Development of Edible Packaging for Selected Food Processing Applications
Dissertation
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy
in the Graduate School of The Ohio State University
By
Shinjie Lin, M.S.
Food Science and Technology Graduate Program
The Ohio State University
2012
Dissertation Committee:
Dr. Melvin A. Pascall, Advisor
Dr. Jianrong Li
Dr. John Litchfield
Dr. Hua Wang
Copyright by
Shinjie Lin
2012
Abstract
Edible packaging (coatings and films) has been used to improve the shelf life, sensory attributes and nutritional content of food products. The manufacturing process for edible packaging depends on the properties of the ingredients and the application end use. The objective of this study was to optimize the method for commercial-scale chitosan film production. This was done by optimizing the raw ingredient selection, and blending sequence. Quality control tests were used to monitor these include viscosity, the drying rate of the ingredient slurries, film solubility, chemical compositions, as well as the thermal properties of the edible films made during the study.
The first part of this dissertation (Chapter 2) focuses on the effects of solvents on the film properties. Edible slurries were prepared by dissolving 1.0-2.0% chitosan in 1.0% food- grade acetic or lactic acids with 0-20% ethanol solutions, then casting them in Teflon plates.
Viscosities of the different film formulations were measured using a viscometer, and changes in drying time were determined using a OHAUS Moisture Determination Balance. Solubility of the films was determined by dissolving the dried films in water for 1 minute then measuring the weight changes. The chemical compositions of the polymeric chains were identified by Attenuated Total Reflectance Fourier Transform Infrared (ATR-FTIR) spectroscopy. Differential scanning calorimetry (DSC) and thermogravimetric analysis
(TGA) were used to characterize changes to the thermal properties of the films as a result of
ii the various treatments. Results indicated that viscosity of the slurries significantly (p< 0.05) increased with increasing chitosan and ethanol concentrations. ATR-FTIR spectra showed that acetic acid/chitosan provided more carboxyl (1442 cm-1) and amine groups (1573 cm-1) within the polymeric network when compared to the lactic acid/chitosan films. The addition of 20% ethanol significantly (p< 0.05) enhanced the drying rate of the film by 30%.
However, additional ethanol did not have a significant influence on the solubility and thermal properties of the films.
The second part of this dissertation (Chapter 3 and Chapter 4) reports two methods for incorporating vitamin E into the chitosan. Edible chitosan slurries were prepared by blending 250 or 500 mg vitamin E into1.0- 2.0% chitosan, and then casting them in Teflon plates. Two blending processes were used to incorporate the vitamin E into the edible slurries: (1) the vitamin E added before lecithin (VE first); and (2) the vitamin E mixed with lecithin (VE mixed) and then added into the slurries. Viscosities of the various formulations, thermal properties, the chemical compositions and solubilities of the film samples were done as mentioned before. High performance liquid chromotagraphy (HPLC) was used to determine the concentration of vitamin E in the films.
This study found that vitamin E addition significantly (p< 0.05) affected the rheology of the edible slurries. In addition, viscosity increased with increasing chitosan and vitamin E concentrations. Low chitosan concentrations in the formulation decreased vitamin E incorporating ability, leading to more vitamin E accumulation on the film surface. Solubility decreased with increasing accumulated vitamin E on the film surface. Furthermore, the drying speed of the film was extended by the addition of vitamin E in the 1.0% and 1.5% chitosan films. Results also showed that 2.0% chitosan blending with 500 mg VE was
iii superior to the other formulations. More than 73% (368.60± 12.40 mg) of the vitamin E was successfully incorporated in the film. TGA analysis indicated that vitamin E decomposition occurred at 430-450°C. However, DSC thermograms showed no significant difference in glass transition temperature (Tg) and the phase changes in the films.
iv
Dedication
To my family, dearest Dad, Mom, and Sis
v
Acknowledgements
I would like to thank my advisor Dr. Melvin Pascall for his support, patience and guidance throughout this graduate study. I appreciate the opportunity he gave me to pursue a doctoral degree at The Ohio State University. I would also like to thank Dr. Shaun Chen, my master degree advisor for his inspiration, without which I would never be in love with food packaging. Also, I would like to extent my sincere gratitude to my committee members Dr.
Jianrong Li, Dr. John Litchfield, Dr. Jiyoung Lee, and Dr. Hua Wang for their guidance and understanding. I am also very grateful to Dr. Luis Rodriguez-Saona for his knowledgeable advice, support and patience.
Thanks to Chongtao Ge, Dr. Xinhui Li, Lizanel Feliciano, Dr. Jeasung Lee, and Mr.
Paul Courtright for their support and help during this 3 years journey. Also thanks to all my good classmates and friends around the world that fulfilled my life.
I want to thank my aunt and uncle (the Cooper Family) for their support and love. Also to thank Run Li, who always stands by my side and gives me the strength to carry on.
vi
VITA
2002-2006 ...... B.S. Food Science and Nutrition
Fu-Jen Catholic University- Taipei, Taiwan, R.O.C.
2006-2008 ...... M.A. Food Science
Fu-Jen Catholic University- Taipei, Taiwan, R.O.C
2009-Present ...... Graduate Research Associate and Teaching Assistant
Food Science and Technology
The Ohio State University, Columbus, OH
Major Field: Food Science and Technology
vii
TABLE OF CONTENTS
ABSTRACT ...... ii
DEDICATIONS ...... v
ACKNOWLEDGEMENTS ...... vi
VITA ...... vii
LIST OF TABLES ...... xiv
LIST OF FIGURES ...... xv
LIST OF EQUATIONS ...... xviii
1. LITERATURE REVIEW ...... 1
1.1. Introduction of edible film ...... 1
1.2. Types of edible film ...... 3
1.2.1. Polysaccharide-based edible film/coating ...... 5
1.2.1.1. Starch ...... 6 1.2.1.2. Cellulose derivatives ...... 6 1.2.1.3. Chitosan ...... 8 1.2.1.4. Carrageenan ...... 11
1.2.2. Lipid-based edible film/coating ...... 12
1.2.3. Protein-based edible film/coating ...... 13
1.2.3.1.Corn zein ...... 13 1.2.3.2. Whey protein ...... 14 1.2.3.3.Collagen ...... 14
viii 1.2.4. Composite edible film/coating ...... 14
1.3. Methods of producing edible films ...... 15
1.3.1. Edible coating ...... 18
1.3.2. Film formation ...... 18
1.3.2.1. Lab-scale casting ...... 18 1.3.2.2. Compression molding ...... 19 1.3.3. Commercial-scale casting ...... 19
1.3.3.1. Continuous casting ...... 20 1.3.3.2. Extrusion ...... 20 1.4. Function and application of edible packaging ...... 21
1.4.1. Barrier ...... 23
1.4.1.1. Moisture barrier ...... 24 1.4.1.2. Oxygen barrier ...... 24 1.4.1.3. Lipid resistance ...... 25 1.4.1.4. Mass transfer ...... 26 1.4.2. Carrier ...... 26
1.4.3. Enhancement ...... 27
1.4.4. Applications of edible packaging ...... 27
1.4.5. Environmental impact ...... 28
1.5. The method of characterizing polymers ...... 30
1.5.1. Permeability ...... 30
1.5.1.1. Oxygen transmission rate (OTR) ...... 31 1.5.1.2. Water vapor transmission rate (WVTR) ...... 32 1.5.1.3. Carbon dioxide transmission rate (CO2TR) ...... 33
ix 1.5.2. Visco-elastic properties ...... 34
1.5.2.1.Viscosity ...... 34 1.5.2.2.Rheology ...... 34 1.5.3. Mechanical strength ...... 35
1.5.4. Optical method ...... 36
1.5.4.1. Attenuated total reflection fourie transform infrared spectroscopy (ATR-FTIR) ...... 38 1.5.4.2. X-ray diffraction (Morphology) ...... 39
1.5.5. Thermal properties ...... 40
1.5.5.1. Differential scanning calorimetry (DSC) ...... 41 1.5.5.2. Thermogravimetric analysis (TGA) ...... 42 1.5.5.3. Dynamic mechanical analysis (DMA) ...... 43
2. THE EFFECT OF ORGANIC SOLVENTS (ACETIC ACID, LACTIC ACID,
AND ETHANOL) ON CHITOSAN EDIBLE FILMS ...... 44
2.1. Abstract ...... 44
2.2. Introduction ...... 45
2.3. Methods and Materials ...... 49
2.3.1. Materials ...... 49
2.3.2. Edible films formation ...... 43
2.3.3. The flow chart of slurry formation ...... 51
2.3.4. Viscosity measurement ...... 52
2.3.5. Drying rate analysis ...... 52
2.3.6. Solubility of the films ...... 52
2.3.7. Functional groups characterization ...... 53
x 2.3.7.1. Attenuated Total Fourier Transform Infrared Spectroscopy (ATR- FTIR) ...... 53 2.3.7.2. Multivariate analysis ...... 53
2.3.8. Thermal analysis ...... 54
2.3.8.1. Differential scanning calometric (DSC) analysis ...... 54 2.3.8.2. Thermogravimetric analysis (TGA) ...... 54
2.3.9. Statistical analysis ...... 55
2.4. Results and Discussion ...... 56
2.4.1. Viscosity of the chitosan dissolved in different solvents ...... 56
2.4.2. Drying rate of chitosan slurries ...... 59
2.4.3. Solubility if chitosan films ...... 64
2.4.4. Attenuated Total Reflection Fourier Transform Infrared Spectroscopy
(ATR-FTIR) characterizations ...... 67
2.4.5. Thermal properties ...... 72
2.4.5.1. Differential scanning calorimetric (DSC) analysis ...... 72 2.4.5.2. Thermogravimetric analysis (TGA) ...... 70
2.5. Conclusion ...... 80
2.6. References ...... 81
3. THE INCORPORATION OF VITAMIN E INTO CHITOSAN FILMS AND ITS
EFFECT ON THE MATERIAL PROPERTIES (DRYING RATE,
SOLUBILITY) ...... 85
3.1. Abstract ...... 85
3.2. Introduction ...... 86
xi 3.3. Methods and Materials ...... 89
3.3.1. Materials ...... 89
3.3.2. Edible films formation ...... 89
3.3.3. The flow chart of slurry formation ...... 91
3.3.4. Drying rate analysis ...... 92
3.3.5. High performance liquid chromatography (HPLC) analysis ...... 92
3.3.5.1. Sample preparation for HPLC analysis ...... 92 3.3.5.2. HPLC analysis ...... 93
3.3.6. Solubility of the films ...... 94
3.3.7. Statistical analysis ...... 94
3.4. Results and Discussion ...... 95
3.4.1. Drying rate of vitamin E chitosan fortified slurries ...... 95
3.4.2. Incorporation of vitamin E in chitosan edible films (HPLC analysis) ... 100
3.4.3. Solubility of vitamin E fortified chitosan films ...... 107
3.5. Conclusion ...... 110
3.6. References ...... 110
4. THE INCORPORATION OF VITAMIN E INTO CHITOSAN FILMS AND ITS
EFFECT ON THE MATERIAL PROPERTIES (VISCOSITY, ATR-FTIR,
THERMAL ANALYSIS) ...... 112
4.1. Abstract ...... 112
4.2. Introduction ...... 113
4.3. Methods and Materials ...... 116
4.3.1. Materials ...... 116
xii 4.3.2. Edible films formation ...... 116
4.3.3. The flow chart of slurry formation ...... 118
4.3.4. Viscosity measurement ...... 119
4.3.5. Attenuated Total Fourier Transform Infrared Spectroscopy (ATR-
FTIR) ...... 119
4.3.5.1.Attenuated Total Fourier Transform Infrared Spectroscopy (ATR- FTIR) ...... 119 4.3.5.2. Multivariate analysis ...... 119 4.3.6. Thermal properties ...... 120 4.3.6.1. Differential scanning calorimetric (DSC) analysis ...... 120 4.3.6.2. Thermogravimetric analysis (TGA) ...... 120 4.3.7. Statistical analysis ...... 121 4.4. Results and Discussion ...... 122 4.4.1. Viscosity of vitamin E fortified edible slurries ...... 122 4.4.2. Attenuated Total Reflection Fourier Transform Infrared Spectroscopy
(ATR-FTIR) characterizations ...... 126
4.4.3. Thermal analysis of the vitamin E fortified edible films ...... 133
4.4.3.1.Differential scanning calorimetric (DSC) analysis ...... 133 4.4.3.2. Thermogravimetric analysis (TGA) ...... 137
4.5. Conclusion ...... 143
4.6. References ...... 143
5. CONCLUSION ...... 146
LIST OF REFERENCES ...... 148
xiii LIST OF TABLE
Table 1.1. Table 1.1. Examples of edible materials for different types of the films ...... 4
Table 1.2. Scientific publications reporting the antimicrobial activity of chitosan …... 10
Table 1.3. Edible film applications ...... 28
Table 1.4. List of commercially used coatings ...... 29
Table 2.1. The drying time of 1.0- 2.0% chitosan in 1.0% acetic or lactic acids with
0-20% ethanol concentrations ...... 63
Table 2.2. The list of functional groups versus wavenumber (cm-1) corresponded to solvents and chitosan parameters ...... 70
Table 3.1. The drying time of 250 and 500 mg vitamin E blended before lecithin or mixed with lecithin in 1.0- 2.0% chitosan ...... 96
Table 3.2. The amount of vitamin E presence in the overall, on the surface, and incorporated chitosan films by HPLC analysis ...... 106
xiv LIST OF FIGURE
Figure 1.1. The structure of polysaccharide (cellulose) and its hydrogen bonds ...... 5
Figure 1.2. The partial structure of (a) amylose; and (b) amylipectin ...... 7
Figure 1.3. The structure of chitosan ...... 9
Figure 1.4. The structure of kappa (κ), iota (ι), and lambda (λ)- carrageenan ……….. 11
Figure 1.5. Structures of composite films ...... 17
Figure 1.6. Steel belt casting line ...... 21
Figure 1.7. The structure of an extruder ...... 22
Figure 2.1. (a) Teflon casting plates; (b) chitosan edible film ………………………... 51
Figure 2.2. The viscosity of 1.0, 1.5, and 2.0% chitosan dissolved in 1.0% lactic acid
(LA) or acetic acid (AA) with variable ethanol concentration (0-20%) ……………… 58
Figure 2.3. The effect of solvent loss on the dry weight of (a) 1.0%; (b) 1.5%; (c)
2.0% chitosan in 1.0% acetic acid with 0-20% ethanol ...... 61
Figure 2.4. The effect of solvent loss on the dry weight of (a) 1.0%; (b) 1.5%; (c)
2.0% chitosan in 1.0% lactic acid with 0-20% ethanol ...... 62
Figure 2.5. The dissolving ability of 1.0- 2.0% chitosan film made by (a) acetic; (b) and lactic acid with 0- 20% ethanol ...... 66
Figure 2.6. (a) SIMCA classification from ATR-FTIR analysis of lactic acid and acetic acid treatments with 1.0-2.0% chitosan; (b) discriminating bands; and (c)
xv PLSR loading plot ...... 68
Figure 2.7. Comparison of overall parameters: acetic acid (AA), lactic acid (LA), 1.0-
2.0% chitosan, and 0- 20% ethanol of ATR-FTIR spectrograms by using SIMCA analysis ...... 71
Figure 2.8. DSC thermograms of (a) 1.0%; (b) 1.5%; (c) 2.0% chitosan films made by acetic acid with 0-20% ethanol addition ...... 74
Figure 2.9. DSC thermograms of (a) 1.0%; (b) 1.5%; (c) 2.0% chitosan films made by lactic acid with 0-20% ethanol addition ...... 75
Figure 2.10. TGA thermograms of (a) 1.0%; (b) 1.5%; (c) 2.0% chitosan films made by acetic acid with 0-20% ethanol addition ...... 77
Figure 2.11. TGA thermograms of (a) 1.0%; (b) 1.5%; (c) 2.0% chitosan films made by lactic acid with 0-20% ethanol addition ...... 78
Figure 3.1. The chemical structure of (a) α-tocopherol; and (b) lecithin ……………... 88
Figure 3.2. The drying rate of vitamin E fortified edible solutions (a) 1.0%; (b) 1.5%; and (c) 2.0% chitosan with 250 mg and 500 mg VE first ...... 97
Figure 3.3. The drying rate of vitamin E fortified edible solutions (a) 1.0%; (b) 1.5%; and (c) 2.0% chitosan with 250 mg and 500 mg VE mixed ...... 98
Figure 3.4. HPLC standard curves of standard α-tocopherol (a) low concentration
(0.25-6.25 mg); (b) high concentration (1.25-125 mg) ...... 102
Figure 3.5. The amount of vitamin E detected by HPLC in 1.0- 2.0% chitosan films with (a) 250 mg; and (b) 500 mg VE first ...... 103
Figure 3.6. The amount of vitamin E detected by HPLC in 1.0- 2.0% chitosan films with (a) 250 mg; and (b) 500 mg VE mixed ...... 104
xvi Figure 3.7. The dissolving (%) of chitosan films with (a) vitamin E added before lecithin; and (b) vitamin E mixed with lecithin ...... 109
Figure 4.1. The viscosity of chitosan slurries with (a) 250 mg; (b) 500 mg vitamin E addition ...... 123
Figure 4.2. The chemical structure of vitamin E bound with chitosan ……………….. 125
Figure 4.3. FTIR analysis of 1.0% chitosan vitamin E fortified films (a) SIMCA analysis; (b) discriminating power ...... 128
Figure 4.4. FTIR analysis of 1.5% chitosan vitamin E fortified films (a) SIMCA analysis; (b) discriminating power ...... 130
Figure 4.5. FTIR analysis of 2.0% chitosan vitamin E fortified films (a) SIMCA analysis; (b) discriminating power ...... 131
Figure 4.6. PLSR analysis of chitosan vitamin E fortified films versus slurry viscosity
(a) linear relationship; and (b) loading plot ...... 132
Figure 4.7. DSC thermograms of (a) 1.0- 2.0% chitosan films (control); and (b) 1.0%;
(c) 1.5%; and (d) 2.0% cihtosan with 250 mg and 500 mg vitamin E addition ...... 139
Figure 4.8. TGA thermograms of (a) 1.0- 2.0% chitosan films (control); and (b)
1.0%; (c) 1.5%; and (d) 2.0% cihtosan with 250 mg and 500 mg vitamin E addition ... 142
xvii LIST OF SCHEME
Scheme 2.1. The experimental design for characterizing solvent variable chitosan films ...... 48
Scheme 2.2. The flow chart of chitosan slurry formation ...... 51
Scheme 3.1. The flow chart of chitosan-vitamin E slurry formation ...... 91
Scheme 4.1. The flow chart of chitosan-vitamin E slurry formation ...... 118
Scheme 4.2. The possible molecular binding sites between vitamin E and chitosan ..... 125
xviii LIST OF EQUATION
1.1. Equation of oxygen permeability ...... 32
1.2. Equation of water vapor permeability ...... 33
2.1. Drying rate analysis ...... 52
2.2. Solubility of the films ...... 53
3.1. Drying rate analysis of vitamin E-chitosan slurries ...... 92
3.2. Solubility of the vitamin E fortified films ...... 94
xix LIST OF EQUATION
1.1. Equation of oxygen permeability ...... 32
1.2. Equation of water vapor permeability ...... 33
2.1. Drying rate analysis ...... 52
2.2. Solubility of the films ...... 53
3.1. Drying rate analysis of vitamin E-chitosan slurries ...... 92
3.2. Solubility of the vitamin E fortified films ...... 94
xix CHAPTER 1
LITERATURE REVIEW
1.1. Introduction of edible films
An edible coating or film could be defined as primary packaging made from edible components. A thin layer of edible material can be directly coated to a product or formed into a film and be used as a food wrap. Edible films have been used to improve barriers, mechanical properties, sensory, convenience, and prolong the shelf life of various products (Krochta 2002; Janjarasskul and Krochta 2010). Another applications of its use include health benefits by incorporating nutrients such as vitamins, minerals and bioflavonoids within the film matrix (Park and others 2001; Larotonda and others 2005;
Park and Zhao 2006). In addition, the biodegradable and environmental friendliness activities of edible films are other desirable benefits associated with their use (Siracusa and others 2008; Janjarasskul and Krochta 2010).
