Sonochemical fabrication of edible fragrant antimicrobial nano coating on textiles and polypropylene cups

Ultrasonics Sonochemistry xxx (2016) xxx–xxx

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Sonochemical fabrication of edible fragrant antimicrobial nano coating on textiles and polypropylene cups O. Tzhayik, A. Lipovsky, A. Gedanken ⇑ Department of Chemistry, Bar-Ilan University, Ramat-Gan 52900, Israel

a r t i c l e

i n f o

Article history: Received 1 May 2016 Received in revised form 10 August 2016 Accepted 14 August 2016 Available online xxxx Keywords: Sonochemistry Edible coating Fragrance Nanoparticles Antimicrobial activity Textile Polypropylene Cotton bandage

a b s t r a c t We report on a simple and effective ultrasound-assisted deposition of vanillin nanoparticles (50 nm in size), raspberry ketone (RK) nanoparticles (40 nm in size) and camphor nanoparticles (width 30 nm, length 40 nm in size) on textiles and on polypropylene surfaces. The excellent antibacterial and antifungal activity of the fragrant coatings on cotton bandages, and polypropylene surface against Escherichia coli (E. coli), Salmonella paratyphi A (S. paratyphi A) and the yeast Candida albicans (C. albicans) cultures was demonstrated. It is worth pointing out that these fragrant materials are edible, making them very useful for packaging. The mechanism of the edible fragrant coating formation and adhesion to the textile was discussed, and finally an up-scaling of the sonochemical process for textile coating was carried out. Ó 2016 Elsevier B.V. All rights reserved.

1. Introduction From ancient times, people were attracted to perfumes and other fragrant materials. Nowadays, there is a growing need for high-quality textiles and packaging materials with antimicrobial properties for food safety, hygienic clothing, active wear, and wound healing [1]. Therefore, properly designed edible fragrant antimicrobial coating on textiles and packaging materials, would provide a significant contribution to the food, textile, medical, cleaning and toiletries industries. Several pathogenic bacteria and fungi were chosen for the antimicrobial testing due to their abundance as a community contaminator. In this work, we have developed a simple new method for the preparation of fragrant cotton bandages with antibacterial and antifungal properties by immobilizing organic edible nanoparticles of vanillin, raspberry ketone (RK) and camphor on the textile’s surface via ultrasound irradiation. The coating process was carried out with a high-intensity ultrasonic horn. Ultrasound radiation is an excellent technique for the formation and adherence of organic [2,3] or inorganic [1] nanoparticles (NPs) to a large variety of substrates and for the deposition of NPs on flat

⇑ Corresponding author. E-mail address: [email protected] (A. Gedanken).

and curved surfaces of ceramic [4–8], polymers [9,10], metals [11], and paper. Using the sonochemical method, nanosized materials can be finely spread on textiles and experience a strong affinity to the textile fibers. Moreover, sonochemistry is a new, effective and environmentally friendly method for the antibacterial treatment of textiles. The edible fragrant solid flavors, Vanillin, RK, and camphor, inheritably possess antimicrobial properties, which have been known and used for centuries. Vanillin (4-hydroxy-3-ethoxybenzal dehyde), a pleasant smelling aromatic compound, is the world’s most popular flavor and fragrance compound [12,13]. It is widely used as flavoring agent in foods, beverages, pharmaceuticals, perfumes and cleaning products [14,15]. Vanillin is slowly oxidized on exposure to air and is slightly water soluble (10 g/L) [12,16]. It displays antioxidant property [17,18] and gives antibacterial and antifungal activity against pathogenic microorganisms [12,18,19]. RK [4-(4-hydroxyphenyl)butan-2-one], is a key flavor molecule with typical raspberry flavor characteristics [20]. RK is widely used as a fragrance in cosmetics and as a flavoring agent in foodstuffs such as soft drinks, ice-cream and sweets [20–24]. RK is insoluble in water, and exhibits anti-androgenic [25], anti-inflammation [26], antiviral and antibacterial activity. Camphor, C10H16O, has a strong mentholic aromatic odor, and it is used for flavoring (mostly sweets), for aromatherapy, medicinal

http://dx.doi.org/10.1016/j.ultsonch.2016.08.020 1350-4177/Ó 2016 Elsevier B.V. All rights reserved.

