells/mL (absorbance at 600 nm equal to 0.6), and this was used for the test. The original cell concentration was determined by the spread-plate method. By appropriate dilution with sterilized 0.9% saline solution, a culture of about 105 cells/mL was prepared and used for antibacterial testing. Films deposited on stainless steel were sterilized by UV irradiation (1 h for each face) and exposed to the E. coli cell suspension (15 mL containing about 105 cells/mL). At a specified time, 0.1 mL of the bacteria culture was added to 0.9 mL of sterilized 0.9% saline solution (the solution was sterilized at 121 °C for 120 min), and several dilutions were carried out. The surviving bacteria were counted by the spread-plate method. At various exposure times, 0.1 mL portions were removed and quickly spread on the nutrient agar. After inoculation, the plates were incubated at 37 °C for 24 h, and the colonies were counted. Counting was tripled for each experiment.
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Figure 1. Hydrodynamic diameter distributions obtained by DLS for the polyelectrolyte micelles formed by the addition of PSS to P(mDOPA)-co-P(DMAEMA+) using the (−/+) charge ratio of 0.3 at 25 °C (detection angle, 90°; temperature, 25 °C; solvent, unfiltered deionized water).
RESULTS AND DISCUSSION Formation and Characterization of Micelles. Since the first reports on micelles formed through electrostatic interaction between two oppositely charged polymers in the mid-1990s,30 the field has attracted considerable interest. Since then, numerous publications on this type of assembly have been found30c,31 in the literature, and this research field has been the topic of reviews.31g,32 To prepare these polyelectrolyte micelles, all of these publications make use of at least one of the partners that is a block copolymer bearing a polyelectrolyte sequence and a nonionic hydrophilic block. This last block is necessary to stabilize the polyelectrolyte micelles in water. To the best of our knowledge, stable aqueous solutions of polyelectrolyte micelles formed without using any block copolymer containing a sequence able to stabilize the micelles in water have not been described to date. Here, we report on the original and simple synthesis of multifunctional micelles by the self-assembly of a positively charged random copolymer with a negatively charged one in water, without using any block copolymer. The first type of micelles is formed through electrostatic interaction between the random copolymer P(mDOPA)-co-P(DMAEMA + ) (1) (Scheme 1; 20 mol % DOPA) and poly(styrene sulfonate) (2) (PSS) in water. Charge ratios and addition modes of the two partners are controlled to form positively charged micelles with DOPA groups at their corona to favor the anchoring of the micelles to the surface through DOPA/metal interactions. For that purpose, the slow addition of PSS (1 g/L) to an aqueous solution of P(mDOPA)-co-P(DMAEMA+) (0.5 g/L) results in the spontaneous formation of positively charged micelles at room temperature provided that the charge ratio (−/+) is between 0.3 and 0.5. Under these conditions, the solutions of micelles are stable for at least 1 month. The polyelectrolytes agglomerate and precipitate for ratios higher than 0.5. Therefore, suspensions with a PSS/P(mDOPA)-co-P(DMAEMA+) charge ratio of 0.3 are systematically considered in this study. Dynamic light scattering (DLS) measurements (Figure 1) evidence the formation of micelles with an average hydrodynamic diameter of 88 nm but with a rather high polydispersity (PDI = 0.2). Transmission electron microscopy (TEM) evidences the presence of spherical micelles whose sizes range from 66 to 100 nm (Figure 2). Figure 3d shows that this aqueous solution of PSS/P(mDOPA)-co-P(DMAEMA+) micelles is clear even without using a filtration step. The second type of micelle is designed to impart strong antibacterial activity to the coatings. They involve the same partners as the micelles described above, namely, PSS and
Figure 2. TEM images of polyelectrolyte micelles formed by the addition of PSS to P(mDOPA)-co-P(DMAEMA+) using the (−/+) charge ratio of 0.3 at 25 °C.
Figure 3. Aqueous solutions of P(mDOPA)-co-P(DMAEMA+) (a) without silver nanoparticles, (b) with silver nanoparticles, and (c) with silver nanoparticles after 5 days. PSS/P(mDOPA)-co-P(DMAEMA+) micelles (d) without silver nanoparticles, (e) with silver nanoparticles, and (f) with silver nanoparticles after 5 days.
