Properties and microstructure of protein-based film from round scad (Decapterus maruadsi) muscle as affected by palm oil and chitosan incorporation

International Journal of Biological Macromolecules 41 (2007) 605–614

Properties and microstructure of protein-based film from round scad (Decapterus maruadsi) muscle as affected by palm oil and chitosan incorporation Thummanoon Prodpran a,∗ , Soottawat Benjakul b , Anuchit Artharn b a

Department of Material Product Technology, Faculty of Agro-Industry, Prince of Songkla University, Hat Yai, Songkhla 90112, Thailand b Department of Food Technology, Faculty of Agro-Industry, Prince of Songkla University, Hat Yai, Songkhla 90112, Thailand Received 21 March 2007; received in revised form 29 July 2007; accepted 30 July 2007 Available online 6 August 2007

Abstract The properties of protein-based film prepared from round scad (Decapterus maruadsi) muscle in the absence and the presence of palm oil and/or chitosan were investigated. Films added with 25% palm oil (as glycerol substitiution) had the slight decrease in water vapor permeability (WVP) and elongation at break (EAB) (p < 0.05). WVP and tensile strength (TS) of films increased but EAB decreased when 10–40% chitosan (as protein substitution) was incorporated (p < 0.05). Hydrophobic interactions and hydrogen bonds, together with disulfide and non-disulfide covalent bonds, played an important role in stabilizing the film matrix. The a* and b*-values increased with increasing chitosan levels (p < 0.05). Films added with chitosan were less transparent and had the lowered transmission in the visible range. The incorporation of 25% palm oil and 40% chitosan yielded the films with the improved TS but decreased water vapor barrier property. Apart from film strengthening effect, chitosan inconjunction with Tween-20 most likely functioned as the emulsifier/stabilizer in film forming solution containing palm oil. © 2007 Elsevier B.V. All rights reserved. Keywords: Protein film; Round scad; Chitosan; Lipid; Muscle proteins

1. Introduction Biodegradable or edible films from biopolymers have currently received increasing attention from both academic and industrial points of view. Due to their biodegradability, they can be used to replace the synthetic packaging films which mainly cause the environmental problems. Proteins are important biopolymers possessing good film-forming ability. Films with varying properties can be prepared from both plant and animal proteins [1]. Generally, films from different protein types and origins exhibit different properties due to the differences in molecular structure and compositions. Among proteins, fish proteins including myofibrillar and sarcoplasmic proteins have been used as film forming materials [2–6]. Different fish muscles with various constituents may affect the characteristic of resulting films differently.

∗

Corresponding author. Tel.: +66 7428 6357; fax: +66 7421 2889. E-mail address: [email protected] (T. Prodpran).

0141-8130/$ – see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.ijbiomac.2007.07.020

Dark muscle fish species, such as round scad, currently make up 40–50% of the total fish catch in the world. There is a great interest in using the large quantities of these low-value fatty pelagic fish to produce the new products with a higher market value such as for surimi production. However, the producing of surimi from the small pelagic species has been facing the problem with difficulty in making the high quality surimi. Therefore, utilization of dark muscle fish for protein-based film production might be a promising alternative. Recently, film from the meat of round scad, a dark-fleshed fish, has been successfully prepared [7]. Muscle types and washing affecting the properties of resulting round scad protein-based film have been reported [7]. Protein-based films have good mechanical properties due to the interactions between protein chains via disulfide (S–S) covalent bonding, hydrogen bonding, electrostatic forces and hydrophobic interactions [4,8]. However, protein-based films are highly water and moisture absorptions, owing to hydrophilicity of amino acids in protein molecules. Additionally, hydrophilic plasticizers (such as polyols and mono-, di- and oligosaccharide) necessarily incorporated into the protein films to