Please cite this article in press as: O. Tzhayik et al., Sonochemical fabrication of edible fragrant antimicrobial nano coating on textiles and polypropylene cups, Ultrason. Sonochem. (2016), http://dx.doi.org/10.1016/j.ultsonch.2016.08.020

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purposes [27], as a rust inhibitor, and moth repellent [28–31]. Camphor has negligible solubility in water (1.6 g/L), and shows antibacterial and antifungal activity against several foodborne pathogens [27,28,32,33]. We have demonstrated that organic materials, biomaterials, as well as soluble inorganic materials can be made into NPs by sonicating their solutions [34–39]. Based on this process and in order to avoid the dissolution of the NPs, they were subsequent to their formation ‘‘thrown” onto a solid substrate creating a coating. The coating containing NPs significantly enhances the properties of the deposited material. In this way UV protection, antibacterial properties were improved [40,41]. Thus vanillin, RK and camphor NPs are apparently potential candidates for antimicrobial coatings as compared to their bulk counterpart. The textiles and polypropylene surfaces functionalized with edible fragrant antimicrobial organic NPs anchored by the sonochemical method were examined for their antibacterial and antifungal activity and showed good biocidal behavior. Finally, the paper demonstrated that the up-scaling of the sonochemical process for coating textiles is a viable possibility. 2. Materials and methods 2.1. Materials Ethanol 99.9%, Vanillin, Raspberry ketone, and D-Camphor were obtained from Sigma–Aldrich. Commercial 100% cotton bandages, 100% cotton textile and synthetic spandex textile were supplied to our laboratory by Delta INC. Carmiel, Israel. Commercial polypropylene disposable cups were purchased from a local grocery. All chemicals were used without any further purification. De-ionized H2O was used in all the experiments. 2.2. Sonochemical fabrication of Vanillin-Coated Cotton Bandage (VCB) Coating of vanillin on a bandage was performed as follows; in a round bottom flask, a piece of 8  8 cm2 cotton bandage (179.0 ± 0.2 mg) was placed and 60 ml of vanillin-ethanol solution [0.1 g/ml] was added. The Vanillin Coated Cotton Bandage (VCB) was fabricated using a high-intensity ultrasonic probe (Sonics and Materials, VC-600, 20 kHz, 0.5 in Ti horn). The horn of the high-intensity ultrasonic was immersed in the solution, employing an intensity of about 150 W cm 2 (using a booster). The sonication lasted for 30 min at 30 ± 2 °C, the temperature was kept constant by using an ice-water bath, thus reducing any external heating effect on the bandage texture. The VCB was dried in a chemical fume hood at room temperature and then washed thoroughly with ethanol/water (e/w) to remove the loosely immobilized particles of vanillin. Finally, the VCB was dried at room temperature and then kept in a Petri dish sealed with parafilm for further use. Control experiments were also carried out excluding the sonication stage, i.e., we immersed a piece of 8  8 cm2 cotton bandage in a vanillin-ethanol solution for 30 min at 30 ± 2 °C (using a warm water bath). The control Vanillin-Coated Cotton Bandage (VCBc) was then washed and dried in the same way used for the solicited samples. Using the same method, Camphor-Coated Cotton Bandage (CCB), RK-Coated Cotton Bandage (RCB), and their control samples CCBc and RCBc respectively, were also fabricated.

