Surface Grafted Antimicrobial Polymer Networks with High Abrasion

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Surface Grafted Antimicrobial Polymer Networks with High Abrasion Resistance Jing Gao, Noah Eric Huddleston, Evan M. White, Jitendra Pant, Hitesh Handa, and Jason Locklin ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.6b00221 • Publication Date (Web): 30 May 2016 Downloaded from http://pubs.acs.org on June 7, 2016

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ACS Biomaterials Science & Engineering

Surface Grafted Antimicrobial Polymer Networks with High Abrasion Resistance Jing Gao,† N. Eric Huddleston,§ Evan M. White, † Jitendra Pant,‡ Hitesh Handa,‡ Jason Locklin *,†,‡

†

‡

Department of Chemistry, University of Georgia, Athens, Georgia, 30602, United States

School of Biological and Biochemical Engineering, College of Engineering, University

of Georgia, Athens, Georgia, 30602, United States §

Department of Chemistry and Biochemistry, University of North Georgia, Dahlonega,

Georgia, 30597

KEYWORDS: quaternary ammonium, benzophenone photocrosslinking, antibacterial, covalent surface attachment, robust polymeric network

Abstract In this work, we have investigated a quaternary ammonium compound that exhibits excellent antimicrobial activity and can be permanently grafted to substrates containing C-H bonds to form a durable polymeric film within one minute. The compound consists of a biocidal component, dodecyl-alkylated quaternary ammonium, and a benzophenone moiety that, under mild UV irradiation, generates a densely crosslinked network and covalently attaches to a variety of substrates, including plastics, fabrics and alkyl-

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modified glass surfaces. The surface attachment is one order of magnitude faster than previously reported benzophenone-associated crosslinkers, due to the electronwithdrawing effect of quaternary ammonium on the benzophenone chromophore. The modified surfaces are non-leaching and exhibit contact-killing and highly effective antimicrobial activity against Staphylococcus aureus (Gram-positive) and Escherichia coli (Gram-negative) using cell count and live/dead staining methods. The charged ammonium group also promotes photoreaction efficiency with respect to network robustness, leading to a thin film that can sustain high shear forces and abrasion when compared to commercially available silane-based quaternary ammonium compounds. The biocidal activity is also retained after exposure to mechanical stress and abrasion.

Introduction Surface microbial contamination associated with medical implants and devices, hospital equipment, food packaging and storing, textiles, and many other materials is one of the most serious global healthcare issues facing humanity.1-5 For example, 1.7 million healthcare-associated infections (HAIs) were estimated for the year 2002 in the United States. The number of associated death was 98,987, of which, 8,205 were form surgical site infections including those associated with orthopedic implants.6 To address surface microbial infection, two major strategies have been developed to incorporate coatings with antimicrobial capabilities: (1) designing surfaces that physically immobilize antimicrobial agents including antibiotics, halogens, heavy metals (silver, copper, or mercury), and quaternary ammonium compounds, that are released or leach from


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coatings in a rate-controlled fashion; and (2) designing surfaces containing covalently bound antimicrobial agents.7-10 Although effective, the first strategy is limited in terms of longevity, mechanical and chemical durability, thermal stability, and the propensity towards environmental contamination due to the nature of the surface interaction (physisorption), which is required by the “leaching” nature of the biocidal action.11 Such agents only interact with the surface through weaker intermolecular forces such as van der Waals, hydrophobic and/or electrostatic interactions.10,

12-13

The second strategy,

which involves preparing permanently effective and durable non-leaching antimicrobial agents covalently immobilized to the surface, has been under intensive study for the past decade.14-17

Among the surface immobilized antimicrobial agents, membrane-disrupting poly“-onium” (ammonium and pyridinium) cations with various alkyl chain lengths have drawn considerable interest because of their facile synthesis, broad application, outstanding antimicrobial activity, low cost, and low bacterial resistance.2,

18-21

The

generally accepted hypothesis for the biocidal mechanism of the surface tethered polycation suggests that as a result of electrostatic interactions, the positively charged “onium” replaces the bacteria’s natural counterions (Mg2+ and Ca2+) which are initially confined within the cell membrane, disrupting the ionic integrity of the membrane.11, 22 Meanwhile, the alkyl chains of polycations intercalate into the phospholipid bilayer structure which disturbs its organization, causing holes to form in the membrane.17, 23 Bieser et al. proposed an alternative mechanism, the “phospholipid sponge effect,” which hypothesized that phospholipids were drawn out of bacterial lipid bilayer where they can