Edible polymers such as polysaccharide, protein, and lipid are the three main ingredients used to produce edible films. In many instances two or all of these ingredients are blended to produce composite edible films (Hernandez-Izquierdo and
Krochta 2008). Polysaccharide based edible films (e.g. chitosan, carrageenan) are hydrophilic and provide strong hydrogen bonding that can be used to cross-link with functional additives such as flavors, colors, and micronutrients. Due to the ability of,
1 Adjacent chains in the polymer to cross-link these films have good oxygen but poor moisture barrier properties. Protein based edible films are also hydrophilic and have good mechanical strength and can be used on fruits to reduce injuries during transportation (Miller and Krochta 1997; Larotonda and others 2005; Saucedo-Pompa and others 2009; Janjarasskul and Krochta 2010). Both polysaccharides and protein films have poor moisture barrier because of their hydrophilic properties. Lipid based edible films have good moisture barrier, but low mechanical properties due to their hydrophobic structures. The manufacture and use of composite films help to minimize the disadvantages of the individual components while making use of the strength in their properties (Hernandez-Izquierdo and Krochta 2008).
Edible films are usually produced by continuous film casting, mould casting or draw- down bar methods. The continuous film casting method is accomplished by coating a wet film onto a belt conveyor and then passing it through a drying chamber. Mould casting and draw down bar are simple and inexpensive methods that can be used as lab- scale edible film production techniques. Edible films are easy to produce and handle since they require less heat and no toxic solvents when compared to traditional petroleum-based food packaging (e.g. polyethylene, polypropylene) (Rossman 2009).
The types of edible films, their properties, end use applications and methods of testing will be discussed in the following sections.
2 1.2. Types of Edible Films
The three main types of edible films (polysaccharides, proteins and lipids based films) are shown in Table 1.1 (Park and others 2002b; Larotonda and others 2005).
Starch, carrageenan, and chitosan are most commonly used to produce polysaccharide films (Park and others 2001; Jansson and Thuvander 2004; Bajpai and others 2009).
Whey, milk and soy proteins are the main sources of protein-based edible films (Seydim and Sarikus 2006; Cho and others 2007; Chai and others 2010). Lipid-based edible compounds such as wax, glycerol esters, and resin are less widely used in the to food industry because of their restrictive application. This is so because lipid oxidation and rancidity of lipid-based films could change the appearance and taste of products during storage. A waxy flavor and nutritional concerns could also affect the acceptance of lipid- based films by consumers (Janjarasskul and Krochta 2010).
3 Table 1.1. Examples of edible materials for different types of the films Types of edible film Examples
Polysaccharide • Cellulose derivatives: methyl cellulose, hydroxypropyl methylcellulose • Starch: amylose starch, corn starch, wheat, tapioca • Chitosan • Pectin: high-methoxyl pectin, low-methoxyl pectin • Alginate • Carrageenan: α-carrageenan, ι-carrageenan, κ-carrageenan • Gums: guar gum, arabic gum, karaya gum, gellan gum
Protein • Gluten: wheat gluten • Collagen and gelatin • Corn zein • Vegetable source: soy protein, rice protein, peanut protein • Animal source: milk protein (whey, casein), egg white protein, fish myofibrillar protein
Lipid • Wax: beeswax, candelilla wax, rice brain wax • Resin: shellac resin, terpene resin • Glycerol ester: acetylated monoglyceride, fatty acid ester (Park and others 2002b; Baldwin 2007; De Moura and others 2009; Janjarasskul and Krochta 2010)
4 1.2.1. Polysaccharide-based edible film/coating
Polysaccharides are composed of monosaccharide and/or disaccharide subunits.
The hydrogen bonds from the subunits interact with the other hydrophilic subunits to form a repeating polymer (Figure 1.1). As a result, polysaccharide-based edible films/ coatings have good oil and oxygen barrier, but the structure is disrupted in the presence of moisture (Janjarasskul and Krochta 2010). It is easy for the food industry to produce polysaccharide films because the sources are numerous, no toxic solvents are needed, they are low cost, and the process is relatively simple.
Figure 1.1. The structure of polysaccharide (cellulose) and its hydrogen bonds.
Polysaccharides and proteins are hydrophilic polymers that generally contain hydroxyl groups and some polyelectrolytes. These types of ingredients also called
“hydrocolloids.” Hydrocolloids have been widely used as edible film-forming solution since their stabilizing effect on emulsions and increasing viscosity of the aqueous phase of edible solutions (Williams and Phillips 2000). Following are some examples of polysaccharide-based ingredients.
5 1.2.1.1. Starch
Starch is commonly used in edible film processing. Amylose and amylopectin provide hydrogen bonding (Figure 1.2) and as a result, starch-based films easily dissolve in water and also bind with other polar functional groups (Park and others 2001; Bravin and others 2006). Advantages of starch as an edible film include: simplicity of preparation, inexpensive, and good barrier to oxygen and lipids; however, it has poor water resistance. Starch-based edible films can be used to package candy and bakery products (Bravin and others 2006). As an example of an application, the sticky surface of candies wrap with a thin starch based film minimizes the inconvenience of the product sticking to a consumer’s fingers (Tharanathan 2003).
1.2.1.2. Cellulose derivatives
Cellulose is an organic compound consisting of (C6H10O5)n units (Figure 1.1).
Highly crystalline cellulose does not dissolve in water, but does so after etherification
(Huber 2009). Etherification helps to separate the intramolecular hydrogen bonding from the crystalline structure. Most of the cellulose derivatives (i.e. methyl cellulose, hydrocypropylmethyl cellulose, and carboxymethyl cellulose) have good film barrier properties especially for decreasing oil uptake and low oxygen transmission (Krochta and others 1994).
6
(a)
(b)
Figure 1.2. The partial structure of (a) amylose; and (b) amylopectin.
7 1.2.1.3. Chitosan
Chitosan (1, 4)-linked 2-amino-deoxy-β-D-glucan, a linear polysaccharide that is derived from chitin (Figure 1.3) and has positive charges on the number 2 carbon of the glucosamine monomer when the condition is below pH 6 (Chen and others 1998). It is extracted by hot water or alkali from the shells of crustaceans such as crabs, shrimps and crawfishes (industry waste). Chitosan is insoluble in water, but soluble in acid solutions such as acetic, citric, and formic acids because of its cationic characteristic. It has been approved for its antimicrobial, biodegradable, biomedical, biocompatible properties, and can be used in food and health related products (Jongrittiporn and others 2001;
Jayakumar and others 2005; Jayakumar and others 2006; Jayakumar and others 2007).
The antimicrobial properties of chitosan is based on the fact that the positive charge on the amino group is attracted to other negatively charged polymers such as the cell membrane of microorganisms, cholesterol, and proteins. When exposed to microorganisms the proteinaceous and other intracellular constituents are induced to leach out from the cell and this causes the death of the microorganism. In support of this, recent studies showed that chitosan successfully inhibited Escherichia coli,
Staphylococcus aureus, Listeria monocytogenes and Bacillus cereus (Coma and others
2002; No and others 2007). This antimicrobial activity enhances the application and usefulness of chitosan to the food industry (Table 1.2). Chitosan has also been used to control the growth of molds (gray mold and blue mold) on post harvest fruits such as grapes and berries while in storage (Han and others 2004; Li and others 2006; Badawy and Rebea 2009).
8 Besides its antimicrobial activities, chitosan has been used on extending the shelf life of bread by retarding starch retrogradation and preventing weight loss in eggs. Park and others (2002a) indicated that 1.0% chitosan brushed on as a coating on baguette significantly increased the shelf life of the baguette by lowering its weight loss, retarded hardness and retrogradation because of its (1.0% chitosan) moisture barrier. This chitosan coating lowered moisture migration from the bread and doubled its shelf life. In the case of eggs, chitosan coating provided a protective layer that reduced moisture and gas transfer from the albumen. This inhibited weight loss and microbial growth in the eggs during storage (Lee 1996).
Figure 1.3. The structure of chitosan (Kumar 2000)
9 Table 1.2. Scientific publications reporting the antimicrobial activity of chitosan Name of microorganism Shelf life extention food types Bacteria Aeromonas hydrophila Sausage, seafoods Bacillus cereus Fruits and vegetables, meat Bacillus subtilis Bread, meat, sausage Clostridium perfrigens Sausage Coliform Meat, soybean sprouts Enterobacter aeromonas Fruits and vegetables Escherichia coli Bread, meat, sausage, seafoods, soybean Lactobacillus fructivorans curd Lactobacillus plantarum Mayonnaise Listeria monocytogenes Fruits and vegetables, kimchi Micrococcus varians Fruits and vegetables, meat, sausage, Pseudomonas fluorescens seafoods Salmonella Typhimurium Meat Staphylococcus aureus Fruits and vegetables, milk Bread, meat, sausage, seafoods Yeast Staphylococci Meat Vibrio cholerae Seafoods Vibrio parahaemolyticus Seafoods Candida albicans Seafoods Saccharomyces cerevisiae Bread, juice, milk Zygosaccharomyces bailii Juice Mold Aspergillus fumigatus Seafoods Aspergillus niger Bread Penicillium digitatum Fruits and vegetables Penicillium italicum Fruits and vegetables Rhizopus nigricans Bread Rhizopus sp. Fruits and vegetables (No and others 2007)
10 1.2.1.4. Carrageenan
Carrageenan is extracted from red seaweed (Rhodophycea). It is a linear chain
polysaccharide with sulphated galactans. The degree of sulphate ester groups in the
structure serves to influence the negative charges and water solubility of carrageenan
(Janjarasskul and Krochta 2010). Based on the degree of sulphate groups present in its
structure (Figure 1.4), carrageenans are divied into three types: kappa (κ), iota (ι), and
lambda (λ)- carrageenan (20%, 33%, 40% w/w, respectively), which are one, two, and
three sulphate esther per dimeric unit (Karbowiak and others 2006). In solution,
carrageenan behaves as a thermoreversible gel if its temperature falls below a critical
point. It can then be easily applied as a coating to selected food items where it can act as
a moisture barrier (prevents dehydration) (Kester and Fennema 1986; Macquarrie 2002).
However, in the presence of a small amount of acid, the thermoreversible system will
change because of extra positive ions that will cause cross-linking between the polymer
chains (Park and others 2001).
Figure 1.4. The structure of kappa (κ), iota (ι), and lambda (λ)- carrageenan.
11 Carrageenan has been used as a thickener, stabilizer, and gelation agent for dairy processing, especially in ice cream making (Karbowiak and others 2006). Several researches have reported that the sulphate group from the carrageenan has antiviral activity against enveloped viruses, (such as Herpes simplex virus), and that this polymer could be studied and developed as an antivirus edible film (Zacharopoulos and others
1997; Coggins and others 2000; Pujol and others 2006). Besides, being a renewable material and safe for humans, economic considerations have greatly increased the application of carrageenan for the food industry.
1.2.2. Lipid-based edible film/coating
Lipid-based edible films have not been well explored because of their structural restrictions. Fatty acids have more covalent bonds than hydrogen bonds and consequently, they do not posses strong mechanical strength. However, they provide higher moisture barrier when compared with hydrocolloid-based edible film (such as rice film). Lipid-based edible materials such as waxes, glycerol esters, and resins are susceptible to lipid oxidation and rancidity and this tends to change the appearance and taste of selected products in contact with coatings/films made from them. In addition to this, these films could introduce a waxy flavor to the product, could cause nutritional concerns, and can affect the acceptance of the product by consumers (Debeaufort and
Voilley 2009; Janjarasskul and Krochta 2010).
Waxes consist of esters long chain fatty acids, and alcohols other than glycerol that show efficient water resistance when applied to packaging materials. Natural waxes such as beeswax, candelilla, rice brain and carnauba waxes are commonly used as edible
12 coating materials, especially on the surface of vegetables and fruits. Additionally, some lipid films have shown antimicrobial properties. Saucedo-Pompa and others (2009) stated that low concentration of ellagic acid (0.01%) mixed with candelilla wax
(extracted from Euphorbia antysyphilitica) could reduce infection of Colletotrichum gloesporioides on avocado, and decrease browning reactions during storage.
1.2.3. Protein-based edible film/coating
Protein-based edible films can be made from both animal and/or plant protein sources. One advantage of these films is their use as an emulsifier to provide a connection between water and oil based additives in a food. Therefore, proteins and lipids could be combined to form a composite-film with increased barrier properties.
Moreover, amino acids provide functional groups, which may cross-link between the chains and help to stabilized the structure of polymers. Protein-based edible films also have longer shelf lives when compared to polysaccharide-based edible films, because they are less sensitive to moisture (Barone and Schmidt 2006). Protein-based edible film are prepared by dissolving protein isolates in a selected solvent and by using a plasticizer to adjust the film properties. However, film properties such as crystallinity and hydrophobicity may also be affect by the type of protein selected. The following are some examples of protein-based edible films.
1.2.3.1. Corn zein
Corn zein is composed of non-polar, alkaline, acidic amino acids. As a result, corn zein does not dissolve in water, but it will in ethanol or in low/ high pH solvents
13 (pH< 4.0 or pH> 11.0) (Park and others 1994). This is helpful when applying corn zein for medical applications. For example, corn zein coated pharmaceutical tablets are used to control the release for certain injected medicines. It has been used to control the ripening of tomatoes by lowering its respiration rate. Also, it can be applied to biodegradable packaging to provide waterproofing characteristics to the materials
(Gennadios and Weller 1990; Park and others 1994).
1.2.3.2.Whey protein
Whey protein is a byproduct of the cheese making process. During this process, the milk protein (casein) coagulates in a pH 4.6 solution and whey protein remains in solution (Morr and Ha 1993). The casein is used to make the cheese and whey protein concentrate (25-80% protein) and whey protein isolate (>90% protein) are produced as a by-product. Whey protein is colorless, tasteless, odorless, heat stable and has good oil and oxygen barriers. The disadvantage of a whey protein isolate edible film is that it shows low mechanical properties, such as tensile strength (Yoo and Krochta 2011). It has good oxygen barrier properties and has been used to minimize lipid oxidation in roasted peanuts (Krochta 2002).
1.2.3.3. Collagen
Collagen is an animal protein consisting of the glycine, proline, hydroxyproline, hydroxylysine and other major amino acid units. It is mostly found in fibrous
(connective) tissue such as skin, tendons, blood vessels and the intestine tract. Collagen is the most representative edible film, and is widely used in sausage making. It shows a
14 thermoplastic behavior and provides extended hydrogen bonding in parallel structures to form into a fiber (Janjarasskul and Krochta 2010). The collagen casing is used to shape the meat into a tubular form. Collagen edible films not only affect the appearance, they also increase the juiciness, texture, and specificity of the product (Janjarasskul and
Krochta 2010).
1.2.4. Composite edible film/coating
Composite edible films can be formulated in such a way that the advantages of their subunits could be used to minimize the disadvantages the individual components.
For example, composite polysaccharide, protein and lipid-based edible films are a combination of different materials blended to develop a multi-functioned structure. For instance, hydrocolloids (made by hydrolyzed gums) and lipid-based materials could be combined to solve the problem of high moisture permeability and structural weakness in a given film (Debeaufort and Voilley 2009). Polysaccharide and protein-based materials are known to have hydrophilic properties and poor moisture barrier, but they are also known for their good oxygen barrier and oil resistance. If a lipid is added to a mixture of polysaccharides and proteins, it provides non-polar groups that can increase the water vapor barrier of the composite film. However, lipid-based materials have poor film structural supporting abilities (mechanical strength) when they are used alone.
Polysaccharide and protein-based materials have better mechanical strength that can compensate for the weakness in lipid-based edible film. The formation of composite edible films can be done either by layer-by-layer (multi-layer) coating or by blending the
15 sub-units (hydrophilic and hydrophobic compounds with emulsifier or solid suspension)
(Figure 1.4) (Debeaufort and Voilley 2009).
Yoo and Krochta (2011) investigated the blending of whey protein isolate with polysaccharides such as methylcellulose, hydroxypropylmethylcellulose, sodium alginate, and corn starch. Results showed that the composition of whey protein and polysaccharides affected oxygen and water vapor permeabilities. Whey protein isolate mixed with methylcellulose or hydroxypropyl methylcellulose had greater tensile strength and elongation. As a result, combination of the different characterized materials improved multi-functional properties to the composite edible films.