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improve the film flexibility decrease water vapor barrier properties of the films [9]. Properties of protein-based films depend on various factors such as the source of protein, pH of protein solution, plasticizers, the preparation conditions and substances incorporated into the film-forming solutions [10,11]. To improve water vapor barrier properties of the protein films, hydrophobic substances such as fats and oils could be added. Fat and lipids of different types have been successfully incorporated into protein- and carbohydrate-based films by means of lamination and dispersion or emulsion [12–14]. Moreover, mechanical properties of protein-based film can be improved by incorporation with other miscible biopolymers such as chitosan. Chitosan derived by deacetylation of chitin is heteropolysaccharide comprising mainly ␤-(1,4)-2-deoxy-2-amino-d-glucopyranose units and partially ␤-(1,4)-2-deoxy-2-acetamido-d-glucopyranose [15,16]. Chitosan can be utilized for biodegradable/edible coating and film preparation. Chitosan films generally have lower water vapor permeability than protein films [17,18]. Chitosan films also exhibited high extensibility as well as excellent mechanical and oxygen barrier properties [15,17–19]. Moreover, chitosan exhibits emulsifying property since its molecules consist of both hydrophilic and hydrophobic portions. It can be used as emulsifier to uniformly stabilize oil droplets in emulsion systems [20]. Thus, chitosan can be used to strengthen the film as well as to emulsify the oil added in film-forming solution. Nevertheless, no information regarding the use of chitosan in fish protein/oil composite films has been reported. Addition of oil and chitosan to the film of round scad protein may improve some physical and physicochemical properties of the films. The objective of this work was to study the properties and morphology of protein-based film from round scad as affected by oil and chitosan incorporation. 2. Experimental procedure 2.1. Fish sample Round scad (Decapterus maruadsi) with an average weight of 85–90 g were obtained from Songkhla–Pattani coast along the Gulf of Thailand. The fish were stored in ice and off–loaded approximately 12 h after capture. Fish were transported in ice to the Department of Food Technology, Prince of Songkla University within 2 h. Upon arrival, fish were immediately washed, filleted and minced using a mincer with the hole diameter of 0.5 cm. Mince obtained were stored on ice until used for film preparation. 2.2. Chemicals Urea, sodium dodecylsulfate (SDS), ␤-mercaptoethanol (␤-ME), high-molecular-weight chitosan and Tween-20 (polysorbate-20) were purchased from Sigma (St. Louis, MO, USA). Acrylamide, N,N,N ,N -tetramethylethylenediamine (TEMED) and bis-acrylamide were procured from Fluka (Buchs, Switzerland). Glycerol was purchased from Wako Pure

Chemical Industries, Ltd. (Tokyo, Japan). Palm oil was obtained from Morakot Industries Co., Ltd. (Bangkok, Thailand). 2.3. Preparation of protein-based film from round scad muscle with and without oil and chitosan incorporation The film-forming solution from round scad muscle was prepared according to the method of Chinabhark et al. [6]. The mince of known protein content, determined by Kjedahl method, was added with 3 vol. of distilled water and homogenized at 13,000 rpm for 1 min using an IKA homogenizer (IKA Labortechnik, Selangor, Malaysia). The protein concentration of the film-forming solution was fixed at 2% (w/v). Film-forming solution was then added with glycerol at 50% (w/w) of protein. The mixture was stirred gently for 30 min at room temperature. Subsequently, to solubilize protein, the pH of the film-forming solution was adjusted to 3 using 1 M HCl. The film-forming solution obtained was filtered through a layer of nylon sheet. The filtrate was used for film casting. To study the effect of oil on film properties, the film-forming solutions were prepared as described previously. The palm oil was then added as a substitute of glycerol at 25% (w/w). To stabilize emulsion, Tween-20 at 15% of added palm oil was mixed thoroughly with the film-forming solution prior to palm oil addition. The mixture was homogenized at 13,000 rpm for 4 min using an IKA homogenizer and allowed to stand for 2 min at room temperature (28–30 ◦ C) before subjected to analyses and film casting. Chitosan was also added into the film-forming solutions in the presence and the absence of 25% oil. Before the addition, chitosan solution at 10% (w/v) in 1% acetic acid was prepared. A designated amount of chitosan solution was then added to film-forming solution to obtain the chitosan concentration of 0, 10, 25 and 40% (w/w) as the protein substitute. For the solutions containing oil, the mixtures were homogenized in the presence of 15% Tween-20 as previously described. To prepare the film, the film-forming solutions/mixtures (4 g) was cast onto a rimmed silicone resin plate (50 × 50 mm2 ) and air blown for 12 h at room temperature prior to further drying at 25 ◦ C and 50% relative humidity (RH) for 24 h in an environmental chamber (WTB Binder, Tuttlingen, Germany). The resulting films were manually peeled off and used for analyses. 2.4. Analysis of film-forming mixtures 2.4.1. Oil particle size and distribution Particle size and distribution of oil droplets in the filmforming mixture were determined using optical microscope equipped with digital camera (Olympus R-2268, DP11, CA, USA). 2.4.2. Emulsion stability Emulsion stability was determined according to the method of Rodriguez et al. [20]. The film-forming mixtures were allowed to stand up to 48 h. The occurrence of phase separation and the elapsed time until the phase separation took place were recorded.

