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Improving functional properties of chitosan films as active food packaging by incorporating with propolis
Food Hydrocolloids 61 (2016) 695e702
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Improving functional properties of chitosan films as active food packaging by incorporating with propolis Ubonrat Siripatrawan*, Waranya Vitchayakitti Department of Food Technology, Faculty of Science, Chulalongkorn University, Bangkok 10330, Thailand
a r t i c l e i n f o
a b s t r a c t
Article history: Received 11 September 2015 Received in revised form 6 March 2016 Accepted 1 June 2016 Available online 8 June 2016
Chitosan films containing 0, 2.5, 5, 10 and 20% w/w propolis extract (PE), high in polyphenols, were developed. The films’ tensile strength, elongation at break, total phenolic content and antioxidant activity increased, while water vapor permeability and oxygen permeability decreased with increasing propolis concentration. Increasing PE concentration resulted in deeper orange color films, as compared to light yellow control films. The ability of the films to inhibit Staphylococcus aureus, Salmonella Enteritidis, Escherichia coli and Pseudomonas aeruginosa was determined using agar diffusion technique. Chitosan films containing PE could inhibit all tested bacteria on contact surface underneath the film discs. Changes in the Fourier Transform Infrared spectra of the films were observed when PE was incorporated, suggesting some interactions occurred between chitosan and propolis polyphenols. The characterization of mechanical properties, oxygen and moisture barrier, and antioxidant and antimicrobial activities revealed the benefits of adding PE into chitosan films and the potential of using the developed film as active food packaging. © 2016 Elsevier Ltd. All rights reserved.
Keywords: Active packaging Antimicrobial Antioxidant Permeability Propolis
1. Introduction Recently, research and developments in active food packaging have focused on bio-based functional packaging materials incorporating natural active compounds and ingredients (Leceta, Guerrero, & de la Caba, 2013; van den Broek, Knoop, Kappen, & Boeriu, 2015; Madureira, Pereira, & Pintado, 2015). Chitosan is a functional biopolymer with intrinsic antimicrobial and antioxidant properties and consequently, it has high potential to be used as an alternative biodegradable active food package (Fernandez-Saiz, Lagaron, & Ocio, 2009; Guoa et al., 2015; van den Broek et al., 2015). Bio-based packaging materials with antioxidant and antimicrobial properties have become popular since oxidation and microbial contamination are major problems affecting food quality and safety. Many studies have been conducted on the utilization of plant polyphenols as alternatives to synthetic antimicrobial and jaro, Alvarez-Casas, antioxidant agents (Evrendilek, 2015; Lores, Pa Domínguez, & García-Jares, 2015; Madureira et al., 2015; SalviaTrujillo, Rojas-Graü, Soliva-Fortuny, & Martín-Belloso, 2015; Siripatrawan, 2016).
* Corresponding author. E-mail address: [email protected] (U. Siripatrawan). http://dx.doi.org/10.1016/j.foodhyd.2016.06.001 0268-005X/© 2016 Elsevier Ltd. All rights reserved.
Propolis, the natural resinous substance collected by honeybees from various plant sources, is considered a good source of natural antioxidants and antibacterials (Bankova, 2005). Propolis contains a variety of chemical compounds such as polyphenols (flavonoid aglycones, phenolic acids and their esters, phenolic aldehydes, alcohols and ketones), sesquiterpene quinines, coumarins, amino ~o et al., 2010). acids and inorganic compounds (Bankova, 2005; Falca Flavonoids (flavones, flavonols and flavonones), aromatic acids and phenolic compounds are the most important active constituents of propolis and appear to be the principal components responsible for the biological activities of propolis samples (Silici & Kutluca, 2005). Propolis has been reported to possess various biological activities, such as antibacterial, antiviral, antitumor, anti-inflammatory, anticancer, antifungal, and antitumoral properties (Falc~ ao et al., 2010). The antimicrobial effects of propolis against Gram-positive (Bacillus cereus, Listeria monocytogenes and Staphylococcus aureus) and Gram-negative (Salmonella Typhimurium, Escherichia coli and Pseudomonas fluorescence) bacteria have been reported (Silici & Kutluca, 2005; Siripatrawan, Vitchayakitti, & Sanguandeekul, 2013). As a good source of polyphenols with multiple biological activities, propolis has high potential to be used as an active agent that can be incorporated into films. Limited research on incorporation of propolis to enhance properties of packaging films and
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nchez-Gonza lez, Cha fer, coatings has been published. Pastor, Sa lez-Martínez (2010) investigated physical and Chiralt, and Gonza antifungal properties of hydroxymethylcellulose based films containing propolis. Bodini, Sobral, Favaro-Trindade, and Carvalho (2013) studied properties of gelatin films added with propolis, and Torlak and Sert (2013) examined antibacterial effectiveness of chitosan-propolis coated polypropylene film. Nevertheless, research providing functional characterizations, including mechanical properties, oxygen and moisture barrier, structural property, antioxidant activity and antimicrobial property, of chitosan films incorporated with propolis extract (PE) has not been fully documented. Hence, this research aimed to develop and characterize chitosan films with enhanced functional properties for potential use as active food packaging by incorporating chitosan with propolis extract. 2. Materials and methods 2.1. Film preparation Propolis was collected from Nan province in northern Thailand and extracted following the methods of Siripatrawan et al. (2013). Three grams of ground propolis were extracted using 100 ml of 30% ethanol aqueous solution. The solution was extracted at 50 C in a water bath shaking incubator (New Brunswick Scientific, Edison, USA) at 200 oscillation/min for 24 h and then filtered through Whatman filter paper No. 1. The extract solution was concentrated using a rotary evaporator (Rotavapor R-200, BÜCHI Laboratory Equipment, Flawil, Switzerland) under reduced pressure at 45 C. Chitosan (Seafresh Industry Public Company Limited, Chumphon, Thailand) with 95% degree of deacetylation was used to prepare chitosan-based films. The chitosan-based film was prepared according to the procedure of Siripatrawan and Harte (2010) with slight modification. A film-forming solution was prepared by dissolving 2 g of chitosan powder into 100 ml of 1% acetic acid solution. Glycerol (Sigma Chemical Co., St. Louis, MO, USA) was added to the solution at 30% w/w of chitosan. The solution was heated at 60 C in a water bath shaking incubator at 100 rpm for 30 min. The PE was dissolved in the film-forming solution to obtain concentrations of 0, 2.5, 5, 10 and 20% w/w of chitosan. The resulting solutions were homogenized using a homogenizer (D79282, Ystral GmbH, Ballrechten-Dottingen, Germany). Air bubbles in the film-forming solutions were removed using a sonicator (Cole-Parmer, Vernon Hills, Illinois, USA). Each film-forming solution was cast on a ceramic plate. The obtained films were conditioned in a chamber at 25 C and 50% relative humidity for 48 h prior to testing. 2.2. Color measurement Color (L*, a*, b*) values were measured using a Minolta Chromameter (CR-300, Minolta Camera Co., Osaka, Japan). Films were cut into 15 2.5 cm and six readings at different positions on each film were measured. Five replications were conducted for each treatment and five film samples were used for each replication. 2.3. Water vapor transmission rate Water vapor transmission rate (WVTR) of the films was determined following the ASTM standard test method (ASTM., 2003). Film samples, previously equilibrated at 50% RH for 48 h, were sealed in glass cups containing silica gel. The film-covered cups were placed in an environmental chamber set at 25 C and 75% RH. The cups were weighed periodically until steady state was reached (±0.0001 g). The WVTR (g m2 day1) of the films was determined
using Eq (1). At least five replications of each film treatment were tested for the permeability coefficient (WVP). The WVTR of the films was then used to calculate the WVP (g mm m2 d1 Pa1) using Eq (2).
WVTR ¼
Dw ADt
WVP ¼ WVTR
(1) x
Dp
(2)
where Dw is the moisture weight gain, A is the tested film area, Dt is the definite time once steady state was reached, x is the film thickness and Dp is the partial water vapor pressure gradient between the inner and outer surface of the film in the chamber.
2.4. Oxygen transmission rate A single film specimen was cut out to have a measurement area of 50 cm2. O2 transmission rate (OTR) was analyzed using MOCON OX-TRAN 2/20 devices (MOCON, Inc., USA) following the ASTM standard method (ASTM., 2003). The specimens were mounted onto the diffusion cells and a mixture of 98% N2 and 2% H2 was used as a carrier gas. OTR was used to measure the amount of oxygen passing through films when exposed to a gradient with partial O2 pressure across the films. As the oxygen permeated through the film sample, it was picked up by the carrier gas and carried through a coulometric sensor. The amount of oxygen contained in the carrier gas at equilibrium was measured. The test conditions used in the OX-TRAN 2/20 system were 25 C and 50% RH. Duplicate film samples were tested. OTR (Eq (3)) is expressed as the quantity (q) of O2 molecules passing through a film surface area (A) during time (Dt) at steady state. Oxygen permeability coefficient (OP) of the films was then calculated using Eq (4):
OTR ¼
q ADt
OP ¼ OTR
(3) x
Dp
(4)
2.5. Mechanical properties Tensile strength (TS) and percentage elongation at break (%E) were measured with an Instron Universal Testing Machine (Model 5655, Instron Corporation, Canton, MA, USA) following the ASTM Standard Test Method D 882-91 (ASTM., 2003). Each film strip (15 2.5 cm) was mounted between the grips of the Instron and tested with an initial grip separation of 5 cm and crosshead speed of 1 mm/s. TS (Eq. (5)) was calculated by dividing the maximum load (Fmax) by the initial cross-sectional area (4) of the film sample expressed as MPa. %E (Eq. (6)) was calculated as the ratio of the film extension (Dl) at the point of sample rupture to the initial length (l0) of a sample and expressed as a percentage. Measurements represent an average of at least nine replications.
