Antimicrobial polymer coatings with efficacy against pathogenic and spoilage microorganisms

Antimicrobial polymer coatings with efficacy against pathogenic and spoilage microorganisms

LWT - Food Science and Technology 97 (2018) 546–554 Contents lists available at ScienceDirect LWT - Food Science and Technology journal homepage: ww...

1MB Sizes 0 Downloads 23 Views

LWT - Food Science and Technology 97 (2018) 546–554

Contents lists available at ScienceDirect

LWT - Food Science and Technology journal homepage: www.elsevier.com/locate/lwt

Antimicrobial polymer coatings with efficacy against pathogenic and spoilage microorganisms

T

Yu-Ting Hunga, Lynne A. McLandsborougha, Julie M. Goddardb, Luis J. Bastarracheac,∗ a

Department of Food Science, University of Massachusetts, Amherst, MA, 01003, USA Department of Food Science, Cornell University, Ithaca, NY, 14853, USA c Department of Nutrition, Dietetics and Food Sciences, Utah State University, Logan, UT, 84322, USA b

A R T I C LE I N FO

A B S T R A C T

Keywords: Antimicrobial coatings N-halamines Cationic polymers Food safety Food waste

Antimicrobial polymer coatings with inherent self-sanitizing properties have been explored to support food safety and preservation. Materials with multiple antimicrobial modes of action represent a novel alternative. Herein, we evaluated the antimicrobial effect and storage stability of coatings with varied molecular weight styrene maleic anhydride (SMA) cross-linkers (6, 8, 120, and 250 kDa) and branched polyethyleneimine (PEI) coated onto polypropylene films. Infrared spectroscopy analyses suggested successful crosslinking between all varieties of SMA and PEI. Coatings were evaluated in their inherent form (cationic), and chlorinated as Nhalamines. Surface concentration of primary amines ranged from 350 to 900 nmol/cm2, and N-halamine concentration ranged from 90 to 130 nmol/cm2, with values varying depending on SMA molecular weight. Surface energy decreased with increasing molecular weight of SMA. Optimal cross-linker molecular weight was determined based on antimicrobial performance, where the coated PP with 6 kDa SMA demonstrated enhanced biocidal effect against Escherichia coli O157:H7 in its chlorinated form. Further, the antimicrobial coating demonstrated efficacy between ∼3 and > 5 logarithmic reductions in its unchlorinated and chlorinated forms against Escherichia coli O157:H7, Listeria monocytogenes, and Pseudomonas fluorescens. Storage studies supported the stability of the chlorinated N-halamines, with full chlorine retention over a 24 h study.

1. Introduction Reducing microbial cross-contamination of pathogenic and spoilage microorganisms from food contact surfaces remains a significant challenge in the food industry. Microorganisms are capable of colonizing solid surfaces and form stable biofilms, which support their viability and growth (Bower, McGuire, & Daeschel, 1996; Di Ciccio et al., 2012; Glinel, Thebault, Humblot, Pradier, & Jouenne, 2012; Kumar & Anand, 1998; Simoes, Simoes, & Vieira, 2010; Srey, Jahid, & Ha, 2013), indicating that food contact surfaces in food processing environments such as containers, working benches, or conveyor belts, are all capable of harboring and potentially transferring microorganisms to the final food product. Contact surfaces contaminated with microorganisms may lead to serious public health problems, which are estimated to cause 48 million illnesses, and of these, 3000 deaths, annually (Centers for Disease Control and Prevention (CDC), 2011). Further, contamination by spoilage microorganisms can reduce product shelf life, contributing to the volume of food wasted due to microbial spoilage (Floros et al., 2010). To reduce the likelihood of microbial cross-contamination from



food contact materials, both physical and chemical strategies are employed in cleaning and sanitation (Kumar & Anand, 1998; Simoes et al., 2010), including application of ultrasound (Baumann, Martin, & Feng, 2009), irradiation (Bintsis, Robinson, & Litopoulou-Tzanetaki, 2000; Mikš-Krajnik et al., 2015), and application of disinfectants like quaternary ammonium compounds (Aase, Sundheim, Langsrud, & Rørvik, 2000; Buffet-Bataillon, Tattevin, Bonnaure-Mallet, & Jolivet-Gougeon, 2012), hydrogen peroxide, and chlorine-based compounds (Aarnisalo et al., 2000; Rutala & Weber, 2008). Yet, despite the implementation of well-established cleaning protocols, clean surfaces can be immediately soiled by a contaminated product, causing cross-contamination to remain a major issue in maintaining food safety and quality. Hence, the application of antimicrobial coatings on food contact materials presents an opportunity to reduce microbial contamination and further support current sanitation practices in food processing environments. Antimicrobial coating materials can reduce, inhibit, or retard microbial growth upon interaction with internal environment or surface contact (Appendini & Hotchkiss, 2002; Bastarrachea, Denis-Rohr, & Goddard, 2015; Bastarrachea, Wong, Roman, Lin, & Goddard, 2015). A

Corresponding author. Department of Nutrition, Dietetics and Food Sciences, Utah State University, 8700 Old Main Hill, Logan, UT, 84322, USA. E-mail address: [email protected] (L.J. Bastarrachea).

https://doi.org/10.1016/j.lwt.2018.07.046 Received 12 April 2018; Received in revised form 11 July 2018; Accepted 24 July 2018 Available online 25 July 2018 0023-6438/ © 2018 Elsevier Ltd. All rights reserved.

LWT - Food Science and Technology 97 (2018) 546–554

Y.-T. Hung et al.

demonstration of the rechargeable characteristic of N-halamines and retention of antimicrobial properties in its unchlorinated form via cationic polymer components (Bastarrachea & Goddard, 2015, 2016). The proposed antimicrobial coatings exhibited the capability of retaining biocidal efficacy after 10 chlorination cycles, showed promising results when challenged in the presence of organic matter, and importantly exhibited no net positive charge. However, as mentioned above, one of the major drawbacks of these antimicrobial technologies lies within the stability of the bond linking the N-halamine moieties to the surface of the coating substrate, which may weaken upon repeated or continuous exposure to halogen sources and can manifest in hydrolysis of the chemical bonds that integrate the coating (Bastarrachea & Goddard, 2015). Introduction of higher molecular weight cross-linking agents with different chemical compositions into the coating system represents a way to modify surface properties. Styrene maleic anhydride (SMA), a polymeric anhydride composed of styrene and maleic anhydride units can cross-link polyethylenimine (PEI) to fabricate antimicrobial polymer coatings due to the high reactivity of the anhydride units (Bastarrachea & Goddard, 2015, 2016). The molecular weight of SMAs with varying maleic anhydride content has been observed to influence rheology and phase behaviors in polymer blends (Chopra, Hatzikiriakos, Kontopoulou, & Vlassopoulos, 2002). Thus, implementation of higher molecular weight SMA with their different chemical composition as cross-linking agents may potentially improve surface properties of the coating system, as high molecular weight molecules imply more active sites for interaction between the component layers. In the present work, we hypothesized that cross-linker molecular weight may influence the surface properties and antimicrobial character of the resulting antimicrobial polymer coating. Therefore, an adaptation of a previously established coating fabrication protocol was followed in this study to determine an optimal combination of crosslinker molecular weight for the proposed antimicrobial polymer coating (Bastarrachea & Goddard, 2016). The surface properties of the coating were characterized, which was followed by an antimicrobial screening assay to evaluate biocidal efficacy. An optimal SMA molecular weight was selected for the coating system based on antimicrobial performance, and was further challenged against pathogenic and spoilage microorganisms including Escherichia coli O157:H7, Listeria monocytogenes, and Pseudomonas fluorescens, in both logarithmic reduction and inactivation kinetics studies, and subjected to an atmospheric storage study to determine practical applicability in the food industry.