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permeate into cationic films.24 Numerous reports have demonstrated different approaches to covalently bind polycations onto a wide variety of surfaces based on various attachment chemistries between surface functional groups and polycations. Some examples include the condensation of poly-quaternary ammonium-silane (PQA-silane) derivatives with hydroxyl groups of cellulose or -OH functionalized glass, ceramics and metals;25-26 formation of an amino-based polyacrylate grown from an alkyl/aryl halide functionalized surfaces via surface initiated atom transfer radical polymerization (SIATRP) followed by subsequent quaternization;14 a vinyl sulfone-PQA derivative reacted with -OH on cellulose via Michael addition;27 and an aryl azide-PQA with -OH on cellulose via photocrosslinking.28 Though these methodologies are effective in killing bacteria, reactive surfaces that contain native –OH or -NH2 groups (i.e., cellulose, chemical or plasma pretreated glass, metal or plastic)29-30 and/or multiple complicated steps are required for the covalent attachments. Therefore, a low-cost, simple, and versatile method to permanently immobilize PQA on common and inert plastic surfaces is highly demanded.

The benzophenone (BP) chromophore is of enduring interest as a photoactive tethering reagent that has been used to functionalize a variety of materials.31-35 The extensive use of BP is due to the following advantages: (1) The BP moiety is able to attach to C-H bonds in a wide range of chemical environments, including commercial plastics and fabrics;36-39 (2) BP can be manipulated in ambient atmosphere and activated with mild UV light, avoiding oxidative damage to the underlying substrate;32 and (3) BP is chemically and thermally more stable and synthetically more versatile than most other


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tethering functionalities, such as sulfonyl azides,40 diazo esters,41 aryl azides,42 and diazirines.43-44 When irradiated, BP absorbs photons, leading to the promotion of an electron from the n-orbital of the carbonyl oxygen to the π*-orbital and the formation of biradicaloid triplet state. The electron-deficient oxygen abstracts a hydrogen atom from neighboring C-H group, followed by the combination of two newly created carbon radicals and subsequent generation of a new C-C bond. This photochemistry has been used to permanently attach polymer films to a broad selection of C-H surfaces in many different applications, such as microfluidic devices,45-46 biosensors,47-48 patterned sheets,49-50 and even organic semiconductors.51 Because of these advantages, we previously developed an antimicrobial copolymer containing hydrophobic N-alkyl and benzophenone moieties on a polyethyleneimine backbone (AmBP) and covalently grafted the copolymer to various plastic and textile surfaces via photocrosslinking.52 The polymer coated surface exhibited high biocidal activity against Gram-positive and Gram-negative bacteria (>98%). However, the photo-crosslinking of the polymer was time consuming and inefficient (i.e. a 30 minute irradiation period with high intensity UV light was necessary), thereby limiting the industrial applicability of such a material as a final coating.

In this work, a benzophenone based antimicrobial small molecule (BPAM) was prepared via a two step synthesis. The photocrosslinking towards both reactive and inert surfaces was both rapid and efficient upon mild irradiation. BPAM coated surfaces also demonstrated excellent biocidal activity against Gram-positive and Gram-negative bacteria on contact. BPAM exhibits high mechanical stability against harsh mechanical abrasion while retaining antimicrobial activity after repeated abrasion cycles.



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Furthermore, its non-leaching killing action was demonstrated, extending its commercial relevance to permanent and durable coatings on a variety of substrates.

Experimental Section Materials.

N-bromosuccinimide (NBS), 2,2’-azobis(2-methylpropiontirle) (AIBN),

N,N-dimethyldodecylamine, octyltrichorosilane, and agar powder were purchased from Alfa-Aesar. Fluorescein sodium salt (Sigma Aldrich), cetyltrimethylammonium bromide (Acros Organic), 4-methylbenzophenone (Oxchem), cyclohexane (Honeywell), tert-amyl alcohol (JT Baker), sodium chloride (EMD Chemical), peptone (HiMedia), yeast extract (Criterion), beef extract (HiMedia), dextrose (BD), and bromopheonol blue (Amresco) were used without further purification. The two-color fluorescent LIVE/DEAD BacLight bacterial viability kit L7012 (Molecular Probes, Life Technologies) which contains SYTO® 9 green-fluorescent nucleic acid stain and propidium iodide red-fluorescent nucleic acid stain was utilized to evaluate the bacterial viability. Silicon wafer with native oxide and glass slides (cut into 2.5 × 2.5 cm pieces) were used as substrates. For fabric and plastic substrate demonstrations, 100% cotton print cloth (black and white), a polyvinyl acetate (PVA) shower curtain (WalMart Inc.), and polyethylene (PE) sheets (Husky) were purchased from Home Depot.