TS ¼
%E ¼
Fmax
F
Dl l0
100
(5)
(6)
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2.6. FTIR analysis Fourier Transform Infrared (FTIR) spectrometry was carried out to observe the structural interactions of chitosan and propolis polyphenols. The FTIR spectra of chitosan films were recorded from 4000 to 650 cm1 using a FTIR spectrometer (PerkinElmer 1760, PerkinElmer Life And Analytical Sciences, Inc., Waltham, Massachusetts USA). 2.7. Total phenolic assay Total phenolic content (TPC) of the chitosan film samples was determined using the Folin-Ciocalteu method (Singleton, Orthofer, & Lamuela-Raventos, 1999) with slight modifications. Briefly, 25 mg of each film sample were dissolved in 3 ml of distilled water; then, 0.1 ml of film extract solution were mixed with 7 ml distilled water and 0.5 ml of Folin-Ciocalteau reagent (Merck Company, Darmstadt, Germany). The mixture was incubated at room temperature (25 ºC) before 1.5 ml of sodium carbonate solution and 0.9 ml of distilled water was added. The mixture was stored in a dark chamber for 2 h. The absorbance of the mixture was then measured at 765 nm using a spectrophotometer (Thermo Scientific GENESYS 20, Thermo Fisher Scientific, Inc., Rochester, NY, USA). Gallic acid solutions (Fluka Chemical Company, St. Louis, MO, USA) were used to prepare a calibration curve. The concentration of TPC in the samples was expressed as gallic acid equivalents (GAE) in mg per gram dry weight of the sample and then calculated using an equation that was obtained from the standard graph (R2 ¼ 0.996) given as:
Abs765 ¼ 0:001 mg gallic acid þ 0:027
(7)
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S. aureus TISTR 118, E. coli TISTR 780 and P. aeruginosa (ATCC 27853) were obtained from Thailand Institute of Scientific and Technological Research (TISTR), Thailand, and S. Enteritidis DMST 17368 was obtained from Culture Collection for Medical Microorganism, Department of Medical Sciences, Thailand. Before use, the test strains were grown in nutrient broth (HiMedia Laboratories Pvt. Ltd., Bombay, India) to give a final density of 107 CFU/ml. The film was cut into a disc form of 6 mm diameter. The film discs were placed on nutrient agar (Merck, Darmstadt, Germany) plates which had been previously seeded with inoculum containing indicator microorganisms. The plates were then incubated at 37 C for 24 h for S. aureus, E. coli and S. Enteritidis and at 30 C for 24 h for P. aeruginosa. The diameter of inhibitory zone surrounding film discs and contact area of film discs with agar surface were measured. If there was no clear zone surrounding the film disc, it was marked as no inhibitory zone. Contact area was also used to evaluate growth inhibition underneath the film discs in direct contact with target microorganisms on the agar (Siripatrawan et al., 2013). Sterile paper discs (6 mm diameter) containing 2.5% PE were also tested against the same strains of bacteria. 2.10. Statistical analysis A completely randomized design (CRD) was used as an experimental design, where film forming solutions containing different concentrations of PE were applied as treatments. Chitosan films without PE were used as the control. The experimental data were subjected to one-way analysis of variance (ANOVA). The statistical significance of differences between mean values was established at p 0.05 and the Duncan’s New Multiple Range Test was applied for all statistical analyses.
This experiment was analyzed in five replications. 3. Results and discussion 2.8. Antioxidant activity 3.1. Water vapor permeability The antioxidant activity of the film samples was evaluated using DPPH (2,2-diphenyl-1-picrylhydrazyl) free radical scavenging assay according to Blois (1958) with slight modification. Briefly, 3 ml of film extract solution (25 mg of film sample were dissolved in 3 ml of distilled water) were mixed with 1 ml of 1 mM methanolic solution of DPPH (Fluka Chemical Company, St. Louis, MO, USA). The mixture was vortexed using a vortex-Genie 2 (Scientific Industries, Inc. Bohemia, New York, USA) and incubated in the dark at room temperature (25 ºC) for 30 min. The absorbance was then measured at 517 nm. The percentage of DPPH free radical quenching activity was determined using Eq (8).
DPPH scavenging effect ð%Þ ¼
AbsDPPH Absextract 100 AbsDPPH
Chitosan films without propolis had the highest (p 0.05) WVP among tested films. Addition of propolis reduced (p 0.05) WVP of the films, but this did not change (p > 0.05) with increasing amounts (Fig. 1). Polyphenolic compounds may be able to fit into the chitosan matrix and establish interactions such as hydrogen or covalent bonding with reactive groups of chitosan (Siripatrawan & Harte, 2010; Wu et al., 2013). The hydrogen and covalent interactions between chitosan network and polyphenolic compounds limit the availability of hydrogen groups to form hydrophilic bonding with
(8)
where AbsDPPH was the absorbance value at 517 nm of the methanolic solution of DPPH and Absextract was the absorbance value at 517 nm for the sample extracts. Each sample was assayed at least five replications. 2.9. Antimicrobial activity assay Antimicrobial activity assay of chitosan films containing propolis was carried out using disc diffusion, following the method of Siripatrawan et al. (2013) with slight modifications. For the antimicrobial assay, Gram-positive bacteria (S. aureus) and Gramnegative bacteria (E. coli, Pseudomonas aeruginosa and Salmonella Enteritidis) were selected from food spoilage or pathogenic bacteria, which are commonly found in food products.
Fig. 1. Water vapor permeability and oxygen permeability of chitosan films containing propolis extract.
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water, leading to a decrease in the affinity of chitosan films toward water and lower WVTR of the films. This study’s results are in agreement with those of Curcio et al. (2009) who observed the formation of covalent bonds between gallic acid and chitosan as verified by FTIR. Similar findings were observed by Park and Zhao (2004) who found that by adding mineral or vitamin into a film matrix, interactions among adjacent molecules increased, resulting in a decrease in diffusivity of water vapor through the film matrix and a decrease in hydrophilic susceptibility of the films. Since propolis is a chemically complex resinous bee product and consists of flavonoids, phenolic acids, waxes, essential oils and ~o et al., 2010), a various organic compounds (Bankova, 2005; Falca decrease in WVP with addition of PE was expected. PE consists of some hydrophobic compounds, and the increase in hydrophobicity of the film matrix would reduce water sorption, leading to an improved water barrier property of the films. This behavior was also reported for addition of lipid fractions (oleic acid, beeswax and oleic acidebeeswax mixture) in chitosan films (Hromis et al., 2015).
3.2. Oxygen permeability The OP of the films decreased (p 0.05) as the concentration of PE increased from 0 to 20% w/w of chitosan (Fig. 1). This decrease was probably due to interactions between chitosan polymer matrix and propolis phenolic compounds. Generally, changes in gas permeability depend on film microstructure, thickness, void volume in the polymer structure, and arrangement of the polymer chain. The addition of PE possibly resulted in increased interactions between chitosan monomers, tightening the polymer chain-to-chain interactions and, consequently, led to a decrease in oxygen permeability. Ojagh, Rezaei, Razavi, and Hosseini (2010) evaluated the physical properties of chitosan-based films containing cinnamon essential oil and suggested that a strong interaction between the chitosan polymer and phenolic components of cinnamon essential oil produced a crosslinking effect leading to a compact structure of the resulting films, which decreased the free volume and the molecular mobility of the polymer. Thus, the structural arrangement of chitosan network induced by the phenolic compounds and other propolis constituents could explain the improved barrier property of the chitosan films. A high oxygen barrier property is required in most food packages to maintain qualities and shelf-life of foods since oxygen permeated through packaging materials may cause rancidity and other oxidative degradation reactions in lipid-containing foods resulting in quality losses. The results in this study suggest that it is possible to obtain chitosan films with improved oxygen barrier when PE is incorporated into the films.
3.3. Film color The color values of the films as affected by propolis concentration are shown in Fig. 2. Adding propolis extract into chitosan films affected (p 0.05) L*, a* and b* values of the film surface. The L* values of the films decreased, whereas a* and b* values increased as the PE concentrations increased from 0 to 20%. Increasing PE concentration resulted in deeper orange color films, as compared to light yellow control films. This is a consequence of the colored substances present in the propolis (Pastor et al., 2010). Ojagh et al. (2010) also reported color changes of chitosan films as a consequence of the addition of cinnamon essential oil. Nevertheless, the color of the developed film may affect the consumer acceptability of the products when applying this film to light color foods.
Fig. 2. Color values of chitosan film containing propolis extract.