number of antimicrobial coating materials have been proposed to address contamination issues in the food industry. Among these, incorporation of antimicrobial substances in the coating matrix which exert their biocidal effect by migration of the antimicrobials represents a main strategy that has been explored (Hanušová et al., 2010; Lee, An, Lee, & Park, 2003; Lee, An, Lee, Park, & Lee, 2004). Although studies have demonstrated the antimicrobial effectiveness of these types of coatings, reliance on leaching of the antimicrobials indicates eventual loss of antimicrobial activity. In addition, active compounds that migrate into food are regulated as direct food additives (Part 170 Title 21 CFR), which limits the range of migratory antimicrobial materials that can be adopted into actual industrial practice (Appendini & Hotchkiss, 2002; Hanušová et al., 2010). A wide range of antimicrobial compounds have been coated on inorganic and plastic surfaces, including metalbased compounds, quaternary ammonium compounds, polycations, essential oils, peptides, and light-activated compounds. Bacteria, molds and yeasts are susceptible to various antimicrobial compounds, with mechanisms of killing occurring primarily through damage to cell membranes and essential biomolecules (Bastarrachea, Denis-Rohr, et al., 2015). More recently, non-migratory N-halamine based polymeric materials have been receiving considerable attention due to their rechargeable antimicrobial character and their efficacy without leaching. N-halamines are antimicrobial moieties characterized by nitrogen functional groups covalently bound to halogens, mostly chlorine (Bastarrachea, Denis-Rohr, et al., 2015; Dong, Wang, Gao, Gao, & Gao, 2017; Hui & Debiemme-Chouvy, 2013; Williams, Elder, & Worley, 1988; Worley & Sun, 1996). Chlorination of the nitrogen atoms in amine, amide, and imide structures establishes the formation of antimicrobial N-halamine moieties, which are then able to regenerate antimicrobial character upon each exposure to a chlorine source, providing long term activity (Qian & Sun, 2003). N-halamines have been reported to exert antimicrobial activity towards a wide range of microorganisms including bacteria, fungi, viruses, and prevent bacterial biofilm formation (Cao & Sun, 2009; Hui & Debiemme-Chouvy, 2013). While their exact inactivation mechanism is not yet confirmed, it is generally agreed that two modes of action are involved, including a direct oxidation of microbial membrane from the bound halogens, and penetration of dissociated halogens, which disrupts vital biomolecules within the microorganism (Chen, Luo, & Sun, 2007; Worley & Williams, 1988). In terms of food-related applications, N-halamine materials represent an opportunity for long-term usage as antimicrobial food contact materials, where their activity can be regenerated via exposure to common chlorine-based sanitizers (e.g. bleach). However, some potential drawbacks, including lack of antimicrobial property in their unchlorinated form, and their chemical integrity and stability after repeated chlorination, remains a significant challenge. Cationic polymers are inherently antimicrobial due to their positive charge, which enables it to retain antimicrobial efficacy after its immobilization onto a substrate matrix. Inactivation mechanism of these antimicrobials is attributed to ionic exchange between the positive charges of the cationic polymer and the critical cations within the microbial membrane, resulting in a loss of membrane integrity and cell disruption (Kügler, Bouloussa, & Rondelez, 2005; Lichter & Rubner, 2009). Cationic polymers have been reported to demonstrate broad spectrum efficacy, exhibiting biocidal effects against Gram positive and Gram negative bacteria, yeasts, and fungi. Yet, despite the reported antimicrobial effectiveness, their cationic nature also makes them more susceptible to fouling by anionic compounds in organic matter (Bastarrachea, Denis-Rohr, et al., 2015). This potential for fouling could further promote bacterial adhesion and establish biofilm formation on the surface of the coating material, which increases the risk of crosscontamination, and represents a major hurdle for application in food contact materials, as food and other biological systems often contain a range of anionic compounds. Previous work has reported successful incorporation of both N-halamine and cationic moieties into the same coating system, with

2. Materials and methods 2.1. Materials Polypropylene (PP) pellets and 6 kDa styrene maleic anhydride (SMA, 50% weight of maleic anhydride) copolymer were purchased from Scientific Polymer Products (Ontario, NY, USA). Molecular weights of 80 kDa (26% weight of maleic anhydride), 120 kDa (26% weight of maleic anhydride), and 250 kDa (8% weight of maleic anhydride) SMA copolymer were purchased from Polyscope (Netherlands). Branched polyethyleneimine (PEI), 2-ethoxy-1-ethoxycarbonyl-1,2-dihydroquinoline (EEDQ) were purchased from SigmaAldrich (St. Louis, MO, USA). 2-(N-Morpholino)ethanesulfonic acid sodium salt (MES) was purchased from GenScript Inc. (Piscataway, NJ, USA). Orange (II) dye (AO7) and sodium hypochlorite solution were purchased from Acros Organics (Fair Lawn, NJ, USA). N,N-Diethyl-pphenylenediamine total chlorine reagent powder (DPD) was purchased from Hach Co. (Loveland, CO, USA). Tryptic soy broth (TSB), Tryptic soy agar (TSA), and neutralizing buffer were purchased from Becton, Dickinson and Company (MD, USA).

547

LWT - Food Science and Technology 97 (2018) 546–554

Y.-T. Hung et al.

2.2. Preparation of PP films

anhydride groups. The surface chemistry and the preparation steps of the reported polymer coating are illustrated in Fig. 1.

PP pellets were cleaned via sonication first with isopropanol, then acetone, followed by deionized water, with two cycles of 10 min cleaning applied for each solution. Cleaned PP pellets were dried and maintained in a desiccator over anhydrous calcium sulfate for 12 h, and then pressed into films on a hot press (Carver Inc., NJ, USA) at 180 °C with a force load of 9000 psi. Pressed PP films (thickness: 0.5 ± 0.1 mm) were cut into 2 × 2 cm squares, and cleaned and dried again following the same cleaning protocol as applied to the PP pellets.