Instrumental Methods. UV-vis spectroscopy was performed on a Cary Bio Spectrophotometer (Varian). Two irradiation wavelengths, 254 and 365 nm, were utilized in this study. The UV light sources were a Compact UV lamp (UVP) and FB-UBXL1000 UV Crosslinker (Fisher Scientic) with bulbs of wavelength at 254 nm for small (1 ×


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1 cm) and larger (2.5 × 2.5 cm) substrates, respectively. The substrates were held at a distance of 0.5 cm from the light source during irradiation to obtain a power of 6.5 mW/cm2. Another UV light source was an OmniCure, Series 1000 with 365 nm bandpass filter, equipped with a liquid filled fiber optic waveguide. The substrates were held 2 cm from the source for a power of 25 mW/cm2. Surface topography was analyzed with a Bruker Multimode atomic force microscope (AFM). All measurements were performed using tapping mode. Infrared spectroscopy studies of coated films were carried out using a Thermo-Nicolet model 6700 spectrometer equipped with a variable angle grazing angle attenuated total reflection (GATR-ATR) accessory (Harrick Scientific). Water contact angles were measured by a DSA 100 drop shape analysis system (KRŰSS) with a computer-controlled liquid dispensing system. Water droplets with volume of 1 µL were used to measure the static contact angle. The thickness of the film was measured using a M-2000 spectroscopic ellipsometer (J.A. Woollam Co., Inc). A Nikon Eclipse Ni-U fluorescent microscope equipped and a 100× objective were used for live/dead bacterial viability photomicrographs. A GFP FITC filter cube (excitation: 490 nm, emission: 503 nm) was used for SYTO® 9 and a Texas Red filter cube (excitation: 577 nm, emission: 620 nm) was used for the propidium iodide.

Synthesis. (4-bromomethyl)benzophenone: 4-methylbenzopheone (6.0 g, 30.6 mmol), NBS (6.0 g, 33.6 mmol), AIBN (1.0 g, 6.1 mmol), and cyclohexane (100 mL) were added to a round bottom flask under nitrogen atmosphere. The suspension was stirred under reflux overnight. After stirring, the mixture was cooled and filtered to remove any solid. The filtrate was concentrated under reduced pressure. The solid mixture was dissolved in



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diethyl ether and washed with water, brine and dried over magnesium sulfate. The mixture was filtered and concentrated under reduced pressure. The recovered solid was recrystallized from absolute ethanol to give fine white crystals. Yield: 7.1 g, 89%. 1H NMR: δ, 7.80 (t, 2H, J = 3.0 Hz); 7.78 (t, 2H, J = 1.4 Hz); 7.60 (t, 1H, J = 7.0 Hz); 7.50 (d, 2H, 8.2 Hz); 7.49 (t, 2H, 7.6 Hz); 4.53 (s, 2H). 13C NMR (CDCl3): δ,

195.93,

142.09, 137.39, 132.54, 130.52, 129.99, 128.92 128.33, 128.16, 32.25.

N-(4-benzoylbenzyl)-N,N-dimethylbutan-1-aminium iodide (BPAM): (4-bromomethyl) benzophenone (1.7 g, 6.2 mmol), N, N-demethyldodecylamine (1.7 mL, 6.2 g), and tertamyl alcohol (15 mL) were added to a sealable pressure flask. The mixture was stirred and heated in the sealed vessel at 95 °C for 24 h. The flask was cooled to room temperature and the solvent was removed under reduced pressure. The resulting brown waxy solid was recrystallized in hexane/ethyl acetate (7:4) to give a waxy white solid. Yield: 1.7 g, 67%. 1H NMR (CDCl3): δ, 7.84 (dd, 4H, J = 8.2, 23.9 Hz); 7.75 (d, 2H, J = 7.0 Hz); 7.59 (t, 1H, J = 7.6 Hz); 7.47 (t, 2H, 7.7 Hz); 3.57 (m, 2H); 3.35 (s, 6H); 1.80 (bs, 2H); 1.31 (bs, 4H); 1.21 (bs, 16H); 0.84 (t, 3H, J= 6.6 Hz).