3.4. Mechanical properties Mechanical properties of chitosan films are associated with intermolecular and intramolecular interaction within the chitosan matrix (Leceta et al., 2013). Tensile strength is the measurement of maximum strength of a film to withstand against applied tensile stress, and percent elongation represents the ability of a film to stretch (Park & Zhao, 2004). The tensile strength increased (p 0.05) when PE concentration increased from 0 to 20% (Fig. 3). The effect of PE incorporation on the strengthening of the corresponding films may be explained by similar assumptions developed for WVP and OP. The improvement in mechanical properties of the films incorporated with PE may be attributed to the interactions between the propolis components (e.g. phenolic acids and their
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699
Fig. 3. Tensile strength and elongation at break of chitosan film containing propolis.
esters) which possess polar characteristics with the hydrophilic groups of the chitosan molecules. These interactions can result in stronger interfacial adhesion between the chitosan molecules and the PE in the film matrix and tightened polymer chain-to-chain interactions leading to a more effective resistance to the mechanical stress (Pastor et al., 2010). The elongation at break of the films increased when propolis concentrations increased from 0 to 10%, but significantly decreased (p 0.05) when 20% propolis was added (Fig. 3). The decrease in the elongation of the films may be ascribed to the crystalline formation of excessive propolis components in the chitosan matrix which leads to a reduction in the film flexibility (Pastor et al., 2010). It is also possible that the interactions between propolis components at high concentration and chitosan would produce a crosslinking effect, which decreases the free volume and molecular mobility of the chitosan polymer, causing a decrease in the elongation (Bodini et al., 2013). Similar findings were also observed by Shen and Kamdem (2015) who found tensile strength increased with increasing citronella essential oil concentration, but elongation decreased at high essential oil concentration and by Ojagh et al. (2010) who reported tensile strength increased when introducing cinnamon essential oil into chitosan films while elongation decreased. Changes in mechanical properties as affected by addition of plant polyphenols were also observed for green tea extract in fish gelatin films (Wu et al., 2013) and microemulsions of cinnamon oil and soybean oil in chitosan films (Ma et al., 2016). Interactions between chitosan and polyphenolic compounds from PE may play a major role on the modification of film properties and will be discussed further based on the FTIR data obtained in the next section.
3.5. Structural properties FTIR spectroscopy was performed to characterize the intermolecular interaction between film matrix and phenolic compounds from PE by measuring the transmittance in the wavenumber range of 4000e650 cm1 at resolution of 1 cm1. The FTIR spectra of propolis extract and chitosan films incorporated with 0, 2.5, 5, 10 and 20% PE are shown in Fig. 4. All film samples show a similar pattern of FTIR spectra, with most of the peaks showing characteristics of chitosan films but with different transmittance intensities at certain peaks. Peaks of NeH stretching in amino group (3500e3300 cm1), OH-stretching (3400e3200 cm1), CeH stretching (3000e2850 cm1), C]O stretching (amide I) (1642 cm1), NeH
Fig. 4. FTIR spectra of chitosan film containing propolis extract.
bending (amide II) (1558 cm1) and CeO (1280e1000 cm1) could be observed and were common to all films. Chitosan spectra presented herein are in line with those presented by Martins, Cerqueira, and Vicente (2012), Silva-Weiss, Bifani, Ihl, Sobral, and mez-Guille n (2013) and Shen and Kamdem (2015). Go For films containing PE, changes taking place at 1700 cm1, ascribable to C]O stretching within the carboxylic group, and peak at 1660 cm1, correlated to C]C stretching within the aromatic ring, indicating functional groups of phenolic compounds in propolis were observed and appeared to be more recognizable with increasing PE concentration. The bands appearing at 2930 and 2870 cm1 showed stronger peak intensity in films containing PE when compared with those of pure chitosan film probably because of stretching vibrations of the CeH bond in eCH2 and eCH3 groups, respectively (Peng, Wu, & Li, 2013; Shen & Kamdem, 2015). The stretching of C]O, -C]CeC]O, -C]C- [(in-ring) aromatic], -C-C- [(in-ring) aromatic], CeO (esters, ethers) and CeO (polyols) found in the phenolic components were observed at 1800e1000 cm1 of the FTIR spectra. Another important change in chitosan films containing PE was found with a peak at 1647 cm1, which was attributed to C]O stretching, indicating an ester linkage and a new peak at 1257 cm1 linked to the bending vibration of OH groups from the phenol group present in the PE. These peak intensities increased with increasing PE concentration, which could be attributed to the more intensive interactions between chitosan and PE. Peaks at 1140e1025 cm1 attributed to phenol group and peak at 870e675 cm1 ascribed to CeH aromatic ring were found in chitosan films containing propolis. It should be noted that CeH stretching at 3000e2840 cm1, -C]C- aromatic ring at 1650 cm1, CeH aromatic ring at 1650e1450 cm1 and 1 870e675 cm observed in the spectra of chitosan containing PE could be found in pure PE (data not shown), indicating the phenolic compounds were incorporated into chitosan film matrix when PE was added. From FTIR, it is evident that intermolecular interaction exists between hydroxyl and amino groups of chitosan matrix and functional groups of polyphenolic compounds from PE. PE polyphenols could form hydrogen bonding and covalent bonding and thus occupy the functional group of the chitosan matrix, subsequently lowering the free hydrogen group which can form hydrophilic bonding with water. These results are comparable to those of Curcio et al. (2009) who observed the formation of covalent bonds
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between gallic acid-chitosan and catechin-chitosan. This information can be used to explain the improved mechanical and barrier properties of the chitosan films containing PE reported in the previous sections. 3.6. Total phenolic content and antioxidant activity Total phenolic content and DPPH in the chitosan films increased (p 0.05) when PE was added and further increased with increasing PE concentration (Fig. 5). The scavenging mechanism of chitosan is related to the fact that free radical can react with the residual free amino (NH2) groups to form stable macromolecule radicals, and the NH2 groups can form ammonium (NHþ 3 ) groups by absorbing a hydrogen ion from the solution (Yen et al., 2008). In the films containing propolis, the antioxidant activity increased in relation to the control samples. The phenolic compounds in Thai propolis extract include rutin, quercetin, naringenin, baicalin, acacetin and galangin with rutin, quercetin and naringenin being the major phenolic compounds (Siripatrawan et al., 2013). Rutin, quercetin and naringenin have been reported to be the major contributors for antioxidant activity in propolis from various geographic locations (Rice-Evans, Miller, & Paganga, 1996). Rutin, quercetin and naringenin are the major phenolic compounds found in Thai propolis extract and, hence, are most likely to contribute to the antioxidant activity of the chitosan films containing PE. The scavenging mechanism of polyphenols in chitosan film containing propolis is related to their reducing properties as hydrogen or electron-donating agents react with free radicals, leading to the inhibition of lipid peroxidation. Due to their antioxidant properties, the chitosan films incorporated with PE are expected to help inhibit detrimental lipid oxidation reaction that may occur in various foods. 3.7. Antimicrobial activity The antimicrobial activity of chitosan incorporated with PE against bacteria was compared with chitosan-alone films, and paper discs containing PE (Fig. 6). Chitosan films without PE did not show any inhibition on the tested bacteria, but when PE was added, antimicrobial activity on contact surface underneath the film discs was evident for all tested bacteria. Although chitosan has been reported to have intrinsic antimicrobial activities against both Gram-positive and Gram-negative ndez-Mun ~ oz, Almenar, Valle, Del Velez, & Gavara, bacteria (Herna 2008), in this research, chitosan film did not show an inhibition of the tested bacteria. It is possible that antimicrobial activity may
Fig. 5. Total phenolic content and DPPH free radical scavenging activity of chitosan film containing propolis extract.
Fig. 6. Antimicrobial activity of paper disc containing propolis extract (P þ PE), chitosan-alone film (CS) and chitosan film containing propolis extract (CS þ PE) against S. aureus, S. Enteritidis, E. coli and P. aeruginosa. Inhibitory zone is clear zone surrounding paper or film discs, calculated by subtracting overall clear zone by diameter of the film disc; colored circular shape indicates no growth in the contact surface area underneath paper or film discs on agar surface; ‘x’ indicates no inhibition of target bacteria.