P(DOPA)-co-P(DMAEMA+), and are also positively charged. The AB activity promoted by the ammonium groups of the polycationic corona is, however, further boosted by doping this polycation with silver-based (nano)particles (Ag0 and AgCl) formed in situ according to our previously reported procedure.18 For that purpose, a silver nitrate solution (1 g/ L) is added to the aqueous solution of P(mDOPA)-coP(DMAEMA+) (0.5 g/L) prior to the formation of the micelles. The addition of AgNO3 in a Ag+/DOPA molar ratio of 1/1 results in the nearly instantaneous formation of a milky 7236
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objects of higher size is also observed between 4000 and 10 000 nm by DLS (Figure 4). Presumably, this fraction is attributed to AgCl nanoparticles formed by ionic exchange between AgNO3 and the polycation.33 TEM analysis (Figures 5d and 5e) of the unfiltered micelles shows the presence of agglomerated AgCl nanoparticles. Because the formation of the polyelectrolyte micelles was imaged by TEM under dry conditions, which are not favorable to the self-assembled systems, AFM measurements in a liquid environment were performed to determine if the procedure for the preparation of the samples for TEM analysis affects the integrity of the micelles. A diluted solution of micelles (loaded or not by silver based nanoparticles) has been left in contact with a freshly cleaved mica surface for 30 min before the surface was thoroughly rinsed with DI water to remove micelles that were weakly attached to the surface. An investigation of the morphology of the micelles in a liquid environment (DI water) was performed by acoustic intermittent contact mode AFM measurements. Disclike structures of different dimensions are present, as already shown in the TEM images (Figure S1, Supporting Information). The procedure for preparing the TEM samples, therefore, does not affect the structure and the integrity of the micelles. Formation and Characterization of Polyelectrolyte Multilayer Films. The first deposited layer is the positively charged micelle (PSS/P(mDOPA)-co-P(DMAEMA+), also defined as Mic+ for clarity)) that does not contain the silverbased nanoparticles in order to benefit from the catechol groups of DOPA for adherence to the substrate. The next layers are then built by alternating the deposition of negatively charged poly(styrene sulfonate) (PSS) with the positively charged hybrid micelles (PSS/P(mDOPA)-co-P(DMAEMA+)/ Ag0/AgCl, also defined as Mic+/Ag/AgCl for clarity) (Scheme 2). This last step is repeated once again. The LbL building is characterized by the zeta potential, quartz crystal microbalance with dissipation (QCMD), and ellipsometry. Because the silver-based nanoparticles interfere with these analytical techniques and preclude the possibility of obtaining reliable results, only the deposition of unloaded micelles (Mic+) is studied. As first evidence of the multilayer formation, ζ-potential measurements are conducted on glass as a model surface (steel was not feasible because of its intrinsic conductivity). This technique offers the opportunity to quantify the overall charge on the surface and so proves the balance between positive and negative values during the LbL building. Figure 6 shows the change in the zeta potential of the multilayer films as a function of the layer number as measured. The initially negative zeta potential of the glass surface becomes positive after the deposition of the cationic Mic+ micelles. The deposition of the anionic homopolymer (PSS) onto this first micellar layer leads to surface charge reversal, producing a negative zeta potential. Such charge reversal is repeatedly observed up to at least the eighth layer. Figure 6 thus gives evidence that both partners are inserted into the multilayer and contribute to the electrostatic counterbalance, which is essential for film growth. As more evidence of the multilayer film buildup, quartz crystal microbalance coupled with dissipation (QCM-D) is used to follow the film growth in real time on SS sensors by measuring the variation in the resonance frequency (Δf) versus time. A decrease in Δf indicates polymer deposition.34 Figure 7 shows that all partners are successfully deposited and remain on the substrate even after rinsing with water. The rinsing steps
yellow-brown suspension (Figure 3b) that is stable for at least 5 days (Figure 3c). According to our previous work,18 the silver cation is reduced by the catechols in Ag0 nanoparticles but also forms AgCl particles by combining with the chlorine counteranion of the polycation (Scheme 1). Both particles are stabilized by the copolymer that prevents their precipitation in water. The hybrid AB micelles are then formed by slowly adding a solution of PSS (1 g/L) to the P(mDOPA)-co-P(DMAEMA+)/ Ag0/AgCl hybrid dispersion in a (−/+) charge ratio of 0.3 (Figure 3e). These suspensions are stable for about 5 days without stirring and have to be used during this period of time (Figure 3f). After the solution is filtered through a 0.45 μm filter, DLS measurements (Figure 4) show that the micelles
Figure 4. Hydrodynamic diameter distributions obtained by DLS for the hybrid polyelectrolyte micelles (PSS/P(mDOPA)-co-P(DMAEMA+)/Ag0/AgCl) at 25 °C with and without filtration (angle of detection, 90°; temperature, 25 °C; and solvent, deionized water).