operating at 5 kV. HRSEM samples were prepared by applying a small piece of dry fragrant coated cotton bandage or of dry fragrant coated polypropylene onto a carbon tape. The samples were then vacuum-coated for 3 min with thin carbon/iridium film. The average size of the edible fragrant coating NPs was evaluated using HRSEM measurements. For further characterization of the coatings, reflection UV spectroscopy was used. Dry fibers of the fragrant coated bandage were used for reflection measurements. Reflection spectra were recorded on a Cary 100 scan Varian UV–vis spectrophotometer. 2.4. Quantification 2.4.1. The amount of deposited fragrance on a cotton bandag The amount of vanillin embedded/deposited on the cotton bandage was determined by weighing dry specimen of 8  8 cm2 cotton piece before and after the deposition. The bandage’s weight gain served us as the vanillin deposited quantity. For this purpose we used an AB135-S Mettler Toledo balance with a linearity of 0.2 mg. Same method was used for RK and camphor quantification. 2.4.2. Leaching measurements For leaching measurements, a piece of 1  1 cm2 of fragrant coated bandage, was placed in a vial, 5 ml of pure ethanol solution were added and the UV–vis spectra of the solution after 24, 48 and 96 h was measured. For vanillin another vial with 5 ml of water was placed and the same measurements were conducted. Since each of the fragrances has a different maximum absorbance wavelength i.e.; vanillin at 308 nm, RK at 278 nm and camphor at 287 nm, UV–vis spectra were recorded on a Cary 100 scan Varian UV–vis spectrophotometer, and the fragrant concentrations were calculated based on the corresponding calibration curves made beforehand. 2.5. Up scale Up-scaling of the sonochemistry process for coating textiles was conducted using a – pilot machine that was designed at our laboratory, which allows roll-to-roll coating of textiles in a continuous mode [1]. Using this pilot machine we preformed our vanillin and RK experiments (separately), and coated two meter long textiles. The concentration of the fragrant in the ethanolic solution was 0.1 g/ml. 2 types of different textiles; 100% cotton and synthetic spandex, were coated, the speed of the rolling machine was 22 cm/min. 2.6. Fabrication of vanillin-coated polypropylene cup (VCPC) For coating of vanillin, 60 ml of vanillin-ethanol solution (0.1 g/ ml) were placed inside a polypropylene cup (Commercial polypropylene disposable cups were purchased from a local grocery store). The vanillin coated cup (VCC) was fabricated using a high-intensity ultrasonic probe (Sonics and Materials, VC-600, 20 kHz, 0.5 in Ti horn). The horn of the high-intensity ultrasonic was immersed in the solution, employing an intensity of about 150 W cm 2. The sonication lasted for 30 min under an ice-water bath. During the sonication, the temperature inside the cup was not higher than 33 ± 2 °C. The VCC was then dried in a chemical fume hood at room temperature. 2.7. Antimicrobial tests

2.3. Characterization For imaging and characterization, high resolution scanning electron microscope (HRSEM) measurements were conducted using a FEI XHR-SEM Magellan 400 L scanning electron microscope

The antimicrobial, both antibacterial and antifungal activity of the active products, were tested against Escherichia coli (E. coli), Salmonella paratyphi A (S. paratyphi A) and the yeast Candida albicans (C. albicans). All strains were obtained from Bacteriological Labora-

Please cite this article in press as: O. Tzhayik et al., Sonochemical fabrication of edible fragrant antimicrobial nano coating on textiles and polypropylene cups, Ultrason. Sonochem. (2016), http://dx.doi.org/10.1016/j.ultsonch.2016.08.020

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tory of Meir Hospital, Kfar Sava, Israel. A typical procedure was as follows: Bacterial cultures were grown overnight on (NA) nutrient agar (Difco, Detroit, MI), these cultures were transferred into an Erlenmeyer flask containing Nutrient Broth (NB) at an initial optical density (OD) of 0.1 at 660 nm and allowed to grow at 37 °C with agitation (190 rpm). When the cultures reached an OD660 of 0.3 (the beginning of the logarithmic phase), they were centrifuged and washed twice with a saline solution (NaCl 0.145 M) at pH 6.5 to yield a final bacterial concentration of approximately 106 CFU/ml, following washing the cells were re-suspended in fresh media (NB). Fresh suspensions of the bacteria were poured into a test tube (d = 1 cm). Each sample of the tested dry product (commercial or coated cotton bandage, 2  3 cm monolayer) was placed in the solution. The tubes were incubated on a rotary shaker at 190 rpm, 37 °C over night. Following the incubation the optical density of the solution was measured (the clear solutions were tested for presence of live bacteria by plating a 100 lL sample on agar plates following serial dilutions in saline). In case of C. albicans medium with 2% of glucose (agar plates) or 4% (liquid media) was used. Washing of the bandages; washing of the bandages for the loosely fixed particles (not coated), was done by placing the bandages in a water or ethanol solution and gently shaking (50 rpm) for 60 s. In the second step the most successful concentration were chosen and tested for a non-contact activity, by placing the bandage on the lid of a bacteria/yeast plated Petri dish approximately 1.5 cm from the growth supporting agar surface. A successful concentration was selected as the lowest concentration that gave the highest antimicrobial activity. Finally, the same active concentration was also coated on polypropylene cups and tested for their activity; a 100 ll drop of bacteria solution (106 cells/ml) was placed on top of polypropylene piece, the number of live bacteria was evaluated prior and after 4 h incubation at 37 °C by plating a sample on agar plates following serial dilutions in saline.

