Multifunctional Polyphenols- and Catecholamines-Based Self

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Multifunctional Polyphenols- and Catecholamines-Based SelfDefensive Films for Health Care Applications Chetna Dhand,† Sriram Harini,† Mayandi Venkatesh,† Neeraj Dwivedi,‡ Alice Ng,† Shouping Liu,† Navin Kumar Verma,§ Seeram Ramakrishna,∥ Roger W. Beuerman,†,⊥ Xian Jun Loh,# and Rajamani Lakshminarayanan*,†,⊥ †

Anti-Infectives Research Group, Singapore Eye Research Institute, Singapore 168751 Department of Electrical and Computer Engineering, National University of Singapore, Singapore 117576 § Lee Kong Chian School of Medicine, Nanyang Technological University, Experimental Medicine Building, 59 Nanyang Drive, Singapore 636921 ∥ Center for Nanofibers and Nanotechnology, Department of Mechanical Engineering, National University of Singapore, Singapore 117576 ⊥ SRP in Neuroscience and Behavioral Disorders, Duke-NUS Graduate Medical School, 8 College Road, Singapore 169857 # Institute of Materials Research and Engineering, A*STAR (Agency for Science, Technology and Research), 3 Research Link, Singapore 117602 ‡

S Supporting Information *

ABSTRACT: In an era of relentless evolution of antimicrobial resistance, there is an increasing demand for the development of efficient antimicrobial coatings or surfaces for food, biomedical, and industrial applications. This study reports the laccase-catalyzed room-temperature synthesis of mechanically robust, thermally stable, broad spectrum antimicrobial films employing interfacial interactions between poly(vinyl alcohol), PVA, and 14 naturally occurring catecholamines and polyphenols. The oxidative products of catecholamines and polyphenols reinforce the PVA films and also alter their surface and bulk properties. Among the catecholamines-reinforced films, optimum surface and bulk properties can be achieved by the oxidative products of epinephrine. For polyphenols, structure−property correlation reveals an increase in surface roughness and elasticity of PVA films with increasing number of phenolic groups in the precursors. Interestingly, PVA films reinforced with oxidized/polymerized products of pyrogallol (PG) and epinephrine (EP) display potent antimicrobial activity against pathogenic Gram-positive and Gram-negative strains, whereas hydroquinone (HQ)-reinforced PVA films display excellent antimicrobial properties against Gram-positive bacteria only. We further demonstrate that HQ and PG films retain their antimicrobial efficacy after steam sterilization. With an increasing trend of giving value to natural and renewable resources, our results have the potential as durable self-defensive antimicrobial surfaces/films for advanced healthcare and industrial applications. KEYWORDS: antimicrobial films, laccase-catalyzed, polyphenols, catecholamines, self-defensive

1. INTRODUCTION Microbial contamination of air, water, surfaces, and medical devices and implants is a major concern for human health and environment, given the fact that hospital-acquired infections are the leading cause of morbidity and mortality among microbialrelated complications.1−3 Together with the rising number of aging populations and the evolution of antimicrobial-resistant microbial strains, there is an increasing demand for medical and industrial materials that can impede or prevent microbial contaminations.4−6 The mechanism of surface microbial colonization is a multifaceted process involving complex interplay among microbes, surfaces, and the surrounding environments. Nevertheless, adhesion of microbes to the surface is the most important primary step that is determined by physicochemical properties of the surfaces such as © XXXX American Chemical Society

roughness, charge density, degree of hydrophobicity, Lewis acid−base character, hydrogen bonding capacity, nanopatterning, and chemical structure, among others.7−14 Thus, several key challenges are associated with designing surfaces that prevent microbial colonization and subsequent biofilm formation.15 Two important strategy types, passive and active, have been employed to design antimicrobial surfaces. Passive strategies involve prevention of microbial attachment on the surface by judiciously tuning the surface features.7−9 Active strategies rely on incorporated antimicrobial compounds that actively promote surface microbial decontamination by Received: October 10, 2015 Accepted: December 24, 2015

A

DOI: 10.1021/acsami.5b09633 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

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ACS Applied Materials & Interfaces

important MRSA and S. epidermis, Gram-negative bacteria, and yeasts strains expanding the potential of our approach for diverse applications where microbial decontamination/decolonization is demanded.

interfering with the biochemical survival strategies of the microbes. Active strategies can be further classified as “with offsurface antimicrobial effect” involving the elution of antimicrobial component into the biological milieu to target the planktonic cells or as “with on-surface antimicrobial effect” in which antimicrobials are grafted on the surface to exercise contact killing of the adhering microbial cells.16−18 Using the latter approach, a number of antimicrobial materials have been prepared for both hard and soft surfaces.19 Recently, nature-inspired adherent polycatecholamines and polyphenols have gained considerable attention because of their inherent potential to form multifunctional surface coatings on wide variety of surfaces such as metals, metal oxides, minerals, polymeric, and superhydrophobic surfaces via simple and inexpensive precleaning strategies and fabrication methods.20−23 The coating method offers a versatile way of tuning the surface hydrophobicity, improving the surface tribological properties as well as enhancing mechanical strength and thermal stability of the materials.21,24−27 Some of the polyphenols are known for their potent antioxidant, antifouling, and antimicrobial properties.28,29 These features evidently indicate their competence toward the development of multifunctional antimicrobial surfaces. Outstanding research efforts have been reported by Messersmith and co-workers to unravel the potential of polyphenol and polycatecholamine coatings toward various applications including their efficacy as antimicrobial surfaces.30−32 In the context of antimicrobial coatings, Sileika et al. exploited the use of polydopamine coating to graft antifouling polymers and induce antimicrobial silver nanoparticles to generate surfaces with multifunctional properties.30 In another work, these authors reported the high ionic strength mediated formation of dopamine−melanin coatings for the release of cationic antimicrobial compounds.31 Inspired by the strong solid−liquid interfacial activity exhibited by plant polyphenols, multifunctional antimicrobial coatings have been demonstrated for low-cost plant phenols, their building blocks, and trihydroxyphenyl-containing molecules.32 In the present study, we utilize the strong interfacial interactions among polycatecholamines/polyphenols and poly(vinyl alcohol) (PVA) to design mechanically robust, thermally stable, broad spectrum antimicrobial films with desirable surface properties. Fourteen different natural catecholamines/polyphenols with potent medicinal properties have been selected for this study. On the basis of the structure of the precursors, the additives are grouped into catecholamines (group I), dihydric/trihydric phenols (group II) and complex polyphenols (group III). A brief summary of various polyphenols/catecholamines used in this report and their properties is presented in Table S1. A 5% PVA solution containing various polyphenols/ catecholamines (2% w/w of PVA) was catalytically oxidized by laccase and cast into films as discussed in detail in section 2. We then investigated the changes in surface and bulk properties of the composite films. PVA was chosen owing to its applications in various food, pharmaceuticals, medicine, and biotechnology industries as well as its excellent film-forming, oxygen permeability, emulsifying, and adhesive properties.33−35 The enzyme-catalyzed oxidative polymerization of polyphenols/ catecholamine is chosen because the method has been used in a number of industrial applications.36,37 The reported approach is effective in improving surface, mechanical, and thermal properties of PVA films. Additionally, some of the reinforced films impart excellent antimicrobial properties against Gram-positive bacteria including clinically

2. MATERIALS AND METHODS 2.1. Materials. Poly(vinyl alcohol) (hydrolyzed: + 99%, Mw: 85 000−124 000), dopamine hydrochloride (DA), norepinephrine hydrochloride (NE), α-methyl norepinephrine (purity: ≥95%; AM), epinephrine hydrochloride (EP), pyrocatechol (PC), resorcinol (RS), hydroquinone (HQ), pyrogallol (PG), 1,3,5-phloroglucinol (purity: 97%; PL), gallic acid (GA), quercitin (QC), tannic acid (TA), and laccase (from Agaricus bisporus) were purchased from Sigma-Aldrich, Singapore. α-Mangostin (purity: >98%; αMG) and γ-mangostin (γMG) were obtained from Chengdu Biopurity Phytochemicals, Ltd., China. 2.2. Preparation of Catecholamines-/Polyphenols-Containing PVA Films. For film fabrication, a homogeneous aqueous solution of 5% (w/v) PVA was first prepared in distilled water at 95 °C for 2 h and cooled to room temperature. Individual catecholamine/polyphenol was separately added to PVA solution to a final concentration of 2% (w/w of PVA). The laccase enzyme was added to the above solutions (0.1% by w/w of catecholamines/polyphenols) and stirred for 24 h at room temperature. Because of the poor solubility of mangostins, their solution concentration was decreased to 1% (w/w of PVA) to maintain solution homogeneity and resultant film uniformity. The final PVA solutions with/without various additives were then poured into 90 mm Petri dishes and allowed to dry for 24 h at room temperature. The PVA films were then removed from the Petri dishes and used for further characterization. For simplicity in data presentation, the PVA films incorporating catecholamine and polyphenol after laccase oxidation were divided into three groups (group I−III), and each film type was labeled using two letter codes as described in the materials section. Group I includes PVA films containing catecholamines and were designated as DA, NE, AM, and EP; Group II PVA films include dihydric and trihydric phenols and were designated as PC, RS, HQ, PG, PL, and GA. Group III PVA films contained complex polyphenols and were represented as QC, TA, αMG, and γMG. 2.3. UV−Visible Spectrometry Analysis. UV−visible spectra of solutions and films were recorded using Shimadzu UV 3600R spectrophotometer (Shimadzu Corporation, Kyoto, Japan) in the wavelength range of 200−600 nm at 25 °C. Pristine PVA film was used as reference for films, and water was used as reference for solutions. 2.4. Fourier Transform Infrared Analysis. FT-IR spectra of all PVA films were collected using a Nicolet 6700 spectrometer (Nicolet Instrument Company, USA) over 64 scans in the region of 400−4000 cm−1 using attenuated total reflectance mode at a resolution of 1 cm−1. The baseline of air was collected before recording the FT-IR of the PVA samples, and baseline correction was done to obtain the final spectrum. 2.5. Contact Angle Measurements. Static contact angle measurements were performed using a VCA optima goniometer (AST Products Inc., MA, USA) at room temperature. Two solvents, MiliQ water (resistance ≈ 18.2 MΩ) and diiodomethane (DIM), were used to determine the wettability of various PVA films. The contact angles for 1 μL droplet of each liquid 10 s after the droplet formation on the film surface were reported. 2.6. Atomic Force Microscopy Studies. The surface topology and roughness of the films were measured via atomic force microscopy (AFM) using a tapping mode Innova AFM (Bruker, Bruker Corporation, MA, USA) equipped with a silicon cantilever having a tip of radius ∼8 nm. Three different locations (10 μm × 10 μm) on each film were scanned. 2.7. Field-Emission Scanning Electron Microscopy Studies. FE-SEM analysis was carried out to analyze the changes in the MRSA DM9808R bacterial morphology after incubating with 100 μL of ∼106 CFU/mL bacterial culture on PVA, EP, PG, and HQ films for 24 h. B