have an average hydrodynamic diameter equal to 123 nm with a rather high polydispersity (PDI = 0.21). TEM analysis evidences the coexistence of spherical micelles with small spherical silver-based nanoparticles in the 5 nm range (Figure 5). These nanoparticles can be found on the micelles and also in the surrounding medium. The latter are certainly stabilized by some excess P(mDOPA)-co-P(DMAEMA+) copolymer that is not involved in the polyelectrolyte micelles, as previously observed.18 When the solution is not filtered, a population of
Figure 5. TEM images (A) of the hybrid micelles (PSS/P(mDOPA)co-P(DMAEMA+)/Ag0/AgCl), (B) at higher magnification, and (C) of Ag0 nanoparticles not fixed to the micelles, and (D, E) of AgCl particles not fixed to the micelles. 7237
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Ag nanoparticles (Mic+/PSS/Mic+/Ag/AgCl) on SS evidence a highly covering coating of the surface by the multilayer film (no naked areas were found). Antibacterial Activity of the Multilayered Polyelectrolyte Films. The time dependence of the antibacterial activity of a series of multilayered films deposited on stainless steel against E. coli is evaluated by using the viable-cell counting method.29 After one night of incubation and the dilution of the samples, the number of viable cells in the suspension in contact with the coated substrate (4 cm2) is counted. Figure 9 is a plot Figure 6. ζ-potential variation of micelles (Mic+/PSS)n coated as a function of the number of deposited layers. Each data point was averaged over 10 runs, and the error bars represent the standard deviations.
Figure 9. Semilogarithmic plot of the number of viable E. coli cells vs the exposure time for (a) bare stainless steel, (b) [P(mDOPA)-coP(DMAEMA+)/[PSS/P(mDOPA)-co-P(DMAEMA+)-silver]45,18 (c) Mic+/(PSS/Mic+/Ag/AgCl)1, (d) Mic+/(PSS/Mic+/Ag/AgCl)2, (e) [P(mDOPA)-co-P(DMAEMA+)/[PSS/P(mDOPA)-co-P(DMAEMA+)silver]60,18 and (f) Mic+/(PSS/Mic+)3. The test was realized at 37 °C.
of the log of the number of survivors against the exposure time when the same number of E. coli cells (105 per mL) is exposed to stainless steel surfaces that are uncoated, coated with three layers of micelles (PSS/Mic+)3, and coated with silver-loaded micelles (PSS/Mic+/Ag/AgCl)n (n = 1, 2). For both coated samples, the first layer is always the anchoring micelle (Mic+) bearing the DOPA groups. Bare stainless steel and non-silverloaded (PSS/Mic+)3 films display no antibacterial activity (Figure 9a,f). When the micelles are silver-loaded, the film shows enhanced antibacterial activity (Figure 9c,d) that increases with the number of bilayers and thus with the amount of released Ag+. No bacteria survive after 2 h of contact with the silver-loaded Mic+/(PSS/Mic+/Ag/AgCl)2 film (Figure 9d). As a comparison,
Figure 7. Variation of frequency (Δf) measured by QCM-D as a function of time at 25 °C for the LbL deposition of Mic+ and PSS, where the overtone number is 11. Mic+ states for the cationic (P(mDOPA)-co-P(DMAEMA+)/PSS) micelle.