3. Results and discussion 3.1. Characterization of the edible fragrant coatings The morphology of the bandage’s fragrant coatings; vanillin, RK and camphor, was characterized using high resolution scanning electron microscopy (HRSEM) (Fig. 1a, b and c, respectively). HRSEM surface analysis of the edible fragrant coatings revealed a nanoparticle layer on the bandage’s fibers (Fig. 1a, b and c), as opposed to the origin 100% cotton bandage, which has a characteristics smooth surface (Fig. 1d). In the case of vanillin and RK, spherical NPs were formed with an average diameter (calculated from 50 random particles) of 50 ± 3 nm and 40 ± 3 nm, respectively (Fig. 1a, inset and b, inset). Whereas in the case of camphor, irregular nano-shapes were formed, with an approximate width of 30 ± 3 nm and 40 ± 3 nm length, as illustrated in Fig. 1c, inset. Overall, the sonication effect is reflected in the creation of NPs, while in the case without sonication, random sized and shaped particles are formed on the bandage. At the end of the sonochemical coating process, VCB, RCB and CCB possess a delicate characteristic scent of the origin fragrant molecule, and their color remained white as a typical cotton bandage. Absorption and reflection UV–vis spectroscopy was used to better characterize the developed edible fragrant coatings and to estimate the influence of the sonication on the vanillin, RK and camphor compound. Fig. 2 displays UV–vis absorption spectra of ethanol solution of vanillin, RK and camphor prior to the sonochemical coating process

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(black lines), and the corresponding diffusion reflection spectra of the fragrant coated bandages after sonication, VCB, RCB and CCB (red lines). As shown in Fig. 2, the obtained UV–vis absorption and the corresponding reflection spectra of each fragrant have a similar profiles, which indicates that there was no significant change in the compound’s structure and that there was no degradation of the molecule during the sonochemical coating process. Minor shift or broadening of 5–10 nm to the red can be observe in the fragrant reflection spectra relative to the corresponding absorption spectra, these shifts fall within the error limit of the measurement, but can also be attribute to the creation of NPs during the sonication process. 3.2. Antimicrobial studies Previous studies [1] indicated that the product yield, and consequently the antimicrobial effectiveness of the coating, is strongly dependent on the rate of inter-particle collisions and on the concentration of the reagents during the sonochemical synthesis. Based on these results, the concentration of the initial fragrant solution was selected as an important factor for the optimization of the sonochemical coating reaction, and for simplification, the other experimental parameters time and sonication intensity were kept unchanged. The aim of this parametization experiments was to reach minimal effective edible fragrant NPs coating that will demonstrate maximal antimicrobial activity (MIC, minimal inhibitory concentration). The antibacterial and antifungal activity of sonicated VCB, CCB and RCB and their corresponding unsonicated control experiments VCBc, CCBc and RCBc, were tested on two bacterial strains (Salmonella paratyphi A and Escherichia coli) and one fungi (Candida albicans), the results are sumarized in Figs. 3–5. Bandages were labeled as follows: V for VCB (Vanillin Coated Bandage); Vc for VCBc (VCB control); V(w)for V washed with water; Vc(w) for Vc washed with water; V(e) for V washed with ethanol; Vc(e) for Vc washed with ethanol. The initial concentration of vanillin is 0.1 g/ml. Regarding Vanillin coating, Fig. 3 reveals that there is no growth of E. coli, S. paratyphi A and C. albicans for both VCB and VCBc (labeled V and Vc respectively). Both bandages achieved a complite inhibition of bacterial and fungal growth, therefore VCB is not preferred over VCBc. After washing the bandages, the antimicrobial activity changes with respect to the washing solvent; ethanol (e) or water (w). In the case of water; washing VCB and VCBc with water (labeled V(w) and Vc(w), respectively) retained the maximum bacterial inhibition ability of the bandage and there was still no growth of E. coli, S. paratyphi A on these bandages. On the other hand, the growth of the fungal C. albicans was inhibited only by the water washed VCB (labeled V(w)) compare to the washed VCBc (labeled Vc(w)), OD = 0 and OD = 0.2, respectively. In the case of ethanol; washing VCB, labeled V(e), reduced the growth inhibition ability of the coating (OD of 0.15 compared to undetectable OD for unwashed VCB, labeled V, Fig. 3). But, washing VCBc with ethanol (labeled Vc(e)), significantly increased the growth of all three (to OD of 0.7–1.2, an OD similar to that of a negative control with no bandage). Thus, the VCB is clearly preffered over VCBc, and the sonication effect is manifested. Looking at Vc(e), (Fig. 3) it is indicated that the E. coli growth was above 1 OD (Fig. 3, blue column), which means that the washing completely removed the active part that was not attached by the sonication method. Non coated bandages did not affect the growth of bacteria and yeast (an OD of 1.1 a.u. and 0.8 a.u. respectively, was measured). By using ethanol we tried to mimic the harshest condition that the product can withstand. Overall, washing VCB with water or ethanol does not affect the bacterial activity and slightly changes the antifungal activity, whereas washing VCBc with ethanol significantly reduces the antimicrobial activity. Keeping in mind that vanillin is highly sol-