2.5. Attenuated total reflectance fourier transform infrared spectroscopy (ATR-FTIR) Surface chemistry of native and modified PP films was analyzed with an IRTracer-100 spectrometer (Shimadzu Corp., Tokyo, Japan) equipped with a diamond ATR crystal. For each set of prepared films, two spots were measured for their absorbance in each sample film for at least 5 randomly selected films. FTIR analysis was conducted using Happ-Genzel apodization at a resolution of 4 cm−1 and with a total of 32 scans applied for each measurement. The obtained spectra then underwent base-line correction and smoothing, and were analyzed with KnowItAll software (Biorad Laboratories, Philadelphia, PA, USA).

2.3. Surface activation of PP The surface of PP films was first activated to facilitate the binding of the polymer coating. Squares of PP (2 × 2 cm) were treated with UVOzone (UVO) irradiation for 15 min on one side using a Jelight Co. Model 42 UVO Cleaner (Irvine, CA, USA), which promotes the formation of carboxylic acids through photo-oxidation. The UVO-treated PP coupons were then subjected to a reaction with EEDQ to create surface anhydride groups, further making the PP surface more reactive towards nucleophilic attack by primary amines groups present in PEI. Briefly, a 0.1 mM solution of EEDQ in 50 mM MES buffer (pH 5.5) was prepared with 50 mL of solution for every UVO-treated coupon (final volume in mL = number of coupons × 50 mL/coupon). This was done by dissolving the appropriate amount of EEDQ first in methanol (1% of final volume), before mixing into the final volume of pH-adjusted MES buffer to make the EEDQ solution. UVO-treated PPs were then immersed and stirred in the EEDQ solution for 2 h followed by rinsing with 1% methanol solution and drying under compressed house air.

2.6. Primary amine and N-halamine quantification The amount of primary amines on the polymer coating surface was determined by the Acid Orange 7 (AO7) colorimetric assay (Uchida, Uyama, & Ikada, 1993), in which it is assumed that the –SO3 functional group of the dye complexes with primary amine units in a 1:1 ratio. Squares of 2 × 2 cm modified PP films were cut into 1 × 1 cm coupons, which were then individually immersed in a 1 mM solution of AO7 dye adjusted to a pH of 3.0 by HCl, and shaken for 3 h. After rinsing in copious pH 3.0 water to remove the unbound dye, the films were shaken in 5 mL of pH 12.0 deionized water (adjusted with NaOH) for 15 min to desorb the bound dye. Absorbances were read at 455 nm, and primary amine content was quantified by comparison to a standard curve prepared with varied concentrations of AO7 in pH 12.0 water. Native PP films treated in the same manner served as negative controls. Assays were performed with at least two replicates (1 × 1 cm coupons) from three separately prepared sets of coated films. N-halamine content of the polymer coating was quantified by a colorimetric DPD assay to measure surface chlorine capacity. Modified PP coupons (1 × 1 cm) were individually exposed to 200 ppm of chlorine prepared from sodium hypochlorite solution, of which the chlorine content was confirmed through standardization by iodometric titration (American Society for Testing and Materials, 2008). After 1 h of chlorination, the chlorinated sample films were rinsed with copious water to remove unbound chlorine, and transferred to individual test tubes with 2 mL of deionized water and 50 μL of DPD reagent (prepared

2.4. Spin coating assembly of polymer coating The three-layer polymer coating was then assembled onto the anhydride functionalized PP coupons by depositing two layers of PEI cross-linked by a layer of SMA using an adaptation of a previously reported method (Bastarrachea & Goddard, 2016). PEI and SMA solutions were prepared with acetone at concentrations of 0.06 g/mL and 0.04 g/ mL, respectively, and sonicated until the polymers were fully dissolved. The polymer solutions were then applied by spin-coating alternating layers of PEI and SMA in the order of PEI, SMA, and PEI, which was operated at 3000 rpm for 1 min per layer. Coated films were cured at 165 °C for 20 min to enable cross-linking between the amines and the

Fig. 1. Schematic of the preparation steps of the antimicrobial N-Halamine and cationic coating. 548

LWT - Food Science and Technology 97 (2018) 546–554

Y.-T. Hung et al.

suspensions were serially diluted first with neutralizing buffer to quench any chlorine present, followed subsequently by 0.9% saline water. Serial dilutions were then plated out in duplicates on TSA plates. To lower the limit of detection to 1 log (CFU/mL), 1 mL of the first dilution was plated onto 3 TSA plates (333 μL per plate). The TSA plates were incubated at the corresponding growth temperature (37 °C for E. coli and L. monocytogenes, and 28 °C for P. fluorescens) for 48 h, and the number of survivors were determined via plate count. Inactivation kinetic assays were conducted three times on different days with independently prepared sets of films, and a representative of the three kinetic curves is presented in the data results.

by mixing 1 packet of DPD total chlorine reagent powder with 1 mL of deionized water). The tubes were shaken for 5 min to allow color formation, and the absorbances were read at 512 nm thereafter. N-halamine content was determined by comparison to a standard curve prepared with varied solutions of sodium hypochlorite in deionized water. Native PP films exposed to 200 ppm chlorine served as negative controls. Disposable test tubes were used throughout the assay to minimize chlorine demand. The assays were performed with at least two coupons (1 × 1 cm) from three separately prepared sets of coated films. 2.7. Water contact angles and surface energy Hydrophobicity of the native and modified PP coupons was evaluated using advancing water contact angles measured with a DSA100 Drop Shape Analyzer (Krüss, Hamburg, Germany). A drop of 5 μL HPLC grade water was dispensed onto the sample films at a rate of 25 μL/min on a 1 × 2 cm film. Measurement data and images were obtained through the Drop Shape Analysis software (Krüss, Hamburg, Germany). For each type of sample, at least two different spots on a 1 × 2 cm film were tested from three separately prepared sets of films. Surface energy of the polymer coating was calculated using the Zisman plot method (Gindl, Sinn, Reiterer, Tschegg, & Gindl, 2001; Kabza, Gestwicki, & McGrath, 2000). Advancing contact angles were obtained in the same protocol as mentioned above, with the following liquids of different surface tension values: water (72.8 mN/m), ethylene glycol (47.7 mN/m), diethylene glycol (44.8 mN/m), and acetone (25.8 mN/m). The measurements and images were obtained through the Drop Shape Analysis software (Krüss, Hamburg, Germany). At least two different spots on a 1 × 2 cm2 film were tested from three separately prepared sets of films for each type of PP and liquid. Cosine values of the obtained advancing contact angles were then plotted against the surface tension value of the corresponding liquid and fitted to a linear regression curve with GraphPad software (GraphPad Software Inc., La Jolla, CA, USA). Surface energy of the native and modified PP films were determined when the advancing contact angle equals 0, giving a cosine value of 1.