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ACS Applied Materials & Interfaces FE-SEM images were collected using a FEI-QUANTA 200F (The Netherlands) FE-SEM instrument at 5 kV. 2.8. Determination of the Mechanical Properties of Films. Mechanical properties of the films were measured using an INSTRON 5542A tensile tester (Instron Inc., MA, USA) at 25 °C and 80% relative humidity. Cast films were cut into rectangular strips (20 mm × 10 mm, thickness 0.15−0.2 mm). Tensile measurements were performed at a loading rate of 1 mm/min for about 5−7 films in each group. The toughness of the films was calculated by integrating the area under the stress−strain curve. 2.9. Thermogravimetric Analysis. Thermogravimetric analysis (TGA) for the films were conducted using a SDT 2960 thermal analyzer (TA Instruments, DE, USA) at a heating rate of 20 °C/min in a dynamic nitrogen atmosphere (flow rate = 70 mL/min) from 25 to 600 °C. Thermal parameters describing various stages of temperatureinduced weight losses were determined directly from the dynamic normalization algorithm provided with the instrument. 2.10. Assessment of Antimicrobial Properties. Antimicrobial properties of the films were evaluated by both Kirby−Bauer disc diffusion method as well as the microbroth dilution method in accordance with Clinical and Laboratory Standards Institute (CLSI).38 The details of various microbial strains used in this study are given in the Supporting Information. For radial disc diffusion assay, freshly cultured Gram-positive or -negative bacterial strains (adjusted to a concentration of 0.5 McFarland standards) were spread onto the surface of sterile Mueller Hilton (MH) agar plates using cotton swabs. For yeast strains (adjusted to a concentration of 0.5 McFarland standards), sterile Sabouraud dextrose (SD) agar (SDA) plates were used. The PVA films from each group (1 cm × 1 cm) were rinsed three times with distilled water, placed on top of the swabbed cultures, and incubated at 37 °C for 24 h. Antimicrobial activity was expressed as the zone of inhibition in millimeters, and the quantitative values were converted into color-coded heat maps using Spotfire software. The assay was performed in two independent duplicate samples. For microbroth growth inhibition assay, films weighing 50 ± 3 mg were incubated for 24 h in 2 mL of MH broth containing bacterial cultures (at 0.5 McFarland standards) or 24 h in 2 mL of SD broth for yeast cultures (at 0.5 McFarland standards). After preparing one-log (10-fold) serial dilutions of the above solutions in PBS, 100 μL of each dilution was pour-plated on MH or SD agar plates and incubated at 37 °C for 48 h. The colony-forming units (CFUs) were counted for 104− 106-fold dilutions for Gram‑positive strains or 104-fold dilution for C. albicans. The reduction factor was estimated using the following equation,39

polymer solution as described in the Materials and Methods. Various catecholamines/polyphenols (2% w/w of PVA) , with or without laccase were mixed with the PVA solution for fabricating their respective hybrid films.36,37 To avoid ambiguity, PVA films containing various catecholamines/ polyphenols are represented by a two-letter code derived from their corresponding generic names of the precursors (Table S1). A number of these compounds are already known to be the substrates of laccase-catalyzed oxidation (Table S2). The presence of laccase caused significant color change from an otherwise optically clear PVA film to pale-yellow, yellow, brown, and dark-brown, depending on the nature of catecholamine/polyphenol incorporated inside the films (Figure 1a).

Figure 1. (a) PVA films prepared by incorporating various catecholamines/polyphenols showing light-yellow to dark-brown coloration due to the laccase-catalyzed polymerization of the additives. (b−d) UV−visible spectra of PVA films incorporating (b) group I (catecholamines) compounds, (c) group II (simple polyphenols) compounds, and (d) group III (complex polyphenols) compounds.

Laccase-supplemented PVA films doped with group I compounds showed marked brown coloration (Figure 1a), indicative of the formation of oligo-/polycatecholamines, without significant loss of transparency (except DA). It has been shown that oxidation of catecholamines by laccase generates corresponding catecholamine o-quinone intermediate that then undergoes intramolecular Michael addition reaction to produce leucocatecholaminechrome (LCAC).40−42 Subsequent oxidation of LCAC produces a more reactive catecholaminechrome, which then polymerizes to form oligo-/polycatecholamines rich in 5,6-hydroxy indole/indoline structures (Table S2). UV−visible spectra of catecholamineloaded PVA films after laccase oxidation are shown in Figure 1b. Compared to laccase-deprived catecholamines-loaded films (Figure S1), a significant enhancement in the intensity of absorption band around 280 nm was observed for all the laccase-containing catecholamine films (except AM), indicating the formation of enzyme-mediated oxidative products with higher degree of conjugation (Figure S2a−d). Appearance of new peak/shoulder around 362, 390, and 327 nm for DA, NE, and EP films, respectively, further corroborates the enzymatic oxidation of catecholamines (Figure 1b). The absorption band around 350 nm corresponds to the formation of quinone groups via laccase-catalyzed oxidation of the catecholamines.

R f = log10 Nc − log10 Nd where Nc is the number of viable cells (CFU) in the inoculum in the presence of control or PVA film and Nd is the number of viable cells (CFU) in the inoculum containing modified PVA films. The antimicrobial activities of catecholamines-/polyphenols-reinforced films were expressed as percent growth inhibition relative to that of pristine PVA. Appropriate amounts of catecholamines or polyphenols dissolved in culture media were used as controls to determine if the oxidized forms retain/lose the antimicrobial activities. For assessing the durability, the PG, HQ, and EP films were autoclaved at 120 °C for 30 min and dried overnight. The antimicrobial activity of the dried films was investigated against the panel of pathogens as before. To study the bacterial growth kinetics with changing composition of the PG, 50 mg of the film was incubated with MRSA DM9808R strain in 2 mL of MH broth containing initial bacterial inoculum of ∼106 CFU/mL at 37 °C, and the optical density at 600 nm (OD600) was measured using an Infinite M200 microplate reader (Tecan group Ltd., Switzerland) at various time points. The growth curve was plotted as OD600 (nm) versus time (h).