Figure 1.5 shows illustrations of composite films. A monolayer of lipid-based edible film is shown (a) in the diagram. Lipid-based ingredients can be blended with macromolecules (polysaccharides, surfactants, and proteins) using (b) the multilayer, (c) solid dispersion or (d) solid emulsion methods to form a composite film. The multilayer illustration of the composite film shows that one layer of hydrocolloid-based film could be coated onto a layer of lipid ingredient. A solid emulsion can result from a stable suspension of fat globules in a macromolecular network. A solid dispersion can be obtained when a small amount of polysaccharides is incorporated into a continuous lipid phase (Debeaufort and Voilley 2009). Lipids that are bound with substrates have greater compatibility with fatty products (e.g. bakery and deep fried food) due to their hydrophobic interaction (Bravin and others 2006). In contrast, the hydrophilic components help to increase the mechanical properties and to decrease the greasiness of monolayer lipid-based edible films. Bravin and others (2006) indicated that polysaccharide-lipid edible films made from corn starch, methylcellulose and soybean oil
16 can help to delay moisture absorption by dry cookies during storage. They also stated that a polysaccharide-lipid edible film coated to commercial crackers could prolong the shelf life by 50% compared to original uncoated crackers stored at 65-85% RH.
Figure 1.5. Structures of composite films.
* “PS” represents polysaccharides; and “Pro” represents proteins.
(Modified from Debeaufort and Voilley 2009)
17 1.3. Methods of producing edible films
The following section will discuss several film-forming techniques, including dipping, spraying/ brushing, and casting.
1.3.1. Edible coatings
Edible films can provide either a clear or colored appearance depending on the application. Most of the time, consumers prefer clear-coated products. Clear coating can be made by a very thin layer of edible film from materials such as waxes, sugar glazing or starch. However, starch coatings give an off-white color depending on the thickness, type of polysaccharides, and the plasticizer used (Chillo and others 2008). A common method of applying coatings is to dip the food products directly into the edible solution, and then dry the coatings until it solidifies. This method has been used for waxing vegetables and fruits, and in seafood processing. Spraying and brushing methods can apply specific coating to one side of a product with a thin layer. It can be applied continuously and it allows for secondary coatings. The advantage of these methods is that the coating can completely wrap products with uneven shapes (Gontard and Guilbert
1994).
1.3.2. Film formation
Wet and dry processes are two methods used for edible film formation. Both these processes begin by dissolving the ingredients in a solvent, and then removing the liquid phase by drying (Peressini and others 2003). Dry processes such as extrusion and compression molding are also used to form edible films. In the dry process, the materials
18 are mixed with less moisture and the temperature is increased beyond the glass transition point (Tg) of the material in an extruder. At this point, the material is transformed from a solid phase to a morphology (molten) phase with an increase in its mobility. Mobile materials are then released from the end of the extruder (die) and cut into specific shape
(Peressini and others 2003). Wet methods such as mould casting, draw down bar, and compression molding are used to form edible films. These wet methods will be described in the next sections.
1.3.2.1. Lab-scale casting
Casting is the most commonly used method for producing edible films. This is so because most edible solutions contain high moisture levels in their polymer matrix.
Therefore, it is more difficult to produce films from edible solutions using the traditional plastic producing method (extrusion) (Peressini and others 2003). Plasticizers and emulsifier are usually needed in the casting process to modify the chemical and mechanical properties of the film (Hernandez-Izquierdo and Krochta 2008). Plasticizers are used to increase the plasticity or flexibility of the material by reducing hydrogen bonding and increasing the spaces between the polymeric chains. Emulsifiers are used to improve compatibility between the edible material and food systems such as oil-in-water or water-in-oil blends (Hernandez-Izquierdo and Krochta 2008).
Mould casting and draw down blade casting are two common methods for lab-scale film production (Du and others 2008; Yoo and Krochta 2011). The adjustable draw down blades (doctor blades) are capable of preparing uniform wet films for lab testing.
The solution is poured onto a substrate (a glass or brass plate) and it is drawn down over
19 the plate to a uniform thickness. The coated plate is placed in a convection oven to dry the film at an equilibrium moisture condition. The film is then peeled off from the substrate and used as desired (Rossman 2009). Draw down bars are usually made with metals such as stainless steel, aluminum or brass.
1.3.2.2. Compression molding
Compression molding involves pouring the liquid ingredients into a mould (mold) then applying pressure and heat to evaporate the solvents and allowing the dried solids to take the shape of the mould (Hernandez-Izquierdo and Krochta 2008). This method works well for protein-based edible film formation because protein provides stronger thermal properties than polysaccharides and lipids. Cunnngham and others (2000) stated that soy protein isolate-glycerol films could be produced at 150°C, 10 MPa and in 2 minutes. This is a rapid method when compared to regular mould casting, which usually takes more than an hour to produce a films (Hernandez-Izquierdo and Krochta 2008).
Silica, Teflon, nylon, polypropylene, and polyethylene have been used as mold materials, and these depend on the polarity of the casting solution (Hernandez-Izquierdo and
Krochta 2008).
1.3.3. Commercial-scale casting
1.3.3.1. Continuous casting
The continuous casting method (e.g. tape casting and steel belt casting) has been used in commercial scale productions. Edible solutions are cast uniformly onto a continuous steel belt or on a designed backer. The solution cast into the belt then passes
20 through a drying chamber to remove water (Figure 1.6). The dry film is stripped from the steel belt and is wound into mill rolls for later conversion. Also, this method can be applied to multilayers casting. After the first layer of dried film exits the drying chamber, it may receive a thin, secondary coating. Furthermore, the end product can be dusted with a powder to prevent sticking or it can be printed with edible inks for decoration. The length of commercial conveyor lines is typically 50–100 ft, and the width of the belts is from 20 to 60 inches (Rossman 2009).
Solvent evaporation
Liquid slurry
Figure 1.6. Steel belt casting line.
(Rossman 2009)
1.3.3.2. Extrusion
Extrusion is another method of producing polymeric films such as low-density polyethylene, polypropylene (Robertson 1993). Extrusion can be divided into three parts: the feeding zone, the kneading zone, and the heating zone upon exit from the equipment
(Figure 1.7) (Hauck and Huber 1989; GSM industries. 2012.). The mixture of film
21 ingredients first enters the feeding zone and is compressed with air. This system works best with minimal water or solvent content. However, plasticizers are needed to increase the flexibility of the film (Peressini and others 2003). As the ingredients move to the kneading zone, the pressure, temperature, and the density of the mixture increase.
Finally, the mixture enters the heating zone, where the highest pressure, temperature, and shears are applied. The screw of the extruder pushes the molten polymer through a die and it cools immediately (Hauck and Huber 1989; Hernandez-Izquierdo and Krochta
2008). The disadvantages of extrusion film making are that only temperature tolerant and low moisture raw material mixes could be processed. Only protein-based edible films made from wheat gluten, soy protein, or whey protein can be used by this method (Redl and others 1999; Zhang and others 2001; Pommet and others 2003; Hernandez-Izquierdo
2007).
Polymer raw materials
Figure 1.7. The structure of an extruder.
(GSM industries 2012)
22 1.4. Function and application of edible packaging
Edible films and coatings provide convenience, protection, and additive-releasing functions without changing the original ingredients and the processing method of food products. They can be completely attached on the product or become a part of the food.
As a result, edible packaging can be used to protect a product from moisture loss, microbial contamination, delayed respiration rate and aging, fortified nutritional value, and improved appearance or mechanical properties (Janjarasskul and Krochta 2010). As an active package, edible films and coatings can be divided into groups based on functional applications. These include: (1) encapsulation or carriage; (2) improvement of mechanical resistance; and (3) individual protection. Depending on the properties of the polymeric matrix, edible film can be used to incorporate flavors and spices that can help to improve the organoleptic properties of the product. Also, natural antimicrobial and antioxidant agents have been incorporated into polysaccharide-based edible films (Park and Zhao 2004). Edible films blended with pigments, light absorbers or other additives have also been used to improve the appearance and the shelf life of various foods.
Secondly, edible packaging helps to maintain the texture of selected products by improving its mechanical strength. Thirdly, edible coatings can be used to separate food into individual portions (Janjarasskul and Krochta 2010). Several functions and applications of edible packaging would be described in the following sections.
23 1.4.1. Barrier
One of the functions of packaging is to act as a barrier that separates and protects the product from exposure to the environment. Quintavalla and Vicini (2002) stated that edible films have been commercially used to protect meat, fruits, and vegetables from pathogenic microbial contamination. Barrier functions include moisture, oxygen and other gases, fats and oils. These barriers can be applied to ready-to-eat food and fresh produce such as fruits and vegetables (Rossman 2009). The extent of the barrier provided by a package depends on the chemical properties of the material used.
However, environmental conditions, such as temperature, relative humility and the stress and handling of the product by consumers also influence the performance of the package
(Krochta 1994).
1.4.1.1. Moisture barrier
The moisture barrier of edible packaging depends on the properties of the ingredients and the hydrophobic nature of the packaging material. Low water activity will significantly decrease microbial deterioration, enzymatic and chemical reactions for both food products and edible films. However, most edible films or coatings cannot completely prevent moisture penetration because of their hydrophilic function. From the edible point of view, edible packaging should at least slightly dissolve in the mouth instead of them being perceived as “chewing a piece of plastic” (Janjarasskul and
Krochta 2010). Also, the moisture-absorbing rate of the film is also important when compared to the intermediate moisture of the food and on how this influences the drying of the product during storage (Park 1999). To solve this problem, research has focused
24 on composite films development. Lipid-based and protein/polysaccharide/lipid composite films could be an alternative for improving moisture barriers (Janjarasskul and
Krochta 2010). However, as mentioned earlier high lipid content of the edible packaging may reduce consumer acceptance due to the waxy taste and the creamy texture of the materials.
1.4.1.2. Oxygen barrier
Oxygen permeability directly affects the respiration of packaged fresh fruits and vegetables. When the oxygen concentration is higher than needed, it will accelerate the aging process of the fruits and vegetables. On the other hand, vegetables and fruits will go into anaerobic respiration if the oxygen content is lower than needed. Anaerobic respiration produces alcohol in the system and causes the plant material to decay.
Moreover, the amount of oxygen present will impact lipid oxidation, enzymatic reaction, and the coloration of myoglobin in meat products (Janjarasskul and Krochta 2010).
According to Delassus (1994) and Mark and Atlas (1997), the polymeric structure of edible materials (e.g. crystallinity and amorphousness) affect the oxygen and aroma permeabilities. As crystallinity increases so will be the gas barrier of the films. Miller and Krochta (1997) and Janjarasskul and Krochta (2010) indicated that protein-based edible films such as collagen, lactic acid casein, and whey protein isolate have greater oxygen and aroma barrier ability than polysaccharide-based films.
25 1.4.1.3. Lipid resistance
The hydrophilic properties of polysaccharide and protein-based edible coatings make them more lipid resistance than others. Balasubramaniam and others (2007) reported that hydrocolloid-based edible coatings decrease oil uptake by 30% in deep fried chicken balls. Hydrocolloid-based edible coatings tend to into a hydrophilic outlayer, and this prevents oil penetration from deep fry processes such as chicken balls. This result indicates that lipid resistant edible coatings could be used to control the calorie content of deep fried foods.
1.4.1.4. Mass transfer
Edible coating and encapsulation have been used to help maintain the aroma and weight of selected food products (Miller and Krochta 1997). Volatile compounds (e.g. aromas, flavors) of packaged foods may decrease during storage by factors such as flavor scalping by the package itself. An edible coating or film that completely coats the food surface can help to decrease the loss of aromas. For instance, Balasubramaniam and others (2007) indicated that hydroxypropyl methylcellulose (HPMC) coated onto chicken balls decreased the moisture loss by 16.4%, and fat absorption by 33.7% after a180 seconds deep fried process. Results showed that the total mass loss, decreased from
18.8% to 14.4%. Dragich and Krochta (2010) also showed that hydrocolloid-based edible materials helped to retain moisture in chicken breast strips. In addition, edible films can act as a protecting layer to the barrier of toxins migrating from traditional packaging material towards the food product. To be of benefit in such cases, the edible film will have to be discarded before the food is consumed.
26 1.4.2. Carriers
During the blending process, active compounds can be added into edible coating solutions. These include antioxidants, antimicrobial agents, flavoring, pigments and nutrients. In such cases, the functional groups from the edible material would cross- linkage with the additives into the polymeric matrix. For example, nisin-added to alginate edible films showed antimicrobial activity against Staphylococcus when applied to beef. Pigment additives carried by edible materials could improve the appearance of selected products during storage. Mei and others (2002) demonstrated that edible films made with a mixture of xanthan gum, calcium lactate and α-tocopherol can decrease the white discoloration, but increase the orange color of baby carrots during 3 weeks of storage. Also, the study showed that the edible film maintained the quantities of β- carotene, and increased the nutrition value of vitamin E and calcium in the carrots (Mei and others 2002; Han and others 2005). An edible coating can act as a part of the food product because it is in direct contact with the product and can be consumed. As a result, if used as a carrier for drug or other medications the dosage and the usage would be subjected to regulations.
1.4.3. Enhancement
The ability of edible coating to improve the mechanical properties of some fragile products has been previously discussed. For example, chitosan coating on strawberries decreases mechanical damage during the storage, processing and transportation of the fruit (Han and others 2004). However, protein-based and carbohydrate-based edible materials have less tensile strength because of their strong cohesive energy density.
27 Because of this, they tend to form brittle films without the addition of plasticizers.
However, this property could be used to provide a hard shell-like protective outer layer to
certain products. Edible coating may also enhance the appearance and flavor of a
product. The wax on fruits (e.g. lemon, orange, apple) polishes the surface and makes
products appear glossy. It also acts as a moisture barrier that reduces wilting of the
product (Mei and others 2002; Han and others 2005).
1.4.4. Applications of edible packaging
According to the previous discussion, edible films have many advantages that can
be used in the food industry. Table 1.3 and Table 1.4 show some commercialized edible
film/coating products and their applications (Pavlath and Orts 2009; Rossman 2009).
The purpose of using edible packaging is to prolong shelf life by delaying respiration
rate, providing barrier, enhancing mechanical strength, and increasing the nutrition value.
Table 1.3. Edible film applications
Categories Application examples
Packaging Vitamins, enzymes, food colors, food additives, beverage mixes, soup
Freestanding films Breath freshener, toothpaste inclusions, confections, labels, nutraceuticals,
over the counter drugs (OTC), contraceptives
Food wraps Vitamins, meat curing, sushi, enrobing, meat glazes, spice blends
(Rossman 2009)
28
Table 1.4. List of commercially used coatings
Commercial name Main component Applications
FreshseelTM Sucrose esters Extending shelf life of melon
Fry ShieldTM Calcium pectinate Reduces fat uptake during frying fish, potatoes, and other vegetables Nature SealTM Calcium ascorbate Apples, avocado, carrot, and other vegetables
NutrasaveTM N,O-Carbocymethyl Reduce the loss of water in avocado, retains firmness Opta GlazeTM chitosan Replaces raw egg based coating to prevent microbial growth Seal gum, Spray gumTM Wheat gluten Prevents darkening of potato during frying
SemperfreshTM Calcium acetate Protect pome fruits from losing water and discoloration Z*CoatTM Corn protein Extends shelf life of nut meats, pecan, and chocolate covered peanut (Pavlath and Orts 2009)
29 1.4.5. Environmental impact
The original concept of edible film was developed by nature “the skin of fruits and vegetables”. For example, grape skin provides a barrier for water loss and the antimicrobial activity of the ripen grape. Since the1920s’, edible wax has been used on vegetables and fruits (US Apple Association, 2012). Currently, many studies have transformed food wastes into edible packaging ingredients such as chitosan from crab shells and whey protein isolate from the dairy industries (Jayakumar and others 2005).
This food waste transformation not only reduces the waste from industries, it also reduces the cost of raw materials (Jayakumar and others 2006; Jayakumar and others 2007;
Siracusa and others 2008). The conversion of edible components into traditional polymers is another prospect for developing biodegradable materials. Some biopolymers are made from biomass such as polysaccharides, proteins, and lipids, and these could be extracted from food wastes. Also, the products from microorganisms (e.g. yeast, lactic acid bacteria) or those synthesized from bio-based monomers are also considered as biodegradable materials (Siracusa and others 2008; Gómez-Estaca and others 2010).
1.5. The methods of characterizing polymers
Characterizing an edible packaging material is important for a determination of its application to the food industry. Different properties of the packaging material affect its function and compatibility with a product. It also influences the mouth feel of the material when consumed by customers. On the other hand, the property characterization provides the relationship between the edible film ingredients and the processing methods when adjusting the formulation (Hernandez-Izquierdo and Krochta 2008). For instance,
30 the rheology of an edible film solution strongly influences the coating process, and the drying speed when applying to continuous belt casting machines. Also, the viscosity of edible film solutions represents the thoroughness of an emulsion system of the ingredient blend of composite film (Peressini and others 2003). Methods that are commonly used to characterize edible packaging are discussed in the following sections.
1.5.1. Permeability
One of the most important functions of food packing is to provide a barrier to the product. Permeability testing can be used to evaluate the types of foods that can be packaged by a material and it can also used to predict the shelf life of the packaged product (Park and Zhao 2004; Prommakool and others 2010). An edible film with good gas barrier properties can be used to prolong the shelf life of the product. For example, a material with high oxygen barrier helps to reduce lipid oxidation and nutrition loss if used to coat peanuts (Han and others 2008). The water vapor permeability of a packaging material indicates the rate of moisture transfer between the storage environment and the internal environment of the package. A film with low water vapor permeability could be used to extend the shelf life of bakery products such as crackers during storage (Bravin and others 2006). Therefore, it is important to understand the oxygen, water vapor, and carbon dioxide permeabilities of edible films in order to predict the shelf life of a product and the end-use application of the materials (Siracusa and others 2008).
31 1.5.1.1. Oxygen transmission rate (OTR)
The oxygen transmission rate (OTR) of a packaging material plays an important role in influencing the shelf life of oxygen sensitive products such as fresh produce
(fruits, vegetables and salads) and high fat (e.g. nuts, donuts) products. In general, higher crystallinity of the polymeric structures the lower the OTR of the materials. Also, the higher the amorphous regions of the material the higher the OTR (Lacroix 2009). Most researches use the ASTM D3985 (2010) method to measure the ITR of films at 23°C and
50% relative humidity (Du and others 2008; Prommakool and others 2011). According to ASTM D3985, oxygen permeance (PO) is the ratio of the OTR to the difference between the partial pressure of O2 on the two sides of the film. The SI unit of PO is the mol/(m2 s Pa). In addition, OTR is the quantity of oxygen gas passing through a unit area of the parallel surfaces of a plastic film per unit time under the conditions of the test (SI unit: mol/(m2s)). These conditions include the environmental temperature and relative humidity. The PO is calculated by average OTR multiplied by the average film thickness and then divided by the pressure differences between the two sides of films (1.1).