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2.5. Determination of properties and protein patterns of film 2.5.1. Film thickness The thickness of film was measured using a micrometer (Gotech, Model GT-313-A, Gotech testing machines Inc., Taiwan). Five random locations around each film of ten film samples were used for thickness determination. 2.5.2. Mechanical properties Prior to testing the mechanical properties, films were conditioned for 48 h at 50 ± 5% relative humidity (RH) at 25 ◦ C. Tensile strength (TS) and elongation at break (EAB) were determined as described by Iwata et al. [5] using the Universal Testing Machine (Lloyd Instrument, Hampshire, UK). Five samples (2 × 5 cm2 ) with initial grip length of 3 cm were used for testing. Cross-head speed was set at 30 mm/min. 2.5.3. Water vapor permeability (WVP) WVP was measured using a modified ASTM method as described by Shiku et al. [3]. The film was sealed on a glass permeation cup (3 cm diameter, 7.5 cm height) containing silica gel (0% RH) with silicone vacuum grease and a rubber gasket to hold the film in place. The cups were placed at 30 ◦ C in a desiccator (30 cm diameter) containing the distilled water (100% RH as measured by a hygrometer). The cups were weighed at 1 h intervals over a 10 h period. WVP of the film was calculated as follows [21]: WVP (g m−1 s−1 Pa−1 ) = wlA−1 t −1 (P2 − P1 )−1 where w is the weight gain of the cup (g), l is the film thickness (m), A is the exposed area of film (m2 ), t is the time of gain (s), (P2 − P1 ) is the vapor pressure difference across the film (Pa). Three films were used for WVP testing and the measurement was run in duplicate. 2.5.4. Color, light transmission and film transparency Color of the film was determined using CIE colorimeter (Hunter associates laboratory, Inc., VA, USA) and expressed as L*, a* and b*. The ultraviolet (UV) and visible light barrier properties of the films were measured at selected wavelengths between 200 and 800 nm using the UV-16001 spectrophotometer (Shimadzu, Kyoto, Japan) as described by Fang et al. [22]. The transparency value of film was calculated by the following equation [23]: transparency value =

−log T600 l

where T600 is the fractional transmittance at 600 nm and l is the film thickness (mm). 2.5.5. Film solubility and protein solubility Film solubility was determined according to the method of Gennadios et al. [24]. The conditioned film samples (2 × 4 cm2 ) were weighed and placed in 50 ml centrifuge tube containing 10 ml of distilled water with 0.1% (w/v) sodium azide. The

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mixture was shaken at a speed of 250 rpm using a shaker (Heidolth Inkubator 10000, Schwabach, Germany) at 30 ◦ C for 24 h. Undissolved debris was removed by centrifugation at 3000 × g for 20 min using a microcentrifuge (MIKRO-20, Hettich Zentrifugan, Germany). The pellet was dried at 105 ◦ C for 24 h and weighed. The weight of solubilized dry matter was calculated by substracting the weight of unsolubilized dry matter from the initial weight of dry matter and expressed as the percentage of total weight. Protein concentration in the supernatant was also determined using the Bradford method [25]. Protein solubility was expressed as the percentage of total protein in the film, which was solubilized with 0.5 M NaOH at 30 ◦ C for 24 h [6]. 2.5.6. Protein patterns Protein patterns of films were determined by SDS-PAGE using 4% stacking gel and 10% running gel according to the method of Laemmli [26]. To solubilize the films prior to SDSPAGE analysis, films were mixed with 20 mM Tris–HCl (pH 8.8) containing 2% SDS and 8 M urea in the presence and the absence of 2% ␤-ME. The mixture was homogenized at 13,000 rpm for 1 min. The homogenate was stirred continuously for 24 h at room temperature (28–30 ◦ C). Then, the sample was centrifuged at 7500 × g for 10 min at room temperature using a Biofuge primo Centrifuge (Sorvall, Hanau, Germany). The supernatant containing 15 ␮g protein determined by the Biuret method [27] was loaded onto the gel and subjected to electrophoresis at a constant current of 15 mA per gel using a Mini-Protean II unit (Bio-Rad Laboratories, Inc., Richmond, CA, USA). After separation, the proteins were stained with 0.02% (w/v) Coomassie Brilliant Blue R-250 in 50% (v/v) methanol and 7.5% (v/v) acetic acid and destained with 50% (v/v) methanol and 7.5% (v/v) acetic acid for 12 min, followed by 5% (v/v) methanol and 7.5% (v/v) acetic acid for 3 h. 2.5.7. Determination of bondings stabilizing the films To elucidate the bondings stabilizing the film, various solvents were used to solubilize the films as described by Chawla et al. [28]. The solvents used included: 0.6 M KCl (S1), 20 mM Tris–HCl (pH 8.0) (S2), 20 mM Tris–HCl (pH 8.0) containing 1% (w/v) SDS (S3), 20 mM Tris–HCl (pH 8.0) containing 1% (w/v) SDS and 8 mM urea (S4), 20 mM Tris–HCl (pH 8.0) containing 1% (w/v) SDS, 8 mM urea and 2% (v/v) ␤-ME (S5). The film samples (0.5 g) were homogenized in 10 ml of various solvents for 1 min at a speed of 13,000 rpm using a homogenizer. The homogenate with S5 was heated in boiled water (100 ◦ C) for 2 min and stirred at room temperature for 4 h. The resulting homogenate was centrifuged at 7500 × g for 30 min using a microcentrifuge. Protein in the supernatant (10 ml) was precipitated by adding 50% (w/v) cold TCA to a final concentration of 10%. The mixture was kept at 4 ◦ C for 18 h and centrifuged at 7500 × g for 30 min. The precipitate was washed with 10% TCA and solubilized in 0.5 M NaOH. The protein content was measured using the Biuret method [27]. To obtain the total amount of protein, films were solubilized in 0.5 M NaOH. The solubility was reported as percentage of total protein.

