Investigation of Antibacterial 1,8-Cineole-Derived Thin Films Formed

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Investigation of Antibacterial 1,8-cineole-derived Thin Films Formed via Plasma Enhanced Chemical Vapor Deposition Michelle Mann, and Ellen R. Fisher ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b09067 • Publication Date (Web): 06 Oct 2017 Downloaded from http://pubs.acs.org on October 9, 2017

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ACS Applied Materials & Interfaces

Submitted to ACS-AMI Investigation of Antibacterial 1,8-cineole-derived Thin Films Formed via Plasma Enhanced Chemical Vapor Deposition Michelle N. Mann1 and Ellen R. Fisher1* 1

Department of Chemistry, Colorado State University, Fort Collins, Colorado 80523-1872

*Author to whom correspondence should be addressed: [email protected] Abstract The need for low-fouling coatings for biomedical devices has prompted considerable interest in antibacterial compounds from natural and sustainable sources, such as essential oils. Herein, a tea tree oil-based precursor, 1,8-cineole, is used to fabricate antimicrobial films (denoted ppCin) by plasma enhanced chemical vapor deposition. Film properties were comprehensively characterized using a variety of surface and bulk analytical tools and the plasma gas phase is assessed using optical emission spectroscopy, which can be correlated to ppCin film properties. Notably, film wettability increases linearly with plasma pressure, yielding water contact angles ranging from ~50° to ~90°. X-ray photoelectron spectroscopy reveals less oxygen is incorporated at higher pressures, likely arising from the lower density of OH(g) species. Further, we utilized H2O(v) plasma surface modification of the ppCin films to improve wettability and find this results in a substantial increase in surface oxygen content. To elucidate the role of film wettability and antibacterial properties, both as-deposited and H2O(v) plasma modified films were exposed to gram-negative Escherichia coli and gram-positive Staphylococcus aureus, using glass slides and hydrocarbon films deposited from 1,7-octadiene as positive controls. Overall, bacteria attach to a similar extent on all films, including controls, yet only essential oil-based films significantly prevent biofilm formation (4-7% coverage compared to ~40% for controls).

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Keywords: plasma enhanced chemical vapor deposition, 1,8-cineole, optical emission spectroscopy, plasma surface modification, biofilm formation, non-fouling surface, antifouling surface

Introduction Many polymers used in biomedical devices have limited biocompatibility because they promote bacterial adhesion and subsequent biofilm formation, ultimately leading to infection and device failure. Indeed, hospital acquired infections (HAIs) are a primary cause of increased morbidity and mortality in patients worldwide. In the United States alone, an estimated 1 out of 25 hospitalized patients are affected by an HAI at any given time, amounting to ~650,000 patients annually.1 Although difficult to accurately calculate the cost of infection-related hospitalization, low estimates place the cost of preventable HAIs at $5.7 – 6.8 billion.2 Approximately 80% of chronic infections are biofilm-related, with Escherichia coli (E. coli) and Staphylococcus aureus (S. aureus) the most frequently isolated strains from these infections.3 Broadly, bacterial infection presents a major challenge to developing advanced biomedical devices, and by extension, treating chronic infections while simultaneously controlling antibiotic resistance and reducing risk to patient health. This, in part, arises from the resilience of bacteria in a biofilm, as they are typically 10 – 1,000 times more resistant to antibiotics than planktonic bacteria.3 Thus, it is critical to develop tailored surfaces to control bacterial adhesion and growth on polymeric biomaterials before biofilms can form, without promoting antibiotic resistance. Ideally, an antibacterial coating could be deposited on the surface of a biomedical device, allowing for the bulk material to remain unmodified, yet discouraging bacterial growth. Alternatively, a biocidal agent could be incorporated in the polymer to actively kill bacteria. A holy grail in antibacterial coatings for biomedical devices is the creation of a surface modification platform to achieve mechan-