OP=OTR×l/ΔP ...... (1.1)
Where l is the thickness of the film (m), ΔP= partial oxygen pressure difference between the two sides of the film (Siracusa and others 2008).
32 1.5.1.2. Water vapor transmission rate (WVTR)
The water vapor transimission (WVT) of a film sample can be determined using the ASTM E96-92 method. In this method, salt solutions or any ingredient with a known relative humidity is placed inside a WVT cup and sealed by the polymer film. The WVT cup is placed into a humidity-controlled chamber and changes in its weight are measured until a steady state weight is obtained (Prommakool and others 2011). The difference in the relative humidity inside the cup and that of the environment is the direction of driving force of the water vapor movement through the sample film. Weight changes in the cup indicate the driving force of the environment (dehydration or absorption). The weight of the ingredient inside the WVT cup decreases in a dehydrating condition, whereas the weight increased occurs in an adsorbing condition. Thus the water vapor transmission rate (WVTR) of the package can be determined by the change in weight over time
(equation 1.2 and 1.3).
WVTR= (C × A × ∆P)/ T ...... (1.2)
WVTR= Q/t ...... (1.3)
Where C is the permeability constant for the sample material (g-m/ day-m2-mmHg).
A is the surface area (m2) of the sample exposed to moisture.
∆P is the driving force (mmHg).
T is the thickness (m) of the film.
Q is the quantity of the water change (g) t is the period of time (day).
33 Henrique and others (2007) indicated that WVTR of edible films could be directly related to the quantity of –OH group on the molecule. It also influences the properties of the polymeric structure (crystallinity, cohesiveness) and the molecular interactions between the polymer chains. For example, increasing the cross-linking between polymeric chains and reducing the impact of solvents and plasticizers are known reported to improve the water vapor barrier of protein-based films (Sabato and others 2001).
1.5.1.3. Carbon dioxide transmission rate (CO2TR)
The carbon dioxide transmission rate is measured using a technique similar to that of the oxygen permeability. It is used to determine the amount of carbon dioxide that is transmitted through a film for a certain period of time, exposed to a specific area, at a given temperature and CO2 partial pressure. Knowing the CO2TR is important when the respiration rate of a product affects the quality of the food. For the modified atmosphere packaging of climacteric fruits (e.g. banana, pineapple, papaya) the speed of carbon dioxide loss from the package is one of the keys to prolonging the shelf life of these products (Siracusa and others 2008).
1.5.2. Visco-elastic properties
1.5.2.1.Viscosity
Viscosity measurement is a simple method used to understand the visco-elasticity properties of edible film solutions. This can be done using a viscometer set at a fixed revolution per minute, at a given temperature, with a probe of known surface area, in order to measure the resistant force from a test solution. Viscosity readings relate to the
34 percentage of the solids and molecular size of the polymeric structures that comprise the sample. The percent solids of an edible film solution has been used to determine how much wet film thickness should be applied to a casting plate before drying in a film processing operation. For example, a 10% solids solution if cast at 0.020 inches (20 mil) would produce a film with 0.002 inches (2 mil) dry film thickness (Rossman 2009). In addition, viscosity also helps in an understanding of the thoroughness of a blended emulsion. Hydrophilic ingredients cross-linked with lipids by the use of a surfactant
(good emulsion) can result in a solution having a higher viscosity than a lipid suspended in a hydrophilic solution without surfactants (Il’ina and Varlamov 2004; Park and Zhao
2004).
Depends on the sample properties, the viscosity of the solution might differ during the stirring (Newtonian or non-Newtonian). Newtonian fluids show a linear curve of shear stress versus strain and maintain a constant viscosity while being stirred. Non-
Newtonian fluids show viscosity changes when exposed to stresses and the duration of the applied force (Gasparoux and others 2008). Defining the behavior of a non-
Newtonian solution is best done by testing the properties of a solution using a rheometer.
In this method, rheology can characterize the minimum number of functions in the solution that relate stresses with rate of change of strains or strain rate (Rao 2007).
1.5.2.2. Rheology
Rheology is the study of the flow of liquids and semisolids that respond to plastic flow under an applied force. A rheology method can be used to identify the deforming and flow properties of a polymeric material at controlled conditions (Rao 2007). It can
35 also be used to determine a product’s texture (sandiness), stability, appearance and how it will behave under processing conditions (Malkin and Isayev 2006). Rheology determines the viscoelasticity of a sample that exhibit both elastic and viscous behaviors (Malkin and
Isayve 2006). A rheometer applies shear stress or shear strain in either steady or oscillatory rate modes. This can be used to monitor changes occurring in a sample and behaviors such as yield stress, kinetic properties, complex viscosity, modulus, creep and recovery. This method helps with an understanding of the internal structure of materials
(Steffe 1996). Rheology has been widely used to describe the behavior of polysaccharide-based edible film solutions since some of them are non-Newtonian fluids.
For instant, some gelatinized starch solutions exhibit shear-thinning behavior at a certain shear rate, and viscosity changes are related to shear time. Therefore, these behaviors can be used to specify the blending process and the coating parameters during an edible film formation (Peressini and others 2003; and El-Hefian and others 2009). For example, if stirring a solution causes conformation changes to the polymeric chains, this could increase or decrease cross-linking of the polymers. This could then cause the viscosity to change in intensity. Khondkar and others (2010) demonstrated that more elastic components had found in gelatinized waxy maize starch/pectin gels that cross-linked and increased in viscosity after exposure to blending treatments.
1.5.3. Mechanical strength
Texture profile analysis (TPA) describes the structure of materials by numerical analytical methods. It provides the stress or strain of samples, and records the force changes when the time increases (Rosenthal 1999). Tensile strength and elongation at
36 break are two commonly used test methods for films and sheets. These methods help to understand the mechanical properties of the material as a force is applied in an opposite direction (Siracusa and others 2008). ASTM D882 (2010) is the standard test method for tensile properties of thin films. In this test method, the sample must be conditioned at
23°C, and 50% relative humidity for over 48 hours and then be cut to a specific length and width. Tensile strength is calculated by dividing the maximum load for breaking the film by the original minimum cross-sectional area. The percent elongation can be calculated by dividing the film elongation at rupture by the initial gauge length.
Elongation is calculated by dividing the change in the dimension by the original dimension (Peressini and others 2003; Ku and others 2008; Prommakool and others
2011). Both rheology and texture profile analyses describe the physical properties of polymers and establish the procedure for handling them during food processing.
The stress/strain curves obtained from mechanical testing provide information about the flexibility, toughness, and elongation that can be used to predict the film performance during handling (Hernandez-Izquierdo and Krochta 2008). However, restrictions in the use of TPA include environment condition and the sample preparation.
Edible films are sensitive to environment conditions such as relative humidity and temperature. During analytical testing, edible films could absorb moisture from the environment or release water to the environment. These may serve to change the mechanical properties of the film and may alter its properties and end use behavior. A small amount of water can act as a plasticizer within the polymeric structure; therefore, the brittleness could increase with reducing moisture concentration as an example
(Rossman 2009).
37 1.5.4. Optical method
1.5.4.1. Attenuated Total Reflection Fourier Transform Infrared spectroscopy
(ATR-FTIR)
Attenuated Total reflection Fourier Transform Infrared spectroscopy (ATR-FTIR) is widely used to characterize the functional groups of materials (Tanaka and others
2001). The infrared beam from an FTIR unit passes through the sample and causes vibration, stretch, bending, and contraction of the chemical bonds. When this occurs, the excited chemical bonds absorb the infrared radiation at specific wavenumbers (cm-1)
(Goddard and Hotchkiss 2007). During the test, the sample is held onto a highly reflective crystal (e.g. diamond) and then exposed to the infrared beam to determine the path of the reflected beam (Tanaka and others 2001). This test produces data in a short time and little sample preparation is needed. In some cases, the test is considered non- destructive (Siracusa and others 2008). The ATR-FTIR spectra obtained can be used to provide information on the biochemical composition and chemical shifts in the sample that occurred within, especially in the fingerprint region (1500-400 cm-1). ATR-FTIR can be used to determine the type of polar bonds and functional groups of hydrocolloids- based films. It can also be used to determine the presence of plasticizers and emulsifiers in composite films. Hassel and Rodriguez-Saona (2011) reported that ATR-FTIR successfully identified non-polar micronutrient (vitamin E) within a polysaccharide-based food matrix. Thus, it can be applied to characterize the functional groups in additives and nutrients used to fortify edible materials.
38 1.5.4.2. X-ray diffraction (Morphology)
The barrier and mechanical properties of films depend on the microstructure of the sample. X-ray diffraction can be used to determine the morphology of polymeric materials (García and others 2000; Yoo and others 2008). X-ray has very strong energy while passing through a material. When X-rays pass/interact with a material, the X-ray photons can be absorbed by the sample and subquently cause excitation within the molecular structure. Ejection of excited electrons from the sample is known as the photoelectric effect. The ionized atoms that result might return to their ground state by either emission of X-photons (fluorescence) or electrons (Auger effect). Some photons may not lose their energy but can separate into course, which is called scatter radiation
(Guinier 1994). X-ray diffraction is capable of providing details of the chemical components and the morphology of the material. It estimates the percent of crystallinity, d-spacing, and the thickness of the crystal (Yoo and others 2008). X-ray diffraction is sensitive to small changes in ingredients used to formulate the material. It is used to determine the degree of crystallinity and amorphous region in the material. Zhong and
Xia (2008) reported that X-ray diffraction helped in an understanding of the interaction and molecular miscibility among the major components of a chitosan/cassava starch/gelatin film. If an X-ray diffraction test produces results from one part of the sample that is different in another part of the same sample, it indicates that the chitosan/cassava starch/gelatin blend was not homogeneous.
39 1.5.5. Thermal properties
When a polymer is heated from a low to a high temperature, it goes through a series of transition zones that could be described as rigid, thermoelastic and thermoplastic
(Mathew and others 2006). In each of these zones, the material will demonstrate certain properties that are characteristic of that material and of each zone. The temperature at which these properties changes are known as transition temperatures. If a polymer is heated from low to high temperatures at a controlled rate, it will go through the glass transition temperature (Tg), then the melt temperature (Tm) (Soraka 2010). Prior to reaching Tg, the material is described as being in a glass state (brittle) at which point there is no movement in its molecular structure. Above Tg, segmental mobility, or vibrational movements of the functional groups and the polymeric chain itself take place.
At this point the polymer is referred to as being in an elastic state. As temperature increases, the polymeric chains become sufficiently disrupted, that it beings to flow. The temperature at which this occurs is the melt temperature (Tm). If the polymer is crystalline, an additional phase, called the crystal melt range occurs at some point between Tg and Tm. In amorphous polymers the Tm is sometimes referred to as the softening point (Mathew and others 2006).
The properties of edible components are significantly affected by the temperature changes. Polysaccharides-based edible materials are heat sensitive and the browning reaction may occur if the temperature is increased too high during drying. Temperature also affects protein degradation and the dehydrations of protein polymeric chains. In addition, increasing temperature may cause lipid oxidation to lipids-based films
(Hernandez-Izquierdo and Krochta 2008).
40 1.5.5.1. Differential scanning calorimetry (DSC)
Differential scanning calorimetry is used to measure changes in the heat flow rate of both sample and a reference sample in a temperature-controlled system. This technique can be used to determine how much heat is needed to maintain the sample and the reference at the same temperature (Sepe 1997; Höhne and others 2003). Exothermic
(heat is given off) and endothermic (heat is absorbed) reactions occur when a material goes through a phase change and it either absorbs or releases energy in the process
(Höhne and others 2003). DSC is a thermo-analytical technique, which detects the phase transitions (physical transformation) in a sample at a controlled temperature rate and environmental condition. The phase transition includes: evaporation, melting, boiling, freezing, crystallization, glass transition, and decomposition. Heat flow versus temperature plots obtained from DSC indicate melting, glass transition, heats of fusion and reactions, purity, thermal stability, and crystallization of a material (Höhne and others 2003). DSC has been used in many areas such as material science to determine the degree of crystallinity, the extent of the amorphous regions, oxidative stability of the material by flushing oxygen into the chamber, and in food science for water distribution in the structure. The advantages of using DSC are that it is fairly fast, easy of sample preparation (all kinds of sample), small quantity of testing specimen, wide range of test temperature (from -170°C to 600°C), and quantitative capability (i.e. to calculate the freezable water in food samples) (Nicula 2002). However, since a small amount of sample is needed during the test, this may not represent the overall components of the larger sample population, especially for non-homogenous materials. Factors to consider when using DSC include non-identification of compounds causing phase changes in a
41 sample, and the degree of phase changes occurring in the heat flow plot depends on the properties of the ingredients present in the sample. Also, the concentration and the interactions between ingredients can affect the transition temperatures. However, depending on the heat flow and phase changes, the transition points may not be easily identified in the DSC thermogram (Höhne and others 2003; Chrissafis and others 2008).
1.5.5.2. Thermogravimetric analysis (TGA)
Thermogravimetric analysis (TGA) is used to measure weight changes in a sample during a temperature program under controlled conditions (TA Instruments
2011). A sensitive scale inside the TGA instrument is designed to precisely detect weight changes in the sample during the test. These weight changes could be due to water loss, release of volatiles or decomposition of the material as the temperature increases (Sepe
1997; TA Instruments 2011). Ogale and others (2000) used TGA to obtain the thermal degradation of soy protein isolate and soy protein isolate-glycerol films. They measured the weight loss between 150-200°C and used the information to establish an optimum temperature (the temperature with less weight loss occured) for compression molding of the film.
The temperature range of a TGA can start from room temperature to high as
1000°C. During the test, the environmental conditions must be precisely controlled and the accuracy of the system, regularly monitored by the use of standards (e.g. for total water measurement, the amount of water in the sample might vary during the waiting time; therefore, sample preloading is not allowed). A TGA thermogram shows changes in the sample weight over the selected temperature range. To make sense of this, it is
42 necessary to integrate the peak transition areas and compare differences in their areas, intensities and shape (Yoo and others 2008). TGA and DSC are usually combined in order to gain a better understanding of the thermal characteristics of a sample.
1.5.5.3. Dynamic mechanical analysis (DMA)
Dynamic mechanical analysis (DMA) illustrates the response of a sample to stress, temperature, and frequency when applying a deformation force to the material
(PerkinElmer Inc., 2011). DMA is based on the fundamental different responses of viscous and elastic elements at a controlled temperature. It is similar to rheometry and it provides a vertical force (controlled stress or a controlled strain) to the sample and monitors the viscoelastic properties of materials. It is also similar to the texture analysis that detects the shape change and recovery of samples (i.e. stiffness) (Lodi and Vodovotz
2008). Besides stiffness, DMA also reports damping and creep-recovery of samples.
Damping is explained as how much energy is absorbed by the material as it is induced to display elastic behavior, which is reported as tan delta. Creep-recovery is a time dependent test that detects the amount of strain recovered in a material after removing an applied stress (Lodi and Vodovotz 2008). DMA is also capable of detecting glass transition temperature by indicating the breadth of the transition and the changes in heat capacity. In a complicated polymeric matrix, DMA shows better sensitivity and more visible transitions necessary for determining Tg when compared with DSC. DMA is a more versatile tool for testing the thermal properties of a sample when compared with the texture analyzer and DSC methods. This is so because it provides a controlled system of both temperature and stress/strain to a detected sample (McFarlan 2001).
43 CHAPTER 2
THE EFFECT OF ORGANIC SOLVENTS (ACETIC ACID, LACTIC ACID, AND
ETHANOL) ON CHITOSAN EDIBLE FILMS
2.1. Abstract
Edible films have been used to extend shelf life and improve the sensory properties of various foods. The objectives of this study were to investigate the effect of solvents on the properties of chitosan edible film. The edible slurries were prepared by dissolving 1.0-2.0% chitosan in 1.0% food-grade acetic or lactic acids with 0-20% ethanol solutions, then cast into Teflon plates. Viscosities of different film formulations were measured using a viscometer, and functional groups on the polymeric chains were identified by Attenuated Total Reflectance Fourier Transform Infrared spectrum (ATR-
FTIR). Differential scanning calorimetry (DSC), and thermogravimetric analysis (TGA) were used to characterize the changes of thermal properties. Result showed that the presence and intensity of hydroxyl, carboxyl, and amide groups were correlated with the thoroughness of the mixing process. The viscosity of the 2.0% chitosan was ten times higher than that of the 1.0% solution. Acetic acid provided more carboxyl (1442 cm-1) and –NH groups (1573 cm-1) and increased the viscosity by 200 cp. The addition of 20% ethanol increased the drying rate and viscosity by 30% and 50% when compared to the control (slurry with 0% ethanol). However, no changes were observed in the solubility
44 and thermal properties. This study showed that increase in chitosan and ethanol concentration positively affected the rheology of the edible slurries.
2.2 Introduction
Polysaccharides are commonly used to develop edible films because of their simplicity of preparation and inexpensive cost. For example, chitosan (1, 4)-linked 2- amino-deoxy-β-D-glucan is a linear polysaccharide deacetylated derivative from chitin with an amine group (Chen and others 1998). Chitin is found in the exoskeleton of insects and crustacean, and in certain plants and fungi (No and others 1989). In addition to its film making ability, chitosan also has antimicrobial and biocompatible properties
(Park and Zhao 2004). To produce edible films, it must first be dissolved in organic acids. Example of these acids include acetic, lactic, and ascorbic acids (Chen and others
2007). These acids are usually diluted in water, but when mixed in ethanol, the speed of dissolving the chitosan is increased. However, the use of ethanol increases the cost of the film production and creates environmental problems when it is evaporated during the film making process (Tropini and other 2004).