13

C NMR (CDCl3): δ,

210.33, 139.75, 136.70, 133.53, 133.24, 131.53, 130.54, 130.24, 120.66, 66.63, 64.12, 49.85, 32.06, 29.69, 29.58, 29.46, 29.37, 26.42, 22.82, 14.29.

Preparation of Surface-Bound BPAM Cross-linked Films. Glass slides were cut into 2.5 × 2.5 cm pieces and sonicated in deionized water, isopropanol, and acetone for 10 min each, followed by argon plasma (Harrick Plasma PDC-32G) for 5 min. A selfassembled monolayer (SAM) of octyltrichlorosilane (OTS) was formed by solution


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deposition in OTS/toluene solution (0.1 mM) overnight. The BPAM film was deposited onto surfaces with two coating techniques: spray coating was utilized for OTS functionalized glass slides, fabric and plastic pieces; and doctor blading was utilized for OTS functionalized quartz slides. BPAM/acetone solution (5 mg/mL) was sprayed using a spray gun from a distance of 20 cm onto a vertically placed substrate to achieve uniform coating upon drying. For doctor blading, BPMA stock solution (5 mg/mL in chloroform, 10 µL) were placed on the substrate under a razor blade. The razor blade was connected to a syringe pump that established a constant linear movement at relative rate of 1.79 µm/s between the razor blade and the substrate, and the solution was spread on the substrate to form a thin layer that resulted in a uniform, thin film upon solvent evaporation. Then BPAM films were irradiated with UV light (254 nm, 6.5 mW/cm2) for 2 min to covalently bind the BPAM to the surfaces. The substrates were sonicated with acetone for 1 min to rinse off unattached BPAM and dried under a stream of nitrogen.

Surface Charge Density Assessment. The charge density of BPAM modified surfaces was analyzed indirectly using UV-Vis spectroscopy by measuring the concentration of the negatively charged fluorescein sodium dye, which complexes with the positively charged -onium salt with 1:1 stoichiometry. A BPAM modifed glass slide (2.5 × 2.5 cm) was dipped in a 1% slution of fluorescein sodium salt/water solution for 10 min to fully adsorb the fluorescein dye. Then the glass slide was rinsed with distilled water, placed in 10 mL of 0.1% cetyltrimethylammonium chloride in distilled water, and shaken for 20 min to completely desorb the fluorescein dye. After adding 0.5 mL aqueous phosphate buffer (pH 8.0), the absorbance of the resultant aqueous solution was measured at 501



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nm. The surface charge density is calculated from the known surface area of the substrate and by determining the the number of dye molecules released into solution dye. Using the Beer Lambert law (extinction coefficient of fluorescein at 501 nm = 77 mM-1 cm-1) the number of cations on the surface may be determined.

Antimicrobial Test Method. The antimicrobial efficacy of the BPAM coated surface was determined by using a modified test method published by Dhende et al52 and Haldar et al. 53 In the antimicrobial test used in this work, a bacterial aerosol was sprayed onto a dry functionalized surface. This method mimics the common mechanism of airborne bacterial infection, which includes infected respiratory droplets produced by sneezing, coughing, or breathing.

The yeast-dextrose broth and 1.5% solid growth agar were prepared using the recipe given by Haldar et al.53 Both Gram-positive, Staphylococcus aureus (S. aureus) (ATCC 5538), and Gram-negative, Escherichia coli (E. coli) (ATCC 11303), were tested. One loopful of bacterial culture was inoculated in 10 mL yeast-dextrose broth in an incubator shaker (New Brunswich Scientific Excella E24) at 37 °C and 200 r.p.m. overnight. The bacteria were revived by inoculating 100 µL of the bacterial suspension in 10 mL fresh both under the same incubating condition for 16 h. The bacterial suspension was centrifuged at 4000 r.p.m. for 10 min at room temperature to harvest bacterial cells. The supernatant was removed and the bacterial pellet was resuspended in 10 mL deionized water using a vortex mixer (Vortex Genie 2). The washing process was repeated once. The bacterial suspension was diluted 100 fold to a concentration of 3 × 106 cfu/mL