2.9. Agar overlay assay To determine if viable bacteria had attached onto the surface of the coating during the antimicrobial evaluation, an agar overlay assay was performed (Tiller, Liao, Lewis, & Klibanov, 2001). In brief, the modified PP coupons were collected after assessing their antimicrobial activity, and rinsed first with neutralizing buffer, followed by three rinses with sterile deionized water to clean and remove any unbound bacteria. The cleaned coupons were then placed individually into sterile petri dishes, and TSA was poured onto the sample films. Plates were incubated at the corresponding growth temperature for each microorganism (37 °C for E. coli and L. monocytogenes, and 28 °C for P. fluorescens). At least two sample coupons (from both unchlorinated and chlorinated treatments) challenged against the selected bacteria were tested for each antimicrobial assay performed. After 24–48 h of growth, the TSA plates with sample coupons were taken out and observed for presence of bacteria, and coupons were imaged using scanning electron microscopy as described below. 2.10. Scanning electron microscopy (SEM) SEM was used to characterize the presence or absence of bacteria attached on the coating surface following antimicrobial activity assays. Modified PP coupons, both unchlorinated and chlorinated, were rinsed three times with sterile deionized water to remove unbound bacteria after antimicrobial evaluation. Coupons were then immersed in absolute ethanol to fix any bound bacteria on the surface and left to dry. The films were sputter-coated with gold using a Cressington Sputter Coater 108auto (Ted Pella, Inc., Redding, CA, USA) under argon for 30 s. Sample films were examined with a scanning electron microscope JCM6000 NeoScope (JEOL, Japan), and at least 9 images were captured at locations distributed across a 3 × 3 grid pattern on each of triplicate samples.

2.8. Microbial inactivation kinetics study Antimicrobial activity of the modified PP coupons was evaluated through inactivation kinetics against Escherichia coli O157:H7 ATCC 43895 (provided by Prof. Lynne McLandsborough, Department of Food Science, University of Massachusetts Amherst), Listeria monocytogenes Scott A, and Pseudonomas fluorescens FSL W5-0203 (both provided by Prof. Martin Wiedmann, Department of Food Science, Cornell University). E. coli and L. monocytogenes were cultured as follows based on previous protocols (Bastarrachea & Goddard, 2015, 2016). Briefly, a single colony of tested bacteria was inoculated in TSB and incubated for 16 h at 37 °C under shaking (125 rpm) overnight to reach stationary phase. A 1% dilution of this bacterial suspension was then prepared with fresh TSB, and incubated at 37 °C again until the bacteria reached mid-exponential phase (2 h for E. coli and 4 h for L. monocytogenes). The resulting broth was diluted 1000 fold with sterile deionized water for a starting inoculum bacterial concentration of ∼6 log (CFU/mL). P. fluorescens was cultured following the same method at an incubation temperature of 28 °C and 300 rpm of shaking to reach mid-exponential phase (∼6 h) after the initial 1% dilution with TSB. The mid-exponential broth was diluted 1000-fold with sterile deionized water to reach ∼6 log (CFU/mL) bacterial suspension. The aqueous suspensions of each bacterium were used for the antimicrobial kinetics study as follows. For each type of bacteria and native or coated PP, four 1 × 1 cm2 coupons were submerged in multiple individual test tubes with 1 mL of aqueous bacterial suspension. The test tubes were then incubated at 32 °C with 60 rpm rotation for E. coli and L. monocytogenes, and 28 °C for P. fluorescens. Test tubes were randomly taken for each bacterium at different points in time for up to 2 h and the bacterial

2.11. Stability of chlorinated N-halamines under storage Stability of the N-halamine moieties were evaluated through a 24 h storage study. Coupons of 1 × 1 cm2 modified PP were individually chlorinated as described above with 200 ppm of chlorine. The chlorinated coupons were then dried with an air gun, and stored at 4 °C and room temperature (∼22 °C), respectively. At different time intervals, the sample films were taken out, and their chlorine capacity was determined with the DPD assay as previously described. For this storage study, at least two sample films from three separately prepared sets of films were tested for each temperature condition. 2.12. Statistical analysis When appropriate, analysis of variance (ANOVA) and Tukey's pairwise comparisons were applied with a 95% confidence interval to determine significant differences between treatments using GraphPad Prism 6 software (GraphPad Software Inc., La Jolla, CA, USA). 549

LWT - Food Science and Technology 97 (2018) 546–554

Y.-T. Hung et al.

Fig. 2. ATR-FTIR spectra of (A) native PP, and PEI-SMA-PEI polymer coatings on PP prepared with (B) 6 kDa SMA, (C) 80 kDa SMA, (D) 120 kDa SMA, (E) 250 kDa SMA crosslinkers.

3. Results and discussion 3.1. Surface analysis of antimicrobial polymer coating ATR-FTIR spectroscopy was used to characterize the surface chemistry of native PP and modified PP (denoted as PP-PEI-SMAMWPEI, where MW indicates molecular weight, Fig. 2). Native PP coupons demonstrated absorbance bands at 2980–2820 cm−1 and ∼1400 cm−1, which are all characteristic of alkane groups from the propylene carbon backbone (C–H bonds). After applying the polymer coatings onto PP films, all PP-PEI-SMAMW-PEI coupons exhibited similar infrared spectra. A noticeable wide absorbance band was detected for each type of polymer coating at around 3600–3000 cm−1, which is most likely contributed to the vibration of the hydroxyl bond from carboxylic acids (3400–3200 cm−1) introduced through the surface activation of PP, and partially by the N–H bond from amines (3500–3300 cm−1) and amides (3320–3270 cm−1). After the curing process to enable crosslinking between SMA and PEI, a more prominent band was observed at ∼1650 cm−1, which can be attributed to the N–H bond from amides (1680–1630 cm−1), and the C=O bond from both amide and imide groups (1670–1630 cm−1). This suggests successful covalent bond formation between the maleic anhydride from SMA and the primary amines of PEI, confirming that the curing step promotes cross-linking. Indeed, spectra of all coating variants (prepared with different molecular weight SMAs) were similar.

Fig. 3. Primary amine and N-halamine content of PEI-SMAMW-PEI coatings prepared with 6, 80, 120, and 250 kDa SMA cross-linkers. PP-PEI-SMAMW-PEI with the same letters are not significantly different (P > 0.05). Values represent means of 6 determinations with error bars indicating standard deviation.