Irrespective of the disadvantages in using ethanol, there is a need to look at how ethanol influences the properties of chitosan when used to make edible films. This is so because of a lack of published data on this topic. Therefore, this research will focus on the relationship of chitosan and solvents (acids and ethanol) on the film properties. These properties include rheology and the drying rate of the edible slurries (chitosan dissolved in solvents), and the solubility, chemical structure, and the thermal characteristics of films made by drying the slurry. Viscosity measurement provides basic information of
45 rheology, which impacts the speed and stability of the drying process during continuous operations (Du and others 2008). It also represents the amount of chitosan hydrolysis and the solids suspended in the solution (Wang and Xu 1994; Hwang and Shin 2000;
Martinez and others 2004; El-Hefian and others 2010). Drying rate of the slurry also affects the film processing time. This is so because of the need to evaporate the solvent from the slurry. The solubility of the edible film affects its application and the mouth feel when being eaten by an individual. In addition, both drying rate and the solubility can also be influenced by the nature of the solvents (acids and ethanol).
To have a deeper insight into the solvent effect on chitosan edible films, ATR-
FTIR, DSC, and TGA ere used to determine the nature of the functional groups in the polymeric matrix and the thermal properties of the films. In this study, ATR-FTIR will be used to obtain information on the biochemical composition of the films (made from chitosan and the selected solvents) from the fingerprint regions (400- 1500 cm-1) of the spectrum (Koca and others 2007). Hassel and Rodriguez-Saona (2011) indicated that the soft independent modeling of class analogy (SIMCA) from the multivariate data analysis could be used to classify differences in film structures caused by different solvent treatments. Thermal analysis such as DSC and TGA are two useful tools that could be used to understand the thermal characteristics of specimens. DSC determines the melting point, glass transition temperature, heats of fusion, purity, thermal stability, and crystallization of a material by showing heat flow versus temperature plots (Höhne and others 2003). It could also be used to study the properties of the melting curve temperature from -170°C to 600°C (Höhne and others 2003). TGA has been used to measure the amount of weight changes in a sample during heating and cooling. Weight
46 changes of the films in this study will occur during solvent evaporation and/or material decomposition when the samples are be heated. Cross validation of the results obtained from the DSC and TGA test will help to identify phase changes in the film samples (Lodi and Vodovotz 2008). The objective of this study was to understand the property changes of chitosan edible film prepared by dissolving the ingredients in acetic or lactic acids with ethanol additions. Also, to optimize the formation of chitosan edible films that could be used to apply to the food industry.
47
Scheme 2.1. The experimental design for characterizing solvent variable chitosan films
Solvent effects
1.0%. 1.5%, 2.0% chitosan film
1.0% acetic acid 1.0% lactic acid
Dissolved in Dissolved in 0, 5, 10, 15, 20% ethanol 0, 5, 10, 15, 20% ethanol
Characterization Functional Properties
1. Viscosity 1. Drying rate 2. ATR-FTIR 2. Solubility
3. DSC 4. TGA
48 2.3. Methods and Materials
2.3.1. Materials
Medium molecular weight (94% purity) chitosan powder was provided by
Huantai Goldenlake Carapace Products Co., Ltd (Tsingtao, China). This was used as the main ingredient in the films. Lactic acid (88% Food Chemical Codex, FCC) manufactured by Birko Corporation (Henderson, CO) and Giant Eagle brand distilled white vinegar containing 5% acetic acid (Pittsburg, PA) were used to dissolve the chitosan powder. Ethanol (190° proof, USP) was purchased from Fisher Scientific
(Decon Labs Inc., King of Prussia, PA) and used as a solvent. Glycerol USP Kosher
99.7% (Chemical direct, online, USA) and soy lecithin (Solec® 100L, The Solae
Company, St. Lousis, MO) were added as a plasticizer and an emulsifier, respectively. A standard 18 mesh (0.0394 inches) fiberglass screen was purchased from an ACE
Hardware Store in Columbus, OH and used for the film solubility test. Differential scanning calorimeter (DSC) stainless steel pans, pan crimper/sealer, O-rings, and thermal gravimetric analysis (TGA) platinum pans were purchased from Perkin Elmer
Instruments LLC (Shelton, CT).
2.3.2. Edible film formation
Edible film solutions were prepared at 22 ± 1°C by dispersing a chitosan powder
(1.0, 1.5 and 2% w/w) in 0, 5, 10, 15, and 20% ethanol-water suspension. Acetic or lactic acid (1%, w/w) was added into the suspension and stirred (300 rpm) for 3 minutes to dissolve the chitosan powder. Glycerol 0.5% (w/w) was added to the solution and mixed for 3 minutes. It was used as a plasticizer to provide flexibility to the films. A 2.0%
49 lecithin solution was then added to the chitosan mixture to produce a 0.1% (w/w) blend as the total dry weight of lecithin in the final films. Lecithin acted as an emulsifier that provided hydrophilic and hydrophobic interactions. The hydrophobic interactions are to help the film provide a good attachment to the food surfaces, especially to the high-fat products. The final solutions were allowed to settle for one hour then tested for viscosity and drying rate.
Edible films were prepared by casting the edible slurry into 10 inch radius Teflon plates (Figure 2.1). Aliquots of 33.0± 0.5 ml, 31.0± 0.5 ml, and 29.0± 0.5 ml of 1.0%,
1.5%, and 2.0% of chitosan slurries were then cast in order to obtain films of uniform thicknesses (50.0 ± 2.5 µm). These were oven dried at 45 ± 2 °C for 2 hours, then the dried films were peeled off from the plate surfaces. The film thickness was controlled by the percent solids in the solution. The thicknesses were measured using a 2804S-10 Agd
Dialindicator Mitutoyo Guage (Aurora, IL) and reported in mil (1/1000 inch).
50 Scheme 2.2. The flow chart of chitosan slurry formation
1.0, 1.5, 2.0% chitosan powder Immersed in 0, 5, 10, 15, and 20% ethanol-water solution Added 1.0% weight of acid solutions Mixed for 3 min Plasticizer added (0.5% w/w glycerol) Mixed for 3 min Added 2% (w/w) of lecithin solution Mixed for 3 min Final edible slurries 33± 0.5 ml 31± 0.5 ml 9.0± 0.5 ml 1.0% chitosan solution 1.5% chitosan solution 2.0% chitosan solution Cast into 10 inches radius Teflon plates
(a) (b)
Figure 2.1. (a) Teflon casting plates; (b) chitosan edible film.
51 2.3.3. Viscosity measurement
Viscosity of the edible slurry was measured using a Brookfield DV-E Viscometer
(Brookfield, MA) fitted with a LV3 spindle at 12 rpm. Each edible slurry sample (with a minimum volume of 200 ml) was tested at 20.0 ± 0.5°C for 5 minutes. The viscosity was recorded every 30 seconds, and the mean viscosity was the average of triplicates.
2.3.4. Drying rate analysis
Drying rate of the slurry was determined using an OHAUS Moisture
Determination Balance (Ohaus Scale Corporation, Parsippany, NJ) fitted with a halogen lamp. For this test, aliquots of edible slurry (5.00 ± 0.01 g) were poured into 2.5 inches wide aluminum dishes, and placed on the balance of the equipment. The distance between the lamp and the balance was adjusted to 2.5 inches. The real-time scale on the balance showed weight changes of the slurry as the solvent evaporated. The halogen lamp temperature was set at 160 ± 5°C during the heating process. The relative humidity around the heated surface area was 0 %. For each sample, the weight was recorded at three-minute intervals until a constant weight was achieved. All results were expressed in percentage of weight by using the following equation:
Weight at each time point ( )× 100% ...... (2.1) Initial weight
2.3.5. Solubility of the films
The water solubility test was conducted to simulate the dissolving property of the film in a liquid system. This method was modified from that reported by Moura and others (2011). Each film sample (100 ± 2 mg) was dried in an oven at 45 ± 2°C for 24
52 hours. The films were then immersed in 30 ml of deionized water and stirred for 1 minute at 25 ± 2°C. The undissolved portions of the film suspension were filtered using the fiberglass # 14 mesh, then oven dried at 45± 2°C for 24 hours. The water solubility of each film was determined from the following equation:
Film initial weight- Undissolved matter weight ( )× 100% ...... (2.1) Initial weight
2.3.6. Functional groups characterization
2.3.6.1. Attenuated Total Reflection Fourier Transform Infrared spectroscopy
(ATR-FTIR)
The ATR-FTIR was used to characterize the functional groups of the films. A
Varian 3100 Infrared Spectroscopy (Varian Inc., Palo Alto CA) with diamond subject at
5 Hz, 4 resolution and 64 scans were applied to all samples. All spectra were collected in the frequency range of 4000-700 cm-1 from four testing points.
2.3.7.2. Multivariate analysis
The spectra from the ATR-FTIR analyses were evaluated using a Pirouette®
Multivariate Analysis Software version 4.0, InfoMetrix, Inc., (Woodville, WA). Soft independent modeling of class analogy (SIMCA) was used to describe and classify the difference between the different edible slurries. The discriminating power was used to compare differences in the wavenumbers versus the intensities of the peaks of each sample. Partial least square regression (PLSR) was obtained by converting the absorbance spectra to secondary derivative spectra. The values of the PLSR showed the linear combination of the variables (Hassel and others 2011).
53 2.3.7. Thermal property analysis
2.3.7.1. Differential scanning calorimetric (DSC) analysis
The thermal stability of the films was evaluated using a 2920 Modulated DSC with Universal Analysis software package v.3.9a (TA Instrument Corp. New Castle,
Delaware). The film samples (11± 0.2 mg) were placed into stainless steel pans and sealed with O-rings. The samples were initially cooled from 25°C (at 20°C/ min cooling rate) to -20°C with 30 ml/ min nitrogen purging. After cooling, the samples were heated to 200°C and then cooled to -20°C at the same condition. Thermal parameters such as glass transition temperature (Tg), enthalpy (ΔH), maximum denaturation temperature
(Tm), and onset of the crystallization temperature (To) of the film samples, corresponding to the endothermic peak areas on the DSC thermogram, were determined by integrating the temperature vs. heat flow curve (Mathew and others 2006). Analysis of the thermograms was done by using the Universal Software that is a part of the equipment software.
2.3.7.2. Thermogravimetric analysis (TGA)
This test was done using a Hi-Res Modulated TGA 2950 Thermogravimetric
Analyzer with Universal Analysis software package v.3.9a (TA Instrument Corp. New
Castle, Delaware). For this test, film samples (11 ± 0.5 mg) were placed in the platinum pans flushed with 90mm Hg of nitrogen, and scanned from 25 to 600ºC at a 30ºC/ min heating rate. Weight loss of the tested samples between 70-100°C was considered as the total moisture in the samples.
54 2.3.9. Statistical analysis
This was conducted using a Minitab Software 15 Microsoft Version (Minitab Inc.,
State College, PA) for both one-way ANOVA with Post Hoc test- Tukey and Student’s t- test to determine the significance effect of acids, chitosan and ethanol concentrations on the properties of the slurries and the films, with a 95% confidence interval. ANOVA analysis provided the overall comparison between treatments, and the Post Hoc test-
Tukey indicated the significant difference between individuals. Multivariate analysis was used to analyze the differences between the ATR-FTIR spectra.
55 2.4. Results and Discussion
2.4.1. Viscosity of the chitosan dissolved in different solvents
Viscosity measurement can provide information on the rheology of a fluid specimen. Figure 2.2 shows the viscosity of chitosan (1.0, 1.5, 2.0%) dissolved in 1.0% acetic acid or 1.0% lactic acid solutions with 0- 20% ethanol concentrations. As the solid content (chitosan) in the solvents increased from 1.0 to 2.0%, the viscosities of the slurries also increased. Also, chitosan dissolved in acetic acid showed a higher viscosity than when dissolved in lactic acid. In the 1.5% chitosan sample, the viscosity increased from 280± 9.5 cp (in lactic acid) to 401± 4.3 cp (in acetic acid). In the 2.0% chitosan samples the viscosity increased from 826.7± 9.8 cp (in lactic acid) to 1023.3± 14.4 cp
(acetic acid) without the addition of ethanol. For all treatments, Figure 2.2 shows that the viscosity increased with increasing concentrations of ethanol in the slurry.
The statistical analysis indicated that viscosity significantly (p<0.05) increased with increasing chitosan concentrations from 1.0% to 2.0%. When the chitosan was dissolved in acetic acid, the viscosities were higher then when dissolved in lactic acid for all treatments. However, in the 1.5% and 2.0% chitosan solutions the viscosity differences were significantly (p< 0.05) higher with increasing ethanol concentrations in the chitosan solutions. For 1.0% chitosan, the effects of the acids were not significant
(p> 0.05). Ethanol also had a significant (p< 0.05) effect on increasing the viscosities of the slurries in both acids.
Viscosity of the film (slurry) is important for producing edible films. Du and others (2008) and Rossman (2009) reported that 1000 cp is the minimal requirement for commercial production of edible films from a slurry using tape-casting methods. Edible
56 film slurries are generally high in moisture and this makes it difficult to produce them by traditional petroleum-based plastic manufacturing methods (extrusion). As a result, casting methods (tape-casting for example) are selected. The use of this method to produce films works well because it allows large quantities of moisture to evaporate from the slurry before fairly dried films are formed (Rossman 2009). Since the heat of drying the slurry is fairly low, burning of the polysaccharide (chitosan) is avoided. The use of traditional petroleum-based plastic extrusion methods to produce edible films from high moisture slurries is not the technique of choice because of the difficulty in evaporating the moisture without burning the ingredients. Extrusion is more suited for protein-based films with low moisture levels in the ingredients.
Several studies have shown that solvent concentration, temperature, molecular weight and acetylation of chitosan can affect the hydrolysis rate and viscosity of the slurry (Il’ina and Varlamov 2004). The amount of chitosan that hydrolyzes in a solvent not only affects the solids content, it also changed the mutual arrangement of two adjacent chitosan polymers in the solution (Wang and others 1991; Rege and Block 1999;
Il’ina and Varlamov 2004). Therefore, the 2.0% chitosan solution in this study had the highest viscosity for all solvents because of the greater number of chitosan molecules it contained, when compared with the 1.0% and 1.5% solutions. Il’ina and Varlamov
(2004) stated that the concentration of hydrogen ions from such solvents directly affect the degree of hydrolysis. Acetic acid showed a better hydrolysis than lactic acid because of the strength of dissociation coefficient. The dissociation coefficient of acetic acid is pKa= 4.756, and lactic acid is pKa= 3.86. Therefore, acetic acid provided more hydrogen ions in the system that could bind with chitosan. The rate in hydrolysis of a given
57 chitosan solution varies depending on the morphology of the polymer (amount and rearrangement of the amorphous and crystalline regions). Viscosity decreases when the polymer orientation becomes more amorphous (Ojovan 2008). As the concentration of ethanol increased in the chitosan polymer, more hydrogen bonding occurred between adjacent chains. Larger macromolecules are known to be produced from the bonding interaction that occurs simultaneously when viscosity increases. As can be seen in Figure
2.2, increasing the ethanol concentration from 0-20% enhanced the viscosity by 61.0%
(lactic acid), 33.3% (acetic acid) in 1.5% chitosan, and 53.2% (lactic acid) and, 59.3%
(acetic acid) in the 2.0% chitosan treatments.
1800.0 1.0% CH-LA 1600.0 1.0% CH-AA 1.5% CH-LA 1400.0 1.5% CH-AA 2.0% CH-LA 1200.0 2.0% CH-AA 1000.0
800.0 iscosity (cp) V 600.0
400.0
200.0
0.0 0% 5% 10% 15% 20% Ethanol
Figure 2.2. The viscosity of 1.0, 1.5, and 2.0% chitosan dissolved in 1.0% lactic acid
(LA) or acetic acid (AA) with variable ethanol concentration (0-20%).
58 2.4.2. Drying rate of chitosan slurries
The drying rate of the slurry helped to determine the optimal process speed for the film making operation during the continuous casting method. Figure 2.3 shows the time versus percentage weight of chitosan dissolved in acetic acid with 0-20% ethanol during the drying process. Results indicated that 1.0% (Figure 2.3a) and 1.5% (Figure 2.3b) chitosan had similar weight changes during the drying process. However, for the 2.0% chitosan samples the differences in the drying rates were more pronounced (Figure 2.3c).
The statistical analysis showed that these differences were significant (p< 0.05) at the 9 minutes time point. At that point, the 0% and 5% treatments were significantly (p< 0.05) different from that of the 15% and 20% treatments. However, for the 1.0% and 1.5% chitosan samples the differences were not significant (p> 0.05) for all time points.
Figure 2-4 shows the time versus percentage weight of chitosan dissolved in lactic acid with 0-20% ethanol during the drying process. Results showed that 1.0% chitosan
(Figure 2.4a) had similar weight changes in all ethanol concentrations. The 1.5% and
2.0% chitosan solutions demonstrated similar drying curves after 9 minutes (Figure 2.4b and 2.4c). The weight reductions of the 1.5% and 2.0% chitosan were significantly (p<
0.05) different in the 15 and 20% ethanol concentrations when compared to the controls.
When the data from Figure 2.3 (acetic acid) and Figure 2.4 (lactic acid) were considered, there were no significant differences (p> 0.05) between acetic and lactic acids on the drying process because of the low quantity of acid (1.0%) added to the system.
Solvent removal is one of the slowest processing step in producing hydrocolloid edible films or coatings (Sano and others 1999). The length of the drying process has potential to increase the cost of the film production if high heat and energy are required.
59 Therefore, it is necessary to optimize the percent solids, acids, and ethanol concentrations on the drying rate. The slope of the drying curves for samples between 0-9 minutes were used to determine the drying rate of the films. This was so because the free water and solvents in the slurries evaporated easily between these time points (0-9 minutes). As the percentage of the solids increased after 9 minutes drying time, the solvent molecules were more tightly bonded to the solid molecules and this made it harder to evaporate the solvents. Martins and others (2009) reported that a strengthing in the cohesive strength of solid-solvent bonding occurs and evaporation becomes more difficult. After 9 minutes, a thin layer of dry film formed on the surface of the slurry and this also increased the barrier against solvent evaporation. The drying times for both acids were shortened from 15 minutes to 13 minutes for the 15% ethanol sample and 12 minutes for the 20% ethanol sample, respectively (Table 2.1). Those times were proximately 13.3% and 20% shorter than the control. The effect of ethanol for increasing the drying rate might be due to its low boiling point and high evaporation properties. To summarize, increasing the concentration of chitosan and ethanol significantly accelerated the drying rate by up to 20%. However, the type of acid did not significantly affect the drying speed.