between each layer are applied to remove excess polymer before the deposition of the next layer. The ellipsometry measurement for a monolayer of Mic+ on stainless steel gives a thickness of 10−12 nm, suggesting that the micelles are flattened on the surface probably because of favorable interactions with the substrate. A height acoustic intermittent contact mode AFM image (Figure 8) with two layers of micelles (Mic+/PSS/Mic+) and two layers loaded with
Figure 8. Height acoustic intermittent contact mode AFM images of the SS surface coated with (a) Mic+/PSS/Mic+ and (b) Mic+/PSS/Mic+/Ag/AgCl. Size of the images: 1.0 μm × 1.0 μm. 7238
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of PSS, Ag0 and AgCl nanoparticles are formed in situ and stabilized by the copolymer without using any other chemical reagent. Catechol groups of DOPA reduce Ag+ to Ag0, and the counteranion of the ammonium group combines with Ag+ to form insoluble AgCl. The addition of PSS to this hybrid solution leads to the formation of novel hybrid polyelectrolyte micelles doped with biocidal Ag0 and AgCl (nano)particles (noted Mic+/Ag/AgCl) provided that excess cationic copolymer is used. The AB coating is formed by the layer-by-layer deposition of Mic+, PSS, and Mic+/Ag/AgCl. High antibacterial activity against gram-negative E. coli bacteria is observed with only two layers of AB micelles (Mic+/Ag/AgCl) deposited onto stainless steel. The film can also be reloaded with silver-based particles by simply dipping it in a AgNO3 aqueous solution, which again boosts the antibacterial activity. As a comparison, we previously showed that when PSS is alternatively codeposited with P(mDOPA)-coP(DMAEMA+) doped with Ag0/AgCl according to a conventional LbL process, a 60-bilayer film was required to reach the same level of AB activity as that obtained by the micelles approach.18 Our new deposition process based on micelles is therefore a straightforward approach to imparting strong antibacterial activity to stainless steel from aqueous solutions under mild conditions and with a limited number of steps.
we previously showed that when the cationic copolymer P(mDOPA)-co-P(DMAEMA+) was silver-loaded and alternatively codeposited with PSS according to a conventional LbL process the deposition of 60 bilayers was required to reach the same level of AB activity18 (Figure 9e). Thus, processing the same partners as hybrid micelles is a straightforward and fast approach to imparting strong antibacterial properties to SS by the deposition of only three bilayers. Indeed, the advantage of the micelle approach over the “conventional” LbL approach is that the different partners (polycation and polyanion) are preassembled in solution prior to deposition onto the surface. Therefore, the deposition of the micelles is much faster and does not require the LbL deposition of 60 bilayers of the polycation and polyanion to get the same AB functionality. After the total depletion of the silver particles, the biocidal effect can be reactivated by dipping the silver-depleted film into a 0.1 M silver nitrate solution for 1 h, followed by extensive rinsing with deionized water to remove excess AgNO3. New AgCl particles form in the polycationic corona of the micelles by ion exchange without needing any other chemical reagent. Ag0 nanoparticles can be reformed only when catechols are still available in the film. When the catechols are fully consumed by oxidation by Ag+, the further addition of AgNO3 cannot form anymore Ag0. Figure 10 shows that the antimicrobial activity is
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ASSOCIATED CONTENT
S Supporting Information *
AFM measurements in a liquid environment. This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*Tel: (+32) 4-3663465. Fax: (+32) 4-3663497. E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
Figure 10. Semilogarithmic plot of the number of viable E. coli cells vs the exposure time for (a) bare stainless steel and for stainless steel coated with hybrid micelles Mic+/(PSS/Mic+/Ag/AgCl)2 for (b) the first test, (c) the second test, and (d) the third test (after reactivation of the film by dipping in a 0.1 M AgNO3 solution). The test was realized at 37 °C.
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ACKNOWLEDGMENTS We are grateful to the “Region Wallonne”, Arcelor-Mittal, and the University of Liège for funding through PPP research program BIOCOAT. We thank the BIOCOAT team for its contribution, in particular, Fabrice Farina for his assistance with the ellipsometric measurements and Christelle Vreuls and Germaine Zocchi for helpful discussions. Christophe Detrem̂ de Recherche” by the F.R.S.-F.NRS (Belgium) bleur is “Maitre and thanks the F.R.S.-F.NRS for financial support. C.D., A.S.D. and C.J. are grateful to BELSPO through IAP VI/27 program for financial support.
restored after the reactivation of the film. Because biocidal silver-based nanoparticles cannot form on SS by simply dipping neat SS into AgNO3, this reactivation shows that most of the organic coating remains on the SS surface after immersing it in water for 19 h.
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CONCLUSIONS In this article, we report on a convenient approach for preparing refillable antibacterial (AB) coatings on stainless steel using aqueous-based solutions. We have first formed stable cationic polyelectrolyte micelles (noted Mic+) bearing DOPA groups in their corona by the addition of a solution of polystyrene sulfonate (PSS) to a solution of a cationic random copolymer of a methacrylate bearing an ammonium group with a methacrylamide bearing DOPA (P(mDOPA)-co-P(DMAEMA+)) in water. Importantly, no block copolymer was necessary to stabilize the micelles in water, making the process simple and economically attractive. When silver nitrate is added to the solution of this copolymer prior to the addition
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