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(colony forming unit). A thin layer chromatography (TLC) sprayer connected to pneumatic dispense regulator (EFD 1500XL) was utilized for bacterial aerosol spraying. The diluted bacterial suspension was transferred into the TLC sprayer and sprayed under onto substrates for 0.1 s at 35 psi. The sprayed side was air dried for 3 min and mounted to the solid growth agar plate. The samples were incubated for 16 h at 37 °C. The colonies grown on the substrate were then counted.

Non-leaching Evaluation. The parallel bacterial streak method (AATCC 147) was conducted to evaluate the non-leaching antimicrobial property of BPAM coated substrates. Cotton was cut into 25 × 50 mm as test specimens and sterilized by sonication in ethanol. Again, S. aureus and E. coli were utilized as test organisms. The bacterial inoculum was prepared by transferring 1 mL of a 24 h broth culture into 9.0 mL of sterile distilled water. By using a 4 mm inoculating loop, the diluted inoculum was transferred to the surface of a sterile agar plate by making five streaks approximately 6 cm in length, spaced 1 cm apart covering the central area of the agar plate. The loop was refilled with the inoculum after each streak. The test substrate was gently pressed transversely across five inoculum streaks to ensure intimate contact with the agar surface and incubated at 37 °C for 24 h.

Live/Dead Bacterial Viability Assay. The antimicrobial capability of surface-bound BPAM was also estimated by examining the bacterial cell viability after direct contact with the coating using the two-color fluorescent live/dead assay. SYTO® 9 dye, which yields green fluorescence, labels all bacteria in population with intact or damaged



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membranes. In contrast, propidium iodide, which yields red fluorescence, penetrates only bacteria with damaged membranes and displaces SYTO® 9 stain, causing a reduction in green fluorescence and the appearance of red fluorescence. Consequently, bacteria with damaged cell membranes can be distinguished from live bacteria. Both S. aureus and E. coli were tested. A 10 mL bacterial culture was grown to late log phase in broth (shaken at 100 r.p.m. for 10 h at 37 °C). The culture was centrifuged at 4000 r.p.m. for 10 min. The supernatant was removed and the pellet was resuspended in sterile distilled water. Before staining, 10 µL bacterial/water suspension with concentration of 108 cfu/mL was placed on the control and BPAM modified glass slides and dried at 37 °C for 5 min to achieve quick and intimate contact. Equal volumes of SYTO® 9 and propidium iodide (1.5 µL) were combined, added to 1 mL of distilled water, and mixed thoroughly. Diluted dye mixture (10 µL) was trapped between the bacterial adhered slide and an 18 mm square coverslip. The sample was incubated in dark for 15 min at room temperature and observed with a fluorescence microscope equipped with proper filter sets to determine cell viability.

Abrasion Testing. The robustness of the surface-bound BPAM on substrate was evaluated by a modified standard rub abrasion test ASTM D6279. A pencil (# 2, HB, Ticonderoga) latex free eraser was pressed against a BPAM coated PVA and PP substrates (2.5 × 2.5 cm) mounted to a scale with 1000 g of force and rubbed back and forth (considered as one rubbing cycle) along the midline over the surface. After given amount of rubbing cycles, the distribution of remaining surface attached BAPM was directly visualized by staining the surface with bromophenol blue (BPB), an anionic dye


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that is able to complex with cationic molecules (i.e. BPAM) via electrostatic interaction. The retention of the BPAM antimicrobial activity was estimated by performing antimicrobial testing on abraded substrates.

Results and Discussions N-(4-benzoylbenzyl)-N,N-dimethylbutan-1-aminium iodide (BPAM) contains a dodecyl substituted quaternary ammonium group that endows the molecule with potent antimicrobial activity, and benzophenone as a photoreactive cross-linker. A long alkyl chain (C12) was chosen because it increases the hydrophobicity of the positively charged network, and consequently maintains the hydrophilic-hydrophobic balance needed for high antimicrobial activity.54 The BP chromophore reacts with any surface that contains a C-H group through hydrogen abstraction followed by C-C recombination under UV irradiation. Besides surface attachment, BP also reacts non-discrimately both intra- and intermolecularly with neighboring alkyl groups, resulting in a densely cross-linked network that is able to resists mechanical stress. The synthesis consists of only two steps and no chromatography was needed for purification, making it ameable to large-scale synthesis. BPAM is soluble in halogenated and non-halogenated organic solvents, alcohols and slightly soluble in water. Its favorable solubility enables coating of hydrophobic (plastic) substrates and hydrophilic (fabric) substrates by dissolving it in compatible solvent with excellent wetting properties.