3. RESULTS AND DISCUSSION 3.1. Structural Characterization of PVA Films by UV− Visible and FT-IR Spectrometry. PVA films were prepared by casting evaporation method from 5% (w/v) aqueous C


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Figure 2. FT-IR spectra of PVA films incorporating (a) group I (catecholamines) compounds, (b) group II (simple polyphenols) compounds, and (c) group III (complex polyphenols) compounds.

no change in the absorption maxima/peak broadening was observed, indicating the absence of any laccase-catalyzed oxidative polymerization. It is likely that the strong interaction between the two dihydric phenols and PVA may prevent their oxidative polymerization or the two may not be the substrate for laccase-catalyzed oxidation under the conditions employed in our experiment. The absence of any laccase-induced changes in the UV spectra of resorcinol/hydroquinone solution (Figure S3b,c) confirmed the fact that the two dihydric phenols were not substrates for laccase oxidation under the conditions used. However, for catechol solution, a strong absorption maxima was observed around 408 nm during early time intervals and disappeared after 6 h, confirming the formation of polypyrocatechols and augmenting previous results (Figure S3a).45 For the three trihydric phenols incorporated films, i.e., PG, PL, and GA, a significant change in the coloration as well as the absorption spectrum were observed for two gallol compounds (pyrogallol and gallic acid) after laccase oxidation (Figure 1c). In the case of PG films, the intense monomeric peak around 264 nm (Figure S2h) due to the aromatic chromophore disappeared, and a new peak at 324 nm appeared with a broad band covering the entire visible range, suggesting the formation of oxidation products.46,47 The presence of intense absorption maximum at 330 nm in the PG films that did not contain laccase (Figure S2h) suggest the plausible laccase-independent pH-induced aerial oxidation. Similarly, in GA films, a decrease in the intensity of monomeric peak at 270 nm with concomitant appearance of a broad hump extending from 370 nm (Figure 1c), due to low energy π−π* transitions for the polymer, is augmented with C−C coupling and formation of polygallic acid.48−50 GA films without laccase did not show appreciable aerial oxidation, suggesting greater stability conferred by the carboxylic acid group (Figure S2j). In the case of PL films with or without laccase, a broad band centered at 350 nm was observed (Figure S2i), indicating the existence of phloroglucin (tautomeric form of phloroglucinol), with the slight redshift of the monomeric

Furthermore, the broad absorption band around 480 nm was due to the characteristic electronic transitions related to the chrome formation by the intramolecular cyclization of the oxidized quinone structures. These results demonstrated that laccase-catalyzed catecholamine polymerization also followed the formation of o-quinone intermediate mechanistic pathway as reported for pH- and oxidant-induced polymerization.40−44 The existing acidic conditions (5.6 ± 0.3) during the preparation of catecholamine-loaded PVA films ruled out the possibility of pH-induced oxidative polymerization of catecholamine, which requires alkaline conditions (pH ∼8.5). The intense brown coloration of DA films was consistent with more rapid kinetics of polymerization and precipitation of polydopamine (pDA) compared to other catecholamines.43 The lack of any coloration (except AM) and absence of characteristic absorption bands around 350 and 480 nm in the UV−visible spectrum of laccase-free catecholamines-incorporated PVA films further supported the enzyme-catalyzed oxidative polymerization of catecholamines in DA, EP, and NE films. However, for AM films, subtle changes in the coloration and UV absorption spectra were observed compared to those of films without laccase, indicating possible laccase-independent aerial oxidative polymerization (Figure S2c). Among group II PVA films containing three dihydric phenols, PC films displayed intense coloration after laccase oxidation, HQ films showed pale-brown coloration, and the RS films did not show any color change (Figure 1a). Comparing the UV spectra of the PVA films incorporated with the three dihydric phenols with or without laccase provides qualitative information on the structural changes upon laccase oxidation (Figure S2e−g). For PC films, a significant peak broadening above 300 nm without any shift in peak maxima at 280 nm was observed, confirming the formation of polypyrocatechol (Figure 1c). The absence of any intermediate peak around 408 nm (due to the formation of o-benzoquinone) suggests complete enzymatic conversion of PC to polypyrocatechols.45 For PVA films containing HQ and RS with or without laccase, D


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Figure 3. Photographs showing the variation in (a) water contact angles and (b) DIM contact angle for PVA films upon incorporation of polycatecholamines/polyphenols. (c) AFM images showing the effect of incorporating different catecholamines/polyphenols on the surface topology of modified PVA films with estimated root-mean-square surface roughness (Rq) values using a 10 μm × 10 μm area.

In nonaqueous solvents, α- and γ-mangostins films showed monomeric absorption peaks at 260, 324, and 370 nm for C C chromophore π to π* transitions and CO chromophore n to π* transitions.53 However, because of their low solubility in water/aqueous buffers, these compounds display poorly resolved spectral characteristics below neutral pH (Figure S2m,n). However, the polyphenolic natural products can be readily solubilized by dispersants such as polyvinylpyrolidone (Figure S2o).53 The spectrum of γMG film was reminiscent of monomeric form of γ-mangostin in aqueous solution containing dispersant, confirming that PVA had a similar stabilizing effect in decreasing the aggregation of mangostins (Figure 1d). The absence of any color change in the films with or without laccase further confirms that neither of the mangostins were substrates for laccase. FT-IR was used to identify the changes in the fine structures of PVA films caused by incorporating polycatecholamines/ polyphenols. The detailed band assignments over the characteristic vibrational changes in the modified PVA films are shown in Table S3 with discussion. Pristine PVA films revealed characteristic bands at 1459, 1563, and 1713 cm−1, which

peak of phloroglucinol. The absence of any color change further confirms the lack of formation of polyconjugated chromophores. UV studies in solution for the three trihydric phenols indicated substantial changes in the spectra of pyrogallol and gallic acid after laccase oxidation but the absence of any significant changes for phloroglucinol, indicating that the latter is not a substrate for laccase oxidation (Figure S 3d−f). Among group III films, QC films showed a blueshift in peaks around 390 nm when compared to films that did not contain laccase and revealed generation of a weak absorption band centered at 336 nm related to o-quinone formation from the catechol ring present in quercetin structure (Figures 1d and S2k). The appearance of less-intense yellow coloration in the laccase-oxidized film confirmed the formation of polyquercetin with less planar structure.51 TA films with or without laccase displayed absorption spectra with a broad band spanning the visible region (Figure S2l), indicating laccase-independent/pHinduced oxidation of TA. A weak amber coloration in TA films indicated the formation of the oxidative products.52 E


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ACS Applied Materials & Interfaces were assigned to δ(CH2−C) bending, δ(C−O−H) bending, and ν(CO) stretching, respectively, for residual acetate groups available on the PVA chains. The presence of a broad band spanning the region from 1042 to 1157 cm−1 was also assigned to ν(C−O) stretching of the alcoholic groups and acetate groups, which constituted the PVA side chains.54 DA and NE films showed intense bands at 1428 and 1443 cm−1, respectively, due to the presence of νring(CC) stretching vibrations for pDA and polynorepinephrine (pNE; Figure 2a). The formation of pDA and pNE was further confirmed by the appearance of additional characteristic bands around 1379, 1659, 1522, and 1331 cm−1 for DA films and 1378, 1655, 1510, and 1333 cm−1 for NE film, respectively, which were designated δ(CH3) bending, δ(N−H) bending, νring(CN) stretching, and νindole ring(CNC) stretching vibrations, respectively. The presence of the indole signatures in the IR spectra further confirmed the formation of pDA and pNE, structurally constituting of 5,6-dihydroxyindole units.55 Interestingly, the presence of weak bands at 1092 and 1239 cm−1 for DA and 1090 and 1239 cm−1 for NE films were indicative of νsymmetric(C−O−C) stretch and νAsymmetric(C−O−C) stretch, respectively, for aryl alkyl ethers, suggesting possible covalent ether bonding between PVA and pDA/pNE. Probably, the existent weak acidic conditions in the preparation of DA and NE films could provide a favorable environment for the ether bond formation between PVA and pDA/pNE. In AM and EP films, polymerization features such as νring(CN) stretching and νindole ring(CNC) stretching were relatively weaker than those of DA/NE films. However, the presence of sharp peaks at 1662 cm−1 for N−H bending and the mild hump around 1339 cm−1 for νindole ring(CNC) stretching indicated the existence of α-methyl norepinephrine and epinephrine in their oligomeric or short polymeric chains as corroborated by UV studies. Among group II compounds, RS and PL films showed no apparent differences in their FT-IR spectrum compared to those of the native PVA films except for the presence of additional peaks at 1603 and 1666 cm−1 for RS and shoulders around 1610 and 1680 cm−1 for PL due to νin‑ring(CC) aromatic stretching vibrations and νquinone(CO) stretching vibrations, respectively (Table S3). Similar to DA and NE films, PC and HQ films also revealed clear vibrational signatures of covalent bonding (Figure 2b). In the case of PG and GA, only subtle changes in the IR spectra were observed, although UV studies indicated significant broadening and apparent color change in the cast films (Figure 2b). Among group III compounds, only αMG and TA films showed the presence of ether linkage; however, the spectrum of QC and γMG did not show substantial changes compared to pristine PVA films (Figure 2c). Altogether, UV and FT-IR results confirmed that DA, NE, AM, EP, PC, PG, PL, GA, QC, and TA films contain the oxidative products of the respective precursors, whereas in HQ, RS, αMG, and γMG films, the phenolic compounds remained in their monomeric forms. The study also revealed that polycatecholamines/polyphenols interacted with PVA matrix both by covalent and noncovalent interactions. Therefore, we investigated the consequences of such interactions by assessing the surface and bulk properties of the films. 3.2. Surface Characterization of the Films by Contact Angle Measurements and Atomic Force Microscopy. We used static contact angle measurements to investigate the change in wettability of PVA films upon incorporating polycatecholamines/polyphenols. Contact angle measurements were performed using two liquids, i.e., water and diiodo-