there appeared to be a practical difference between these and that of the 250 kDa SMA, which may explain the observed deviation. Between PEI-SMA6-PEI, PEI-SMA80-PEI, and PEI-SMA120-PEI, the increasing primary amine content may be attributed to the lower ratio of maleic anhydride with higher molecular weight SMA, implying a lower degree of cross-linking between the polymer layers. Chlorine capacity of the coatings was determined via DPD dye assays after the PEI-SMAMW-PEI coatings were exposed and chlorinated with 200 ppm chlorine (Fig. 3). This concentration was chosen because federal regulations (Part 178 Title 21 CFR) limits the use of higher chlorine concentrations (> 200 ppm available chlorine) when sanitizing food contact surfaces and processing equipment. No measurable amount of N-halamines was observed in native PP chlorinated under those conditions, while the PEI-SMAMW-PEI coatings with 6, 80, 120, and 250 kDa SMA cross-linkers exhibited 133.3 ± 14.9, 134.6 ± 14.2, 87.7 ± 9.4, and 115.7 ± 3.9 nmol of N-halamines per cm2, respectively. Although statistical differences were observed, there appears to be no practical difference between coating variants, which exhibited an equivalence of 1.5–2.5 ppm of chlorine. These results supported the ATR-FTIR analysis which indicates successful introduction of cationic and N-halamine antimicrobial moieties in the polymer coating. Hydrophobicity of the polymer coating was quantified via water contact angle (Table 1). As expected, native PP exhibited a hydrophobic surface (θA = 107.8 ± 2.3°) due to its nonpolar hydrocarbon backbone, which provides inherent hydrophobicity (Farris et al., 2014). Comparable results of advancing water contact angles have been

3.2. Primary amine and N-halamine quantification The PEI-SMAMW-PEI polymer coatings were further characterized using AO7 dye assay to quantify the surface concentration of primary amines (Fig. 3). No measurable primary amines were detected on the surface of native PP, while the PEI-SMAMW-PEI coatings prepared with 6, 80, 120, and 250 kDa SMA cross-linkers contained 351.8 ± 37.3, 750.1 ± 64.7, 941.9 ± 147.2, and 348.9 ± 53.9 nmol of primary amines per cm2, respectively. It is interesting to note that although the concentration of applied PEI (which provides the cationic amine groups) is constant for each molecular weight of SMA, statistically significant difference exists between the different molecular weight cross-linker coatings, with PEI-SMA6-PEI and PEI-SMA250-PEI showing comparable amounts of primary amine. A correlation was observed with PEI-SMA6-PEI, PEI-SMA80-PEI, and PEI-SMA120-PEI, where primary amines increased with higher molecular weight cross-linker used. According to the provider's technical information, SMA with 6, 80, and 120 kDa had similar maleic anhydride to styrene molar ratios, while 550

LWT - Food Science and Technology 97 (2018) 546–554

Y.-T. Hung et al.

Table 1 Advancing water contact angles and surface energy of PEI-SMAMW-PEI polymer coatings prepared with 6, 80, 120, and 250 kDa SMA cross-linkers. Samples with the same letters for each column are not significantly different (P > 0.05). Values represent means of 6 and 3 determinations for contact angles and surface energy studies respectively, with error bars indicating standard deviation. Sample

θA (Degrees)

Critical surface tension (mN/m)

Native PP PP-PEI-SMA6-PEI PP-PEI-SMA80-PEI PP-PEI-SMA120-PEI PP-PEI-SMA250-PEI

107.8 ± 2.3A 88.8 ± 1.3AB 82.8 ± 3.7B 83.9 ± 3.6B 83.9 ± 3.8B

20.7 23.3 21.8 20.7 19.5

± ± ± ± ±

0.9A 0.2A 3.7A 0.8A 1.8A

reported for PP in prior work (Denis-Rohr, Bastarrachea, & Goddard, 2015). An increase in hydrophilicity was observed for the modified PP coupons prepared with varied molecular weight SMA cross-linkers (PPPEI-SMAMW-PEI). This is likely attributed to the surface activation of PP, a necessary step to enhance surface reactivity of polyolefin (Singh, 1992), which introduced polar carboxylic groups. In addition, amine groups from PEI also introduced more polarity. While coated PP presented significantly higher hydrophilicity compared to native PP, there was no statistical difference between contact angle values of coatings prepared with different cross-linker molecular weight, suggesting that SMA molecular weight does not significantly affect wetting behavior. Zisman's plot approach was used to calculate surface energy of the polymer coating, by extrapolation from a linear regression curve plotted with surface tensions of different liquids and cosine values of their corresponding contact angles. According to this methodology, surface tension of the PEI-SMAMW-PEI coatings is defined as that of a liquid when it is just able to spread completely across the film surface (θ = 0, as in cosθ = 1) (Combe, Owen, & Hodges, 2004; Owens & Wendt, 1969). Critical surface tension values (an empirical approximation of surface energy) of native PP and PP-PEI-SMAMW-PEIs are presented in Table 1, with native PP showing low surface energy values, in agreement with reported literature (Bastarrachea & Goddard, 2015; Navaneetha Pandiyaraj, Selvarajan, Deshmukh, & Gao, 2008). No significant difference (P > 0.05) was observed between the values of critical surface tension of native PP and the different PP-PEI-SMAMWPEI treatments, in spite of the increasing number of styrene units with increasing molecular weight of SMA, which was expected to bring more hydrophobicity to the modified PP coupons. It is worth noting that PPPEI-SMAMW-PEI still demonstrated critical surface tension values similar to native PP despite the introduction of higher energy groups (amine, hydroxyl, and carboxyl groups) onto the surface, indicating the importance of low polar styrene groups from SMA. 3.3. Microbial inactivation kinetics study

Fig. 4. Microbial inactivation kinetics study. Filled markers represent unchlorinated PP-PEI-SMA6-PEI, and empty markers represent chlorinated PP-PEISMA6-PEI. Dashed lines represent the limit of detection.

Inactivation kinetic studies were conducted with PP-PEI-SMA6-PEIs against E. coli O157:H7, L. monocytogenes, and P. fluorescens, as model pathogenic and spoilage microorganisms. PP-PEI-SMA6-PEIs was used for the microbial inactivation kinetics study because this MW of SMA showed more reproducibility (data not shown). P. fluorescens is an obligate aerobe, and was cultured at 28 °C under intensive shaking (300 rpm) to ensure sufficient aeration (Van Tassell et al., 2012). As shown in Fig. 4, chlorinated PP-PEI-SMA6-PEI inactivated E. coli O157:H7 below the limit of detection, providing a > 5 log reduction (> 99.999%) after 45 min of contact. An approximate 3 log reduction (∼99.9%) was observed for the coated PP after 30–45 min in its unchlorinated state. Biocidal functions of N-halamines and cationic polymers both work by targeting the membrane of microorganisms (Bastarrachea, Denis-Rohr, et al., 2015). Literature has reported N-halamines to exhibit two modes of inactivation, including a direct oxidation of vital components through microbial cell membrane upon contact with the intact N-halamines, or via penetration from the

dissociated free chlorine which disrupts cell functionality (Hui & Debiemme-Chouvy, 2013; Liang et al., 2006). On the other hand, cationic polymers exert antimicrobial activity via an ionic exchange between its positive charges and that of the cell membrane surface, causing destabilization (Gilbert & Moore, 2005; Ikeda, Hirayama, Yamaguchi, Tazuke, & Watanabe, 1986; Lichter & Rubner, 2009). According to our results, inactivation kinetics of the unchlorinated and chlorinated PP-PEI-SMA6-PEI showed similar behavior, demonstrating an initial fast decrease in microbial population which then plateaus. For the unchlorinated PP-PEI-SMA6-PEI, this is in accordance with kinetics observed by other studies (Milovic, Wang, Lewis, & Klibanov, 2005), which also indicated biocidal properties imparted by the cationic moieties of the polymer coating. Chlorinated PP-PEI-SMA6-PEI required a longer contact time (45 min) to reach a plateau during its inactivation 551