become negligible when chitosan is in a form of insoluble films. Torlak and Sert (2013) reported that chitosan films did not exhibit antibacterial activity by agar diffusion because chitosan in a film form is unable to diffuse through the adjacent agar media. Tripathi, Mehrotra, and Dutta (2008) investigated the antibacterial activity of chitosan-starch film and the solution of the chitosan-starch film against E. coli, S. aureus and B. subtilis using agar plate diffusion method, and they found that the film solution showed an inhibitory effect against all bacteria while the chitosan-starch film showed negative results. This result suggests that addition of PE enhanced the antimicrobial activity of the chitosan films. Chitosan films containing PE exhibited antimicrobial activity on the contact surface underneath films, but this was not observed for chitosan-alone films. The antimicrobial effect of chitosan containing PE was mainly attributed to propolis flavonoids (e.g. rutin, quercetin, and naringenin) (Mirzoeva et al., 1997; Siripatrawan et al., 2013). Nazzaro, Fratianni, De Martino, Coppola, and De Feo (2013) have suggested that the antimicrobial action of natural plant extracts depends on their chemical composition, and their antimicrobial actions involve biochemical mechanisms at various sites of the entire bacterial cell. ~o Since a variety of molecules are present in propolis extracts (Falca et al., 2010; Rice-Evans et al., 1996), the antibacterial activity of chitosan films containing propolis polyphenols may be explained by different actions, including alteration of membrane permeability, inhibition of DNA and RNA synthesis, chemical modification of the cell membrane, cytoplasm, enzymes and proteins, and deformation of the microbial cell, consequently leading to cell death (Fig. 7) (Hilliard, Krause, & Bernstein, 1995; Mirzoeva et al., 1997; Nazzaro et al., 2013). Chitosan films containing PE showed only antimicrobial activity on the contact surface underneath the films, whereas paper discs containing PE showed inhibitory zone for S. aureus. These results are comparable to those of Altiok, Altiok, and Tihminlioglu (2010) who studied the antimicrobial properties of films containing thyme oil and found no clear zone of E. coli, P. aeruginosa and S. aureus, but no bacterial growth under the films. The possible reasons for the decrease in activity of the PE incorporated in the chitosan films when compared with that of pure PE may be due to potential partial loss of phenolic compounds which have tightly interacted with functional groups of chitosan in the film matrix and thus be incapable of diffusing through the adjacent agar media, preventing expression of the antimicrobial action, or it may be due to slower/controlled release of active compounds from the chitosan
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701
Fig. 7. Antibacterial action of chitosan film containing propolis extract.
film than from cellulose filter paper (Torlak & Sert, 2013). It is also possible that the interaction between propolis polyphenols and chitosan caused diminished contact between bacterial cells and polyphenolic molecules and this could lead to lower antibacterial activity. Moreover, besides flavonoids and phenolic acids and their esters, propolis also consists of waxes, essential oils, and various organic compounds. Chemical interaction between phenolic compounds and such chemical constituents may result in hindering the diffusion or release of antimicrobial phenolic compounds from the film matrix to inhibit bacteria surrounding film discs during agar diffusion assay.
This study showed that propolis extract was more effective against Gram-positive bacteria (S. aureus) than Gram-negative bacteria (E. coli, P. aeruginosa and S. Enteritidis). This may be explained by the structural differences of the bacterial cell wall of Gram-positive and Gram-negative bacteria. Gram-negative bacteria, apart from the cell membrane, possess an additional outer layer membrane, which consists of phospholipids, proteins and lipopolysaccharides, and this membrane is impermeable to most molecules (Inouye, 1986).
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4. Conclusions An active film from chitosan incorporated with PE could be achieved with enhanced functional properties. Incorporation of PE into chitosan films improved mechanical and barrier properties, as well as antimicrobial and antioxidant activities. The modification of film properties could be attributed to the interactions between functional groups of chitosan and polyphenols and other constituents of propolis as verified by FTIR analysis. The developed film has potential to be used as antimicrobial and antioxidant packaging materials which will have a broad application in the food industry. Acknowledgements The authors thank the National Research Council of Thailand for awarding the research grant (No. 2556NRCT53492). The propolis used in this research was supported by the Science for Locale Project under Chulalongkorn University Centenary Academic Development. References Altiok, D., Altiok, E., & Tihminlioglu, F. (2010). Physical, antibacterial and antioxidant properties of chitosan films incorporated with thyme oil for potential wound healing applications. Journal of Material Science, Materials in Medicine, 21, 2227e2236. ASTM.. (2003). Annual book of ASTM standards. Pennsylvania: American Society for Testing and Materials. Bankova, V. (2005). Chemical diversity of propolis and the problem of standardization. Journal of Ethnopharmacology, 100, 114e117. Blois, M. S. (1958). Antioxidant determinations by the use of a stable free radical. Nature, 181, 1199e1200. Bodini, R. B., Sobral, P. J. A., Favaro-Trindade, C. S., & Carvalho, R. A. (2013). Properties of gelatin-based films with added ethanolepropolis extract. LWT - Food Science and Technology, 51, 104e110. van den Broek, L. A. M., Knoop, R. J. I., Kappen, F. H. J., & Boeriu, C. G. (2015). Chitosan films and blends for packaging material. Carbohydrate Polymers, 116, 237e242. Curcio, M., Puoci, F., Iemma, F., Parisi, O. I., Cirillo, G., Spizzirri, U. G., et al. (2009). Covalent insertion of antioxidant molecules on chitosan by a free radical grafting procedure. Journal of Agricultural and Food Chemistry, 57, 5933e5938. Evrendilek, G. A. (2015). Empirical prediction and validation of antibacterial inhibitory effects of various plant essential oils on common pathogenic bacteria. International Journal of Food Microbiology, 202, 35e41. ~o, S. I., Vilas-Boas, M., Estevinho, L. M., Barros, C., Domingues, M. R. M., & Falca Cardoso, S. (2010). Phenolic characterization of northeast portuguese propolis: Usual and unusual compounds. Analytical and Bioanalytical Chemistry, 396, 887e897. Fernandez-Saiz, P., Lagaron, J. M., & Ocio, M. J. (2009). Optimization of the biocide properties of chitosan for its application in the design of active films of interest in the food area. Food Hydrocolloids, 23, 913e921. Guoa, M., Maa, Y., Wanga, C., Liub, H., Li, Q., & Fei, M. (2015). Synthesis, anti-oxidant activity, and biodegradability of a novel recombinant polysaccharide derived from chitosan and lactose. Carbohydrate Polymers, 118, 218e223. ndez-Mun ~ oz, P., Almenar, E., Valle, V. D., Del Velez, V., & Gavara, R. (2008). Herna Effect of chitosan coating combined with postharvest calcium treatment on strawberry (Fragaria ananassa) quality during refrigerated storage. Food Chemistry, 110, 428e435. Hilliard, J. J., Krause, H. M., & Bernstein, J. I. (1995). A comparison of active site binding of 4-quinolones and novel flavones gyrase inhibitors to DNA gyrase. Advances in Experimental Medicine and Biology, 390, 59e69. Hromis, N. M., Lazic, V. L., Markov, S. L., Vastag, Z. G., Popovic, S. Z., Suput, D. Z., et al. (2015). Optimization of chitosan biofilm properties by addition of caraway essential oil and beeswax. Journal of Food Engineering, 158, 86e93. Inouye, M. (1986). Bacterial outer membranes as model systems. New York: Wiley. Leceta, I., Guerrero, P., & de la Caba, K. (2013). Functional properties of chitosanbased films. Carbohydrate Polymers, 93, 339e346. Lores, M., P ajaro, M., Alvarez-Casas, M., Domínguez, J., & García-Jares, C. (2015). Use
of ethyl lactate to extract bioactive compounds from Cytisus scoparius: Comparison of pressurized liquid extraction and medium scale ambient temperature systems. Talanta, 140, 134e142. Madureira, A. R., Pereira, A., & Pintado, M. (2015). Current state on the development of nanoparticles for use against bacterial gastrointestinal pathogens. Focus on chitosan nanoparticles loaded with phenolic compounds. Carbohydrate Polymers, 130, 429e439. Martins, J. T., Cerqueira, M. A., & Vicente, A. A. (2012). Influence of a-tocopherol on physicochemical properties of chitosan-based films. Food Hydrocolloids, 27, 220e227. Ma, Q., Zhang, Y., Critzer, F., Davidson, P. M., Zivanovic, S., & Zhong, Q. (2016). Physical, mechanical, and antimicrobial properties of chitosan films with microemulsions of cinnamon bark oil and soybean oil. Food Hydrocolloids, 52, 533e542. Mirzoeva, O. K., Grishanin, R. N., & Calder, P. C. (1997). Antimicrobial action of propolis and some of its components: The effects on growth, membrane potential and motility of bacteria. Microbiological Research, 152, 239e246. Nazzaro, F., Fratianni, F., De Martino, L., Coppola, F., & De Feo, V. (2013). Effect of essential oils on pathogenic bacteria. Pharmaceuticals (Basel), 6, 1451e1474. Ojagh, S. M., Rezaei, M., Razavi, S. H., & Hosseini, S. M. H. (2010). Development and evaluation of a novel biodegradable film made from chitosan and cinnamon essential oil with low affinity toward water. Food Chemistry, 122(1), 161e166. Park, S., & Zhao, Y. (2004). Incorporation of a high concentration of mineral or vitamin into chitosan-based films. Journal of Agricultural and Food Chemistry, 52, 1933e1939. nchez-Gonz fer, M., Chiralt, A., & Gonza lez-Martínez, C. Pastor, C., Sa alez, L., Cha (2010). Physical and antifungal properties of hydroxypropylmethylcellulose based films containing propolis as affected by moisture content. Carbohydrate Polymers, 82, 1174e1183. Peng, Y., Wu, Y., & Li, Y. (2013). Development of tea extracts and chitosan composite films for active packaging materials. International Journal of Biological Macromolecules, 59, 282e289. Rice-Evans, C. A., Miller, N. J., & Paganga, G. (1996). Structure-antioxidant activity relationship of flavonoids and phenolic acids. Free Radical Biology and Medicine, 20, 933e956. Salvia-Trujillo, L., Rojas-Graü, A., Soliva-Fortuny, R., & Martín-Belloso, O. (2015). Physicochemical characterization and antimicrobial activity of foodgrade emulsions and nanoemulsions incorporating essential oils. Food Hydrocolloids, 43, 547e556. Shen, Z., & Kamdem, D. P. (2015). Development and characterization of biodegradable chitosan films containing two essential oils. International Journal of Biological Macromolecules, 74, 289e296. Silici, S., & Kutluca, S. (2005). Chemical composition and antibacterial activity of propolis collected by three different races of honeybees in the same region. Journal of Ethnopharmacology, 99, 69e73. mez-Guille n, M. C. (2013). Silva-Weiss, A., Bifani, V., Ihl, M., Sobral, P. J. A., & Go Structural properties of films and rheology of film-forming solutions based on chitosan and chitosan-starch blend enriched with murta leaf extract. Food Hydrocolloids, 31, 458e466. Singleton, V. L., Orthofer, R., & Lamuela-Raventos, R. M. (1999). Analysis of total phenols and other oxidation substrates and antioxidants by means of FolinCiocalteu reagent. Methods in Enzymology, 299, 152e178. Siripatrawan, U. (2016). Active food packaging from chitosan incorporated with plant polyphenols. In A. M. Grumezescu (Ed.), Novel approaches of nanotechnology in food (pp. 465e507). London: Academic Press. Siripatrawan, U., & Harte, B. R. (2010). Physical properties and antioxidant activity of an active film from chitosan incorporated with green tea extract. Food Hydrocolloids, 24, 770e775. Siripatrawan, U., Vitchayakitti, W., & Sanguandeekul, R. (2013). Antioxidant and antimicrobial properties of Thai propolis extracted using ethanol aqueous solution. International Journal of Food Science and Technology, 48, 22e27. Torlak, E., & Sert, D. (2013). Antibacterial effectiveness of chitosanepropolis coated polypropylene films against foodborne pathogens. International Journal of Biological Macromolecules, 60, 52e55. Tripathi, S., Mehrotra, G. K., & Dutta, P. K. (2008). Chitosan based antimicrobial films for food packaging applications. e-Polymers, 093, 1e7. Wu, J., Chen, S., Ge, S., Miao, J., Li, J., & Zhang, Q. (2013). Preparation, properties and antioxidant activity of an active film from silver carp (Hypophthalmichthys molitrix) skin gelatin incorporated with green tea extract. Food Hydrocolloids, 32, 42e51. Yen, M., Yang, J., & Mau, J. (2008). Antioxidant properties of chitosan from crab shells. Carbohydrate Polymers, 74, 840e844.