Please cite this article in press as: O. Tzhayik et al., Sonochemical fabrication of edible fragrant antimicrobial nano coating on textiles and polypropylene cups, Ultrason. Sonochem. (2016), http://dx.doi.org/10.1016/j.ultsonch.2016.08.020

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a

b

c

d

Fig. 1. HRSEM images of cotton bandages; (a) VCB, magnified by 50,000 (inset); (b) RCB, magnified by 100,000 (inset); (c) CCB, magnified by 50,000 (inset); (d) 100% cotton bandage, magnified by 200,000 (inset).

uble in ethanol (1:2 vanillin: ethanol 95%) [42], our last result can be attributed to the fact that the sonochemically prepared vanillin NPs are strongly embedded inside the textile and cannot be removed (even in ethanol), thus showing the high advantage of sonochemically-assisted coating. Moreover, the fabrication of vanillin NPs significantly enlarged the vanillin surface area, and the washing step reveals new surface which gives the VCB its antimicrobial superiority over VCBc. In all cases where the OD was bellow the detection limit a sample was taken for verification of bacteria presence. In all the samples with un-detectible OD, live bacteria or fungi were not found. Bandages were labeled as follows: R for RCB (Raspberry Coated Bandage); Rc for RCBc (RCB control); R(w) for R washed with water; Rc(w) for Rc washed with water. The numbers 2, 3 and 4 are given to differentiate between RK series R2, R3 and R4 i.e. the initial concentration of RK is 0.011 g/ml for R2 series, 0.033 g/ml for R3 series, and 0.066 g/ml for R4 series. Regarding RK coating, Fig. 4 reveals that the RK MIC (minimum inhibitory concentration) is on RCB labeled as bandage R4 is 0.066 g/ml. R4, possess a coating that completely prevents the growth of the bacteria E. coli and S. paratyphi A, (the OD created by the growth of bacteria is not detected, OD = 0), whereas the unsonicated RCBc, labeled R4c, has no effect on the bacteria growth, and doesn’t inhibit bacteria (OD = 1–1.2), as indicated by the blue and green columns for E. coli and S. paratyphi A, respectively (Fig. 4). Thus, it can be concluded that the sonicated RCB, labeled R4, effectively prevents bacteria growth, and clearly preferred over the unsonicated bandage, labeled R4c. As in the case of vanillin, this phenomenon can be attributed to the sonochemically creation of RK NPs, these NPs prevent the bacteria growth, and by doing this, enable their use as an antibacterial bandage coating. Moreover, washing RCB with water, (labeled R4(w),

Fig. 4), didn’t change the coating’s antibacterial properties. Furthermore, it was found out that the antifungal RK MIC (orange columns in Fig. 4) is on the sonicated RCB, labeled as bandage R3, which corresponds to initial RK concentration of 0.033 g/ml. Here, it is assumed that C. albicans is more sensitive towards RK NPs than E. coli and S. paratyphi A, and this is the reason why the C. albicans’s fungal growth prevention is possible at lower RK starting concentration (0.033 g/ml). Washing RCB with water, (labeled R3(w), Fig. 4), didn’t change the coating’s antifungal properties. Non coated bandages did not affect the growth of bacteria and yeast (an OD of 1.15 a.u. and 2.4 a.u. respectively, was measured). Overall, from the antimicrobial perspective, RCB is clearly preffered over RCBc, and the sonication effect is again manifested. As in the case of vanillin; in all the samples with un-detectible OD, live bacteria or fungi were not present. Bandages were labeled as follows: C for CCB (Camphor Coated Bandage); Cc for CCBc (CCB control); C(w) for C washed with water; Cc(w) for Cc washed with water. The numbers 1,2,3 were given to differentiate between camphor series C1, C2 and C3 i.e. The initial concentration of camphor is 0.11 g/ml for C1 series, 0.22 g/ml for C2 series, and 0.44 g/ml for C3 series. Regarding camphor coating, Fig. 5 reveals that the MIC of camphor measured for S. paratyphi A (green) and C. albicans (orange), is on CCB and CCBc (labeled C2 and C2c respectively), both have the initial camphor concentration 0.22 g/ml. At this concentration the sonochemical creation of NPs coating on bandage C2 did not show any advantage over C2c, and in both cases there was no growth of C. albicans or S. paratyphi A. After washing with water to remove loosely embedded particles, CCB and CCBc completely lost their activity and the C. albicans and S. paratyphi A flourish, as shown by the orange and green columns, labeled as bandages C2(w) and C2c(w), Fig. 5. It is important to note that in all camphor