methane. Because water is a solvent for PVA, we determined the contact angles of the films using a nonsolvent, diiodomethane (DIM).56 Figure 3a,b reveals the water contact angle measurements in water (θwater) and in diiodomethane (θDIM), respectively, for the PVA films containing various polycatecholamines/polyphenols after induction of laccase-catalyzed oxidative polymerization. Pristine PVA films showed a θwater of 82 ± 5°, which was similar to the value reported during the early stages on PVA membrane.56 Oxidative polymerization of group I compounds within the PVA films decreased θwater significantly (p < 0.01 for DA and p ≤ 0.05 for NE, EP, and AM films); DA displayed the maximum decrease in θwater, suggesting pDA reinforcement increased the wettability of PVA films remarkably. These results suggest possible burial of the hydrophobic heteroaromatic ring and exposure of −OH groups upon formation of polycatecholamine aggregates.41 In support of these observations, θDIM values were unaltered for all the polycatecholamines (except AM)-reinforced films (Figure 3b). For the group II polyphenols, no significant changes in θwater (p > 0.05) were observed among the dihydric phenols reinforced PVA films, despite marked structural changes in PC films. Among the trihydric phenols, the PG and GA films displayed higher water wettability (p < 0.05). However, reinforcement of group III polyphenols in PVA either maintained or enhanced the water contact angle, indicating maintained or increased hydrophobic nature of PVA. This is consistent with the previous studies that showed that complex polyphenols did not alter the surface chemistry of substrates significantly.57 To infer the changes in surface topography, we imaged the films and quantified their surface roughness (defined as the standard deviation of the height value in the image) by AFM (Figure 3c). Pristine PVA film displayed a smooth appearance with a root-mean-square surface roughness (Rq) of 4.2 nm. However, the surface topography of PVA films changed substantially, depending on the nature of catecholamines/ polyphenols incorporated. For all group I compounds, surface roughness increased significantly upon polycatecholamine formation except for the AM films. Among the three dihydric phenols reinforced films, PC showed the maximum (6.3 nm) increase in Rq when compared to that of pristine PVA films, whereas HQ and RS films decreased the surface roughness, suggesting that polymerization of catechol may have contributed to the increased surface roughness of the PVA films (Figure 3c). However, no clear trend was observed among the three trihydric phenols because PG and GA films displayed marginal changes in Rq whereas PL films showed more than twofold increases in Rq values compared to those of PVA films. Of all the complex polyphenolic compounds, TA films showed the maximum surface roughness of 19.6 nm. On the basis of these analyses, we attempted to correlate the changes in surface properties of polyphenols-/polycatecholamines-reinforced films with the structure of the precursors. Because the presence of aromatic vicinal diols is common among catechol and catecholamines and all the catecholamines differ by an additional −OH (norepinephrine) and C- or Nmethyl groups (epinephrine or α-methyl norepinephrine), we compared the changes in surface roughness of catecholamine films with those of PC-reinforced films (Figure S4a). When compared to pristine PVA, PC films displayed no substantial changes in Rq values. The values increased significantly upon formation of polycatecholamines that contained aminoethyl group (as in dopamine) or 2-hydroxy aminoethyl group (as in F


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Table 1. Mechanical Properties of Various Catecholamines-/Polyphenols-Loaded PVA Films after Laccase Oxidation for 24 ha sample

group I

group II

group III a

PVA DA NE AM EP PC RS HQ PG PL GA QC TA αMG γMG

σy (MPa) 11.1 24.0 17.8 16.3 17.5 20.7 26.1 24.1 10.9 22.5 21.8 18.1 18.1 17.6 17.1

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.9 6.0**** 3.7* 0.5 0.1* 2.0**** 2.7**** 2.4**** 1.7 1.0**** 1.4**** 2.8**** 2.0**** 1.0**** 1.6****

24 66.3 34.4 40.8 30.1 45.3 44.8 41.9 38.4 46.9 58.6 28.6 27.4 37.9 26.1

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

2.7 19.0**** 2.9 4.4 2.1 12.9** 2.0** 3.9* 9.6 5.4** 13.1**** 3.5 1.0 6.4** 4.3

Jlc (MJ m−3)

εb (%)

E′ (MPa) 303.2 263.8 378.0 254.9 421.3 375.4 473.2 476.7 257.6 376.5 166.8 549.5 693.4 481.8 587.2

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

36.8 11.2 43.7** 14.5 36.7**** 11.3 23.7**** 68.3**** 58.5 12.8 7.5 20.2**** 45.2**** 16.8*** 93.9****

22.0 ± 3.0 45.8 ± 11.8** 49.7 ± 13.3*** 29.4 ± 2.1 49.34*** 53.0 ± 6.4**** 84.6 ± 9.0**** 76.6 ± 11.9**** 21.3 ± 5.5 62.0 ± 4.3**** 42.3 ± 3.1** 68.0 ± 10.4**** 87.3 ± 10.8**** 62.8 ± 7.4**** 71.8 ± 11.7****

Significance values: *, p ≤ 0.05; **, p < 0.01; ***, p < 0.001; and ****, p < 0.0001, as compared to PVA by t test or 1-way ANOVA.

norepinephrine) in the precursor. These results suggest that the polyheteroaromatic catechols formed by oxidative polymerization of dopamine or norepinephrine increased the surface roughness. However, the increase in Rq can be decreased by the presence of epinephrine or α-methylnorepinephrine, which had an extra N-methyl or α-methyl groups, respectively. Similarly, we compared the surface properties of PVA and PC films with PG, GA, and TA films due to structural similarity of the precursors and their ability to undergo laccase-catalyzed oxidative polymerization (Figure S4b). TA was also included as it contained 5 catechol and 5 pyrogallol units. The results suggested that as the number phenolic −OH groups increased in the precursor, surface roughness remained unchanged until a particular number of phenolic groups in the precursor beyond which a large increase was observed. These observations were supported by comparing the surface properties of polyphenols that remained monomeric wherein a large increase in surface roughness was observed with increase in number of phenolic −OH groups (Figure S4c). It is likely that the oxidative products formed by tannic acid may form insoluble aggregates thus generating a heterogeneous surface. In support of these observations, SEM image of the TA films displayed the presence of nm-sized particulates on the film surface (Figure S4d). On the basis of these observations, for polyphenolsreinforced films we conclude that the number of phenolic −OH groups in the precursor must exceed ≥5 to achieve high surface roughness. Similarly, the decrease in θwater caused by oxidative polymerization of dopamine/norepinephrine can be increased by N- or C-methylation of catecholamine precursors. 3.3. Tensile Properties of Catecholamines-/Polyphenols-Reinforced PVA Films. Furthermore, we examined the effect of reinforcing catecholamines/polyphenols on bulk mechanical properties of PVA films in terms of four fundamental tensile properties (Table 1): yield stress (σy), Young’s modulus (E′), failure strain (εb), and toughness (Jlc). Of the four catecholamines-reinforced PVA films, statistically significant improvement in σy, E′, and Jlc was observed for DA films (Figure 4a). NE and EP films showed significant improvement in σy, εb, and Jlc values, whereas AM-modified films did not show significant alteration of the mechanical properties (p > 0.05). HQ and RS films displayed marked increase in all four mechanical properties, whereas PC, PL, and GA films showed significant improvement of σy, E′, and Jlc

Figure 4. Representative stress−strain curves showing the influence of (a) group I, (b) group II, and (c) group III compounds on the mechanical properties of PVA films. (d) Correlation between numbers of phenolic −OH groups in the precursors versus the mechanical properties of the PVA film. Elongation at break (εb) values are plotted in log10 scale for better clarity.

without affecting the εb values when compared to those of pristine PVA (Figure 4b). However, PG films did not display any significant changes in the mechanical properties compared to those of pristine PVA films. In the case of group III complex phenols containing PVA films, αMG alone increased all four mechanical properties, whereas other polyphenols had significantly altered values of σy (p < 0.0001), εb (p < 0.0001), and Jlc (p < 0.0001) but unaltered E′ values (Figure 4c). To obtain a better structure−properties relationship, we combined the σy and E′ values and εb and Jlc values and compared the effect of polycatecholamines-reinforced films with those of PC films as before. For DA films, a significant increase in E′ (p < 0.05) and substantial decrease in εb (p < 0.0001) was observed, suggesting that polyheteroaromatic catechol structure increased the stiffness but decreased the elasticity (Figure S5a,b). These changes in mechanical properties could be reversed by reinforcing with polynorepinephrine or polyepinephrine because no apparent changes G


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Figure 5. TG and DTG curves for the PVA films incorporated with (a and b) group I, (c and d) group II, and (e and f) group III compounds.

reached a plateau after ≥3 phenolic −OH groups in the precursor compounds (Figure 4d). 3.4. Thermogravimetric Analysis. To investigate the effect of incorporating polycatecholamines/polyphenols on the thermal stability of the PVA films, TGA was carried out. The representative thermogravimetric (TG) and differential thermogravimetric (DTG) curves for PVA films incorporating group I−III compounds is shown in Figure 5. Pristine PVA film displayed three well-defined temperature-induced weight-loss regions associated with three different physicochemical changes, consistent with published literature values (Figure 5).58 For PVA, an initial weight loss (∼4−10%) occurred between 30 and 140 °C because of the removal of weakly bound water or dehydration from the films. The second weight loss (∼65%) occurred between 200 and 400 °C and is due to thermal degradation of the PVA structure by chain scissoring and formation of volatile products. The final stage occurs above 400 °C and is due to the decomposition of the products formed in the second stage. For a better understanding, we reported the thermal properties in terms of four parameters that describe the various stages of temperature-induced weight losses, viz., onset temperature of degradation (Ti, temperature in which second stage weight loss begins), peak temperatures for the second stage (1Tmax) and third stage (2Tmax) of degradation, and half decomposition temperature (T1/2). Table 2 summarizes the changes in thermal parameters for pristine PVA films and films reinforced with various polycatecholamines/polyphenols. For group I compounds, a marked increase in Ti, T1/2, and 1 Tmax values and decrease in the rate of degradation (indicated by the broadening of DTG curve) were observed (Figure 5a,b).