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Improving functional properties of chitosan films as active food packaging by incorporating with propolis Ubonrat Siripatrawan*, Waranya Vitchayakitti Department of Food Technology, Faculty of Science, Chulalongkorn University, Bangkok 10330, Thailand
a r t i c l e i n f o
a b s t r a c t
Article history: Received 11 September 2015 Received in revised form 6 March 2016 Accepted 1 June 2016 Available online 8 June 2016
Chitosan films containing 0, 2.5, 5, 10 and 20% w/w propolis extract (PE), high in polyphenols, were developed. The films’ tensile strength, elongation at break, total phenolic content and antioxidant activity increased, while water vapor permeability and oxygen permeability decreased with increasing propolis concentration. Increasing PE concentration resulted in deeper orange color films, as compared to light yellow control films. The ability of the films to inhibit Staphylococcus aureus, Salmonella Enteritidis, Escherichia coli and Pseudomonas aeruginosa was determined using agar diffusion technique. Chitosan films containing PE could inhibit all tested bacteria on contact surface underneath the film discs. Changes in the Fourier Transform Infrared spectra of the films were observed when PE was incorporated, suggesting some interactions occurred between chitosan and propolis polyphenols. The characterization of mechanical properties, oxygen and moisture barrier, and antioxidant and antimicrobial activities revealed the benefits of adding PE into chitosan films and the potential of using the developed film as active food packaging. © 2016 Elsevier Ltd. All rights reserved.
Keywords: Active packaging Antimicrobial Antioxidant Permeability Propolis
1. Introduction Recently, research and developments in active food packaging have focused on bio-based functional packaging materials incorporating natural active compounds and ingredients (Leceta, Guerrero, & de la Caba, 2013; van den Broek, Knoop, Kappen, & Boeriu, 2015; Madureira, Pereira, & Pintado, 2015). Chitosan is a functional biopolymer with intrinsic antimicrobial and antioxidant properties and consequently, it has high potential to be used as an alternative biodegradable active food package (Fernandez-Saiz, Lagaron, & Ocio, 2009; Guoa et al., 2015; van den Broek et al., 2015). Bio-based packaging materials with antioxidant and antimicrobial properties have become popular since oxidation and microbial contamination are major problems affecting food quality and safety. Many studies have been conducted on the utilization of plant polyphenols as alternatives to synthetic antimicrobial and jaro, Alvarez-Casas, antioxidant agents (Evrendilek, 2015; Lores, Pa Domínguez, & García-Jares, 2015; Madureira et al., 2015; SalviaTrujillo, Rojas-Graü, Soliva-Fortuny, & Martín-Belloso, 2015; Siripatrawan, 2016).
* Corresponding author. E-mail address: [email protected] (U. Siripatrawan). http://dx.doi.org/10.1016/j.foodhyd.2016.06.001 0268-005X/© 2016 Elsevier Ltd. All rights reserved.
Propolis, the natural resinous substance collected by honeybees from various plant sources, is considered a good source of natural antioxidants and antibacterials (Bankova, 2005). Propolis contains a variety of chemical compounds such as polyphenols (flavonoid aglycones, phenolic acids and their esters, phenolic aldehydes, alcohols and ketones), sesquiterpene quinines, coumarins, amino ~o et al., 2010). acids and inorganic compounds (Bankova, 2005; Falca Flavonoids (flavones, flavonols and flavonones), aromatic acids and phenolic compounds are the most important active constituents of propolis and appear to be the principal components responsible for the biological activities of propolis samples (Silici & Kutluca, 2005). Propolis has been reported to possess various biological activities, such as antibacterial, antiviral, antitumor, anti-inflammatory, anticancer, antifungal, and antitumoral properties (Falc~ ao et al., 2010). The antimicrobial effects of propolis against Gram-positive (Bacillus cereus, Listeria monocytogenes and Staphylococcus aureus) and Gram-negative (Salmonella Typhimurium, Escherichia coli and Pseudomonas fluorescence) bacteria have been reported (Silici & Kutluca, 2005; Siripatrawan, Vitchayakitti, & Sanguandeekul, 2013). As a good source of polyphenols with multiple biological activities, propolis has high potential to be used as an active agent that can be incorporated into films. Limited research on incorporation of propolis to enhance properties of packaging films and
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nchez-Gonza lez, Cha fer, coatings has been published. Pastor, Sa lez-Martínez (2010) investigated physical and Chiralt, and Gonza antifungal properties of hydroxymethylcellulose based films containing propolis. Bodini, Sobral, Favaro-Trindade, and Carvalho (2013) studied properties of gelatin films added with propolis, and Torlak and Sert (2013) examined antibacterial effectiveness of chitosan-propolis coated polypropylene film. Nevertheless, research providing functional characterizations, including mechanical properties, oxygen and moisture barrier, structural property, antioxidant activity and antimicrobial property, of chitosan films incorporated with propolis extract (PE) has not been fully documented. Hence, this research aimed to develop and characterize chitosan films with enhanced functional properties for potential use as active food packaging by incorporating chitosan with propolis extract. 2. Materials and methods 2.1. Film preparation Propolis was collected from Nan province in northern Thailand and extracted following the methods of Siripatrawan et al. (2013). Three grams of ground propolis were extracted using 100 ml of 30% ethanol aqueous solution. The solution was extracted at 50 C in a water bath shaking incubator (New Brunswick Scientific, Edison, USA) at 200 oscillation/min for 24 h and then filtered through Whatman filter paper No. 1. The extract solution was concentrated using a rotary evaporator (Rotavapor R-200, BÜCHI Laboratory Equipment, Flawil, Switzerland) under reduced pressure at 45 C. Chitosan (Seafresh Industry Public Company Limited, Chumphon, Thailand) with 95% degree of deacetylation was used to prepare chitosan-based films. The chitosan-based film was prepared according to the procedure of Siripatrawan and Harte (2010) with slight modification. A film-forming solution was prepared by dissolving 2 g of chitosan powder into 100 ml of 1% acetic acid solution. Glycerol (Sigma Chemical Co., St. Louis, MO, USA) was added to the solution at 30% w/w of chitosan. The solution was heated at 60 C in a water bath shaking incubator at 100 rpm for 30 min. The PE was dissolved in the film-forming solution to obtain concentrations of 0, 2.5, 5, 10 and 20% w/w of chitosan. The resulting solutions were homogenized using a homogenizer (D79282, Ystral GmbH, Ballrechten-Dottingen, Germany). Air bubbles in the film-forming solutions were removed using a sonicator (Cole-Parmer, Vernon Hills, Illinois, USA). Each film-forming solution was cast on a ceramic plate. The obtained films were conditioned in a chamber at 25 C and 50% relative humidity for 48 h prior to testing. 2.2. Color measurement Color (L*, a*, b*) values were measured using a Minolta Chromameter (CR-300, Minolta Camera Co., Osaka, Japan). Films were cut into 15 2.5 cm and six readings at different positions on each film were measured. Five replications were conducted for each treatment and five film samples were used for each replication. 2.3. Water vapor transmission rate Water vapor transmission rate (WVTR) of the films was determined following the ASTM standard test method (ASTM., 2003). Film samples, previously equilibrated at 50% RH for 48 h, were sealed in glass cups containing silica gel. The film-covered cups were placed in an environmental chamber set at 25 C and 75% RH. The cups were weighed periodically until steady state was reached (±0.0001 g). The WVTR (g m2 day1) of the films was determined
using Eq (1). At least five replications of each film treatment were tested for the permeability coefficient (WVP). The WVTR of the films was then used to calculate the WVP (g mm m2 d1 Pa1) using Eq (2).