Please cite this article in press as: O. Tzhayik et al., Sonochemical fabrication of edible fragrant antimicrobial nano coating on textiles and polypropylene cups, Ultrason. Sonochem. (2016), http://dx.doi.org/10.1016/j.ultsonch.2016.08.020

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Vanillin

1.2

F(R) (Bandage)

OD (a.u)

Abs (Soluon)

0.9 0.6 0.3 0 V

V(w)

Vc

Vc(w)

V(e)

Vc(e)

Bandage 230

250

270

290

310

330

350

370

S. Parratyphi A

E. coli

C. albicans

Wavelength [nm] Fig. 3. Antimicrobial activity of VCB and VCBc against S. paratyphi A (green columns), E. coli (blue columns), and C. albicans (orange columns). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

Raspberry Ketone Abs (Soluon)

F (R) (Bandage) 1.5

OD (a.u.)

1

230

250

270

290

310

330

350

0.5

0 R3

Wavelength [nm]

R3c

R3(w)

R3c(w)

R4

R4c

R4(w)

R4c(w)

R3(w)

R3c(w)

Bandage S. paratyphi A

E. coli

Camphor Abs (Soluon)

3

F(R) (Bandage)

OD (a.u.)

2.5 2 1.5 1 0.5

230

250

270

290

310

330

350

0 R2

R2c

R2(w)

Wavelength [nm] Fig. 2. (black lines) UV–vis absorption spectra of the ethanol solutions of vanillin, RK and camphor, prior to sonication; (red lines) the corresponding diffused reflection spectra of the fragrant coated bandages VCB, RCB and CCB. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

experiments, as opposed to vanillin, a characteristic odor of camphor was barely distinguished on the bandage after the water washing and the subsequent drying. This is not surprising knowing that the vapor pressure of camphor is 5500 times larger than vanillin (at 25 °C) [43,44]. Non coated bandages did not affect the growth of bacteria and yeast (an OD of 1.5 a.u. and 2 a.u. respectively, was measured). On top of that, apparently the camphor is easily detaches from the bandage and was washed out by the water during the washing step, thus the few embedded left NPs evaporated or were not enough to inhibit the bacteria S. paratyphi A or the fungi C. albicans growth. In the case of E. coli, its growth was successfully prevented at higher camphor concentration relative to the concentration that was enough to prevent the growth of C. albicans and S. paratyphi A. The Camphor MIC (orange color in Fig. 5) for E. coli is on CCB and CCBc labeled as bandages C3 and C3c, respectively, in Fig. 5, both

R2c(w)

R3

R3c

Bandage C. albicans Fig. 4. Antimicrobial activity of RCB and RCBc against S. paratyphi A (green columns), E. coli (blue columns), and C. albicans (orange columns). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

have the initial concentration 0.44 g/ml. Once again, after washing with water, CCB and CCBc lost their activity and E. coli flourished, as depicted by C3(w) and C3c(w), respectively, blue columns in Fig. 5. In all cases where the OD was bellow the detection limit, live bacteria or fungi were not detected. Furthermore, all antibacterial and antifungal measurements showed identical results even 8 months after their preparation. During this period all the fragrant coated bandages were kept in a sealed Petri dish and stored at room temperature. 3.3. Leaching measurements As expected, after the sonochemical coating process, the cotton bandages gained weight. In the case of vanillin, the bandage gained 26 ± 5% (0.7 ± 0.1 mg/cm2), and in the case of RK and camphor the