LWT - Food Science and Technology 97 (2018) 546–554

Y.-T. Hung et al.

also non-viable.

as compared to its unchlorinated state (30–45 min). One possible explanation might be because the dissociation of free chlorine and subsequent penetration of cells occurred at a slower rate in contrast to unchlorinated films, which draws bacteria to the surface faster due to opposite charges (Kügler et al., 2005). Both unchlorinated and chlorinated PP-PEI-SMA6-PEI were able to achieve a > 5 logarithmic reductions (> 99.999%) in L. monocytogenes within the observed 2 h time frame (Fig. 4). However, in contrast to the results observed for E. coli O157:H7 inactivation, in which chlorinated coatings demonstrated enhanced biocidal activity over the unchlorinated form, unchlorinated PP-PEI-SMA6-PEI coatings inactivated L. monocytogenes below limit of detection in 15 min, while chlorinated PP-PEI-SMA6-PEIs took 30 min to obtain the same reduction level of > 5 logarithmic reductions. It was expected that the chlorinated coating, with both cationic and N-halamine characters, would provide a more pronounced antimicrobial effect than its unchlorinated counterpart as was observed with E. coli O157:H7. Yet susceptibility to antimicrobials is highly dependent on the nature of each type of microorganism (Timofeeva & Kleshcheva, 2011). The observed results may be a result of Gram positive bacteria being generally more susceptible to antimicrobials due to lack of an outer membrane, which limits penetration of antibacterial substances into the cell (Maillard, 2002). As a result, unchlorinated coatings were more effective against L. monocytogenes than their chlorinated counterparts. Inactivation kinetics against P. fluorescens are presented Fig. 4. Both unchlorinated and chlorinated PEI-SMA6-PEI coatings exhibited similar kinetic behavior, reducing microbial populations by ∼3 logarithmic cycles (99.9%) by a gradual reduction over 2 h. As mentioned earlier, different types of bacteria may react differently towards the same antimicrobial (Maillard, 2002; McDonnell & Russell, 1999). The tested P. fluorescens strain was isolated from cheese curds in a commercial cheese manufacturing plant (Martin, Murphy, Ralyea, Wiedmann, & Boor, 2011; Trmcic, Martin, Boor, & Wiedmann, 2015), which may explain its durability against chlorine, as sodium hypochlorite is a common food sanitizer in industrial practice.

3.5. Stability of chlorinated N-halamines under storage In addition to effective antimicrobial functionality, stability and consistency of its activity is equally important in terms of practical applications in the food manufacturing environment. The U.S. Food and Drug Administration established several guidelines regarding the cleaning frequency of equipment food contact surfaces and utensils. According to Food Code (2013), food contact surfaces should be cleaned between usage of different types of raw animal food, from raw food to ready-to-eat food, raw fruits and vegetables with time/temperature control for safety food (formerly “potentially hazardous food”), and any time when contamination is suspected. Aside from the conditions stated above, equipment food contact surfaces should be cleaned at least every 4 h if time/temperature control for safety food is used, with less cleaning frequencies allowed for lower processing temperatures (Food and Drug Administration, 2013). Based on these recommendations, chlorinated PP-PEI-SMA6-PEI coupons were tested for their chlorine capacity at different time points under 4 °C and room temperature (22 °C) to evaluate if the antimicrobial N-halamine moieties could sustain until the recommended cleaning time (recharged with sanitizers). The results demonstrated no significant difference (P > 0.05) between each time point and temperature condition tested under atmospheric storage (Fig. 6), indicating the stability of the chlorine bond in PP-PEI-SMA6-PEI coating and sustained antimicrobial activity without reduction over storage. 4. Conclusions Successful implementation of cationic and N-halamine moieties within a polymeric antimicrobial coating applied to PP was confirmed through infrared spectroscopy, with primary amine and N-halamine contents were characterized via colorimetric dye assays, confirming the cationic nature and the ability to bind chlorine. All PP-PEI-SMAMW-PEI coatings demonstrated low surface energy comparable to that of native PP, but showed no statistical differences between different molecular weight of SMA cross-linker groups. In the end, optimal molecular weight SMA was selected in terms of antimicrobial performance, where PP-PEI-SMA6-PEI exhibited an enhanced biocidal effect against E. coli O157:H7 in its chlorinated form. Inactivation kinetics results showed that antimicrobial performances of the modified PP coupons varied in their unchlorinated and chlorinated state for each type of bacteria, indicating that the nature of the microorganism plays an important factor in biocidal efficacy. Although traces of bacteria were observed on the unchlorinated PP-PEISMA6-PEIs after contact with L. monocytogenes and P. fluorescens, an agar overlay assay confirmed the absence of viable cells, supporting our conclusion that the observed antimicrobial effects were contributed by both unchlorinated and chlorinated forms of the polymer coating. Storage study of the chlorinated PP-PEI-SMA6-PEI under atmospheric conditions demonstrated stability of the chlorinated N-halamine moieties in between simulated cleaning frequency. Overall, this cationic and N-halamine incorporated polymer coating with the determined optimal molecular weight SMA cross-linker exhibited solid antimicrobial effectiveness and potential of retaining its antimicrobial activity under recommended sanitizing cycles. Such antimicrobial coatings can support routine sanitization standard operating procedures in reducing cross-contamination of pathogenic and spoilage microorganisms in food handling and processing facilities.

3.4. Scanning electron microscopy (SEM) and agar overlay assay Although cationic polymers owe their antimicrobial character to their charged nature, this charge has been suggested to promote fouling by anionic compounds and subsequent establishment of microbial biofilms (Brooks & Flint, 2008). Prior work suggested that this coating's chemical heterogeneity effectively neutralizes the expected positive charges, thus in theory reducing the likelihood of fouling (Bastarrachea & Goddard, 2016). Unchlorinated and chlorinated PP-PEI-SMA6-PEIs were analyzed with SEM after the microbial inactivation kinetics studies to evaluate the presence of bacterial attachment. Direct electron microscopy was coupled with agar overlay assays to determine viability of any attached cells. In the case of L. monocytogenes (Fig. 5) after the antimicrobial evaluation there was no evidence of bacterial adhesion for the chlorinated coatings. A few cells of L. monocytogenes, were observed for the unchlorinated coated PPs (∼12 organisms observed over 9 images acquired on 3 independently prepared samples, fewer than 3 organisms per observed field of view). To confirm that the observed adhered cells were not viable, an agar overlay test was conducted, in which agar were laid atop films and incubated for 48 h at 37 °C. Importantly, no growth was observed, confirming that no viable organisms were present on PPPEI-SMA6-PEI for either unchlorinated or chlorinated forms. As with L. monocytogenes, no evidence of bacterial adhesion was observed for the chlorinated coatings, while a few cells of P. fluorescens was captured for the unchlorinated PP-PEI-SMA6-PEI (∼5 organisms observed over 9 images acquired on 3 independently prepared samples, fewer than 3 organisms per observed field of view) (Fig. 5). The agar overlay test indicated no growth of bacteria, implying that the attached bacteria observed on the unchlorinated PP-PEI-SMA6-PEI coatings were

Acknowledgements This work was supported in part by the National Institute of Food and Agriculture, U.S. Department of Agriculture, under Agriculture and Food Research Initiative Grant No. 2018-67017-27874. The authors 552

LWT - Food Science and Technology 97 (2018) 546–554

Y.-T. Hung et al.