60 120 0% 1.0% chitosan 5% 100 10% 80 15% 20% 60 eight (%)
W 40
20
0 0 3 6 9 12 15 (a) Time (min)
120 0% 1.5% chitosan 5% 100 10% 80 15% 20% 60 eigh (%)
W 40
20
0 0 3 6 9 12 15 (b) Time (min)
120 0% 2.0% chitosan 5% 100 10% 80 15% 20% 60
eight (%) 40 W
20
0 0 3 6 9 12 15 (c) Time (min) Figure 2-3. The effect of solvent loss on the dry weight of (a) 1.0%; (b) 1.5%; (c) 2.0% chitosan in 1.0% acetic acid with 0-20% ethanol.
61 120 0% 1.0% chitosan 5% 100 10% 80 15% 20% 60 eight (%)
W 40
20
0 0 3 6 9 12 15 (a) Time (min)
120 0% 1.5% chitosan 5% 100 10% 80 15% 20% 60
eight (%) 40 W
20
0 0 3 6 9 12 15 (b) Time (min)
120 0% 2.0% chitosan 100 5% 10% 80 15% 60 20% eight (%)
W 40
20
0 0 3 6 9 12 15 (c) Time (min) Figure 2-4. The effect of solvent loss on the dry weight of (a) 1.0%; (b) 1.5%; (c) 2.0% chitosan in 1.0% lactic acid with 0-20% ethanol.
62
Table 2.1. The drying time of 1.0- 2.0% chitosan in 1.0% acetic or lactic acids with 0-
20% ethanol concentrations.
1.0% chitosan Ethanol concentration 0% 5% 10% 15% 20% Acids End time (minutes) Acetic acid 15.5± 0.7 15.0± 0.0 12.5± 0.4 13.0± 0.0 12.0± 0.0 Lactic acid 15.5± 0.7 14.5± 0.4 13.0± 0.0 13.0± 0.0 12.0± 0.0
1.5% chitosan Ethanol concentration 0% 5% 10% 15% 20% Acids End time (minutes) Acetic acid 15.0± 0.0 14.5± 0.4 13.5± 0.4 13.0± 0.0 12.0± 0.0 Lactic acid 15.0± 0.0 14.5± 0.4 13.5± 0.4 13.0± 0.0 12.0± 0.0
2.0% chitosan Ethanol concentration 0% 5% 10% 15% 20% Acids End time (minutes) Acetic acid 15.0± 0.0 15.0± 0.0 13.5± 0.7 13.0± 0.0 12.0± 0.0 Lactic acid 15.0± 0.0 15.0± 0.0 13.0± 0.7 13.0± 0.0 12.0± 0.0
63 2.4.3. Solubility of chitosan films
Figure 2.5 shows the percentage of the chitosan films that dissolved and that were made with acetic acid (Figure 2.5a) and lactic acids (Figure 2.5b) dissolved in water. As could be seen in Figure 2.5a, the dissolving percentage (%) increased with increasing chitosan and ethanol concentrations. Figure 2.5b demonstrates that the dissolving (%) decreased with increasing chitosan concentrations. However, no trend could be found with increasing ethanol concentrations versus solubility.
The solubility of chitosan made with acetic acid significantly (p< 0.05) increased with increasing chitosan concentrations. No significant (p> 0.05) differences could be found between the solubilities of all chitosan films made with acetic acid when they were exposed to increasing ethanol concentrations. On the contrary, chitosan films prepared by lactic acid showed significant (p< 0.05) reductions in solubility with increasing chitosan concentrations. However, the addition of ethanol did not have a significant (p>
0.05) effect on the solubilities of all chitosan films made with lactic acid. The chitosan films made with acetic acid had a significantly (p< 0.05) higher solubility when compared with the chitosan films made with lactic acids.
In the lactic acid made films, the solubility decreased with increasing chitosan concentrations and this might have been due to the total solids present within the films.
In general, a larger number of polysaccharide molecules would need more solvents or a longer time for saturation of the polymer in order to melt the film. Bonilla and others
(2011) reported that, as chitosan concentration increased in a polymer its crystallinity increased and this made it more difficult for solvents to penetrate and dissolve the film.
However, chitosan films made with acetic acid demonstrated an opposite trend when
64 compared with lactic acid. The solubility increased of chitosan films made with acetic acid may have been caused by the presence of a larger quantity of acetic acid trapped within the matrix of the film. The ATR-FTIR spectra shown in Figure 2.6b illustrated that the chitosan films made with acetic acid had more C-H, COO-, CO-H and CO functional groups when compared with the films made by lactic acid (ATR-FTIR results will be discussed in the next section). Therefore, when exposed to water, the films made with acetic acid had a higher dissolving (%) because they had a faster rate of hydrolysis.
The addition of ethanol did not show any effect on the solubility. This may be so because most of the ethanol might have evaporated during the drying process. However, more research is needed to understand the interaction between chitosan and ethanol.
65 80 Acetic acid 0% 70 5% 60 10% 15% 50 20% 40
30 Dissolving (%) 20
10
0 1.0% 1.5% 2.0% Chitosan (a)
80 Lactic acid 70 0% 5% 60 10% 50 15% 20% 40
30 Dissolving (%) 20
10
0 1.0% 1.5% 2.0% Chitosan (b)
Figure 2.5. The dissolving ability of 1.0- 2.0% chitosan film made by (a) acetic; (b) and lactic acid with 0- 20% ethanol.
66 2.4.4. Attenuated Total Reflection Fourier Transform Infrared spectroscopy (ATR-
FTIR) characterization
Soft independent modeling of class analogy (SIMCA) analysis was first set up to classify the films made with acetic and lactic acids, with three levels of chitosan. It separated the results into two main groups. These were acetic acid (left side) and lactic acid (right side) in Figure 2.6a, and there were three subgroups (1.0, 1.5, and 2.0% chitosan) in each of them. Also, the discriminating power generated from the ATR-FTIR
(Figure 2.6b) showed bands at 862 cm-1, 1120-1160 cm-1, 1338 cm-1 and 1442 cm-1. The cross-validated regression (PLSR) based on ATR-FTIR spectral information and the viscosities of the slurries are showed in Figure 2.6c. The PLSR loading spectra indicated that the 990 cm-1 and 1260- 1290 cm-1 regions were associated with viscosity changes within the chitosan films.
Interclass distances generated from the SIMCA analysis between all the sub groups were greater than 3, which indicated significant (p< 0.05) differences existed between each of the groups. This means that the ATR-FTIR spectra had been successfully characterized the changes in the functional groups from each edible film.
Also, the discriminating power generated from ATR-FTIR (Figure 2.6b) demonstrated that 1442 cm-1 and 1120-1160 cm-1 had the greatest differences between acetic and lactic acids chitosan films, and these corresponded to COO-, C-OH stretching or C-O-C bridge from the acids. Also, ATR-FTIR identified the presence of ethanol at 862 cm-1 (Et-OH) and chitosan at 1138 cm-1 (C-N). This conclusion is supported from research published by Paluszkiewicza and others (2011).
67 Acetic acid Lactic acid PC 2 1.0% chitosan 1.0% chitosan
PC 3 1.5% chitosan PC 1 1.5% chitosan
2.0% chitosan
2.0% chitosan (a)
Figure 3a. SIMCA classification from ATR-FTIR analysis of lactic acid and acetic acid treatments with 1.0-2.0% chitosan.
(b)
(c) Figure 2.6. (a) SIMCA classification from ATR-FTIR analysis of lactic acid and acetic acid treatments with 1.0-2.0% chitosan; (b) discriminating bands; and (c) PLSR loading plot.
68 The cross-validated regression (PLSR) based on ATR-FTIR spectral information and the viscosities of the slurries showed fairly good correlation at r> 0.87. This indicated that differences in the functional groups obtained from ATR-FTIR directly correlated with the changes in the viscosities. Examination of the PLSR loading spectra
(Figure 2.6c) indicated that the 990 cm-1 and 1260- 1290 cm-1 regions were associated with CH and OH bonds from the ethanol. These deductions are supported from research published by Tanka and others (2001). As the amount of chitosan increased, the edible films showed higher C-OH stretching at 1120- 1160 cm-1 in the discriminating power plot
(Figure 2.6b) and these exactly matched the results of the PLSR loading (Figure 2.6c).
However, the linear relationship between viscosity and IR spectra was not very high.
Chen and others (2007) reported that chitosan dissolution might be affected by acid interactions, the protonation of amino groups, and other unknown factors.
In Figure 2.7a, SIMCA analysis showed three main groups comparing the films made with acetic and lactic acids with three levels of chitosan, and the 0-20% ethanol.
Chitosan films made with acetic acid did not show any separating group in the SIMCA analysis. However, The interclass distance of lactic acid with and without ethanol was greater than 3. This result revealed that ethanol addition significantly influenced the biochemical composition and the chemical shift of chitosan films made with lactic acid.
Functional groups, C-O (1220 cm-1) and amide II (1338 cm-1) stretching became significantly visible in the discriminating plots when the ethanol concentration increased
(Figure 2.7a and 2.7b) (Saarakkala and others 2010).
The changes in biochemical composition of the film samples and viscosities of slurries may have caused by ethanol cross-linking with the chitosan polymer. The cross-
69 linking interactions seem to influence the formation of a larger macro polymer. When measuring the viscosity, the resistance in stirring increased as the polymer units became larger, and seem to directly affect the viscosity of the slurries. According to Sano and others (1999) the solubility of chitosan in ethanol is higher than in water. This means that ethanol has a better affinity than water for attachment to the chitosan functional groups. Table 2.2 shows the contribution of the functional groups versus wavenumbers
(cm-1) from all treatments including acids, chitosan, and ethanol.
Table 2.2. The list of functional groups versus wavenumber (cm-1) corresponding to solvents and chitosan parameters.
Functional groups Contributed by Et OH (860- 900 cm-1) Ethanol C-H (990 cm-1) Acids, ethanol C-OH (1141 cm-1) Acids, ethanol C-N (1263 cm-1) Chitosan Amide II (1338 cm-1) Chitosan COO- (1442 cm-1) Acids
70 (a)
4000 1120- 1140 cm-1 C-O
3000
1338 cm-1 Amine 2000 860- 990 cm-1
Discriminating Power EtOH
1000
0
1082.063232 1467.825562 1853.587769 Wavenumber (cm-1) (b) •! 1141.85 1120.63 1338.59 860.24 1091.70 Figure 2.7. Comparison of overall parameters: acetic acid (AA), lactic acid (LA), 1.0-
2.0% chitosan, and 0- 20% ethanol of ATR-FTIR spectrograms using SIMCA analysis.
71 2.4.5. Thermal properties
2.4.5.1. Differential scanning calorimetric (DSC) analysis
Differential scanning calorimetric (DSC) curves displaying the thermally-induced endothermic transitions of chitosan edible films at -20°C to 200°C are shown in Figure
2.8 and Figure 2.9. The major endothermic peaks in the DSC curves of the film samples were observed over a temperature range of 60- 80°C. The DSC curves of the chitosan films made by acetic acid are shown in Figure 2.8. The phase changes for 1.0%, 1.5%, and 2.0% chitosan films occurred at 67.98- 69.16°C (Figure 2.8a); at 67.57- 69.94°C
(Figure 2.8b); and at 67.57- 69.94°C (Figure 2.8c), respectively.
Figure 2.9 shows the DSC curves of chitosan films made by lactic acid. The phase changes of 1.0%, 1.5%, and 2.0% chitosan films are demonstrated at 68.22- 71.58°C
(Figure 2-9a); 69.12- 70.17°C (Figure 2.9b); and 63.50- 71.62°C (Figure 2.9c), respectively.
The endothermic peaks for chitosan films made by acetic acid shows more consistency than these made in lactic acid; however, no significant (p> 0.05) differences were found between those results. As chitosan concentration increased, the transitions in the DSC curve of lactic acid treated samples tended to decrease. Ethanol addition did not show significant (p> 0.05) influence on the endothermic transitions; however, some transitions shifted slightly to higher temperatures.
Heat flow changes in the transitions occurring at 64.0± 0.5 to 71.0± 0.5°C were related to water evaporation associated with the hydrophilic groups from the polymeric structure as described in research reported by Dhanikula and Panchangnula (2004) and
El-Hefian and others (2010). As the chitosan concentrations increased, the phase
72 transitions for water evaporation became less significant because of the increasing solid- water interaction. The solid-water interaction is the bound water associated with chitosan solids. More energy is required to remove the bound water from the chitosan films in such cases. The endothermic peaks for the 1.5% and 2.0% chitosan samples were less sharp when compared with the 1.0% chitosan. Some of the endothermic peaks that shifted slightly to higher temperatures may have been caused by ethanol evaporation, as reported by El-Hefian and others (2010). However, there was no evidence showing how much ethanol was presented in the films. No glass transition temperatures (Tg) could be identified in the DSC thermograms for all films. This might have occurred because of the rigid structure and tight inter- and intra-hydrogen bonding associated with the chitosan chemical structure (Don and others 2005).
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-+#! !+ "+ %+ (a) FG52H> /?01@?/27&'; HCIJ/?K042L)#"M2 (b) (c) Figure 2.8. DSC thermograms of (a) 1.0%; (b) 1.5%; (c) 2.0% chitosan films made by acetic acid with 0-20% ethanol addition. 74 (a) (b) (c) Figure 2.9. DSC thermograms of (a) 1.0%; (b) 1.5%; (c) 2.0% chitosan films made by lactic acid with 0-20% ethanol addition. 75 2.4.5.2. Thermogravimetric analysis (TGA) Thermogravimetric analysis (TGA) was used to understand the correlations between temperature and sample weight changes. In Figure 2.10 and Figure 2.11, the smooth decreasing lines indicate the percent weight changes percent (%; Y-1) and sharp peaks indicate the derivative of the weight percentage per degree (%/°C; Y-2) versus temperature (X). The derivative of the weight loss curves identify the temperature point where weight loss is most apparent. The chitosan films prepared with acetic acid and 0- 20% ethanol are demonstrated in Figure 2.10. The first peak from the left in all samples occurred at 65-75°C. The second peak from the left is located at 100- 150°C; however, a wide temperature range was observed for the 1.0% chitosan films. The third peak from the left is located at 150- 210°C. The highest weight loss (the fourth peak from the left) occurred at 270- 320°C. Figure 2.11 shows the TGA thermograms for the chitosan films made with lactic acid and 0-20% ethanol concentrations. Similar weight loss curves were obtained for both acetic and lactic acids. The first derivative weight peak from the left in the lactic acid treatment occurred at 65-75°C, and the second peak region was located at 140- 210°C. The third peak from the left had the largest weight loss and occurred at 270- 320°C. The statistical analysis indicated that there were no significant (p> 0.05) differences in temperatures ranges versus chitosan concentrations, type of acids, and ethanol additions. Moreover, no significant difference could be found in the weight losses during increasing temperatures for all samples. As a result, changes in solvents and the chitosan concentrations did not significantly (p< 0.05) affect the thermal properties of the films. 76 (a) (b) (c) Figure 2.10. TGA thermal graph of (a) 1.0%; (b) 1.5%; (c) 2.0% chitosan films made by acetic acid with 0-20% ethanol addition. * Black line: Y-1 weight (%); blue line: Y-2 Derived weight (%/ °C). 77 (a) !"% ++++++++++++++++++!1+'/.=?@ABBBBBBB ++++++++++++++++++91+'/.=?@AB+B+B+B ++++++++++++++++++#!1+'/.=?@ABBBBB+C ++++++++++++++++++#91+'/.=?@ABBB+B+B #!! ++++++++++++++++++$!1+'/.=?@ABBB+BBB !"$ 8! ,')-./+015 &'()*"+,')-./+012345 !"# 7! 6! !"! ! #!! $!! %!! 6!! 9!! 7!! (b) :';<'(=/>('+0345 D?)*'(E=A+F6"GH+:H+I?E/(>;'?/E ##! !"% ++++++++++++++++++#1+'/.?ABCDDDDDDD ++++++++++++++++++61+'/.?ABCD+D+D+D ++++++++++++++++++#!1+'/.?ABCDDDDD+E ++++++++++++++++++#61+'/.?ABCDDD+D+D ++++++++++++++++++$!1+'/.?ABCDDD+DDD #!! :! !"$ 9! ,')-./+015 &'()*"+,')-./+012345 8! !"# 7! 6! !"! ! #!! $!! %!! ;!! 6!! 7!! (c) <'=>'(?/@('+0345 FA)*'(G?C+H;"8I+ Saarakkala S, Rieppo L, Rieppo J, Jurvelin JS. 2010. Fourier transform Infarared (FTIR) microscopy of immature mature and degenerated articular cartilage. Microscopy: Sci Technol Appl Educ. 403-414. Sano M, Hosoya O, Taoka S, Seki T, Kawaguchi T, Sugibayashi K, Juni K, Morimoto Y. 1999. Relationship between solubility of chitosan in alcoholic solution and its gelation. Chem Pharm Bull. 47: 1044-1046. Sepe MP. 1997. Thermal Analysis of Polymers, RAPRA Technology LTD, UK, p.13. 83 Tanaka T, Nagao S, Ogawa H. 2001. Attenuated total reflection fourier transform infrared (ATR-FTIR) spectroscopy of functional groups of humic acid dissoving in aqueous solution. Anal Sci. i1081-1084. Tapia-Blácido D, Sobral PJ, Menegalli FC. 2005. Effects of drying temperature and relative humidity on the mechanical properties of amaranth flour films plasticized with glycerol. Braz J Chem Eng. 22:249-256 Tropini V, Lens J-P, Mulder WJ, Silvestre F. 2004. Wheat gluten films cross-linked with 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide and N-hydroxysuccunumide. Ind Crop Prod. 20: 281-289. Wang W, Bo S, Li S, Qin W. 1991. Determination of the Mark-Houwink equation for chitosans with different degrees of deacetylation. Int J Macromol. 13: 281-317. Wang W, Xu D. 1994. Viscosity and flow properties of concentrated solutions of chitosan with different degrees of deacetylation. Int J Biol Macromol. 16: 471-152. 84 CHAPTER 3 THE INCORPORATION OF VITAMIN E INTO CHITOSAN FILMS AND ITS EFFECT ON THE MATERIAL PROPERTIES (DRYING RATE, SOLUBILITY) 3.1. Abstract Nutrient fortified edible films have been used to improve the health benefit and extend the shelf life of various foods. The effect of vitamin E on the properties of chitosan edible film was evaluated in this study. The edible ingredient slurries were made with 1.0-2.0% chitosan blended with 250 or 500 mg vitamin E by two formats: vitamin E added before lecithin (VE first) and vitamin E mixed with lecithin (VE mixed) then added to the slurry. Changes in drying rate and vitamin E concentrations in the various formulations were determined using a OHAUS Moisture Determination Balance, and high performance liquid chorography (HPLC), respectively. Solubility was determined by dissolving the film in water for 1 minute then measuring the weight changes. Results showed that vitamin E addition significantly (p< 0.05) increased the drying rate of the formulation by 3- 9 minutes. The smallest quantity of vitamin E incorporating uptake in the films occurred in the 1.0% chitosan materials, which caused vitamin E accumulation on the surface instead of in the matrix of the film. The blending process for vitamin E and lecithin influenced the stability of the chitosan-vitamin E networks. The 2.0% chitosan had the best homogeneity with the blended vitamin E- 85 lecithin mixture. This also decreased the film solubility, especially in samples with higher chitosan and VE first concentrations. 