WVTR ¼
Dw ADt
WVP ¼ WVTR
(1) x
Dp
(2)
where Dw is the moisture weight gain, A is the tested film area, Dt is the definite time once steady state was reached, x is the film thickness and Dp is the partial water vapor pressure gradient between the inner and outer surface of the film in the chamber.
2.4. Oxygen transmission rate A single film specimen was cut out to have a measurement area of 50 cm2. O2 transmission rate (OTR) was analyzed using MOCON OX-TRAN 2/20 devices (MOCON, Inc., USA) following the ASTM standard method (ASTM., 2003). The specimens were mounted onto the diffusion cells and a mixture of 98% N2 and 2% H2 was used as a carrier gas. OTR was used to measure the amount of oxygen passing through films when exposed to a gradient with partial O2 pressure across the films. As the oxygen permeated through the film sample, it was picked up by the carrier gas and carried through a coulometric sensor. The amount of oxygen contained in the carrier gas at equilibrium was measured. The test conditions used in the OX-TRAN 2/20 system were 25 C and 50% RH. Duplicate film samples were tested. OTR (Eq (3)) is expressed as the quantity (q) of O2 molecules passing through a film surface area (A) during time (Dt) at steady state. Oxygen permeability coefficient (OP) of the films was then calculated using Eq (4):
OTR ¼
q ADt
OP ¼ OTR
(3) x
Dp
(4)
2.5. Mechanical properties Tensile strength (TS) and percentage elongation at break (%E) were measured with an Instron Universal Testing Machine (Model 5655, Instron Corporation, Canton, MA, USA) following the ASTM Standard Test Method D 882-91 (ASTM., 2003). Each film strip (15 2.5 cm) was mounted between the grips of the Instron and tested with an initial grip separation of 5 cm and crosshead speed of 1 mm/s. TS (Eq. (5)) was calculated by dividing the maximum load (Fmax) by the initial cross-sectional area (4) of the film sample expressed as MPa. %E (Eq. (6)) was calculated as the ratio of the film extension (Dl) at the point of sample rupture to the initial length (l0) of a sample and expressed as a percentage. Measurements represent an average of at least nine replications.
TS ¼
%E ¼
Fmax
F
Dl l0
100
(5)
(6)
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2.6. FTIR analysis Fourier Transform Infrared (FTIR) spectrometry was carried out to observe the structural interactions of chitosan and propolis polyphenols. The FTIR spectra of chitosan films were recorded from 4000 to 650 cm1 using a FTIR spectrometer (PerkinElmer 1760, PerkinElmer Life And Analytical Sciences, Inc., Waltham, Massachusetts USA). 2.7. Total phenolic assay Total phenolic content (TPC) of the chitosan film samples was determined using the Folin-Ciocalteu method (Singleton, Orthofer, & Lamuela-Raventos, 1999) with slight modifications. Briefly, 25 mg of each film sample were dissolved in 3 ml of distilled water; then, 0.1 ml of film extract solution were mixed with 7 ml distilled water and 0.5 ml of Folin-Ciocalteau reagent (Merck Company, Darmstadt, Germany). The mixture was incubated at room temperature (25 ºC) before 1.5 ml of sodium carbonate solution and 0.9 ml of distilled water was added. The mixture was stored in a dark chamber for 2 h. The absorbance of the mixture was then measured at 765 nm using a spectrophotometer (Thermo Scientific GENESYS 20, Thermo Fisher Scientific, Inc., Rochester, NY, USA). Gallic acid solutions (Fluka Chemical Company, St. Louis, MO, USA) were used to prepare a calibration curve. The concentration of TPC in the samples was expressed as gallic acid equivalents (GAE) in mg per gram dry weight of the sample and then calculated using an equation that was obtained from the standard graph (R2 ¼ 0.996) given as:
Abs765 ¼ 0:001 mg gallic acid þ 0:027
(7)
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S. aureus TISTR 118, E. coli TISTR 780 and P. aeruginosa (ATCC 27853) were obtained from Thailand Institute of Scientific and Technological Research (TISTR), Thailand, and S. Enteritidis DMST 17368 was obtained from Culture Collection for Medical Microorganism, Department of Medical Sciences, Thailand. Before use, the test strains were grown in nutrient broth (HiMedia Laboratories Pvt. Ltd., Bombay, India) to give a final density of 107 CFU/ml. The film was cut into a disc form of 6 mm diameter. The film discs were placed on nutrient agar (Merck, Darmstadt, Germany) plates which had been previously seeded with inoculum containing indicator microorganisms. The plates were then incubated at 37 C for 24 h for S. aureus, E. coli and S. Enteritidis and at 30 C for 24 h for P. aeruginosa. The diameter of inhibitory zone surrounding film discs and contact area of film discs with agar surface were measured. If there was no clear zone surrounding the film disc, it was marked as no inhibitory zone. Contact area was also used to evaluate growth inhibition underneath the film discs in direct contact with target microorganisms on the agar (Siripatrawan et al., 2013). Sterile paper discs (6 mm diameter) containing 2.5% PE were also tested against the same strains of bacteria. 2.10. Statistical analysis A completely randomized design (CRD) was used as an experimental design, where film forming solutions containing different concentrations of PE were applied as treatments. Chitosan films without PE were used as the control. The experimental data were subjected to one-way analysis of variance (ANOVA). The statistical significance of differences between mean values was established at p 0.05 and the Duncan’s New Multiple Range Test was applied for all statistical analyses.
This experiment was analyzed in five replications. 3. Results and discussion 2.8. Antioxidant activity 3.1. Water vapor permeability The antioxidant activity of the film samples was evaluated using DPPH (2,2-diphenyl-1-picrylhydrazyl) free radical scavenging assay according to Blois (1958) with slight modification. Briefly, 3 ml of film extract solution (25 mg of film sample were dissolved in 3 ml of distilled water) were mixed with 1 ml of 1 mM methanolic solution of DPPH (Fluka Chemical Company, St. Louis, MO, USA). The mixture was vortexed using a vortex-Genie 2 (Scientific Industries, Inc. Bohemia, New York, USA) and incubated in the dark at room temperature (25 ºC) for 30 min. The absorbance was then measured at 517 nm. The percentage of DPPH free radical quenching activity was determined using Eq (8).
DPPH scavenging effect ð%Þ ¼
AbsDPPH Absextract 100 AbsDPPH
Chitosan films without propolis had the highest (p 0.05) WVP among tested films. Addition of propolis reduced (p 0.05) WVP of the films, but this did not change (p > 0.05) with increasing amounts (Fig. 1). Polyphenolic compounds may be able to fit into the chitosan matrix and establish interactions such as hydrogen or covalent bonding with reactive groups of chitosan (Siripatrawan & Harte, 2010; Wu et al., 2013). The hydrogen and covalent interactions between chitosan network and polyphenolic compounds limit the availability of hydrogen groups to form hydrophilic bonding with
(8)
where AbsDPPH was the absorbance value at 517 nm of the methanolic solution of DPPH and Absextract was the absorbance value at 517 nm for the sample extracts. Each sample was assayed at least five replications. 2.9. Antimicrobial activity assay Antimicrobial activity assay of chitosan films containing propolis was carried out using disc diffusion, following the method of Siripatrawan et al. (2013) with slight modifications. For the antimicrobial assay, Gram-positive bacteria (S. aureus) and Gramnegative bacteria (E. coli, Pseudomonas aeruginosa and Salmonella Enteritidis) were selected from food spoilage or pathogenic bacteria, which are commonly found in food products.
Fig. 1. Water vapor permeability and oxygen permeability of chitosan films containing propolis extract.
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water, leading to a decrease in the affinity of chitosan films toward water and lower WVTR of the films. This study’s results are in agreement with those of Curcio et al. (2009) who observed the formation of covalent bonds between gallic acid and chitosan as verified by FTIR. Similar findings were observed by Park and Zhao (2004) who found that by adding mineral or vitamin into a film matrix, interactions among adjacent molecules increased, resulting in a decrease in diffusivity of water vapor through the film matrix and a decrease in hydrophilic susceptibility of the films. Since propolis is a chemically complex resinous bee product and consists of flavonoids, phenolic acids, waxes, essential oils and ~o et al., 2010), a various organic compounds (Bankova, 2005; Falca decrease in WVP with addition of PE was expected. PE consists of some hydrophobic compounds, and the increase in hydrophobicity of the film matrix would reduce water sorption, leading to an improved water barrier property of the films. This behavior was also reported for addition of lipid fractions (oleic acid, beeswax and oleic acidebeeswax mixture) in chitosan films (Hromis et al., 2015).