in mechanical properties were observed for NE and EP films when compared to PC films, suggesting that the presence of an −OH group (as in norepinephrine) and an N-methyl group (as in epinephrine) increased the elastic properties with concomitant decrease in stiffness. As a result, no substantial change in overall toughness (p > 0.05) was observed. However, the presence of a C-methyl group (as in α-methylnorepinephrine) decreased elasticity considerably (p < 0.0001) when compared to that of PC, resulting in a substantial decrease (p < 0.001) in toughness of AM films. These results imply that the presence of substituents in the polyheteraromatic catecholamines (formed by the oxidative polymerization) could modulate the mechanical properties of PVA, depending on the nature of substituents in the precursor catecholamines. Among the polyphenols that were substrates of laccase oxidation, a significant decrease in tensile strength (p < 0.0001) without notable changes in elastic modulus (p > 0.05) was observed for PG films. The presence of a carboxylic acid group (gallic acid) reversed the loss of mechanical stiffness and strength as indicated by an increase (p < 0.05) in E′ with GA (Figure S5c,d). However, as the structural complexity of precursors increased, a decrease in stiffness (p < 0.05) and profound increase in elasticity (p < 0.0001) was observed, as shown by QC and TA films, which resulted in increased toughness values. Consistent with these results, significant increase in elasticity and a concomitant decrease in tensile stiffness was observed for complex polyphenols that were not substrates for laccase oxidation (Figure S5e,f). Thus, the elasticity of PVA films increased with the number of phenolic groups, whereas the tensile strength, after an initial increase, H


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AM films indicates weaker interaction of the PVA with the oxidative products of α-methylnorepinephrine. Among group II compounds, PL and RS did not show any improvement in thermal stability, as indicated by the absence of substantial changes in 1Tmax and T1/2 values and rate of degradation (Figure 5c,d). However, PC, HQ, PG, and GA films displayed enhanced 1Tmax and T1/2 values, with PC conferring maximum thermal stability among group II cross-linkers. The TGA/DTG profile of PC-reinforced PVA films displayed a merger of second and third transitions, similar to that of catecholamines, indicating greater improvement in thermal stability upon polypyrocatechol formation. It is interesting to note that pyrogallol showed a marked improvement in thermal stability of PVA films but did not show alteration of the mechanical properties. Among group III compounds, QC- and TAcontaining films showed prominent effect on the thermal properties with improvement in the 1Tmax and T1/2 values (Figure 5e,f). Broadening and merger of the second and third thermal transitions were also observed in the DTG curve of QC and TA films. Among the two natural products, αMG lowered the thermal stability, whereas γMG showed a higher T1/2 values. To understand the structure−function relationship, we calculated the ΔT1/2 (= T1/2add − T1/2PVA), which is defined as the difference in half decomposition temperature of PVA film with polycatecholamine/polyphenols (T1/2add) and pristine PVA (T1/2PVA). The analysis indicated that the heteroaromatic catechol structures formed by norepinephrine conferred the maximum shift among all catecholamines. Presence of a Cmethyl group decreased the ΔT1/2 values, indicating that increasing the complexity of catecholamine precursors moder-

Table 2. Thermal Analysis of PVA Films Reinforced with Various Catecholamines/Polyphenols

group I

group II

group III

sample

Ti (°C)

PVA DA NE AM EP PC RS HQ PG PL GA QC TA αMG γMG

215.4 225 227 222.5 221.6 221.6 224.2 196.5 200.0 216.3 200.7 217.2 220.7 226.0 207.6

1

Tmax (°C)

T1/2 (°C)

289.8 365.6 385 304.8 370.7 352.2 304.1 362 315.6 295.6 301.0 338.6 342.2 296.8 290.7

303.6 362.3 379.1 325.4 366.2 355.2 312.0 350.3 337.3 299.5 328.4 347.4 346.3 302.0 322.4

2

Tmax (°C) 450.8 456.6 449.2 439.1 462.0 460.1 442.6 459.9 450.8 432.4 450.6 437.3 447.0 434.4 442.1

Among the catecholamines, NE films showed a maximum increase, whereas AM film displayed the lowest increase in thermal stability. The effect of group I compounds can further be supported by the fact that the second (i.e., the decomposition of side chains) and third transitions (decomposition of backbone) overlap in the TGA/DTG curves. The merger of second and third stage transition in the TGA/DTG indicates strong covalent or noncovalent interactions between the −OH groups of PVA with −OH moieties of polycatecholamines.59−63 The lowest increase in the thermal properties for

Figure 6. (a) Heat map showing the zone of inhibition of PG, HQ, and EP films against 20 differents Gram-positive bacterial strains. (b−c) Photographs of the disc diffusion assay showing antimicrobial activity of (b) PG films and (c) HQ films before and after steam sterilization at 120 °C for 30 min. I


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Table 3. Reduction Factor (Rf) for S. aureus and C. albicans after 24 h Exposure to Polycatecholamine-/Polyphenol-Reinforced Filmsa Reduction factor (Rf)

a

strains

PG

HQ

EP

pyrogallol

hydroquinone

epinephrine

S. aureus 29213 S. aureus DM 4808R MRSA 700699 MRSA DM9808R P. aeroginosa 18531 C. albicans 2672R C. albicans 2091

2.3 (99.5%) 2.3 (99.5%) 3.2 (99.94%) 1.9 (98.7%) 3.7 (99.98%) 3.2 (99.9%) 2.14 (99.27%)

1.9 (98.7%) 2.3 (99.5%) 1.1 (92.6%) 1.7 (98%) ND ND ND

0.82 (84.9%) 3.0 (99.9%) 2.1 (99.1%) 0.7 (80.0%) 1.7 (97.9%) ND ND

1.6 (97.6%) 1.3 (95.0%) 2.4 (99.6%) 1.8 (98.2%) 5.44 (99.999%) −0.4 0.04 (8.5%)

1.3 (95.5%) 1.0 (90.0%) 2.6 (99.3%) −0.3 ND ND ND

−0.02 −3.3 0.4 (62%) 0.1 (7%) ND ND ND

The percent microbial lethality is shown in parentheses.

at 120 °C for 30 min.65 PG films retained antimicrobial activity against S. aureus, S. saphrophyticus, P. aeruginosa, and C. albicans strains, whereas a loss of activity was observed against S. epidermis and MRSA strains (Figure 6b). HQ films, however, retained antimicrobial activity against all the Gram-positive bacterial strains tested (Figure 6c). In contrast, a complete loss of activity was observed for the EP films after autoclaving, suggesting that the film could not withstand the harsh conditions used during industrial processes (data not shown). The ability of PG, HQ, and EP films to decrease the bacterial bioburden was further verified by microbroth growth inhibition assay under proliferating conditions. Briefly, the films were immersed in bacterial medium (∼0.5−1 × 105−1 × 108 CFU/ mL of bacterial suspension) for 24 h at 37 °C, and the amount of viable bacteria that remained in the culture medium were enumerated by colony counting. For a comparison, we also determined the inhibitory effect of pyrogallol, hydroquinone, or epinephrine dissolved in broth solution to determine if the inhibitory activity was maintained or abrogated after laccase oxidation. It should be noted that the amount of inoculum taken for these experiments far exceeds the microbial bioburden observed in clinical environments.66,67 In addition, more stringent conditions were employed (full-strength bacterial culture medium) to determine if the films retard/promote the growth of bacteria in the surroundings. The viability of bacteria/yeasts after 24 h exposure of the films was quantified by estimating the reduction factor (Rf). A positive Rf value indicates bactericidal efficacy, and a negative value implies poor bactericidal properties. Table 3 shows the results obtained from cell suspension experiments for PG, HQ, and EP films and their precursors. For a better understanding, the amount of bacteria/yeasts killed (expressed as percent lethality) upon exposure is shown in parentheses. Against both bacterial and yeasts strains, PG films displayed the highest Rf values when compared to those of HQ and EP films as well as those of their precursors, confirming that the oxidative product of pyrogallol conferred the broad spectrum microbicidal properties. Except for one MRSA strain, >99% killing efficacy was achieved in the presence of PG films. HQ films displayed excellent bactericidal activity against two S. aureus strains (>98%), and the results were comparable to those of hydroquinone in solution, indicating that the antimicrobial properties of hydroquinone were retained in the PVA films. In contrast, EP films displayed complete reversal of antimicrobial activity compared to that of epinephrine in solution, suggesting enhanced antimicrobial activity of oxidative products of epinephrine against S. aureus and MRSA strains. In addition, PG and EP films displayed a substantial decrease in the viability of P. aeruginosa 18531 strains, with PG films showing