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2.5.8. Surface morphology Surface morphology of selected films was examined using scanning electron microscope (SEM) (JEOL JSM-5800 LV, Tokyo, Japan). The film was mounted on a bronze stub and sputter-coated with gold using Sputter coater (SPI-Module, PA, USA). The surface and cross-section of the films were observed at an acceleration voltage of 10 kV. 2.6. Statistical analysis Data were subjected to analysis of variance (ANOVA) and mean comparisons were carried out by Duncans’ multiple range test [29]. Analysis was performed using the SPSS package (SPSS 11.0 for windows, SPSS Inc., Chicago, IL). 3. Results and discussion 3.1. Mechanical properties Round scad protein based-film had a slight increase in thickness as the levels of chitosan added increased, particularly in the presence of 25% palm oil (p < 0.05) (Table 1). Tensile strength (TS) of film increased when chitosan at a level of 40% was incorporated (p < 0.05). Nevertheless, elongation at break (EAB) decreased when the amount of chitosan increased (Table 1). At the same level of chitosan added, lower EAB was found in films containing palm oil (p < 0.05). On the other hand, TS tended to increase as the palm oil was incorporated into the film. From the results, it was suggested that the functional groups (hydroxyl and amino groups) of chitosan could interact with peptide chains of protein, leading to the increase in TS of resulting films [30,31]. Moreover, at the acidic pH used for muscle protein solubilization, the amino groups became positively charged. As a result, ionic interaction between chitosan and protein intermolecularly could occur. The increase in TS of films suggested that polymers were arranged and cohered in the way which could strengthen the film matrix. This resulted in tighter and more compact structure. From the result, it was noted that the increase in TS of films was found when palm oil was incorporated in the film containing of 40% chitosan (p < 0.05). This was associated with the decrease

in glycerol amount added. Glycerol is a polar plasticizer, which could distribute uniformly throughout the film matrix. Since oil was not dissolved in the film forming solution thoroughly, the protein molecules could interact each other to a greater extent. As a consequence, higher TS was observed in comparison with that found in the film without oil. Rolodziejska et al. [32] reported that the increase in strength of gelatin film was found after chitosan was added, indicating that the amine groups of chitosan participated in cross-linking reactions with gelatin. Jagannath et al. [33] blended the starch with different proteins to increase tensile strength of film. From the result, chitosan film (2%) showed the highest TS and EAB, compared with protein film and protein-chitosan composite films. Chitosan with high chain length could undergo cross-linking with more inter-junctions. As a consequence, the more rigid and stronger network could be formed. Physical and chemical properties of chitosan generally depend strongly on the molecular weight and the degree of deacetylation [32]. Srinivasa et al. [34] reported that the molecular weight of chitosan has a profound influence on the thermal, mechanical and permeability properties of the films. The cationic property of chitosan offers an opportunity to take advantage of its electrostatic interaction properties which can enhance the film properties [35,36]. EAB of round scad protein-based films incorporated without and with chitosan at different levels (as protein substitution) and palm oil at 25% glycerol substitution is shown in Table 1. In general, EAB decreased when chitosan was added up to 20% (protein substitution) (p < 0.05). No differences in EAB were noticeable between film added with 20% and 40% chitosan (p > 0.05). Srinivasa et al. [34] reported that EAB of polyvinyl alcohol film decreased when chitosan concentration increased. Chitosan can interact with –OH and NH3 + of protein via hydrogen bonding [34]. This led to the decrease in EAB of resulting films. At the same level of chitosan added, film incorporated with palm oil had the slight decrease in EAB (p < 0.05). The increasing palm oil with concomitant decreasing glycerol content might lessen the integrity of films network structure. The higher aggregation of protein molecules in the protein-rich phase of the film was presumed. The lower continuity and coherence (cohesiveness) of protein network in the presence of lipid glob-

Table 1 Tensile strength (TS), elongation at break (EAB), water vapor permeability (WVP) and thickness of round scad protein-based films as affected by the addition of chitosan and palm oil EAB* (%)

WVP* (×10−10 g m−1 s−1 Pa−1 )

Thickness* (mm)