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ically and chemically robust films with highly tailorable surface properties for the desired application. One broad class of compounds being investigated for their medicinal properties are essential oils, many of which have antimicrobial properties supported by extensive clinical trials,4-8 but are difficult to immobilize by conventional coating methods. Researchers have employed a few techniques to fabricate solid materials with essential oils, including incorporating polysaccharide films with essential oils,9-11 microencapsulation,12-13 and plasma-enhanced chemical vapor deposition (PECVD). In PECVD, a nonthermal rf plasma induces radicalization in a monomeric feed gas, creating reactive chemical species that polymerize to form a smooth, conformal coating on a substrate.14 This technique allows for the fabrication of soft matter thin films from solid, liquid, or gaseous precursors where conventional methods may fail (e.g., monomers with only C-C functionality or ring structures). Plasma polymerized films from essential oils are often optically transparent and smooth, making them suitable for a range of applications, including medical coatings and dielectric layers for electronics.15-19 PECVD is ideal for depositing thin coatings on biomedical devices, as it provides a universal dry, single-step and sterile platform for device fabrication at low temperatures (2 h before and during analysis. For all spectra, atmospheric signals were suppressed and baseline was corrected using the onboard software (Omnic v8.2). Surface analysis For all surface characterization described herein, films were analyzed immediately after deposition (unless otherwise noted), and measurements were repeated for a minimum of n = 3 to gauge reproducibility of the plasma deposition processes. Water contact angle (WCA) was measured using a Krüss DSA30S contact angle goniometer (Matthews, NC, USA). A 2 µL drop of ultrapure water (Millipore, 18 mΩ cm) was applied to the film surface and the static WCA was measured using high-speed video recording for 10 s at 64 frames per second. Data were analyzed with onboard Krüss software; the circle fitting method was used for low WCA (>30°) and the tangent line fitting method used for higher WCA values.41 Deposited films (ppCin and ppOct) and H2O(v) plasma treated ppCin films were aged under ambient laboratory conditions to assess changes in WCA after 24 h, 1 week, 2 weeks, 1 month, and 2 months. X-ray photoelectron spectroscopy (XPS) was performed using a Physical Electronics PE5800 ESCA/AES system (Chanhassen, MN, USA) equipped with an Al Kα monochromatic X-ray source (1486.6 eV), hemispherical electron analyzer, and multichannel detector. Samples were


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mounted to the sample holder using double sided carbon tape. Spectra were collected using a 45° take off angle, and a low energy electron flood gun (5 – 15 eV) was used to minimize surface charging. Survey spectra were collected for 5 min over 50 – 1100 eV to obtain approximate elemental composition. High-resolution spectra were collected for all elements present in >1% amounts including C1s, O1s, and, in some cases, N1s. Three different spots on each sample were analyzed to elucidate surface homogeneity. CasaXPS software (Casa Software Ltd., Cheshire, UK) was used to process all high-resolution spectra with Gaussian-Lorentzian (30:70) fits and FWHM constrained to ≤2.0 eV.33 High-resolution C1s spectra were charge corrected by setting the C-C/C-H component to 285.0 eV for all samples.33, 39 Film stability The quality of film adhesion was assessed by testing film stability in biologically-relevant media for both glass and PS substrates. Each film was immersed in sterile physiological saline (10 mL; 0.85% w/v NaCl, certified ACS grade, Fisher Scientific, Fair Lawn, NJ, USA) and incubated at room temperature for 18 h, before carefully rinsing 3 times with copious amounts of Millipore water. After drying, films were then visually inspected to observe film integrity, and, if film presence was not visually clear, WCA analysis was performed to confirm or deny film presence/integrity. Biological assays Freshly prepared ppCin, H2O(v) plasma treated ppCin, and ppOct films, as well as the cineole and octadiene monomers, were analyzed for biocidal performance against both gram-negative E. coli and gram-positive S. aureus. Towards our goal of determining the role of wettability on bacterial attachment and growth, we chose to interface bacteria with two ppCin films of disparate wettabilities: (1) ppCin deposited at 100 mTorr and 100 W, denoted ppCin (high p,P) and (2)