ately altered the thermal stability (Figure S6a). However, polyphenols which were substrates of laccase oxidation, after an initial decrease in ΔT1/2 for trihydric phenols, reached values that were closer to that of PC film for QC and TA films (Figure S6b). In contrast, no clear relationship was observed among films that remained in their monomeric forms (Figure S6c). Among the simple and complex polyphenols, HQ and PC films displayed maximum thermal stability. 3.5. Antimicrobial Properties of Polymer Films. Because a number of polyphenols displayed antimicrobial activity against microbial pathogens, we evaluated the antimicrobial efficacy of laccase-oxidized PVA films against a panel of Gram-positive (20 strains) bacteria and Gram-negative bacteria (13 strains) and yeasts (5 strains) by Kirby−Bauer disc diffusion assay (Supporting Information). The selection of various pathogenic bacteria and yeasts is based on the fact that these microorganisms are frequently found in the infected tissue surfaces and are major etiological agents in deviceassociated infections.64 Colonies of microorganisms grew all over the plates containing PVA films (data not shown). However, PG films displayed a broad spectrum of inhibition compared to other polyphenol cross-linked films. A clear zone of inhibition was observed against 18 Gram-positive strains, whereas two strains (S. aureus DM4299, a clinical isolate of eye, and a reference strain of E. faceium) displayed resistance to the PG films (Figures 6a and S7). Significant inhibitory activity was observed against pathogenic S. epidermis, S. saphrophyticus, and both reference and clinical isolates of S. aureus and MRSA. When tested against Gram-negative bacteria, PG films displayed inhibitory activity against five P. aeruginosa strains, which includes drug-resistant strains, whereas no activity was observed against E. coli and K. pneumoniae (Figure S8a). Of the five yeasts strains tested, two strains of C. albicans were inhibited by PG films (Figure S8b). These results confirm the potent broad spectrum activity of the oxidative products of PG against pathogenic bacteria and yeasts. HQ film was effective against 19 Gram-positive strains tested (Figures 6a and S9), and no activity was observed against Gram-negative and yeasts strains. Interestingly, EP films showed a clear zone of inhibition against 14 Gram-positive strains, and inhibitory activity was observed against two strains of P. aeruginosa (Gram-negative; Figure S10). However, EP did not inhibit the growth of pathogenic C. albicans. In addition, in all those films that displayed clear zone of inhibition no growth was observed on either side of the films confirming that the films could also effectively prevent the adhesion of microorganisms. To assess the durability of the films, we tested the antimicrobial activity of PG, HQ, and EP films after autoclaving J


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altered the surface and bulk properties of the PVA films depending on the structure of the precursor compounds. The presence of other functional groups in catecholamine precursors decreased the surface roughness of the films, whereas increasing the number of phenolic groups enhanced the surface roughness. For the polyphenols-loaded films, a significant correlation was observed between enhancement in elastic properties and the number of phenolic groups in the precursors. The oxidative products of pyrogallol and epinephrine displayed potent antimicrobial properties, inhibiting the growth of pathogenic bacteria and yeasts. Among the various catecholamines-reinforced films, EP displayed optimum surface, mechanical, thermal, and antimicrobial properties, providing excellent potential in multitude of applications. We have demonstrated a simple method for the preparation of multifunctional PVA films by reinforcing with catecholamines/ polyphenols, and the process can be tuned to fulfill the end-use requirements. In addition, the method offers a simple and versatile platform that can be readily coupled/integrated with industrial processes. The identication of oxidative products of pyrogallol and epinephrine with inherent antimicrobial properties would further expand the diverse applications of musselinspired surface coatings.73 We speculate that the method can be extended to produce films/hydrogels with outstanding mechanical, thermal, and antimicrobial properties with advanced health care applications, including medical device coatings, antimicrobial contact lens wares, food packaging, clean-room surfaces, and antimicrobial window protective films, among others.

antimicrobial activity superior to that of EP films. These results indicate that in situ laccase oxidation of pyrogallol and epinephrine generates oxidative products that display pronounced antimicrobial activity compared to their precursors. To determine the concentration-dependent changes, we decreased the amount of pyrogallol in the films by about fourfold ([email protected]; PG with 0.5% of pyrogallol incorporated) and followed growth kinetics of MRSA DM9808R strains by monitoring the OD600 values. The results indicated a substantial decrease in the growth and increase in lag time for [email protected] when compared to those of pristine PVA films (Figure S11a). However, no bacterial growth was detected in the presence of PG films (containing 2% pyragallol), thus confirming the concentration-dependent bactericidal activity. To obtain further insight, we determined the morphology of bacterial cells after expsosure of the cells to various PVA films using FE-SEM. Bacterial cells grown on pristine PVA films displayed smooth morphology devoid of any deformation and release of intracellular components (Figue S11b). However, films exposed to PG, HQ, and EP displayed considerable deformation and truncated morphologies with ill-defined surfaces (Figure S11c− e). These results probably suggest membrane lytic action of the modified films upon contact with the bacterial cells.68,69 It should be noted that α- and γ-mangostins which possess potent antimicrobial activity against Gram-positive pathogens68 lacked inhibitory activity because we could not observe a clear zone of inhibition in films containing these natural products. This can probably be attributed to the tight complex formed between mangostins and PVA that would have compromised their antimicrobial properties. In support of this, we determined the MIC of α- and γ-mangostins in the presence of PVA against four Gram-positive strains, and a complete loss of activity was observed as the values shifted from 1 to >50 μg/mL (Table S4). On the basis of these findings, we propose that a strong interaction between mangostins and PVA abrogates its antimicrobial properties, though it increased the mechanical properties of the films. 3.6. Possible Healthcare Applications of Antimicrobial PVA Films. Owing to their optical transparency and high mechanical strength, these antimicrobial PVA films could play a promising role in designing antimicrobial ophthalmic devices such as contact lens wares, ocular inserts, ocular bandages, and intraocular lenses.70 This catecholamine-/polyphenol-based coating strategy can be extended to wide variety of substrates and may prove their potential as antimicrobial coatings for various consumers and medical products. Because of their high thermal stability, these films can be used to develop active foodpackaging materials to extend the shelf life of the food products by preventing their microbial contamination and biofouling. Because of the broad spectrum antimicrobial properties, these films can be used to design self-defensive surfaces, having applications in constructing clean-room and anti-infective window protective surfaces. The strategy can be further advanced to prepare PVA-based antimicrobial hydrogels that could be used extensively in a wide variety of medicinal applications including contact lenses, wound dressings, antiadhesive tissue barriers, and artificial cartilages.56,71,72

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.5b09633. List of microbial strains used for antimicrobial assessment of the films, discussion over the FT-IR band assignments, additional tables (molecules used in this study, chemical reactions of investigated molecules as substrates, FT-IR features, minimum inhibitory concentrations), and additional figures (modified PVA films, UV−vis spectra, surface roughness data, correlation of structure and mechanical properties, half decomposition temperature data, disc diffusion assay data, and data on variation in bacterial growth and morphology). (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This research is supported by the Singapore National Research Foundation under its Translational and Clinical Research Flagship Programme (NMRC/TCR/008-SERI/2013) and administered by the Singapore Ministry of Health’s National Medical Research Council. R.L. acknowledges the funding support from National Medical Research Council’s Cooperative Basic Research Grant (NMRC/CBRG/0048/2013), and C.D. thanks the funding support from SingHealth Foundation Research Grant (SHF/FG637S/2014). N.K.V. acknowledges funding support from the NTU Lee Kong

4. CONCLUSIONS In this work, we attempted to discern the differences in surface and bulk properties of PVA films brought about by reinforcing the oxidative products of polyphenols and polycatecholamines. The oxidative products of polyphenols or catecholamines K