3.31 ± 0.67e 3.65 ± 0.92 de

120.47 ± 9.93a 89.61 ± 3.33c

1.08 ± 0.05c 0.92 ± 0.03d

0.034 ± 0.001f 0.038 ± 0.002de

0 25

3.52 ± 0.37de 4.61 ± 0.61de

102.65 ± 11.95b 86.08 ± 6.53c

1.09 ± 0.10c 0.90 ± 0.01d

0.035 ± 0.002f 0.036 ± 0.002ef

20

0 25

4.13 ± 0.93de 4.95 ± 0.54d

94.56 ± 6.96bc 72.68 ± 4.90d

1.14 ± 0.02c 1.09 ± 0.03d

0.041 ± 0.001c 0.047 ± 0.001b

40

0 25

9.11 ± 1.44c 11.35 ± 1.68b

87.03 ± 5.53c 75.86 ± 6.36d

1.54 ± 0.06b 1.47 ± 0.05b

0.039 ± 0.001cd 0.047 ± 0.001b

14.64 ± 1.14a

112.63 ± 1.75a

1.86 ± 0.08a

0.057 ± 0.001a

Chitosan concentration (%)

Palm oil concentration (%)

0

0 25

10

Chitosan film * Means ± S.D.

TS* (MPa)

from five determinations. The same superscript in the same column indicates the non-significant difference (p > 0.05).

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ules might result in the decrease in EAB as pointed out by Anker et al. [37] and Peroval et al. [38]. Since absorbed water can act as plasticizer in hydrophilic films [39], lower water content of film containing lipids may also cause the decreased elongation of the film. 3.2. Water vapor permeability (WVP) WVP is considered crucial because most natural biopolymers are very prone to water absorption [40]. WVP of round scad protein-based films incorporated with different amounts of chitosan in the absence and the presence of 25% palm oil is shown in Table 1. WVP of films increased when chitosan at 40% was incorporated (p < 0.05). In the presence of chitosan, a hydrophilic polymer, the increase in hydrophilicity of film was obtained. This led to the increased WVP of resulting film. From the result, the increase in WVP of film containing 40% chitosan could decrease the barrier efficiency against moisture transfer of film. The water vapor permeability is dependent on the relative polarity of the carbohydrate polymers [41]. Butler et al. [19] reported that chitosan films had low water vapor barrier properties because of their hydrophilic nature. The polysaccharide films generally possessed weak moisture barrier properties [42]. If the films are cationic and strongly hydrophilic, water interacts with the polymer matrix and the permeation for water vapor is increased [30]. At the same chitosan level incorporated, WVP of films decreased when palm oil at 25% (glycerol substitution) was added (p < 0.05). From the result, the palm oil added into film-forming solution might distribute uniformly as emulsion. Chitosan added possibly emulsified the oil. Chitosan was reported to exhibit as emulsifier [20]. As a result, the oil could be dispersed throughout the film and moisture barrier property was enhanced. Shellhammer and Krochta [43] reported that permeability and mechanical properties of film were affected by lipid types and amounts. 3.3. Color, light transmission and transparency of film L*, a* and b*-values of round scad protein-based films added with chitosan at different levels in the presence and the absence of palm oil at 25% glycerol substitution are shown in Table 2. In

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general, the films became darker when 40% chitosan was incorporated as evidenced by the decrease in L*-value (p < 0.05). The films had the increases in a* and b*-values with increasing chitosan levels (p < 0.05). This indicated that the film became more yellowish as chitosan was incorporated. At the same chitosan level, the addition of palm oil at 25% caused no marked effect on L* and a*-values. However, the addition of 25% palm oil in combination of 40% chitosan resulted in the increase in b*-value (p < 0.05). The increase in chitosan used possibly resulted in the increase in free amino group. Thus, the browning via Maillard reaction might be enhanced as shown by the increased yellowness (b*-value). Srinivasa et al. [34] reported that chitosan films were transparent with a white to slight yellow tint and the film became darker as thickness of the film increased. The transparency of round scad protein-based films added with chitosan at different levels in the presence and the absence of palm oil is shown in Table 2. Films added with chitosan were less transparent as indicated by the greater transparency values, compared with the control film regardless of oil incorporation (p < 0.05). However, marked differences were found among films added with different levels of chitosan (p < 0.05). Therefore, an increase in chitosan levels resulted in more opacity as indicated by the increase in transparency value. The increase in opacity was possibly caused by the interaction between chitosan and protein, which resulted in the greater cross-linking intermolecularly. Round scad protein-based films containing chitosan were more transparent when the palm oil was incorporated (p < 0.05). Liquid dispersed in film matrix might affect the transparency of resulting film. Yang and Paulson [44] reported that the differences in opacity of film were determined by the optical properties of lipids incorporated. Solid fat was reported to increase the film opacity compared with liquid lipid [39]. Light transmission of protein-based film containing different chitosan levels in the presence and the absence of 25% palm oil for both UV and visible ranges is shown in Table 3. In general, the protein-based film showed much lower transmission to light in the UV ranges (200–280 nm), while chitosan film had the higher transmission in that range. For the visible range, chitosan film had the highest transmission, followed by proteinbased film (without chitosan). The addition of chitosan led to the lowered transmission in the visible range. The interaction of