Photoreactive Cross-linked BPAM Thin Films. To investigate the surface physical properties, crosslinking kinetics, and morphology of the BPAM coatings, OTS modified



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silicon wafer and quartz slides were utilized (results shown in Table 1). To the SAM modified surfaces a thin layer of BPAM was deposited by spray coating. Covalent attachment was generated by exposure to UV light at 254 nm for 2 min (Scheme 1). The cross-linked film was then sonicated in acetone to remove any residual, unbound material. The film thickness was measured before and after sonication and was observed to be 48 and 42 nm respectively, indicating that ~85% of the applied BPAM was covalently tethered to the surface. After BPAM attachment, the surface is more hydrophilic with a static contact angle of 69° compared to the initial OTS SAM with contact angle of 101°, due to the positively charged ammonium functionality exposed on the surface.21 The hydrophilicity implies a high charge density, which was calculated to be 35.7 nm-2. The studies of Murata et. al. indicate that for a thin film ( 95% bacterial cells on BPAM coated surfaces are stained red, indicating the cell membrane disruption caused by contact with surface immobilized BAPM. It is noteworthy to state that we observed an aggregation of bacterial cells after exposure to BPAM. This phenomenon could be explained by the disturbance of localized cell membrane electrostatic integrity, which might cause bacterial cells to deform and collapse together. The plausible explanation for the observed aggregation might be attributed to quorum sensing of bacteria in adverse conditions.61-64 Along with culture-based viable count assays, the fluorescent microscopic images of the Live/Dead assay demonstrate that surface-coated BPAM damage the bacterial cell membrane structure in short reaction times and efficiently kills bacteria at a very high concentration (108 cfu/mL). By comparison, the diagnostic criterion for urinary tract infection and bacterial lowrespiratory tract infection is the presence ≥ 105 cfu.65-66



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Figure 5. Fluorescent images of bacteria on glass substrates after live-dead staining: S. aureus in contact with (A) control and (B) BPAM functionalized substrates sprayed; E. coli. in contact with (C) control and (D) BPAM functionalized substrates.

BPAM Coated On Commercially Available Substrates. To investigate the versatility of BPAM coating on commercially available plastics and textiles, polyethylene (PE), polyvinyl acetate (PVA), and cotton swatches were photochemically modified with BPAM and tested using the same bacterial test methods described above. After irradiation and removal of unbound BPAM by sonication and drying, the substrates, particularly cotton swatches, retain their flexibility (do not become stiff) due to the ultrathin nature of the coating. The coated plastic and fabric substrates were challenged against S. aureus and E. coli aerosol. Figure 6 demonstrate the antimicrobial activity of surface coated BPAM on plastic and textiles against S. aureus. On uncoated samples, biofilms formed underneath the substrates. For clarity, the edges of the biofilms are


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shown in the insets of each image. With cotton, the formed biofilms are prominent, as a result of high porosity and affinity between bacteria and cotton (Figure 6A). For PE and PVA, the bacteria colonies form a continuous layer over the sample surface (Figure 6C and 6E). On the other hand, for BPAM coated cotton, PE, and PVA, no bacterial colonies were observed on the bacterial-exposed area or around the edges of the samples. The same antibacterial activity against E. coli is presented in Figure S3. The results demonstrate the high crosslinking reactivity between BPAM and the surrounding C-H groups lead to the effective film formation on a variety of materials, including both those with reactive functional groups such as cotton, PVA, as well as inert plastics such as PE.

Figure 6. Digital pictures of the plastic and fabric substrates sprayed with S. aureus suspension and incubated overnight at 37 °C: (A) uncoated cotton, (B) BPAM coated



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cotton, (C) uncoated PVA, (D) BPAM coated PVA, (E) uncoated PE, (F) BPAM coated PE. Insets: zoom-in images of squared areas.