Fig. 5. SEM images of coated coupons after microbial inactivation assays.

References Aarnisalo, K., Salo, S., Miettinen, H., Suihko, M.-L., Wirtanen, G., Autio, T., et al. (2000). Bactericidal efficiencies of commercial disinfectants against Listeria monocytogenes on surfaces. Journal of Food Safety, 20(4), 237–250. Aase, B., Sundheim, G., Langsrud, S., & Rørvik, L. M. (2000). Occurrence of and a possible mechanism for resistance to a quaternary ammonium compound in Listeria monocytogenes. International Journal of Food Microbiology, 62, 57–63. American Society for Testing and Materials (2008). Standard test methods of sampling and chemical analysis of chlorine-containing bleaches. D 2022 – 89. (Philadelphia, PA, USA). Appendini, P., & Hotchkiss, J. H. (2002). Review of antimicrobial food packaging. Innovative Food Science & Emerging Technologies, 3, 113–126. Bastarrachea, L. J., Denis-Rohr, A., & Goddard, J. M. (2015a). Antimicrobial food equipment Coatings: Applications and challenges. Annual Review of Food Science and Technology, 6(6), 97–118. Bastarrachea, L. J., & Goddard, J. M. (2015). Antimicrobial coatings with dual cationic and N-Halamine Character: Characterization and biocidal efficacy. J.Agric. Food Chem. Journal of Agricultural and Food Chemistry, 63(16), 4243–4251. Bastarrachea, L. J., & Goddard, J. M. (2016). Self-healing antimicrobial polymer coating with efficacy in the presence of organic matter. Applied Surface Science, 378, 479–488. Bastarrachea, L. J., Wong, D. E., Roman, M. J., Lin, Z., & Goddard, J. M. (2015b). Active packaging coatings. Coatings, 5(4), 771–791. Baumann, A. R., Martin, S. E., & Feng, H. (2009). Removal of Listeria monocytogenes biofilms from stainless steel by use of ultrasound and ozone. Journal of Food Protection, 72(6), 1306–1309. Bintsis, T., Robinson, R. K., & Litopoulou-Tzanetaki, E. (2000). Existing and potential applications of ultraviolet light in the food industry - a critical review. Journal of the Science of Food and Agriculture, 80(6), 637–645. Bower, C. K., McGuire, J., & Daeschel, M. A. (1996). The adhesion and detachment of bacteria and spores on food-contact surfaces. Trends in Food Science & Technology, 7(5), 152–157. Brooks, J. D., & Flint, S. H. (2008). Biofilms in the food Industry: Problems and potential solutions. International Journal of Food Science and Technology, 43(12), 2163–2176. Buffet-Bataillon, S., Tattevin, P., Bonnaure-Mallet, M., & Jolivet-Gougeon, A. (2012). Emergence of resistance to antibacterial Agents: The role of quaternary ammonium compounds-a critical review. International Journal of Antimicrobial Agents, 39(5), 381–389.

Fig. 6. Storage stability of chlorinated PP-PEI-SMA6-PEI films. N-Halamine content at varied temperature conditions under atmospheric storage. Values represent means of 6 determinations with error bars indicating standard deviation.

gratefully acknowledge Brenda Werner from Cornell University (Food Science department) and Prof. María Corradini from University of Massachusetts (Food Science department) for technical guidance and mentorship during the project execution.

553

LWT - Food Science and Technology 97 (2018) 546–554

Y.-T. Hung et al.

Technology and Science, 16(3), 99–106. Lee, H. C., An, D. S., Lee, S. C., Park, H. J., & Lee, D. S. (2004). A coating for use as an antimicrobial and antioxidative packaging material incorporating nisin and α-tocopherol. Journal of Food Engineering, 62, 323–329. Liang, J., Chen, Y., Barnes, K., Wu, R., Worley, S. D., & Huang, T. S. (2006). N-Halamine/ Quat siloxane copolymers for use in biocidal coatings. Biomaterials, 27(11), 2495–2501. Lichter, J. A., & Rubner, M. F. (2009). Polyelectrolyte multilayers with intrinsic antimicrobial Functionality: The importance of mobile polycations. Langmuir, 25(13), 7686–7694. Maillard, J. Y. (2002). Bacterial target sites for biocide action. Journal of Applied Microbiology, 92, 16S–27S. Martin, N. H., Murphy, S. C., Ralyea, R. D., Wiedmann, M., & Boor, K. J. (2011). When cheese gets the blues: Pseudomonas fluorescens as the causative agent of cheese spoilage. Journal of Dairy Science, 94, 3176–3183. McDonnell, G., & Russell, A. (1999). Antiseptics and Disinfectants: Activity, action, and resistance. Clinical Microbiology Reviews, 12(1), 147–179. Mikš-Krajnik, M., Yuk, H.-., Kumar, A., Yang, Y., Zheng, Q., KIM, M., Ghate, V., Yuan, W., & Pang, X. (2015). Ensuring food security through enhancing microbiological food safety. Cosmos, 11(1), 69–97. Milovic, N. M., Wang, J., Lewis, K., & Klibanov, A. M. (2005). Immobilized N-Alkylated polyethylenimine avidly kills bacteria by rupturing cell membranes with no resistance developed. Biotechnology and Bioengineering, 90(6), 715–722. Navaneetha Pandiyaraj, K., Selvarajan, V., Deshmukh, R. R., & Gao, C. (2008). Adhesive properties of polypropylene (PP) and polyethylene terephthalate (PET) film surfaces treated by DC glow discharge plasma. Vacuum, 83, 332–339. Owens, D. K., & Wendt, R. C. (1969). Estimation of the surface free energy of polymers. Journal of Applied Polymer Science, 13(8), 1741–1747. Qian, L., & Sun, G. (2003). Durable and regenerable antimicrobial Textiles: Synthesis and applications of 3-methylol-2,2,5,5-tetramethylimidazolidin-4-one (MTMIO). Journal of Applied Polymer Science, 89(9), 2418–2425. Rutala, W., & Weber, D. (2008). Guideline for disinfection and sterilization in healthcare facilities, 2008. Washington, DC: Centers for Disease Control (U.S.). Simoes, M., Simoes, L. C., & Vieira, M. J. (2010). A review of current and emergent biofilm control strategies. Lwt-food Science and Technology, 43(4), 573–583. Singh, R. P. (1992). Surface grafting onto polypropylene - a survey of recent developments. Progress in Polymer Science, 17(2), 251–281. Srey, S., Jahid, I. K., & Ha, S. (2013). biofilm formation in food industries: A food safety concern. Food Control, 31(2), 572–585. Tiller, J. C., Liao, C. J., Lewis, K., & Klibanov, A. M. (2001). Designing surfaces that kill bacteria on contact. Proceedings of the National Academy of Sciences of the United States of America, 98(11), 5981–5985. Timofeeva, L., & Kleshcheva, N. (2011). Antimicrobial Polymers: Mechanism of action, factors of activity, and applications. Applied Microbiology and Biotechnology, 89(3), 475–492. Trmcic, A., Martin, N. H., Boor, K. J., & Wiedmann, M. (2015). A standard bacterial isolate set for Research on contemporary dairy spoilage. Journal of Dairy Science, 98, 5806–5817. Uchida, E., Uyama, Y., & Ikada, Y. (1993). Sorption of low-molecular-weight anions into thin polycation layers grafted onto a film. Langmuir, 9(4), 1121–1124. Van Tassell, J. A., Martin, N. H., Murphy, S. C., Wiedmann, M., Boor, K. J., & Ivy, R. A. (2012). Evaluation of various selective media for the detection of Pseudomonas species in pasteurized milk. Journal of Dairy Science, 95, 1568–1574. Williams, D. E., Elder, E. D., & Worley, S. D. (1988). Is free halogen necessary for disinfection? Applied and Environmental Microbiology, 54(10), 2583–2585. Worley, S. D., & Sun, G. (1996). Biocidal polymers. Trends in Polymer Science, 4(11), 364–370. Worley, S. D., & Williams, D. E. (1988). Halamine water disinfectants. CRC Critical Reviews in Environmental Control, 18(2), 133–175.