Table 2 L*, a*, b*-values and transparency value of round scad protein-based films as affected by the addition of chitosan and palm oil Chitosan concentration (%)

L*

a*

0

0 25

86.62 ± 0.32bcd 85.92 ± 0.34d

−2.04 ± 0.03a −2.02 ± 0.02d

7.78 ± 0.21e 7.81 ± 0.37e

1.68 ± 0.01e 1.74 ± 0.03d

10

0 25

87.38 ± 0.24a 86.70 ± 0.44abc

−1.97 ± 0.06d −1.73 ± 0.25b

9.40 ± 0.29d 9.48 ± 0.96d

2.37 ± 0.01bc 2.27 ± 0.09c

20

0 25

86.35 ± 0.32cd 84.14 ± 0.87e

−1.80 ± 0.06b −1.83 ± 0.05bc

12.28 ± 0.46c 12.79 ± 0.40bc

2.46 ± 0.02a 2.31 ± 0.02c

40

0 25

84.45 ± 0.80e 84.62 ± 0.76e

−1.94 ± 0.04cd −1.93 ± 0.03cd

13.08 ± 0.50b 14.31 ± 0.47a

2.38 ± 0.06b 2.32 ± 0.02bc

87.15 ± 0.15ab

−1.44 ± 0.04a

2.27 ± 0.12f

1.31 ± 0.02f

Chitosan film * Means ± S.D.

b*

Transparency value*

Palm oil concentration (%)

from five determinations. The same superscript in the same column indicates the non-significant difference (p > 0.05).

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Table 3 Light transmission of round scad protein-based films as affected by the addition of chitosan and palm oil Chitosan concentration (%)

Palm oil concentration (%)

Wavelength (nm) 200

280

350

400

500

600

700

800

28.99 28.37

43.28 40.86

51.17 48.69

53.76 51.70

55.87 53.22

55.31 53.82

0

0 25

0 0

0 0

10

0 25

0 0

0 0

0 0

7.66 10.72

9.66 14.09

10.55 15.79

11.02 16.59

11.31 17.14

20

0 25

0 0

0 0

0 0

4.82 5.34

6.76 7.88

7.48 9.28

8.02 10.26

8.58 11.15

40

0 25

0 0

0 0

0 0

5.55 5.67

8.32 8.35

10.16 10.12

11.58 11.51

12.65 12.69

81.25

83.97

84.68

84.79

84.30

Chitosan film

1.57

molecules between chitosan and protein might prevent the light transmission at selected wavelength. When 10–20% chitosan was incorporated, films added with palm oil at 25% glycerol substitution showed the higher light transmission in the visible ranges. In the presence of oil, protein-based film had the decreased transmission. The droplet of dispersed palm oil in round scad protein-based film might prevent the light transmission through the film. From the result, the amounts of chitosan and lipid used were found to affect the light transmission of the films differently. Therefore, the addition of chitosan into the protein-based film was an effective means to improve the barrier property to the light. 3.4. Film solubility and protein solubility Table 4 shows film solubility of the round scad protein-based films with and without chitosan and lipid addition. Generally, solubility of film decreased to some extent with increasing chitosan content irrespective of oil incorporation (p < 0.05). From the result, the loss in film solubility was most likely caused by interaction between proteins and chitosan, leading to greater molecular weight cross-links. Additionally, glycerol could bind Table 4 Film solubility and protein solubility of round scad protein-based films as affected by the addition of chitosan and palm oil Chitosan concentration (%)

Palm oil concentration (%)

Film solubility* (%)

Protein solubility* (%)

0

0 25

52.64 ± 0.43b 56.88 ± 0.58a

11.99 ± 1.19c 12.21 ± 0.43bc

10

0 25

51.53 ± 0.22b 52.20 ± 0.71b

13.13 ± ± 0.71bc 13.48 ± 0.07bc

20

0 25

48.91 ± 1.34c 48.68 ± 0.87c

13.98 ± 1.40b 14.04 ± 0.94b

40

0 25

41.76 ± 1.69d 39.31 ± 0.61e

20.61 ± 2.00a 21.33 ± 0.85a

Chitosan film

27.44 ± 0.21f

–

* Means ± S.D. from triplicate determinations. The same superscript in the same

column indicates the non-significant difference (p > 0.05).