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(18) Rochford, E. T.; Richards, R. G.; Moriarty, T. F. Influence of Material on the Development of Device-Associated Infections. Clin. Microbiol. Infect. 2012, 18, 1162−1167. (19) Noimark, S.; Dunnill, C. W.; Wilson, M.; Parkin, I. P. The Role of Surfaces in Catheter-Associated Infections. Chem. Soc. Rev. 2009, 38, 3435−3448. (20) Lee, H.; Dellatore, S. M.; Miller, W. M.; Messersmith, P. B. Mussel-Inspired Surface Chemistry for Multifunctional Coatings. Science 2007, 318, 426−430. (21) Kang, S. M.; You, I.; Cho, W. K.; Shon, H. K.; Lee, T. G.; Choi, I. S.; Karp, J. M.; Lee, H. One-Step Modification of Superhydrophobic Surfaces by a Mussel-Inspired Polymer Coating. Angew. Chem., Int. Ed. 2010, 49, 9401−9404. (22) Kang, S. M.; Hwang, N. S.; Yeom, J.; Park, S. Y.; Messersmith, P. B.; Choi, I. S.; Langer, R.; Anderson, D. G.; Lee, H. One-Step Multipurpose Surface Functionalization by Adhesive Catecholamine. Adv. Funct. Mater. 2012, 22, 2949−2955. (23) Barrett, D. G.; Sileika, T. S.; Messersmith, P. B. Molecular Diversity in Phenolic and Polyphenolic Precursors of Tannin-Inspired Nanocoatings. Chem. Commun. (Cambridge, U. K.) 2014, 50, 7265− 7268. (24) Zhang, W.; Yang, F. K.; Han, Y.; Gaikwad, R.; Leonenko, Z.; Zhao, B. Surface and Tribological Behaviors of the Bioinspired Polydopamine Thin Films Under Dry and Wet Conditions. Biomacromolecules 2013, 14, 394−405. (25) Lee, W.; Lee, J. U.; Jung, B. M.; Byun, J. H.; Yi, J. W.; Lee, S. B.; Kim, B. S. Simultaneous Enhancement of Mechanical, Electrical and Thermal Properties of Graphene Oxide Paper by Embedding Dopamine. Carbon 2013, 65, 296−304. (26) Feng, L.; Li, J. F.; Ye, J. R.; Song, W.; Jia, J.; Shen, Q. Enhancing the Mechanical and Thermal Properties of Polyacrylonitrile through Blending with Tea Polyphenol. J. Appl. Polym. Sci. 2014, 131, 40411. (27) Shen, J.; Gao, G.; Liu, X.; Fu, J. Natural Polyphenols Enhance Stability of Crosslinked UHMWPE for Joint Implants. Clin. Orthop. Relat. Res. 2015, 473, 760−766. (28) Daglia, M. Polyphenols as Antimicrobial Agents. Curr. Opin. Biotechnol. 2012, 23, 174−181. (29) Scalbert, A.; Johnson, I. T.; Saltmarsh, M. Polyphenols: Antioxidants and Beyond. Am. J. Clin. Nutr. 2005, 81, 215S−217S. (30) Sileika, T. S.; Kim, H. D.; Maniak, P.; Messersmith, P. B. Antibacterial Performance of Polydopamine-Modified Polymer Surfaces Containing Passive and Active Components. ACS Appl. Mater. Interfaces 2011, 3, 4602−4610. (31) Kuang, J.; Guo, J. L.; Messersmith, P. B. High Ionic Strength Formation of DOPA-Melanin Coating for Loading and Release of Cationic Antimicrobial Compounds. Adv. Mater. Interfaces 2014, 1, 1400145. (32) Sileika, T. S.; Barrett, D. G.; Zhang, R.; Lau, K. H. A.; Messersmith, P. B. Colorless Multifunctional Coatings Inspired by Polyphenols Found in Tea, Chocolate, and Wine. Angew. Chem., Int. Ed. 2013, 52, 10766−10770. (33) Yang, S. H.; Lee, Y. S.; Lin, F. H.; Yang, J. M.; Chen, K. S. Chitosan/Poly(Vinyl Alcohol) Blending Hydrogel Coating Improves the Surface Characteristics of Segmented Polyurethane Urethral Catheters. J. Biomed. Mater. Res., Part B 2007, 83B, 304−313. (34) Walker, J.; Young, G.; Hunt, C.; Henderson, T. Multi-Centre Evaluation of Two Daily Disposable Contact Lenses. Contact Lens Anterior Eye 2007, 30, 125−133. (35) Alves, M. H.; Jensen, B. E.; Smith, A. A.; Zelikin, A. N. Poly(Vinyl Alcohol) Physical Hydrogels: New Vista on a Long Serving Biomaterial. Macromol. Biosci. 2011, 11, 1293−1313. (36) Riva, S. Laccases: Blue Enzymes for Green Chemistry. Trends Biotechnol. 2006, 24, 219−226. (37) Madhavi, V.; Lele, S. S. Laccase: Properties and Applications. BioResources 2009, 4, 1694−1717. (38) Clinical and Laboratory Standard Institute (CLSI). Performance Standards for Antimicrobial Susceptibility Testing, 17th Information Supplement; CLSI document M100-S18; Clinical and Laboratory Standard Institute: Wayne, PA, 2008.

Chian School of Medicine (L0412130 and L0412290) and the Ministry of Education Singapore AcRF-Tier I (2014-T1-001141) grants.

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REFERENCES

(1) Edmiston, C. E., Jr; Seabrook, G. R.; Cambria, R. A.; Brown, K. R.; Lewis, B. D.; Sommers, J. R.; Krepel, C. J.; Wilson, P. J.; Sinski, S.; Towne, J. B. Molecular Epidemiology of Microbial Contamination in the Operating Room Environment: Is There a Risk for Infection? Surgery 2005, 138, 573−582. (2) Kulakov, L. A.; McAlister, M. B.; Ogden, K. L.; Larkin, M. J.; O’Hanlon, J. F. Analysis of Bacteria Contaminating Ultrapure Water in Industrial Systems. Appl. Environ. Microbiol. 2002, 68, 1548−1555. (3) Silbergeld, E. K.; Graham, J.; Price, L. Industrial Food Animal Production, Antimicrobial Resistance and Human Health. Annu. Rev. Public Health 2008, 29, 151−169. (4) Koyonos, L.; Zmistowski, B.; Della Valle, C. J.; Parvizi, J. Infection Control Rate of Irrigation and Déb ridement for Periprosthetic Joint Infection. Clin. Orthop. Relat. Res. 2011, 469, 3043−3048. (5) Mizan, M. F.; Jahid, I. K.; Ha, S. D. Microbial Biofilms in Seafood: A Food-Hygiene Challenge. Food Microbiol. 2015, 49, 41−55. (6) Hargreaves, J.; Shireley, L.; Hansen, S.; Bren, V.; Fillipi, G.; Lacher, C.; Esslinger, V.; Watne, T. Bacterial Contamination Associated with Electronic Faucets: A New Risk for Healthcare Facilities. Infect. Control Hosp. Epidemiol. 2001, 22, 202−205. (7) Leslie, D. C.; Waterhouse, A.; Berthet, J. B.; Valentin, T. M.; Watters, A. L.; Jain, A.; Kim, P.; Hatton, B. D.; Nedder, A.; Donovan, K.; Super, E. H.; Howell, C.; Johnson, C. P.; Vu, T. L.; Bolgen, D. E.; Rifai, S.; Hansen, A. R.; Aizenberg, M.; Super, M.; Aizenberg, J.; Ingber, D. E. A Bioinspired Omniphobic Surface Coating on Medical Devices Prevents Thrombosis and Biofouling. Nat. Biotechnol. 2014, 32, 1134−1140. (8) Fadeeva, E.; Truong, V. K.; Stiesch, M.; Chichkov, B. N.; Crawford, R. J.; Wang, J.; Ivanova, E. P. Bacterial Retention on Superhydrophobic Titanium Surfaces Fabricated by Femtosecond Laser Ablation. Langmuir 2011, 27, 3012−3019. (9) Hochbaum, A. I.; Aizenberg, J. Bacteria Pattern Spontaneously on Periodic Nanostructure Arrays. Nano Lett. 2010, 10, 3717−3721. (10) Gristina, A. G.; Naylor, P.; Myrvik, Q. Infections from Biomaterials and Implants: A Race for the Surface. Med. Prog. Technol. 1988, 14, 205−224. (11) Timofeeva, L.; Kleshcheva, N. Antimicrobial Polymers: Mechanism of Action, Factors of Activity and Applications. Appl. Microbiol. Biotechnol. 2011, 89, 475−492. (12) Lichter, J. A.; Van Vliet, K. J.; Rubner, F. Y. M. Design of Antibacterial Surfaces and Interfaces: Polyelectrolyte Multilayers as a Multifunctional Platform. Macromolecules 2009, 42, 8573−8586. (13) Sanni, O.; Chang, C. Y.; Anderson, D. G.; Langer, R.; Davies, M. C.; Williams, P. M.; Williams, P.; Alexander, M. R.; Hook, A. L. Bacterial Attachment to Polymeric Materials Correlates with Molecular Flexibility and Hydrophilicity. Adv. Healthcare Mater. 2015, 4, 695−701. (14) Murata, H.; Koepsel, R. R.; Matyjaszewski, K.; Russell, A. J. Permanent, Non-Leaching Antibacterial Surface–2: How High Density Cationic Surfaces Kill Bacterial Cells. Biomaterials 2007, 28, 4870− 4879. (15) Moriarty, T. F.; Grainger, D. W.; Richards, R. G. Challenges in Linking Preclinical Anti-Microbial Research Strategies with Clinical Outcomes for Device-Associated Infections. Eur. Cells Mater. 2014, 28, 112−128. (16) Salwiczek, M.; Qu, Y.; Gardiner, J.; Strugnell, R. A.; Lithgow, T.; McLean, K. M.; Thissen, H. Emerging Rules for Effective Antimicrobial Coatings. Trends Biotechnol. 2014, 32, 82−90. (17) Bazaka, K.; Jacob, M. V.; Chrzanowski, W.; Ostrikov, K. AntiBacterial Surfaces: Natural Agents, Mechanisms of Action and Plasma Surface Modification. RSC Adv. 2015, 5, 48739−48759. L