3.2. Introduction The demand for nutraceutical and functional foods has increased in the past decade along with an awareness of healthy eaten. Smart packaging such as edible films is an alternative method for providing functional additives without changing the original ingredients and the processing method of food products. Rossman (2009) indicated that polysaccharide-based edible coating or films could be simply incorporated with colors, flavors, sweeteners, and micronutrients due to the chemical structure and the polarity of these materials. The addition of active ingredients may vary depends on the functions and the applications of the developed films. Polysaccharides such as chitosan has been used as the main ingredient in some edible films. The cationic property of chitosan supplies the electrostatic interaction that can be easily bounded to other anionic compounds. Also, the high density of amino groups and hydroxyl groups provide coagulation ability with minerals, lipids, and proteins (Park and Zhao 2004). As a result, chitosan is an ideal ingredient for the manufacture of vitamin E fortified edible films. Vitamin E (tocopherol) is an essential fat-soluble vitamin that protects body tissue against free radical damage. It is widely used as a food additive to improve the stability of lipids and extend the shelf life of oxygen sensitive products (Gonçalves and others 2010). According to a recent dietary intake survey, the prevalence of vitamin E intake by elderly Americans (above the age of 71) is 75% lower than the Estimated Average 86 Requirement stated by the United States National Academy of Science (Troesch and others 2012). The healthy youth program office at Linus Pauling Institute (LPI) also reported that none of the children from 172 elementary schools surveyed met the recommended intake level for vitamin E (Frei 2012). Therefore, there is a need to develop methods aimed at solving this problem. Vitamin E fortified edible film can be used in school cafeterias on ready-to-eat products as a means of improving vitamin E intake in elementary school as a possible solution to this problem. Vitamin E is a lipophilic compound containing a hydroxyl group on the phenolic ring (Figure 3.1a). It can be used to quench free radicals and has one hydroxyl group available for hydrophilic bonding (Park and Zhao 2004). This weak hydrophilic bonding is not strong enough to bind with chitosan in an acidic solution. Therefore, an emulsifier is needed to connect the chitosan molecules with that of vitamin E. An emulsifier such as lecithin is a natural component of phospholipids and it is an economical food additive (Rossman 2009). The molecular structure of lecithin shows both hydrophilic and hydrophobic properties. As a result, lecithin is an excellent surface-active agent that can be used in various food products (Figure 3.1b) (The Solae Company 2012). Also, Liu and Park (2009) reported that lecithin helps to stabilize vitamin E in encapsulation technology. 87 (a) (b) Figure 3.1. The chemical structure of (a) α-tocopherol; and (b) lecithin. The objective of this study was to determine the optimal method for incorporating vitamin E into a chitosan polymer and to investigate how it influences the properties of the edible film. Two blending processes were used. Vitamin E was blended into the edible film mixture in two different steps: (1) vitamin E added before lecithin (VE first); and (2) vitamin E mixed with lecithin (VE mixed) and then added to the chitosan solutions. 88 3.3. Materials and Materials 3.3.1 Materials Medium molecular weight (94% purity) chitosan powder was provided by Huantai Goldenlake Carapace Products Co., Ltd (Tsingtao, China). This was used as the main ingredient in the films. Lactic acid (88% Food Chemical Codex, FCC) manufactured by Birko Corporation (Henderson, CO) and Giant Eagle brand distilled white vinegar containing 5% acetic acid (Pittsburg, PA) were used to dissolve the chitosan powder. One hundred percent natural vitamin E with mixed tocopherol (400 IU per softgel) was purchased from Whole Foods Market (365™ Everyday Value, Austin, TX). Glycerol USP Kosher 99.7% (Chemical direct, online, USA) and soy lecithin (Solec® 100L, The Solae Company, St. Lousis, MO) were added as a plasticizer and an emulsifier, respectively. A standard 18 mesh (0.0394 inches) fiberglass screen was purchased from an ACE Hardware Store in Columbus, OH and used for the film solubility test. High-performance liquid chromatography (HPLC) grade methanol, α- tocopherol standard, nylon membrane filter (13 mm diameter, 0.45 µm pore size) (Fisher Scientific Inc., Fairlawn, NJ); Allsphere ODS-2 analytical column (250 mm× 4.6 mm, 5 um particle size) with a guard column (Alltech Associates, Deerfield, IL) were used for the HPLC analysis. 3.3.2. Edible Films Formation Edible film solutions were prepared at 22 ± 1°C by dispersing the chitosan powder (1.0, 1.5 and 2% w/w) in distilled water. Lactic acid (1%, w/w) was added to the suspension and stirred (300 rpm) for 3 minutes to dissolve the chitosan powder. Glycerol 89 0.5% (w/w) was added to the solution and mixed for 3 minutes. It was used as a plasticizer to provide flexibility to the films. A 2.0% lecithin solution was then added to the chitosan mixture to produce a 0.1% (w/w) blend as the total dry weight of lecithin in the final films. Lecithin acted as an emulsifier that provided hydrophilic and hydrophobic interactions. The hydrophobic interactions were to help the film provide a good attachment to the food surfaces, especially to the high-fat products. Final slurry samples were prepared by the addition of Vitamin E purchased from the Whole Foods Market in Columbus, OH (250 and 500 mg) and added to the chitosan slurries with or without premixing with 2.0g of lecithin solution. The lecithin or vitamin E- lecithin/chitosan solutions were allowed to settle for one hour prior to further analyses (viscosity and drying rate measurement). Edible films were prepared by casting the edible slurry into 10 inch radius Teflon plates. Aliquots of 33.0± 0.5 ml, 31.0± 0.5 ml, and 29.0± 0.5 ml of 1.0%, 1.5%, and 2.0% chitosan slurries were then cast in order to obtain films of uniform thicknesses (50.0 ± 2.5 µm). This was oven dried at 45 ± 2 °C for 2 hours, then the dried films peeled off from the plate surfaces. The thickness was controlled by the percent solids in the solution. The thicknesses were measured using a 2804S-10 Agd Dialindicator Mitutoyo Guage (Aurora, IL) and reported in mil (1/1000 inch). 90 Scheme 3.1. The flow chart of chitosan-vitamin E slurry formation 1.0, 1.5, 2.0% Chitosan powder Dissolved in 1% (w/w) lactic acid solution Mixed for 3 min Plasticizer added (0.5% w/w glycerol) Mixed for 3 min Added 250/500 mg Vitamin E 250/500 mg Vitamin E blended with Mixed for 3 min 2.0 g (5% w/w) lecithin solution Added 2.0 g (5% w/w) lecithin then added into the chitosan slurries solution Mixed for 3 min Final chitosan-vitamin E slurry 33± 0.5 ml 31± 0.5 ml 9.0± 0.5 ml 1.0% chitosan solution 1.5% chitosan solution 2.0% chitosan solution Cast into Teflon plates 91 3.3.4. Drying rate analysis Drying rate of the slurry was determined using an OHAUS Moisture Determination Balance (Ohaus Scale Corporation, Parsippany, NJ) fitted with a halogen lamp. For this test, aliquots of edible slurry (5.00 ± 0.01 g) were poured into 2.5 inches wide aluminum dishes, and placed on the balance of the equipment. The distance between the lamp and the balance was adjusted to 2.5 inches. The real-time scale on the balance showed weight changes of the slurry as the solvent evaporated. The halogen lamp temperature was set at 160 ± 5°C during the heating process. The relative humidity around the heated surface area was 0 %. For each sample, the weight was recorded at three-minute intervals until a constant weight was achieved. All results were expressed in percentage of weight using the following equation: Weight at each time point ( )× 100% ...... (3.1) Initial weight 3.3.5. High performance liquid chromatography (HPLC) analysis 3.3.5.1. Sample preparation for HPLC analysis This experiment was designed to determine the amount of vitamin E incorporated in the film. Two sets of samples were prepared for this test. On set was used to examine the concentrations of vitamin E on surface of the film. In the other set the vitamin E in the matrix of film was determined. For the film surface test (test 1), 100 ± 5 mg of the sample were immersed in 5.0 ml of 100% methanol, and vortexed for 2 minutes. The solution was filtered using a 0.45 µm nylon membrane, then injected into the HPLC system. For the overall vitamin E quantification (test 2), 50± 2 mg samples were briefly frozen by liquid nitrogen and grounded into a powder. The powder was mixed with 4.0 92 ml 100% methanol and vortexed 2 min for 5 times (with 2 min interval during each time). All solutions were filtered and then directly injected into the HPLC system. 3.3.5.2. HPLC Analysis The HPLC analysis was done to determine the amount of vitamin E in the film samples using a method reported by Hassel and Rodriguez-Saona (2011). The system had a photodiode array detector. The vitamin E was detected at λmax 290 nm at ~7 min retention time with 25 µl sample injection. The analysis was conducted using an HP- 1050 reverse-phase HPLC system with an Allsphere ODS-2, 5um analytical column. The flow rate of the mobile phase (100% of methanol) was 1 ml/min. Five-minute intervals for the column wash were used between every sample injection. The concentration of vitamin E was calculated using a linear equation of HPLC peak area of standard (α- tocopherol) in methanol. Each measurement was done in duplicate and the mean value of vitamin E incorporated into the film was calculating by vitamin E concentration of the overall (test 2) subtracting the vitamin E concentration on the surface (test 1). 3.3.6. Solubility of vitamin E fortified chitosan films The water solubility test was conducted to simulate the dissolving property of the film in a liquid system. This method was modified from that reported by Moura and others (2011). Each film sample (100 ± 2 mg) was dried in an oven at 45 ± 2°C for 24 hours. The films were then immersed in 30 ml of deionized water and stirred for 1 minute at 25 ± 2°C. The undissolved portions of the film suspension were filtered using 93 the fiberglass # 14 mesh, then oven dried at 45± 2°C for 24 hours. The water solubility of each film was determined from the following equation: Film initial weight- undissolved matter weight ×100% ...... (3.2) Film initial weight 3.3.7. Statistical analysis This was conducted using a Minitab Software 15 Microsoft Version (Minitab Inc., State College, PA) for both one-way ANOVA with Post Hoc test- Tukey and Student’s t- test to determine the significant effect of chitosan, vitamin E concentrations and the blending processes on the properties of the films with a 95% confidence interval. ANOVA provided the overall comparison between treatments, and the Post Hoc test- Tukey indicated significant differences between individual treatments. The statistical analysis was used to: (1) compare chitosan concentrations versus 2 levels of vitamin E; (2) evaluate chitosan, vitamin E and the blending process on the drying rate, incorporated vitamin E concentration, and solubility of the films. The student’s t-test was used to compare the two levels of vitamin E and the two blending processes on the drying rate, incorporated vitamin E concentration, and solubility of the samples. Two replicates were tested for each treatment. 94 3.4. Results and Discussion 3.4.1. Drying rate of vitamin E fortified chitosan slurries The drying rate of the slurry helps to determine the optimal process speed for film making operations during continuous casting methods. Table 3.1 shows the drying times for 1.0-2.0% chitosan blended with 250 mg and 500 mg vitamin E. The drying time ended within 15.0-24.0 minutes in the 1.0% chitosan; 15.5-19.5 minutes in the 1.5% chitosan, and 15-18 minutes in the 2.0% chitosan samples. The longest drying time was showed to be with the 1.0% chitosan having 500 mg vitamin E added before the lecithin (VE first). In the 1.0% and the 1.5% chitosan slurries, both 250 and 500 mg vitamin E samples demonstrated significantly (p< 0.05) longer drying times than the control (without vitamin E) either in the VE first or in the VE mixed (vitamin E mixed with lecithin). In the 2.0% chitosan slurries, only the 500 mg VE first showed significantly (p< 0.05) longer drying time when compared with the other treatments. Figure 3.2 shows the curves for drying times versus percentage weights of 1.0- 2.0% chitosan blended with 250 mg and 500 mg VE first. Figure 3.3 demonstrates the drying curves of 1.0-2.0% chitosan blended with the 250 mg and 500 mg VE mixed. Similar drying curves were observed for all samples, but slight differences were seen between the 6-9 minute time points. Approximately 50% weight remained in the controls after 6 minutes drying, which was 10% less weight when compared with the vitamin E samples (both 250 mg and 500 mg addition). However, the chitosan slurries made by VE mixed indicated a faster drying rate than the VE first. 95 Table 3.1. The drying time of 250 and 500 mg vitamin E blended before lecithin or mixed with lecithin in 1.0- 2.0% chitosan solutions. 1.0% chitosan Vitamin E concentration 0% 250 mg 500 mg Process Control First Mixed First Mixed Drying time (minute) 15.0± 0 22.5± 2.1 21.0± 0 24.0± 0 21.0± 0 1.5% chitosan Vitamin E concentration 0% 250 mg 500 mg 96 Process Control First Mixed First Mixed Drying time (minute) 15.5± 0.7 19.5± 2.1 18.0± 0.8 18.0± 0 18.0± 0 2.0% chitosan Vitamin E concentration 0% 250 mg 500 mg Process Control First Mixed First Mixed Drying time (minute) 15.0± 0 15.0± 0 15.0± 0 18.0± 0 15.0± 0 120 100 Control 250 mg 80 500 mg 60 eight (%) W 40 20 0 0 3 6 9 12 15 18 21 (a) Time (minutes) 120 Control 100 250 mg 80 500 mg 60 eight (%) W 40 20 0 0 3 6 9 12 15 18 21 (b) Time (minutes) 120 Control 100 250 mg 80 500 mg 60 eight (%) W 40 20 0 0 3 6 9 12 15 18 21 (c) Time (minutes) Figure 3.2. The drying rate of vitamin E fortified edible solutions (a) 1.0%; (b) 1.5%; and (c) 2.0% chitosan with 250 mg and 500 mg VE first. 97 120 Control 100 250 mg 500 mg 80 60 eight (%) W 40 20 0 0 3 6 9 12 15 18 21 (a) Time (minutes) 120 Control 100 250 mg 500 mg 80 60 eight (%) W 40 20 0 0 3 6 9 12 15 18 21 (b) Time (minutes) 120 Control 100 250 mg 500 mg 80 60 eight (%) W 40 20 0 0 3 6 9 12 15 18 21 (c) Time (minutes) Figure 3.3. The drying rate of vitamin E fortified edible solutions (a) 1.0%; (b) 1.5%; and (c) 2.0% chitosan with 250 mg and 500 mg VE mixed. 98 Results of the drying rate indicated that the slurries without vitamin E dried faster than the slurries with vitamin E additions. The 2.0% chitosan blended with vitamin E showed a faster drying speed than in the 1.0% chitosan. This could be explained by the amount of amine groups that were in the chitosan solution. The amine groups with cation properties in the chitosan structure are essential for binding with other compounds such as emulsifiers and vitamin E (Park and Zhao 2004). Thus, more vitamin E remained within the networks instead of dissociating from the mixture. This is supported in reports published by Martins and others (2012). Moreover, the higher moisture contents of the 1.0% chitosan slurry caused a stronger hydrophilicity of the system. The hydrophobic vitamin E probavly became immiscible in this highly hydrophilic solution and may have been pushed out of the chitosan network. Also, the higher the moisture content, the greater the competition with vitamin E. This is so because the chitosan molecules would have had a higher partitioning for the water molecules when compared with the vitamin E. When vitamin E dissociated from the polymeric matrix, the overall emulsion system became unstable. Unincorporated vitamin E then clustered into an upper layer in the slurry. This formed into a hydrophobic layer and acted as a barrier to water evaporation. As a result, the 1.0% chitosan slurries showed longer drying time than the 1.5% and the 2.0% chitosan slurries. The dissociation of vitamin E can also explain the drying rate differences between the VE first and the VE mixed. According to Lu and others (2008) the blending processes of emulsifiers and vitamin E affect the stability of emulsions. The vitamin E slurries prepared by the VE first in this present study, showed less stability when compared with VE mixed. Therefore, the drying times of the VE first chitosan slurries were longer than the VE mixed (more discussion will be included in HPLC section). 99 In summary, the drying rate was significantly affected by the amount of chitosan and the degree of incorporated vitamin E within the films. A faster drying speed was observed in the 2.0% chitosan with VE mixed slurry. 3.4.2. Incorporation of vitamin E in chitosan edible films High performance liquid chromatography (HPLC) was used to determine the vitamin E concentration on the film surface and the total (overall) amount incorporated into the matrix. The standard curves for α-tocopherol are shown in Figure 3.4a and Figure 3.4b. The linear equation (y=2275.1612x- 314.9707) from Figure 3.4a was used to calculate the vitamin E concentrations within the 200-1000 peak areas. The linear equation (y= 19938.0328x- 101033.5765) from Figure 3.4b was used to calculate vitamin E concentrations within 1000-100000 peak areas. Two samples (250 and 500 mg) of vitamin E were analyzed for their initial concentrations using HPLC before adding them to the edible slurries (represented as “before process” in Figure 3.5 and Figure 3.6). The result from the HPLC analysis showed that the concentration on the label of the product was 6.5% less than what was reported. HPLC determined concentrations of vitamin E were for all the calculations in this study. To further explain the results obtained from drying rate, vitamin E fortified films were divided by two types of measurements: (1) to analyze vitamin E concentration on the film surface (surface) and; (2) to detect the total vitamin E concentration in the entire films (overall). The amount of incorporated vitamin E was calculated by subtracting the concentration on surface from the overall (Incorporated vitamin E= Overall- Surface). 