46.09

86.73

with chitosan, especially at amino group or OH groups. As a consequence, glycerol was most likely imbibed in the film matrix and could not be leached out. For protein-based films added with palm oil, film solubility markedly increased when palm oil was incorporated. Lipid droplets might insert in the film matrix and the proteins underwent the cross-linking to the lower extent. Hydrophobic lipid was rarely dispersed throughout the very polar protein-rich phase. Without the sufficient glycerol, a hydrophilic plasticizer, the more aggregation took place, leading to a great loss in film solubility. From the result, chitosan film showed the lowest film solubility (27.44%). Cabodevila et al. [45] reported that the decrease in water solubility was due to the formation of crystalline of chitosan during the drying process. It was noted that film solubility of film containing 40% chitosan was lowered when oil was incorporated. At high chitosan concentration, chitosan could interact with protein effectively and the glycerol could be embedded in the matrix. Also, the oil could be emulsified potentially by chitosan with the sufficient amount. Therefore, the film matrix was strengthened by “filler effect” as well as the pronounced cross-linking of both polymers. In contrast, protein solubility using water as the solubilizing medium increased with increasing chitosan concentration (p < 0.05). When chitosan content increased from 0% to 40%, protein solubility increased from 11.99% to 21.33%. However, no difference in protein solubility was noticeable when the oil was incorporated, regardless of chitosan levels used (p > 0.05). At 40% chitosan added, the protein solubility was increased (p < 0.05), suggesting that chitosan preferably interact each other, rather than with proteins. The coincidental lower protein cross-linking might result in the loosened structure. Those proteins might be washed out easily. 3.5. Effect of chitosan on emulsion stability To verify the role of chitosan and/or protein as the emulsifier of film-forming solution containing oil, the emulsion stability and distribution of oil droplet of film-forming solution were determined. For emulsion stability, it was found that no phase separation was visually observed in emulsion of all samples after 48 h at room temperature. In the absence of chitosan, emulsion was still retained, suggesting the ability of muscle protein in

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Fig. 1. Oil droplet size distribution in film-forming mixture incorporated with different levels of chitosan (magnification: 100×).

stabilizing the oil droplet in the system together with Tween-20, which was added as the surfactant. When chitosan was added, it might play a role in stabilizing emulsion of film forming solution in combination with Tween-20 and muscle protein. The emulsifying ability of chitosan in oil-in-water emulsion system has been reported in the literatures, due to its amphiphilic property [20,46]. The distribution of oil droplets is the most important parameter in characterizing any emulsion [20]. The emulsions were characterized for particle size distribution using an optical microscope. The particle size distribution is shown in Fig. 1. For the film-forming solution containing no chitosan, the larger droplet size of oil was obtained. The addition of chitosan caused a reduction in a particle size of emulsions. It was noted that the oil droplet became smaller as the amounts of chitosan added increased. This is because the chitosan can function as an emulsifier due to it amphiphilic property [20]. Once the shearing force was applied to breakdown the droplets, chitosan as emulsifier could migrate to interface and functioned as surface active agent. The smallest droplets were found in film-forming solution containing 40% chitosan (protein substitution). The average particle sizes of oil droplet were 5.97, 4.06 and 3.54 ␮m for emulsions having 10%, 20% and 40% (w/w) chitosan as protein substitution, respectively. The unimodal distribution of droplet is coincidental with stability of the emulsion. The effect of chitosan on emulsion stability can be attributed to an increase in surface activity of a continuous phase [47]. It was noticed that the film-forming solution without chitosan had the small droplet of oil on the surface of films after drying. Therefore, film matrix containing chitosan could imbed or coat the oil more effectively rather than muscle proteins. 3.6. Protein pattern Protein patterns of round scad protein-based films added with chitosan at different levels in the absence and the pres-

Fig. 2. Protein patterns of round scad protein-based films containing different levels of chitosan without (N) and with (O) 25% oil in the absence (A) and the presence (B) of ␤-ME. Numbers designate the level of chitosan as substitute of protein (%).

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ence of 25% palm oil (glycerol substitution) are depicted in Fig. 2. Generally, myosin heavy chain (MHC) and actin bands of films almost disappeared. The lower band intensity of MHC and actin was noticeable in films added with chitosan. Amino group of chitosan possibly reacted with carbonyl groups of protein via Maillard reaction, leading to the covalent cross-linking of proteins. The reaction was most likely enhanced during the drying process of film. From the result, the addition of oil had no much impact on protein patterns of films. Also, no differences in protein patterns were observed between reducing and non-reducing conditions. From the result, interaction between proteins intermolecularly or between proteins and chitosan would contribute to the good mechanical properties of the films. 3.7. Effect of palm oil and chitosan on protein interaction The distribution and extents of inter- and intra-molecular interactions, which give rise to a three-dimensional network structure of the films, should affect their mechanical properties. The main associative forces involved in fish muscle protein-based films may be inter-molecular covalent bonds with secondary hydrophobic and hydrogen interactions [2–4,8]. Selected films from round scad protein including control film, film added with 25% palm oil and film added with 25% palm oil and 40% chitosan were solubilized in four different solutions which can disrupt ionic bonds, hydrogen bonds, hydrophobic interactions, and disulfide bonds. The protein solubilities (percent of total protein content) in four solubilizing solutions are listed in Table 5. The proteins in all films were solubilized at very low extent in S1 (0.6 M KCl) and S2 (20 mm Tris–HCl, pH 8.0). It was noted that film containing oil had the high solubility in S1. Oil droplet might reduce the integrity of the film matrix as well as the interaction between protein molecules, leading to the ease of solubilization. When dissolved with S3, hydrogen bonds as well as ionic interaction stabilizing film matrix were disrupted and the marked increase in solubility was observed except for those containing both chitosan and oil. The solubility markedly increased to more than 40% for the control film and film incorporated with palm oil at 25% glycerol substitution. The considerable increase in solubility of films was observed when S4 was used as the solubilizing medium. With the addition of S4, hydrophobic interaction was destroyed. The increase in the urea-soluble fraction indicated the formation of hydrogen and hydrophobic bonds. The increase in protein solubility in S3 and S4 suggested non-covalent intermolecular interactions