Please cite this article in press as: O. Tzhayik et al., Sonochemical fabrication of edible fragrant antimicrobial nano coating on textiles and polypropylene cups, Ultrason. Sonochem. (2016), http://dx.doi.org/10.1016/j.ultsonch.2016.08.020

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O. Tzhayik et al. / Ultrasonics Sonochemistry xxx (2016) xxx–xxx 3 2.5

OD (a.u)

2 1.5 1 0.5 0 C1

C1c

C1(w)

C1c(w)

C2

C2c

C2(w)

C2c(w)

Bandage S. paratyphi A

C. albicans

3

from the bandage (leading to its eventual evaporation) which correlates with the lack of significant antibacterial and antifungal activity of CCB after the washing step, as is reported in detail in Section 3.2. For Vanillin and RK only 13% and 23%, respectively, leached to ethanol, and most of the coatings were left on the bandage. This confirms that these sonochemically prepared edible fragrant NPs coatings are not easily detached or dissolve in ethanol, indicating that most of the edible fragrant coating is left on the bandage through the short water wash step, especially since the solubility in water is low in the case of vanillin and negligible in the case of RK. This later correlates with our antibacterial and antifungal activity results, which demonstrated no significant change in the antibacterial and antifungal activity of the VCB and RCB after the water washing step. 3.4. Up scaling

OD (a.u.)

2.5 2 1.5 1 0.5 0 C2

C2c

C2(w)

C2c(w)

C3

C3c

C3(w)

C3c(w)

Bandage E. coli Fig. 5. Antimicrobial activity of CCB and CCBc against S. paratyphi A (green columns), E. coli (blue columns), and C. albicans (orange columns). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

% Leached from bandage

bandage gained 40 ± 5% (1.1 ± 0.1 mg/cm2) and 60 ± 5% (1.7 ± 0.1 mg/cm2), respectively. Fig. 6 summarizes the leaching measurements of the fragrant coated bandages, immersed in pure ethanol for 24 h, in the case of vanillin leaching to water was also measured. The values marked on each column represent the percentage of the coating that was leached from the bandage to the ethanol/water solution. As depicted in Fig. 6, it is clear that of the three edible fragrant, the camphor has the greatest tendency to leach, and in 24 h 90% of the coating leached out. In same conditions, only 23% of RK and 13% of vanillin were leached out to ethanol. After 48 h and 96 h there was no change in data and all the values obtained after 24 h remained unchanged, which implies that an equilibrium is reached after 24 h. The results depicted in Fig. 7, may also imply that the camphor coating is easily detached

100 90 80 70 60 50 40 30 20 10 0

90%

23% 13% 1% VCB(water)

VCB(ethanol)

RCB(ethanol)

CCB(ethanol)

Bandage Fig. 6. The relative amount (in percentage) of fragrant that leached out of the overall amount of coating on 100% cotton bandages; VCB, RCB and CCB, after 24 h of immersion in pure ethanol. In the case of vanillin we also calculated the data after 24 h of immersion in water VCB (labeled VCB (water)). The initial concentration of the fragrance is 0.1 g/ml for vanillin, 0.066 g/ml for RK, and 0.44 g/ml for camphor. The values marked on each column are the percentage of the coating that was leached.