Non-leaching Killing Action. The parallel bacterial streak test (AATCC 147) was performed on BPAM coated cotton strips to further confirm its non-leaching biocidal action. Cotton strips were spray-coated with BPAM/isopropyl alcohol (iPA) solution, photo-crosslinked, and sonicated sufficiently in solvent (iPA) to guarantee no diffusion of unbound BPAM in the bacterial streaking test. Bacterial inoculum with a concentration of 107 cfu/mL was prepared for a proper streaking density. Five fine streaks were made on an agar plate and a cotton strip was carefully pressed across the inoculum streaks to prevent interruption between each streak. Figure 7 shows the control and BPAM coated cotton strips on agar plates streaked with S. aureus and E. coli after incubation overnight. On the control plates (Figure 7A and 7C), bacterial colonies grow in well-defined lines across the cotton strips. On the coated plates (Figure 7B and 7D), no microorganisms were observed underneath the BPAM coated strip. Importantly, no zone of inhibition beyond the edge of the coated samples was observed (insets of Figure 7B and 7D, arrowed), suggesting effective, localized, and non-leaching antimicrobial capability of the BPAM coated samples.


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Figure 7. Digital pictures of parallel bacterial streaking test on cotton strips after incubated overnight at 37 °C: S. aureus streaks on (A) control and (B) BPAM coated substrates; E. coli streaks on (C) control and (D) BPAM coated substrates. Inset: zoom-in images of the substrate edge.

Robustness of BPAM Coated Surfaces. The robustness of the BPAM polymer networks that are covalent bonded to the underlying substrates against mechanical stress was evaluated by conducting an eraser-rubbing test on BPMA coated PE and PVA. As a



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comparison,

we

subjected

identical

samples

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coated

with

(trihydroxylsilyl)propyldimethyloctadecyl ammonium chloride (Si-Quat), which is the active ingredient in many commercially available antimicrobial products that have label claims of permanent attachment. Before abrasion, both antimicrobial coatings were stained with BPB, a color indicator that easily indicates the distribution of QAM on surfaces. A desktop scanner (brother MFC-8860DN) was utilized as a colorimeter to quantify the amount of quaternary ammonium on the surface by measuring the dye absorption. Stained substrates were scanned and the resultant images were analyzed using Photoshop (Adobe, Inc.). Of the L, a, b value of the color, b was used to quantify the intensity of the blue color resulting from BPB staining. The more negative the b value, the higher the BPB adsorption. Then both surfaces were subjected to the same rubbing conditions. The cleavage of the surface bonding of the quaternary ammonium compound due to mechanical abrasion was then visualized by the depletion of the blue colored stain. The samples were scanned and analyzed after every two abrasion cycles. The relative change of the average b value at 10 different spots on the abraded area is plotted versus abrasion cycles in Figure 8.

Next, we compared the retention of these two surface-bound quaternary ammoniums tethered via C-C and Si-O crosslinks to PE after mechanical abrasion. Figure 8A demonstrates the percentage of remaining BAPM and Si-Quat versus abrasion cycle, and Figure 8B presents the optical images of BPAM (top) and Si-Quat (bottom) coated PE before and after abrasion. With the BPMA coated substrate, after 15 abrasion cycles, 99% of the dye remained on the PE surfaces (Figure 8A). Also, the abraded area of the BPAM


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coated substrates remains its original color. In order to exclude the possibility that the reduction of dye intensity is a result of breakage of electrostatic attraction between dye and QAM, the substrate was re-stained in BPB solution. The intensity of surface adsorbed dye was not recovered (increased by only 2%), confirming that the loss of the dye is due removal of the coating and not removal of the dye. In contrast, on the Si-Quat coated PE, the dye intensity decreases drastically after first three cycles (by 60%), and after 15 cycles only 5% of the coating remained on surface, with bare PE exposed. Upon re-staining, the dye intensity was not restored, ascertaining that the loss of the dye intensity on Si-Quat coated surface is attributed to the removal of the quaternary ammonium from the plastic surface. The abrasion resistances of the two QAM coated PVA samples were also determined, and likewise BPAM showed similar remarkable robustness against abrasion (Figure S4). After the first three cycles, the dye intensity deceases by 12 %, with no further decrease over the following 12 cycles. The initial prominent reduction is most likely a result of loosely or non-covalently bounded BPAM. However, with the Si-Quat, 70% was eliminated after abrasion. Therefore, it can be concluded that the surface attachment introduced by generation of new carbon bonds between the plastic and the BPAM polymer network during cross-linking is more robust than commercially available silane-based quaternary ammonium compounds. The latter compound requires a hydroxyl group on surface to undergo hydrolysis and condensation to form siloxane bonds. For surfaces such as PE and PP that lack -OH groups, silanebased material tends to crosslink only with itself and form a physisorbed layer on the substrate surface. Though densely crosslinked, this layer is not durable against severe