3.2. Oxygen permeability The OP of the films decreased (p 0.05) as the concentration of PE increased from 0 to 20% w/w of chitosan (Fig. 1). This decrease was probably due to interactions between chitosan polymer matrix and propolis phenolic compounds. Generally, changes in gas permeability depend on film microstructure, thickness, void volume in the polymer structure, and arrangement of the polymer chain. The addition of PE possibly resulted in increased interactions between chitosan monomers, tightening the polymer chain-to-chain interactions and, consequently, led to a decrease in oxygen permeability. Ojagh, Rezaei, Razavi, and Hosseini (2010) evaluated the physical properties of chitosan-based films containing cinnamon essential oil and suggested that a strong interaction between the chitosan polymer and phenolic components of cinnamon essential oil produced a crosslinking effect leading to a compact structure of the resulting films, which decreased the free volume and the molecular mobility of the polymer. Thus, the structural arrangement of chitosan network induced by the phenolic compounds and other propolis constituents could explain the improved barrier property of the chitosan films. A high oxygen barrier property is required in most food packages to maintain qualities and shelf-life of foods since oxygen permeated through packaging materials may cause rancidity and other oxidative degradation reactions in lipid-containing foods resulting in quality losses. The results in this study suggest that it is possible to obtain chitosan films with improved oxygen barrier when PE is incorporated into the films.
3.3. Film color The color values of the films as affected by propolis concentration are shown in Fig. 2. Adding propolis extract into chitosan films affected (p 0.05) L*, a* and b* values of the film surface. The L* values of the films decreased, whereas a* and b* values increased as the PE concentrations increased from 0 to 20%. Increasing PE concentration resulted in deeper orange color films, as compared to light yellow control films. This is a consequence of the colored substances present in the propolis (Pastor et al., 2010). Ojagh et al. (2010) also reported color changes of chitosan films as a consequence of the addition of cinnamon essential oil. Nevertheless, the color of the developed film may affect the consumer acceptability of the products when applying this film to light color foods.
Fig. 2. Color values of chitosan film containing propolis extract.
3.4. Mechanical properties Mechanical properties of chitosan films are associated with intermolecular and intramolecular interaction within the chitosan matrix (Leceta et al., 2013). Tensile strength is the measurement of maximum strength of a film to withstand against applied tensile stress, and percent elongation represents the ability of a film to stretch (Park & Zhao, 2004). The tensile strength increased (p 0.05) when PE concentration increased from 0 to 20% (Fig. 3). The effect of PE incorporation on the strengthening of the corresponding films may be explained by similar assumptions developed for WVP and OP. The improvement in mechanical properties of the films incorporated with PE may be attributed to the interactions between the propolis components (e.g. phenolic acids and their
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699
Fig. 3. Tensile strength and elongation at break of chitosan film containing propolis.
esters) which possess polar characteristics with the hydrophilic groups of the chitosan molecules. These interactions can result in stronger interfacial adhesion between the chitosan molecules and the PE in the film matrix and tightened polymer chain-to-chain interactions leading to a more effective resistance to the mechanical stress (Pastor et al., 2010). The elongation at break of the films increased when propolis concentrations increased from 0 to 10%, but significantly decreased (p 0.05) when 20% propolis was added (Fig. 3). The decrease in the elongation of the films may be ascribed to the crystalline formation of excessive propolis components in the chitosan matrix which leads to a reduction in the film flexibility (Pastor et al., 2010). It is also possible that the interactions between propolis components at high concentration and chitosan would produce a crosslinking effect, which decreases the free volume and molecular mobility of the chitosan polymer, causing a decrease in the elongation (Bodini et al., 2013). Similar findings were also observed by Shen and Kamdem (2015) who found tensile strength increased with increasing citronella essential oil concentration, but elongation decreased at high essential oil concentration and by Ojagh et al. (2010) who reported tensile strength increased when introducing cinnamon essential oil into chitosan films while elongation decreased. Changes in mechanical properties as affected by addition of plant polyphenols were also observed for green tea extract in fish gelatin films (Wu et al., 2013) and microemulsions of cinnamon oil and soybean oil in chitosan films (Ma et al., 2016). Interactions between chitosan and polyphenolic compounds from PE may play a major role on the modification of film properties and will be discussed further based on the FTIR data obtained in the next section.
3.5. Structural properties FTIR spectroscopy was performed to characterize the intermolecular interaction between film matrix and phenolic compounds from PE by measuring the transmittance in the wavenumber range of 4000e650 cm1 at resolution of 1 cm1. The FTIR spectra of propolis extract and chitosan films incorporated with 0, 2.5, 5, 10 and 20% PE are shown in Fig. 4. All film samples show a similar pattern of FTIR spectra, with most of the peaks showing characteristics of chitosan films but with different transmittance intensities at certain peaks. Peaks of NeH stretching in amino group (3500e3300 cm1), OH-stretching (3400e3200 cm1), CeH stretching (3000e2850 cm1), C]O stretching (amide I) (1642 cm1), NeH
Fig. 4. FTIR spectra of chitosan film containing propolis extract.
bending (amide II) (1558 cm1) and CeO (1280e1000 cm1) could be observed and were common to all films. Chitosan spectra presented herein are in line with those presented by Martins, Cerqueira, and Vicente (2012), Silva-Weiss, Bifani, Ihl, Sobral, and mez-Guille n (2013) and Shen and Kamdem (2015). Go For films containing PE, changes taking place at 1700 cm1, ascribable to C]O stretching within the carboxylic group, and peak at 1660 cm1, correlated to C]C stretching within the aromatic ring, indicating functional groups of phenolic compounds in propolis were observed and appeared to be more recognizable with increasing PE concentration. The bands appearing at 2930 and 2870 cm1 showed stronger peak intensity in films containing PE when compared with those of pure chitosan film probably because of stretching vibrations of the CeH bond in eCH2 and eCH3 groups, respectively (Peng, Wu, & Li, 2013; Shen & Kamdem, 2015). The stretching of C]O, -C]CeC]O, -C]C- [(in-ring) aromatic], -C-C- [(in-ring) aromatic], CeO (esters, ethers) and CeO (polyols) found in the phenolic components were observed at 1800e1000 cm1 of the FTIR spectra. Another important change in chitosan films containing PE was found with a peak at 1647 cm1, which was attributed to C]O stretching, indicating an ester linkage and a new peak at 1257 cm1 linked to the bending vibration of OH groups from the phenol group present in the PE. These peak intensities increased with increasing PE concentration, which could be attributed to the more intensive interactions between chitosan and PE. Peaks at 1140e1025 cm1 attributed to phenol group and peak at 870e675 cm1 ascribed to CeH aromatic ring were found in chitosan films containing propolis. It should be noted that CeH stretching at 3000e2840 cm1, -C]C- aromatic ring at 1650 cm1, CeH aromatic ring at 1650e1450 cm1 and 1 870e675 cm observed in the spectra of chitosan containing PE could be found in pure PE (data not shown), indicating the phenolic compounds were incorporated into chitosan film matrix when PE was added. From FTIR, it is evident that intermolecular interaction exists between hydroxyl and amino groups of chitosan matrix and functional groups of polyphenolic compounds from PE. PE polyphenols could form hydrogen bonding and covalent bonding and thus occupy the functional group of the chitosan matrix, subsequently lowering the free hydrogen group which can form hydrophilic bonding with water. These results are comparable to those of Curcio et al. (2009) who observed the formation of covalent bonds
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between gallic acid-chitosan and catechin-chitosan. This information can be used to explain the improved mechanical and barrier properties of the chitosan films containing PE reported in the previous sections. 3.6. Total phenolic content and antioxidant activity Total phenolic content and DPPH in the chitosan films increased (p 0.05) when PE was added and further increased with increasing PE concentration (Fig. 5). The scavenging mechanism of chitosan is related to the fact that free radical can react with the residual free amino (NH2) groups to form stable macromolecule radicals, and the NH2 groups can form ammonium (NHþ 3 ) groups by absorbing a hydrogen ion from the solution (Yen et al., 2008). In the films containing propolis, the antioxidant activity increased in relation to the control samples. The phenolic compounds in Thai propolis extract include rutin, quercetin, naringenin, baicalin, acacetin and galangin with rutin, quercetin and naringenin being the major phenolic compounds (Siripatrawan et al., 2013). Rutin, quercetin and naringenin have been reported to be the major contributors for antioxidant activity in propolis from various geographic locations (Rice-Evans, Miller, & Paganga, 1996). Rutin, quercetin and naringenin are the major phenolic compounds found in Thai propolis extract and, hence, are most likely to contribute to the antioxidant activity of the chitosan films containing PE. The scavenging mechanism of polyphenols in chitosan film containing propolis is related to their reducing properties as hydrogen or electron-donating agents react with free radicals, leading to the inhibition of lipid peroxidation. Due to their antioxidant properties, the chitosan films incorporated with PE are expected to help inhibit detrimental lipid oxidation reaction that may occur in various foods. 3.7. Antimicrobial activity The antimicrobial activity of chitosan incorporated with PE against bacteria was compared with chitosan-alone films, and paper discs containing PE (Fig. 6). Chitosan films without PE did not show any inhibition on the tested bacteria, but when PE was added, antimicrobial activity on contact surface underneath the film discs was evident for all tested bacteria. Although chitosan has been reported to have intrinsic antimicrobial activities against both Gram-positive and Gram-negative ndez-Mun ~ oz, Almenar, Valle, Del Velez, & Gavara, bacteria (Herna 2008), in this research, chitosan film did not show an inhibition of the tested bacteria. It is possible that antimicrobial activity may
Fig. 5. Total phenolic content and DPPH free radical scavenging activity of chitosan film containing propolis extract.