Research Article

ACS Applied Materials & Interfaces (39) Muller, G.; Kramer, A. Biocompatibility Index of Antiseptic Agents by Parallel Assessment of Antimicrobial Activity and Cellular Cytotoxicity. J. Antimicrob. Chemother. 2008, 61, 1281−1287. (40) Dreyer, D. R.; Miller, D. J.; Freeman, B. D.; Paul, D. R.; Bielawski, C. W. Elucidating the Structure of Poly(Dopamine). Langmuir 2012, 28, 6428−6435. (41) Hong, S.; Na, Y. S.; Choi, S.; Song, I. T.; Kim, W. Y.; Lee, H. Non-Covalent Self-Assembly and Covalent Polymerization CoContribute to Polydopamine Formation. Adv. Funct. Mater. 2012, 22, 4711−4717. (42) D’Ischia, M.; Napolitano, A.; Ball, V.; Chen, C. T.; Buehler, M. J. Polydopamine and Eumelanin: from Structure-Property Relationships to a Unified Tailoring Strategy. Acc. Chem. Res. 2014, 47, 3541−3550. (43) Li, Y.; Tan, Y.; Deng, W.; Xie, Q.; Zhang, Y.; Chen, J.; Yao, S. Electropolymerization of Catecholamines after Laccase-Catalyzed preoxidation to Efficiently Immobilize Glucose Oxidase for Sensitive Amperometric biosensing. Sens. Actuators, B 2010, 151, 30−38. (44) Wei, Q.; Zhang, F.; Li, J.; Li, B.; Zhao, C. Oxidant-Induced Dopamine Polymerization for Multifunctional Coatings. Polym. Chem. 2010, 1, 1430−1433. (45) Sun, X.; Bai, R.; Zhang, Y.; Wang, Q.; Fan, X.; Yuan, J.; Cui, L.; Wang, P. Laccase-Catalyzed Oxidative Polymerization of Phenolic Compounds. Appl. Biochem. Biotechnol. 2013, 171, 1673−1680. (46) Gao, R.; Yuan, Z.; Zhao, Z.; Gao, X. Mechanism of Pyrogallol Autoxidation and Determination of Superoxide Dismutase Enzyme Activity. Bioelectrochem. Bioenerg. 1998, 45, 41−45. (47) Roy, J. R.; Abraham, T. E. Continuous Biotransformation of Pyrogallol to Purpurogallin Using Cross-Linked Enzyme Crystals of Laccase as Catalyst in a Packed-Bed Reactor. J. Chem. Technol. Biotechnol. 2006, 81, 1836−1839. (48) Lopez, J.; Hernandez-Alcantara, J. M.; Roquero, P.; Montiel, C.; Shirai, K.; Gimeno, M.; Barzana, E. Trametes Versicolor Laccase Oxidation of Gallic Acid Toward a Polyconjugated Semiconducting Material. J. Mol. Catal. B: Enzym. 2013, 97, 100−105. (49) Bozic, M.; Strancar, J.; Kokol, V. Laccase-Initiated Reaction between Phenolic Acids and Chitosan. React. Funct. Polym. 2013, 73, 1377−1383. (50) Tóth, I. Y.; Szekeres, M.; Turcu, R.; Sáringer, S.; Illés, E.; Nesztor, D.; Tombácz, E. Mechanism of In Situ Surface Polymerization of Gallic Acid in an Environmental-Inspired Preparation of Carboxylated Core−Shell Magnetite Nanoparticles. Langmuir 2014, 30, 15451−15461. (51) Bruno, F. F.; Trotta, A.; Fossey, S.; Nagarajan, S.; Nagarajan, R.; Samuelson, L. A.; Kumar, J. J. Enzymatic Synthesis and Characterization of PolyQuercetin. J. Macromol. Sci., Part A: Pure Appl.Chem. 2010, 47, 1191−1196. (52) Lukes, V.; Darvasiova, D.; Furdikova, K.; Hubertova, I.; Rapta, P. Solvent Effect on the Anodic Oxidation of Tannic Acids: EPR/UV– Vis Spectroelectrochemical and DFT Theoretical Study. J. Solid State Electrochem. 2015, 19, 2533−2544. (53) Aisha, A. F.; Ismail, Z.; Abu-Salah, K. M.; Majid, A. M. Solid Dispersions of α-Mangostin Improve its Aqueous Solubility through Self-Assembly of Nanomicelles. J. Pharm. Sci. 2012, 101, 815−825. (54) Mansur, H. S.; Sadahira, C. M.; Souza, A. N.; Mansur, A. A. P. FTIR Spectroscopy Characterization of Poly (Vinyl Alcohol) Hydrogel with Different Hydrolysis Degree and Chemically Crosslinked with Glutaraldehyde. Mater. Sci. Eng., C 2008, 28, 539−548. (55) Zangmeister, R. A.; Morris, T. A.; Tarlov, M. J. Characterization of Polydopamine Thin Films Deposited at Short Times by Autoxidation of Dopamine. Langmuir 2013, 29, 8619−8628. (56) Weis, C.; Odermatt, E. K.; Kressler, J.; Funke, Z.; Wehner, T.; Freytag, D. Poly(Vinyl Alcohol) for membrane prevention. J. Biomed. Mater. Res. 2004, 70B, 191−202. (57) Hong, S.; Yeom, J.; Song, I. T.; Kang, S. M.; Lee, H.; Lee, H. Pyrogallol 2-Aminoethane: a Plant Flavonoid-Inspired Molecule for Material-Independent Surface Chemistry. Adv. Mater. Interfaces 2014, 1, 1400113. (58) Holland, J. N.; Hay, J. N. The Thermal Degradation of Poly(vinyl alcohol). Polymer 2001, 42, 6775−6783.

(59) Yang, C. C. Synthesis and Characterization of the Cross-linked PVA/TiO2 Composite Polymer Membrane for Alkaline DMFC. J. Membr. Sci. 2007, 288, 51−60. (60) Figueiredo, K. C. S.; Alves, T. L. M.; Borges, C. P. Poly(vinyl alcohol) Films Crosslinked with Glutaraldehyde under Mild Conditions. J. Appl. Polym. Sci. 2009, 111, 3074−3080. (61) Jia, X.; Li, Y.; Cheng, Q.; Zhang, S.; Zhang, B. Preparation and Properties of Poly(vinyl alcohol)/Silica Nanocomposites from Copolymerization of Vinyl Silica Nanoparticles and Vinyl Acetate. Eur. Polym. J. 2007, 43, 1123−1131. (62) Lim, M.; Kim, D.; Seo, J.; Han, H. Preparation and Properties of Poly(vinyl alcohol)/Vinyltrimethoxysilane (PVA/VTMS) Hybrid Films with Enhanced Thermal Stability and Oxygen Barrier Properties. Macromol. Res. 2014, 22, 1096−1103. (63) Lim, M.; Kwon, H.; Kim, D.; Seo, J.; Han, H.; Khan, S. B. Highly-Enhanced Water Resistant and Oxygen Barrier Properties of Cross-linked Poly(vinyl alcohol) Hybrid Films for Packaging Applications. Prog. Org. Coat. 2015, 85, 68−75. (64) Darouiche, R. O. Device-Associated Infections: A Macroproblem that Starts with Microadherence. Clin. Infect. Dis. 2001, 33, 1567−1572. (65) McKeen, L. W. The Effect of Sterilization on Plastics and Elastomers, 3rd ed.; William Andrew: Norwich, NY, 2012. (66) Schmidt, M. G.; Attaway, H. H.; Sharpe, P. A.; John, J.; Sepkowitz, K. A.; Morgan, A.; Fairey, S. E.; Singh, S.; Steed, L. L.; Cantey, J. R.; Freeman, K. D.; Michels, H. T.; Salgado, C. D. Sustained Reduction of Microbial Burden on Common Hospital Surfaces through Introduction of Copper. J. Clin. Microbiol. 2012, 50, 2217− 2223. (67) Szczotka-Flynn, L. B.; Pearlman, E.; Ghannoum, M. Microbial Contamination of Contact Lenses, Lens Care Solutions, and Their Accessories: A Literature Review. Eye Contact Lens 2010, 36, 116−129. (68) Koh, J. J.; Qiu, S.; Zou, H.; Lakshminarayanan, R.; Li, J.; Zhou, X.; Tang, C.; Saraswathi, P.; Verma, C.; Tan, D. T.; Tan, A. L.; Liu, S.; Beuerman, R. W. Rapid Bactericidal Action of Alpha-Mangostin against MRSA as an Outcome of Membrane Targeting. Biochim. Biophys. Acta, Biomembr. 2013, 1828, 834−844. (69) Kim, H.; Jang, J. H.; Kim, S. C.; Cho, J. H. De novo generation of short antimicrobial peptides with enhanced stability and cell specifity. J. Antimicrob. Chemother. 2014, 69, 121−132. (70) Hyon, S. H.; Cha, W. I.; Ikada, Y.; Kita, M.; Ogura, Y.; Honda, Y. Poly(vinyl alcohol) hydrogels as soft contact lens material. J. Biomater. Sci., Polym. Ed. 1994, 5, 397−406. (71) Masters, K. S. B.; Leibovich, S. J.; Belem, P.; West, J. L.; PooleWarren, L. A. Effects of nitric oxide releasing poly(vinyl alcohol) hydrogel dressings on dermal wound healing in diabetic mice. Wound Repair Regen. 2002, 10, 286−294. (72) Kobayashi, M.; Chang, Y. S.; Oka, M. A two year in vivo study of polyvinyl alcohol-hydrogel (PVA-H) artificial meniscus. Biomaterials 2005, 26, 3243−3248. (73) Madhurakkat Perikamana, S. K.; Lee, J.; Lee, Y. B.; Shin, Y. M.; Lee, E. J.; Mikos, A. G.; Shin, H. Materials from Mussel-Inspired Chemistry for Cell and Tissue Engineering Applications. Biomacromolecules 2015, 16, 2541−2555.

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