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ppCin deposited at 15 mTorr and 50 W, denoted ppCin (low p,P). Films chosen for biological assessment not only had disparate wettabilities, but also varied in stability in biological media after 18 h of immersion. As such, all ppCin films reported herein were deposited for 5 min at ~10 cm downstream, conditions that resulted in films that remained stable after >1 week in physiological solution. All bactericidal activity assays were performed according to a modified protocol from the National Committee for Clinical Laboratory Standards.33, 42 Preparation of bacterial culture. E. coli (ATCC 25922) and S. aureus (ATCC 29213) were obtained from American Type Culture Collection (ATCC, Manassas, VA, USA). Lyophilized bacteria were grown in warm nutrient broth media (NBM, Oxoid™ OXCM0001B, Fisher Scientific, Fair Lawn, NJ, USA) overnight at 37 ○C and 150 rpm. The overnight culture was diluted 1:1 in 30% v/v glycerol (≥99.5%, Sigma-Aldrich, St. Louis, MO, USA) solution and stored at −80 ○C until use. For each trial, a tube of culture was thawed at room temperature and centrifuged at 4700 rpm for 10 min. The resulting pellet was dispersed in warm NBM and incubated overnight at 37 ○C and 150 rpm. The optical density at 600 nm (O.D.600nm) of the overnight culture was analyzed via UV-vis spectroscopy (Thermo Scientific, Genesys 20), and the culture was diluted with fresh warm NBM to an O.D.600nm of ~0.1 a.u. The culture was incubated at 37 ºC and 100 rpm until it reached the logarithmic growth phase (O.D.600nm ~0.3 a.u.) prior to exposing the bacterial culture to the films or liquid monomer. Bacterial attachment and growth. To determine the propensity for E. coli and S. aureus to attach to plasma polymerized films, bacterial attachment and growth assays were performed. In addition to ppCin and H2O(v) plasma treated ppCin films, ppOct films were used as a hydrophobic control and glass slides were used as positive, hydrophilic, controls. Films were placed in 12 well tissue culture (TC) plates (VWR, Arlington Heights, IL, USA). and 2 mL of prepared


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bacterial culture were added to each well. After 24 h of incubation at 37 °C, NBM was removed and each film was rinsed 3 times with sterile 0.85% w/v NaCl without being allowed to dry out. Propidium iodide (PI, 1.0 mg mL-1in water) and SYTO9 (5 mM in DMSO) fluorescence stains were purchased from Life Technologies Corporation (Carlsbad, CA). A working solution of stain was prepared with concentrations of 3 µM PI and 0.5 µM SYTO9; this solution was refrigerated between uses for a maximum of 1 month. Each sample was covered with 500 µL of staining solution containing PI and SYTO9 and allowed to incubate protected from light for 20 min before stain was removed and samples were rinsed 3 times with Millipore water. Films were air dried covered from light. Images were obtained using an Olympus IX73 fluorescence microscope and processed using Olympus CellSens software (v1.14). PI fluoresces red, with excitation and emission wavelengths of 543 nm and 617 nm, respectively, whereas SYTO9 appears green, with excitation and emission wavelengths of 488 nm and 500 nm, respectively. The redfluorescent PI stain intercalates with DNA and cannot permeate intact cell membranes; thus, PI only stains non-viable bacterial cells. In contrast, green-fluorescent SYTO9 enters both viable and non-viable cells. Images displayed herein depict signals from PI and SYTO9 overlaid to observe both viable and nonviable bacteria. Yellow signal sometimes arises when red and green filters overlap because SYTO9 was not completely displaced from the cell by PI, thus both stains remain in the cell, and we may assume the cells are nonviable.43 Minimal processing was performed when necessary to eliminate background auto-fluorescence of the film, likely arising from unsaturated carbon bonds within the plasma polymer.44 Biofilm formation. The use of viability assays in combination with biofilm formation assays allow for a more comprehensive assessment of antibiofouling behavior. Biofilm formation assays commenced in the same manner as the bacterial attachment assays described above, but the