Fig. 6. Antimicrobial activity of paper disc containing propolis extract (P þ PE), chitosan-alone film (CS) and chitosan film containing propolis extract (CS þ PE) against S. aureus, S. Enteritidis, E. coli and P. aeruginosa. Inhibitory zone is clear zone surrounding paper or film discs, calculated by subtracting overall clear zone by diameter of the film disc; colored circular shape indicates no growth in the contact surface area underneath paper or film discs on agar surface; ‘x’ indicates no inhibition of target bacteria.
become negligible when chitosan is in a form of insoluble films. Torlak and Sert (2013) reported that chitosan films did not exhibit antibacterial activity by agar diffusion because chitosan in a film form is unable to diffuse through the adjacent agar media. Tripathi, Mehrotra, and Dutta (2008) investigated the antibacterial activity of chitosan-starch film and the solution of the chitosan-starch film against E. coli, S. aureus and B. subtilis using agar plate diffusion method, and they found that the film solution showed an inhibitory effect against all bacteria while the chitosan-starch film showed negative results. This result suggests that addition of PE enhanced the antimicrobial activity of the chitosan films. Chitosan films containing PE exhibited antimicrobial activity on the contact surface underneath films, but this was not observed for chitosan-alone films. The antimicrobial effect of chitosan containing PE was mainly attributed to propolis flavonoids (e.g. rutin, quercetin, and naringenin) (Mirzoeva et al., 1997; Siripatrawan et al., 2013). Nazzaro, Fratianni, De Martino, Coppola, and De Feo (2013) have suggested that the antimicrobial action of natural plant extracts depends on their chemical composition, and their antimicrobial actions involve biochemical mechanisms at various sites of the entire bacterial cell. ~o Since a variety of molecules are present in propolis extracts (Falca et al., 2010; Rice-Evans et al., 1996), the antibacterial activity of chitosan films containing propolis polyphenols may be explained by different actions, including alteration of membrane permeability, inhibition of DNA and RNA synthesis, chemical modification of the cell membrane, cytoplasm, enzymes and proteins, and deformation of the microbial cell, consequently leading to cell death (Fig. 7) (Hilliard, Krause, & Bernstein, 1995; Mirzoeva et al., 1997; Nazzaro et al., 2013). Chitosan films containing PE showed only antimicrobial activity on the contact surface underneath the films, whereas paper discs containing PE showed inhibitory zone for S. aureus. These results are comparable to those of Altiok, Altiok, and Tihminlioglu (2010) who studied the antimicrobial properties of films containing thyme oil and found no clear zone of E. coli, P. aeruginosa and S. aureus, but no bacterial growth under the films. The possible reasons for the decrease in activity of the PE incorporated in the chitosan films when compared with that of pure PE may be due to potential partial loss of phenolic compounds which have tightly interacted with functional groups of chitosan in the film matrix and thus be incapable of diffusing through the adjacent agar media, preventing expression of the antimicrobial action, or it may be due to slower/controlled release of active compounds from the chitosan
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Fig. 7. Antibacterial action of chitosan film containing propolis extract.
film than from cellulose filter paper (Torlak & Sert, 2013). It is also possible that the interaction between propolis polyphenols and chitosan caused diminished contact between bacterial cells and polyphenolic molecules and this could lead to lower antibacterial activity. Moreover, besides flavonoids and phenolic acids and their esters, propolis also consists of waxes, essential oils, and various organic compounds. Chemical interaction between phenolic compounds and such chemical constituents may result in hindering the diffusion or release of antimicrobial phenolic compounds from the film matrix to inhibit bacteria surrounding film discs during agar diffusion assay.
This study showed that propolis extract was more effective against Gram-positive bacteria (S. aureus) than Gram-negative bacteria (E. coli, P. aeruginosa and S. Enteritidis). This may be explained by the structural differences of the bacterial cell wall of Gram-positive and Gram-negative bacteria. Gram-negative bacteria, apart from the cell membrane, possess an additional outer layer membrane, which consists of phospholipids, proteins and lipopolysaccharides, and this membrane is impermeable to most molecules (Inouye, 1986).
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4. Conclusions An active film from chitosan incorporated with PE could be achieved with enhanced functional properties. Incorporation of PE into chitosan films improved mechanical and barrier properties, as well as antimicrobial and antioxidant activities. The modification of film properties could be attributed to the interactions between functional groups of chitosan and polyphenols and other constituents of propolis as verified by FTIR analysis. The developed film has potential to be used as antimicrobial and antioxidant packaging materials which will have a broad application in the food industry. Acknowledgements The authors thank the National Research Council of Thailand for awarding the research grant (No. 2556NRCT53492). The propolis used in this research was supported by the Science for Locale Project under Chulalongkorn University Centenary Academic Development. References Altiok, D., Altiok, E., & Tihminlioglu, F. (2010). Physical, antibacterial and antioxidant properties of chitosan films incorporated with thyme oil for potential wound healing applications. Journal of Material Science, Materials in Medicine, 21, 2227e2236. ASTM.. (2003). Annual book of ASTM standards. Pennsylvania: American Society for Testing and Materials. Bankova, V. (2005). Chemical diversity of propolis and the problem of standardization. Journal of Ethnopharmacology, 100, 114e117. Blois, M. S. (1958). Antioxidant determinations by the use of a stable free radical. Nature, 181, 1199e1200. Bodini, R. B., Sobral, P. J. A., Favaro-Trindade, C. S., & Carvalho, R. A. (2013). Properties of gelatin-based films with added ethanolepropolis extract. LWT - Food Science and Technology, 51, 104e110. van den Broek, L. A. M., Knoop, R. J. I., Kappen, F. H. J., & Boeriu, C. G. (2015). Chitosan films and blends for packaging material. Carbohydrate Polymers, 116, 237e242. Curcio, M., Puoci, F., Iemma, F., Parisi, O. I., Cirillo, G., Spizzirri, U. G., et al. (2009). Covalent insertion of antioxidant molecules on chitosan by a free radical grafting procedure. Journal of Agricultural and Food Chemistry, 57, 5933e5938. Evrendilek, G. A. (2015). Empirical prediction and validation of antibacterial inhibitory effects of various plant essential oils on common pathogenic bacteria. International Journal of Food Microbiology, 202, 35e41. ~o, S. I., Vilas-Boas, M., Estevinho, L. M., Barros, C., Domingues, M. R. M., & Falca Cardoso, S. (2010). Phenolic characterization of northeast portuguese propolis: Usual and unusual compounds. Analytical and Bioanalytical Chemistry, 396, 887e897. Fernandez-Saiz, P., Lagaron, J. M., & Ocio, M. J. (2009). Optimization of the biocide properties of chitosan for its application in the design of active films of interest in the food area. Food Hydrocolloids, 23, 913e921. Guoa, M., Maa, Y., Wanga, C., Liub, H., Li, Q., & Fei, M. (2015). Synthesis, anti-oxidant activity, and biodegradability of a novel recombinant polysaccharide derived from chitosan and lactose. Carbohydrate Polymers, 118, 218e223. ndez-Mun ~ oz, P., Almenar, E., Valle, V. D., Del Velez, V., & Gavara, R. (2008). Herna Effect of chitosan coating combined with postharvest calcium treatment on strawberry (Fragaria ananassa) quality during refrigerated storage. Food Chemistry, 110, 428e435. Hilliard, J. J., Krause, H. M., & Bernstein, J. I. (1995). A comparison of active site binding of 4-quinolones and novel flavones gyrase inhibitors to DNA gyrase. Advances in Experimental Medicine and Biology, 390, 59e69. Hromis, N. M., Lazic, V. L., Markov, S. L., Vastag, Z. G., Popovic, S. Z., Suput, D. Z., et al. (2015). Optimization of chitosan biofilm properties by addition of caraway essential oil and beeswax. Journal of Food Engineering, 158, 86e93. Inouye, M. (1986). Bacterial outer membranes as model systems. New York: Wiley. Leceta, I., Guerrero, P., & de la Caba, K. (2013). Functional properties of chitosanbased films. Carbohydrate Polymers, 93, 339e346. Lores, M., P ajaro, M., Alvarez-Casas, M., Domínguez, J., & García-Jares, C. (2015). Use
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