Cao, Z., & Sun, Y. (2009). Polymeric N-Halamine latex emulsions for use in antimicrobial paints. ACS Applied Materials & Interfaces, 1(2), 494–504. Centers for Disease Control and Prevention (CDC) (2011). Vital Signs: Incidence and trends of infection with pathogens transmitted commonly through food - foodborne diseases active surveillance network, 10 U.S. Sites, 1996-2010. MMWR.Morbidity and Mortality Weekly Report, 60(22), 749–755. Chen, Z., Luo, J., & Sun, Y. (2007). Biocidal efficacy, biofilm-controlling function, and controlled release effect of chloromelamine-based bioresponsive fibrous materials. Biomaterials, 28, 1597–1609. Chopra, D., Hatzikiriakos, S. G., Kontopoulou, M., & Vlassopoulos, D. (2002). Effect of maleic anhydride content on the rheology and phase behavior of poly(styrene-Comaleic anhydride)/poly(methyl methacrylate) blends. Rheologica Acta, 41(1), 10–24. Combe, E. C., Owen, B. A., & Hodges, J. S. (2004). A protocol for determining the surface free energy of dental materials. Dental Materials, 20(3), 262–268. Denis-Rohr, A., Bastarrachea, L. J., & Goddard, J. M. (2015). Antimicrobial efficacy of NHalamine coatings prepared via dip and spray layer-by-layer deposition. Food and Bioproducts Processing, 96, 12–19. Di Ciccio, P., Conter, M., Zanardi, E., Ghidini, S., Vergara, A., Paludi, D., et al. (2012). Listeria monocytogenes: Biofilms in food processing. Italian Journal of Food Science, 24(3), 203–213. Dong, A., Wang, Y.-., Gao, Y., Gao, T., & Gao, G. (2017). Chemical insights into antibacterial N-Halamines. Chemical Reviews, 117(6), 4806–4862. Farris, S., Pozzoli, S., La Vecchia, S., Biagioni, P., Bianchi, C. L., & Piergiovanni, L. (2014). Mapping physicochemical surface modifications of flame-treated polypropylene. Express Polymer Letters, 8(4), 256–266. Floros, J. D., Newsome, R., Fisher, W., Barbosa-Cánovas, G. V., Chen, H., Dunne, C. P., et al. (2010). Feeding the world today and Tomorrow: The importance of food science and technology. Comprehensive Reviews in Food Science and Food Safety, 9(5), 572–599. Food and Drug Administration (2013). Food Code 2013: Rcommendations of the United States public health service food and Drug administration. College Park, MD, USA. Gilbert, P., & Moore, L. E. (2005). Cationic Antiseptics: Diversity of action under a common epithet. Journal of Applied Microbiology, 99(4), 703–715. Gindl, M., Sinn, G., Reiterer, A., Tschegg, S., & Gindl, W. (2001). A comparison of different methods to calculate the surface free energy of wood using contact angle measurements. Colloids and Surfaces A: Physicochemical and Engineering Aspects, 181(1–3), 279–287. Glinel, K., Thebault, P., Humblot, V., Pradier, C. M., & Jouenne, T. (2012). Antibacterial surfaces developed from bio-inspired approaches. Acta Biomaterialia, 8(5), 1670–1684. Hanušová, K., Šťastná, M., Votavová, L., Klaudisová, K., Dobiáš, J., Voldřich, M., et al. (2010). Polymer films releasing nisin and/or natamycin from polyvinyldichloride lacquer Coating: Nisin and natamycin migration, efficiency in cheese packaging. Journal of Food Engineering, 99, 491–496. Hui, F., & Debiemme-Chouvy, C. (2013). Antimicrobial N-Halamine polymers and coatings: A review of their synthesis, characterization, and applications. Biomacromolecules, 14(3), 585–601. Ikeda, T., Hirayama, H., Yamaguchi, H., Tazuke, S., & Watanabe, M. (1986). Polycationic biocides with pendant active groups - molecular - weight dependence of antibacterial activity. Antimicrobial Agents and Chemotherapy, 30(1), 132–136. Kabza, K., Gestwicki, J. E., & McGrath, J. L. (2000). Contact angle goniometry as a tool for surface tension measurements of solids, using Zisman plot method - a physical chemistry experiment. Journal of Chemical Education, 77(1), 63–65. Kügler, R., Bouloussa, O., & Rondelez, F. (2005). Evidence of a charge-density threshold for optimum efficiency of biocidal cationic surfaces. Microbiology, 151(5), 1341–1348. Kumar, C. G., & Anand, S. K. (1998). Significance of microbial biofilms in food industry: A review. International Journal of Food Microbiology, 42(1–2), 9–27. Lee, H. C., An, D. S., Lee, D. S., & Park, H. J. (2003). Wide-spectrum antimicrobial packaging materials incorporating nisin and chitosan in the coating. Packaging

554