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incubation period was extended to 5 days to allow for observation of biofilm growth. Warm NBM was replaced each day until the end of the assay, after which NBM was removed from each sample well and slides were rinsed 3 times with 2 mL of physiological saline solution, without being allowed to dry out. Crystal violet stain (750 µL, 1% aqueous solution, SigmaAldrich, St. Louis, MO, USA, diluted 1:10 in ultrapure water) was added to each well. After well plates were incubated at room temperature for 20 min, the stain was removed from each well, and samples were rinsed 3 times with copious amounts of Millipore water (taking care not to disturb film integrity). Finally, samples were dried at room temperature, and bright field images were obtained using an Olympus IX73 fluorescence microscope and processed using Olympus CellSens software (v1.14). ImageJ image analysis software (v1.6; National Institutes of Health, USA) was used to assess extent of biofilm surface coverage for control samples and films.33, 45 Bright field images were converted to 8-bit and binary format. The threshold was adjusted so that each pixel of the converted image accurately represented the original image. A minimum of 9 images were analyzed per sample type and the mean ± standard deviation is reported. Kill rate. 2 mL aliquots of bacterial culture in logarithmic growth phase were added to wells containing thin films deposited onto glass slides. To test the bactericidal activity of the cineole and octadiene monomers, the liquid monomer (0.5 % v/v) was added to vials containing 6 mL of bacterial solution in logarithmic growth phase (O.D.600nm ~0.3 a.u.) and vortexed to disperse the monomer. For all kill rate assays, bacterial culture (2 mL) was placed in empty wells as positive controls. Samples were placed in a static incubator at 37 ºC for the duration of the assay (72 h). Bacterial population was quantified as the number of colony forming units per mL (CFU mL-1) and monitored over time (2, 4, 24, and 72 h). At these times, 100 µL aliquots were removed


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from each well to undergo serial 10-fold dilutions to a factor of 106 or 107 using sterile 0.85% (w/v) NaCl. Aliquots (50 µL) of diluted bacterial solution were then dispensed onto nutrient agar (NA, OXCM0003B, Fisher Scientific, Fair Lawn, NJ, USA). Plates were left overnight in a static incubator at 37 ºC, counted the following day, and CFU mL-1 of bacterial culture was calculated using equation 1. colony forming units (CFU) volume (mL)

=

number of colonies dilution factor ⨯ plated volume (mL)

(1)

When no colonies were observed on plates, an undiluted aliquot was plated to confirm that no colonies grew. Only thereafter was the limit of detection of 1 CFU mL-1 applied.46-48 To determine the log reduction caused by the liquid monomer or the plasma polymerized films, the CFU mL-1 in each well was compared to that of the positive control.49 All biological assays were completed for n ≥ 9 and data are expressed as mean ± one standard deviation. Statistical significance was determined using a one-tailed Student’s t-test and considered at p < 0.05.

Results Gas phase spectroscopy Raw OES spectra for 100% cineole plasmas ignited under two different conditions are shown in Figure 1. In general, the plasmas contained similar emission signals regardless of the plasma parameters used. The most intense peaks in the plasmas appear at 282.4, 309.1, 485.9, and 656.1 nm, corresponding to emission from CO, OH, Hβ and Hα, respectively.50-52 Other spectral features include H2 molecular bands centered at ~610 and ~750 nm.53 We have previously established that steady state conditions in our systems are reached in 0.9) for all treatments except ppCin (low p,P ). Deposition rates vary from 5 ±

Investigation of Antibacterial 1,8-Cineole-Derived Thin Films Formed

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