Up scaling of the sonochemical coating process was carried out by using a pilot scale machine (Viatech, Russia), as displayed in Fig. 7, and fabricating long pieces of yarns with vanillin and RK NPs coated on a 100% cotton textile and synthetic spandex textile. The morphology of the coatings obtained in the pilot machine is the same morphology as in the lab (small scale); i.e. spherical NPs where formed. Antimicrobial measurements in the small scale (100% cotton bandage) and large scale (100% cotton textile and synthetic spandex textile/pilot machine) gave the same results, and in both cases there was no growth of E. coli S. paratyphi A or C. albicans. An OD of 0.2 a.u. and 0.1 was measured in bacteria and fungi solution respectively. The uncoated textile did not affect the pathogens concentration (OD of 1.5 a.u. for all three pathogens was measured). Thus, sonicating textile (cotton or synthetic spandex) immersed in 0.1 g/ml ethanolic solution of vanillin or RK, can easily produce a fragrant textile with an effective antimicrobial edible nanoparticle coating. 3.5. Vanillin coated polypropylene cup (VCPC), characterization and antimicrobial activity In an attempt to extend our methodology to other useful surfaces that can be sonochemically coated, disposable materials were immobilized by vanillin. The idea was to choose a surface that is used commonly worldwide (such as polypropylene), and to coat it with an edible (and nontoxic) fragrant layer that will impart antimicrobial properties to its surface. Initially a disposable cup (200 ml) was chosen, since the aim was to upgrade food containers or packaging materials such as dair3y products, baking products, storage boxes etc. For this, vanillin used as an edible fragrant model, and an ethanol-vanillin solution [0.1 g/ml] was sonicated inside an open polypropylene cup. Looking at the HRSEM image of VCPC piece, as depicted in Fig. 8, it is shown (again) that using sonication the polypropylene surface can be homogeneously coated with spherical vanillin NPs having an average diameter of 70 ± 5 nm (averaging 50 random particles). The vanillin NPs on the cup appeared to be slightly larger relative to the NPs on the cotton bandages, this is probably due to the different surface properties, and can be also attributed to the different distance from the sonication source i.e. the bandages were placed closer to the sonication source than the cup, thus, in the case of the bandage, the NPs did not had sufficient distance to further grow during the sonication process. For the antibacterial tests, a 100 lL sample (of bacteria solution) was placed on a 1.5 cm2 cup piece, bacteria number was determined after growing the plated bacteria in the control (uncoated cup piece) and the coated sample. VCPC was found able to prevent the growth of all model bacteria and yeast (E. coli, S. paratyphi A and C. albicans) thus adding to the polypropylene surface both the fragrance and the antimicrobial properties.

Please cite this article in press as: O. Tzhayik et al., Sonochemical fabrication of edible fragrant antimicrobial nano coating on textiles and polypropylene cups, Ultrason. Sonochem. (2016), http://dx.doi.org/10.1016/j.ultsonch.2016.08.020

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Fig. 7. Photos of the pilot machine. The Left picture is a photo of the machine with the outline of the functionality of the various parts; (1) the rod on which the textile is rolled initially (2) a bath in which the fragrant solution is placed (3) sonication unit (4) the part on which the textile is finally rolled after the sonication step. The Right picture is a photo of the pilot machine in operation.

4. Conclusions

Fig. 8. HRSEM image of vanillin coated polypropylene cup (VCPC), magnified by 15,000 and 100,000 (inset).

Knowing that synthetic or natural fibers are not resistant to bacteria and pathogenic fungi [46], it is clearly demonstrated that the fragrance coated bandages prevented or inhibited the growth of contaminating bacteria and yeast. The increased surface area of these edible NPs promoted the release of the active molecules preventing or inhibiting the growth of the pathogens. This phenomenon was even more stringing in the non-contact inhibition test, in which the vanillin coated surface prevented the growth of bacteria placed approximately 1.5 cm from the growth supporting agar surface. All three fragrances were successfully coated onto a surface and showed antimicrobial activity. Using the sonication method the coating was very strongly embedded and was not removed by washing of the bandages. This sonochemical coating method can be employed to a variety of surfaces, thus may establish a significant contribution to the textile, food packaging, medical, cleaning and toiletries industries. References

3.6. Proposed mechanism for the formation and deposition of nanosized edible fragrances on textiles and solid surfaces by the sonochemical method The proposed mechanism for the sonochemical formation and deposition of edible fragrant NPs on textile and solid surfaces is similar to that described and suggested by Gedanken et al. [1]. During sonication, a process of acoustic cavitation is initiated, that is the formation, growth, and explosive collapse of bubbles. According to the hot spot theory, upon the collapse of the acoustic bubble, which occurs in less than a nanosecond, very high temperatures and very high cooling rates are obtained. In each collapsing bubble, due to the fast kinetics, few nucleation centers are formed and their growth is limited by the implosive collapse. Cavity collapse in a homogeneous liquid is very different from cavitation near a liquid-solid interface [45], and when the microscopic cavitation bubbles collapse near the surface of the solid substrate, they generate powerful shock waves and micro-jets. In our experiments the solid substrate (cotton or polypropylene) was introduced into the sonication cell containing the ethanolic fragrant solution. The micro-jets formed after the collapse of the bubble throw the just-formed edible fragrant NPs at the surface of the substrate at such a high speed that they strongly adhere to the surface, either by physical or chemical interactions, depending on the nature of the substrate. In the case of textile, this phenomenon might cause a local melting of the fibers at the contact sites, which may be the reason why the particles strongly adhere to the textile.

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