(hydrogen bonds and/or hydrophobic interactions) involved in stabilizing the protein structure [48]. In the presence of 2% ␤ME (S5), the increase in protein solubility of all films except for film containing 25% oil was observed (p < 0.05). In addition, the higher solubility of films in the presence of 2% ␤-ME (S5), compared to that found in S3 and S4, indicated the contribution of disulfide bonds to the film network. The lower protein solubility in S5 of film incorporated with chitosan at 40% protein substitution and palm oil at 25% glycerol substitution in comparison with other samples was possibly due to the involvement of non-disulfide covalent bonds in stabilizing the film network. Surimi film was formed via the cross-links stabilized by various bondings including hydrogen bond, hydrophobic interaction, ionic bonds as well as disulfide bond [2]. Inter-molecular disulfide bonds can be formed during the drying of the protein solution [3]. From the results, it was elucidated that various bondings such as hydrogen bonds, hydrophobic interactions and disulfide bonds played an important role in the formation of round scad protein-based films. Non-disulfide covalent bonds could be formed, especially in the film added with chitosan. 3.8. Film microstructure SEM micrographs of round scad protein-based films incorporated with chitosan at different amounts with and without 25% oil (glycerol substitution) are shown in Fig. 3. The control film (without chitosan and oil) had the smooth and continuous surface without grainy and porous structure. This indicated that film with ordered matrix was formed. With the addition of chitosan, the surface of film became rougher, especially with increasing chitosan content. The entanglement of long chains of chitosan together with muscle protein molecules via ionic interaction would enhance the roughness of film surface. Nevertheless, chitosan film had the smooth surface (Fig. 3). For films added with palm oil, the surface of film had the irregular surface with the distribution of oil droplets. The oil droplets were less intense on the surface as the amount of chitosan incorporated increased. For film incorporated with 40% chitosan (protein substitution) and palm oil at 25% glycerol substitution, no oil droplet on the surface of film was observed. Those oil droplets were localized inside the film matrix as observed in cross-section. The film structure as well as the distribution of oil droplets might be associated with the properties of film, particularly water vapor permeability of resulting film [38,43].

Table 5 Protein solubility (%)* of round scad protein-based films as affected by the addition of chitosan and palm oil Scad films

S1**

Control Palm oil 25% Chitosan 40% and palm oil 25%

8.75 ± 13.73 ± 1.10a 8.87 ± 0.54b

S2 0.64b***

S3

9.83 ± 21.72 ± 0.66a 5.95 ± 0.88c

0.56b

S4

57.14 ± 41.68 ± 0.39b 5.38 ± 0.73c 0.63a

S5

85.32 ± 79.81 ± 2.08b 14.25 ± 2.23c 1.37a

91.04 ± 2.74a 79.92 ± 3.06b 47.99 ± 0.31c

Mean ± S.D. from triplicate determinations. (S1) 0.6 M KCl; (S2) 20 mM Tris–HCl, pH 8.0; (S3) 20 mM Tris–HCl, pH 8.0 containing 1% (w/v) SDS; (S4) 20 mM Tris–HCl, pH 8.0 containing 1% (w/v) SDS and 8M urea; (S5) 20 mM Tris–HCl, pH 8.0 containing 1% (w/v) SDS, 8 M urea, and 2% (v/v) ␤-mercaptoethanol. *** The different superscript in the same column indicates the significant differences (p < 0.05). *

**

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Fig. 3. SEM micrograph of round scad protein-based films. Surface (A) and cross-section (B). Control film (without chitosan and palm oil) (1), film incorporated with palm oil at 25% glycerol substitution (without chitosan) (2), film incorporated with chitosan at 10% protein substitution (without palm oil) (3), film incorporated with chitosan at 10%protein substitution and palm oil at 25% glycerol substitution (4), film incorporated with chitosan at 40% protein substitution and palm oil at 25% glycerol substitution (5), chitosan film (6).

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