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mechanical stress. On the other hand, even for a thin layer, the BPAM coating leaves a surface with potent antimicrobial activity and excellent mechanical stability.

Figure 8. Plots of percentage of remaining BAPM (black squares) and Si-Quat (red circles) on PE over mechanical abrasion cycles (A) and optical images of the samples before and after 15 abrasion cycles.

The biocidal activity of the abraded substrates were further investigated using the bacterial aerosol test described above. Two PE substrates were spray-coated with BPAM and one of them was cured with UV light. Both BPAM treated substrates underwent 15 abrasion cycles with a load of 500 g along the midline, and were exposed to S. aureus inoculum and incubated. Figure 9 shows the resultant photographs after bacterial testing. As expected, bacterial colonies grew over the entire control substrate (see biofilm in Figure 9A). In direct contrast, the BPAM treated, UV cured, and abraded substrate (Figure 9B) maintained 100% killing efficacy, illustrating that the covalently bounded


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BPAM retains remarkable antimicrobial activity even after harsh mechanical stress. In an effort to further demonstrate the robustness of the covalent linkage, a BPAM treated substrate without UV curing (Figure 9C), was exposed to bacteria after abrasion. In this case, colonies were found in the area that endured abrasion, where the coating was removed.

Figure 9. Digital pictures of the PE substrates sprayed with S. aureus suspension and incubated overnight at 37 °C: (A) uncoated, (B) BPAM coated, cured and abraded, (C) BPAM coated, uncured and abraded (dotted box: abraded area).

Conclusions In conclusion, we have demonstrated that BPAM can be used as a potent biocide that can be photochemically grafted to surfaces containing C-H bonds. The formation of surface grafted polymeric networks is rapid (less than 1 min) when illuminated with very low UV intensity, attributed to electron-withdrawing ammonium substituent on the benzophenone chromophore. BPAM functionalized glass, plastic, and textile substrates provide coatings that are effective, non-leaching, and have rapid biocidal activity against S. aureus and E. coli on contact. Most importantly, although the BPAM coating is less than 50 nm, it resists harsh mechanical abrasion with retention of its high killing efficiency. The BPAM



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molecule itself is thermally stable (degradation temperature is above 140 °C) and chemically stable under ambient conditions. After the attached bacteria are killed by BPAM functionalized surface, the dead cells can be washed or wiped away using water, and the antimicrobial activity of the coatings restored. We have also insulted the BPAM functionalized surface with household cleaning products, such as Windex®, Clorox® and Glass Plus®. The surface-bound positive charges are not affected by the cleaners, implying that the antimicrobial activity remains after washing with common household cleaning solutions. It is noteworthy that washing with anionic detergent can decrease the antimicrobial efficacy as the negatively charge surfactants complex to the positively charged “-oniums” on surface via electrostatic interactions. The facile and economical synthesis, the versatile and robust surface attachment, and the outstanding antimicrobial activity of BPAM make it an ideal candidate for a wide array of potential domestic, medical and industrial coating applications.

Associated Content Supporting Information Available The following files are available free of charge on the ACS publications website at DOI: UV-Vis spectroscopy of AmBP, AFM with scan size of 10 µm, antimicrobial test of BPAM coated samples against E. coli, and abrasion test on PVA (PDF).

Author Information Corresponding Author


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*E-mail: [email protected]. Tel.: +1 706 542 2359. Fax: +1 706 542 3804.

ACKNOWLEDGMENT This work was supported by the Georgia Research Alliance through GRA Ventures program. Special thanks to Dr. Vincent J. Starai for use of the fluorescence microscope.

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Surface Grafted Antimicrobial Polymer Networks with High Abrasion

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