100 Figure 3.5 shows the amount of vitamin E detected by HPLC for (a) 250; and (b) 500 mg VE first in 1.0- 2.0% chitosan. Results indicated that in the 250 mg VE first (Figure 3.5a), the overall amount of vitamin E significantly (p< 0.05) increased with increasing chitosan concentrations. However, the concentration of vitamin E on the surface was significantly (p> 0.05) reduced as chitosan concentrations increased. In the 500 mg VE first (Figure 3.5b), no specific trend between chitosan concentrations (1.0-2.0%) and the amount of vitamin E (250/500 mg) was found in the overall or on the surface of the films. On the other hand, the 1.5% chitosan showed the highest overall and the surface concentrations when compared to other VE first samples. The statistical analysis indicated that the presence of vitamin E in the overall amount and on the surface of the film significantly (p< 0.05) increased with increasing the vitamin E concentrations. The 250 mg VE mixed is shown in Figure 3.6a and it demonstrated that the 1.5% chitosan had the highest vitamin E concentration for the overall and amount on the surface of the films. In the 500 mg VE mixed (Figure 3.6b), the 2.0% chitosan showed the highest overall vitamin E concentration. The presence of vitamin E on the surface significantly (p< 0.05) decreased with increasing chitosan concentrations. The statistical analysis also demonstrated that the vitamin E concentrations in the overall films were significant (p< 0.05) greater when increasing the initial VE mixed concentrations. Table 3.2 shows the vitamin E concentrations (overall, surface, and the incorporated) of all the films. In the 250 mg vitamin E additions, 2.0% chitosan VE first showed the highest incorporated vitamin E concentration when compared with other 250 mg samples. In the 500 mg treatment, 2.0% VE mixed showed the highest incorporated vitamin E concentration when compared with other 5o0 mg samples. However, 1.0% chitosan had the 101 lowest presence of vitamin E incorporation, and only 5- 20% of initial concentrations of vitamin E were incorporated into the films. Also, a large amount of vitamin E was present on the surface of the film in the 1.0% chitosan films, and this could explain why the 1.0% chitosan films had the slowest drying rate (Table 3.1). 1.60E+04 1.40E+04 1.20E+04 1.00E+04 ea y = 2275.1612x - 314.9707 8.00E+03 R² = 0.9960 Peak ar 6.00E+03 4.00E+03 2.00E+03 0.00E+00 0 2 4 6 8 (a) α-tocopherol (mg) 3.00E+06 2.50E+06 2.00E+06 ea 1.50E+06 y = 19938.0328x - 101033.5765 Peak ar 1.00E+06 R² = 0.9999 5.00E+05 0.00E+00 0 50 100 150 (b) α-tocopherol (mg) Figure 3.4. The HPLC standard curves with linear equations of α-tocopherol (a) low concentration (0.25-6.25 mg); (b) high concentration (1.25-125 mg). 102 250mg VE first 250 Before process Overall * 200 Surface * 150 100 Concentration (mg) 50 0 Before process 1.0% 1.5% 2.0% Chitosan (a) 500mg VE first 500 450 Before process Overall * 400 Surface * 350 300 250 200 150 Concentration (mg) 100 50 0 Before process 1.0% 1.5% 2.0% Chitosan (b) Figure 3.5. The concentrations of vitamin E quantified by the HPLC in 1.0- 2.0% chitosan films with (a) 250 mg; and (b) 500 mg VE first. *Overall: vitamin E incorporated inside the film and on the surface *Surface: vitamin E residual on the film surface. 103 250mg VE mixed 250 Before process Overall * 200 Surface * 150 100 Concentration (mg) 50 0 Before process 1.0% 1.5% 2.0% Chitosan (a) 500mg VE mixed 550 500 Before process * 450 Overall Surface * 400 350 300 250 200 150 Concentration (mg) 100 50 0 Before process 1% 1.5% 2.0% chitosan (b) Figure 3.6. The concentrations of vitamin E quantified by the HPLC in 1.0- 2.0% chitosan films with (a) 250 mg; and (b) 500 mg VE mixed. *Overall: vitamin E incorporated inside the film and on the surface *Surface: vitamin E residual on the film surface. 104 A greater incorporation of vitamin E was found in the VE mixed. The 250 mg VE mixed with 2.0% chitosan showed 44.5% (111.34± 0.22 mg) of the vitamin E incorporated within the films. This was because of the presence of lecithin. Lecithin acted as an emulsifier that provided both hydrophilic and hydrophobic characteristics. Vitamin E combined with lecithin while being mixed together; consequently, the vitamin E- lecithin mixture had stronger hydrophilicity than vitamin E by itself. As the vitamin E-lecithin mixture was introduced into the chitosan slurries, the covalent cross-linking and hydrogen bonds from the chitosan provided sites for chemical bonding (Matsuda and others 2005; Lu and others 2008). Also, the 500 mg VE mixed in the 2.0% chitosan showed 73.6% (368.60± 12.40 mg) vitamin E incorporated within the polymers, which was the highest vitamin E incorporated concentration in all treatments. A similar trend demonstrated in Figure 3.5 and Figure 3.6, showed that vitamin E incorporation increased with increasing chitosan concentrations. This could be explained by the cation connection within the chitosan polymeric structures. The presence of cation bonding increased with increasing chitosan concentrations. Therefore, more vitamin E- chitosan connections were formed in the 2.0% chitosan slurries than in the 1.0% solution. 105 Table 3.2. The amount of vitamin E presence in the overall, on the surface, and incorporated chitosan films by HPLC analysis. Treatment 250 mg 500 mg Concentration (mg) Film making process Film making process 9 6 Chitosan Before process VE first VE mixed Before process VE first VE mixed Overall 1.0% 87.66± 0.87 106.22± 5.62 146.21± 5.92 266.38± 8.54 1.5% 234.89± 6.81 124.85± 1.41 127.31± 4.20 469.80± 10.91 360.44± 1.52 227.38±10.73 2.0% 159.54± 0.96 88.45± 1.65 275.54± 14.45 407.68± 13.38 Surface 1.0% 77.69± 2.83 55.70± 0.12 106.84± 2.50 133.69± 7.38 106 1.5% 234.89± 6.81 66.87± 0.39 61.44± 0.30 469.80± 10.91 168.34± 0.25 43.48± 0.28 2.0% 48.19± 0.95 34.61±0.30 64.91± 0.38 39.07± 0.78 Incorporated 1.0% 9.96± 2.67 50.52± 4.42 39.37± 2.80 132.69± 12.00 (Overall- 1.5% 234.89± 6.81 57.98± 0.92 65.88± 5.65 469.80± 10.91 211.79± 0.27 183.90± 10.38 surface) 2.0% 111.34± 0.22 53.84± 1.28 210.63± 14.93 368.60± 12.40 3.4.3. Solubility of vitamin E fortified chitosan films The solubility of the films could be extrapolated to simulate the mouth feel and the speed of dissolving when consuming the edible films. Figure 3.7a shows the percentage of the edible films that dissolved in water. This represent the chitosan film made by blending vitamin E before adding the lecithin solution (VE first). In the 1.0% chitosan films, the VE first addition did not show any difference in vitamin E concentration. However, in the 1.5% and the 2.0% chitosan, the solubility decreased significantly (p< 0.05) with increasing vitamin E concentrations. Figure 3.7b demonstrates the percentage of edible films dissolved in water versus the concentrations of the chitosan made by blending vitamin E with lecithin (VE mixed). As could be seen, the solubility of the films significantly (p< 0.05) decreased when vitamin E concentrations increased. In the controls (without vitamin E), solubility decreased with increasing chitosan concentrations. This was due to the amount of solids in the system. Bonilla and others (2011) reported that, as the chitosan concentration increases in a polymer it crystallinity increases and this makes it more difficult for water to penetrate and dissolve the film. However, the films made with VE first showed less dissolving ability than the films made with VE mixed. The HPLC results revealed that the chitosan films made by VE first obtained more vitamin E on the film surface. Hence, the hydrophobicity of vitamin E repelled significant water attachment and this reduced the solubility of the films. On the other hand, the VE mixed demonstrated a good dissolving ability in all chitosan concentrations (Figure 3.7b). As could be seen, the solubility in the 2.0% chitosan increased by 37.7% in 250 mg VE mixed, and 54.7% in 500 mg VE mixed when 107 compared to the control. However, the mechanism that caused this outcome is still unclear. One possibility might be the chitosan-vitamin E-lecithin-water interaction. However, the exact mechanism is unknown and could not be found in the literature. Thus, more research is needed to address this issue. Martins and others (2012) reported that a large amount of α-tocopherol addition could increase the water vapor permeability of chitosan films. This occurred because the hydrophobic vitamin E affected the cohensive density of the chitosan networks and tended to push the chitosan polymeric chains apart. Therefore, the polymer became less crystalline and this caused the change in water permeability (Bonilla and others 2011). In summary, the solubility of the vitamin E incorporated chitosan film was affected by the chitosan and vitamin E concentrations and the blending processes of vitamin E. Results indicated that, VE mixed had the greatest dissolving ability than other treatments. 108 (a)! VE first 100 Control 90 250 mg 80 500 mg 70 60 50 40 Dissolving (%) 30 20 10 0 1.0% 1.5% 2.0% Chitosan ! (a) VE mixed 100 Control ! "#$! 90 250 mg 80 500 mg 70 60 50 40 Dissolving (%) 30 20 10 0 1.0% 1.5% 2.0% Chitosan (b) Figure 3.7. The dissolving (%) of chitosan films with (a) vitamin E added before lecithin (VE first); and (b) Figure 3.7. The dissolving (%) of chitosan films with (a) vitamin E added before lecithin vitamin E mixed with lecithin (VE mixed). (VE !first); and (b) vitamin E mixed with lecithin (VE mixed). ! """! 109 3.5. Conclusion This work demonstrated that two concentrations of vitamin E had been successfully introduced into edible film system. The best incorporation was showed in the VE mixed treatment. Also, increasing chitosan concentrations enhanced the vitamin E incorporation due to the amount of amine groups present in the chitosan molecule. The vitamin E addition not only affected the drying rate of the edible slurries, it also influenced the solubility of the films by decreasing the crystallinity of the polymeric structure. 3.6. References Bonilla J, Atarés L, Vargas M, Chiral A. 2011. Effect of essential oils and homogenization conditions on properties of chitosan based films. Food Hydrocolloids. 26: 9-16. De Nardo, T., Shiroma-Kian, C., Halim, Y., Francis, D., Rodriguez-Saona, L. 2009. Rapid and simultaneous determination of lycopene and b-carotene contents in tomato juice by infrared spectroscopy. J Agric Food Chem. 57:1105–1112. Gonçalves CMB, Tomé LC, Coutinho AP, Marrucho IM. 2011 Addition of α-Tocopherol on Poly(lactic acid): thermal, mechanical, and sorption properties. J Appl Polym Sci. 119: 2468- 2475. Frei S. Linus Pauling Institute’s Healthy Youth Program. Linus Pauling Institute Research Newsletter Spring/Summer 2012. Lu Y, Wang YJ, Tang X. 2008. Formulation and thermal sterile stability of a less painful intravenous clarithromycin emulsion containing vitamin E. Int J Pharm. 346: 47-56. 110 Mathew S, Brahmakumar M, Abraham TE. 2006. Microstructureal imaging and characterization of the mechanical, chemical, thermal, and swelling properties of starch-chitosan blend films. Biopolymers. 82: 176-187. Matsuda A, Kobayashi H, Itoh S, Kataoka K, Tanaka J. 2005. Immobilization of laminin peptide in molecularly aligned chitosan by covalent bonding. Biomaterials. 26: 2273- 2279. Marins JT, Cerqueria MA, Vincete AA. 2012. Influences of α-tocopherol on physicochemical properties of chitosan-based films. Food Hydrocolloids. 27: 220-227. Park S-I, Zhao Y. 2004. Incorporation of high concentration of mineral or vitamin into chitosan-based films. J Agric Food Chem. 52: 1933-1939. Troesch B, Eggersdorfer M, Weber P. 2012. 100 Years of Vitamins: Adequate intake in the elderly is still a matter of concern. J Nutr. 142: 979-980. United States Department of Agriculture, National agriculture online library. 2012. http://fnic.nal.usda.gov/dietary-guidance/dietary-reference-intakes Urano S, Yano K, Matsuo M. 1988. Membrane-stabilizing effect of vitamin E: effect of - tocopherol and its model compounds on fluidity liposomes. Biochem Biophys Res Commun. 150: 469-475. Wessling C, Nielsen T, Giacin JR. 2001. Antioxidant ability of BHT- and alpha- tocopherol-impregnated LDPE film in packaging of oatmeal. J Sci Food Agric. 81: 194-201. 111 CHAPTER 4 THE INCORPORATION OF VITAMIN E INTO CHITOSAN FILMS AND ITS EFFECT ON THE MATERIAL PROPERTIES (VISCOSITY, ATR-FTIR, THERMAL ANALYSIS) 4.1. Abstract In recent times, increased attention has focused on the development of edible films because of increasing food safety and health benefits needs. The objective of this study was to determine the effect of vitamin E on its viscosity of chitosan slurry, and the thermal properties of the resultant films. Edible slurries were prepared by blending 250 or 500 mg vitamin E into 1.0-2.0% chitosan, and then casting them in Teflon plates. In one blending process vitamin E was added to the slurry before the lecithin (VE first). In a second blending process vitamin E was mixed with lecithin (VE mixed). Viscosities of the chitosan-vitamin E slurries were measured using a viscometer, and functional groups on the polymeric chains were identified by Attenuated Total Reflectance Fourier Transform Infrared (ATR-FTIR) with multivariate analysis. Differential scanning calorimetry (DSC) and thermogravimetric analysis (TGA) were used to characterize changes in the thermal properties of the films. The results indicated that the second vitamin E-lecithin blending process had the greatest effect on the viscosity. This viscosity significantly (p< 0.05) increased from 108± 9.5 cp (control) to 491± 6.2 cp 112 (VE mixed) in the 1.5% chitosan slurry, and from 827± 9.8 cp to 1200± 20 cp (VE mixed) in the 2.0% slurry. Functional groups related to vitamin E were identified at 827 and 2900-3200 cm-1in the ATR-FTIR analysis. The linear correlations between the functional groups and viscosity (r>0.98) were obtained from the multivariate analysis. No significant difference (p> 0.05) was seen in the DSC thermograms for all treatments. However, TGA analysis showed that the decomposition of vitamin E occurred at 430- 450°C. Therefore, vitamin E fortified chitosan film prepared by VE mixed had the best incorporation ability. 4.2. Introduction The demand for developing edible packaging has increased due to its simple preparation method and multi function uses. Edible packaging is capable of providing quality improvement, shelf life extension, sensory enhancement, and functional additives to food products (Janjarassku and Krochta 2010). The incorporation of functional ingredients such as vitamins, minerals and antioxidants into edible films have enhanced the applications of edible packaging in the food industry (Park and Zhao 2006; Janjarasskul and Krochta 2010; Martins and others 2012). Chitosan is an optimal component for incorporating food additives or micronutrients since it provides bonding sites for electrostatic interaction within the polymeric network. Also, it is known for its antimicrobial, biodegradable, biocompatible, and film-forming properties, which can be used in food and health related products (Jongrittiporn and others 2001; Jayakumar and others 2005; Jayakumar and others 2007). 113 Chitosan can be easily dissolved in a weak organic acid without heating; thus, it is an ideal matrix of combining heat sensitive nutrients and antioxidants. Vitamin E is a well-known antioxidant and an essential nutrient for human health. It is used as a nutraceutical (antioxidant) supplement for human consumption because of its free radicals scavenging ability (Wessling and others 2000). As a result, developing vitamin E fortified chitosan films will not only help to improve the nutrition content of products, it will also to extend the shelf life of packaging materials by preventing oxidation. In developing vitamin E incorporated chitosan films it is essential to consider the chemical structures of vitamin E and chitosan. Chitosan dissolved in acids shows high hydrophilic properties. Therefore, vitamin E (lipophilic compound) will have a low solubility in chitosan slurries (chitosan dissolved in solvents) (Martins and others 2012). As a result, surfactant such as lecithin will be needed in this study. Lecithin supplies both hydrophilic and hydrophobic properties that can help to bond vitamin E and chitosan together to become a stable emulsion system (Mathew and others 2006). Several analyses were performed in this research. For example, the viscosities of the edible solutions, the chemical structure and thermal properties of the films were used to characterize the effect of vitamin E enrichment. The viscosities of the edible solutions revealed the stability and rheology of the blended ingredients. A minimum viscosity is required to ensure that insoluble compounds are uniformly suspended in the slurries (Rossman 2009). However, the viscosity of the edible solutions cannot excess the limit of mixing, pumping, filtering, spreading, and transferring during the film casting process. 114 Therefore, a knowledge for the viscosity of the slurries will determine the application and the manufacturing requirements of vitamin E incorporated chitosan films. ATR-FTIR combined with multivariate data analysis is a rapid test that could be used to differentiate the functional groups of the film samples. ATR-FTIR provided qualitative information of biochemical composition of compounds such as chitosan and vitamin E (Halim and others 2006; De Nardo and others 2009). Soft independent modeling of class analogy (SIMCA) from the multivariate data analysis has been used to classify the differences between experimental treatments and the intensity of ATR-FTIR reading. Partial least square regression (PLSR) is a cross-validate analysis that is used to find the fundamental relations (linear regression) between two variables (De Nardo and others 2009). It can be used to estimate the relationship of chemical and physical properties; for example, the degradation temperature and viscosity from the IR spectra (Bjorsvik and Martens 1992). Thermal analysis such as differential scanning calorimetry (DSC) and thermogravimetric analysis (TGA) are two useful tools that could be used to determine the thermal properties of the samples. DSC examines phase changes of a material by showing heat flow versus temperature plot (Höhne and others 2003). The DSC thermograms show exothermic and endothermic peaks (from -170°C to 600°C) of the ingredients, and it also provides quantitative capability (Höhne and others 2003). TGA measures the amount of weight changes in a sample during controlled heating. In this study, the weight changes of the films occurred by solvent evaporation, material degradation, and decomposition when the samples were heated. 115 The objective of this study was to investigate the effect of vitamin E fortified chitosan on the rheology of the slurries, functional groups of the chemical structure, and the thermal properties of the films. 4.3. Methods and Materials 4.3.1. Materials Medium molecular weight (94%) purity chitosan powder was provided by Huantai Goldenlake Carapace Products Co., Ltd (Tsingtao, China). This was used as the main ingredient in the films. Lactic acid (88% Food Chemical Codex, FCC) manufactured by Birko Corporation (Henderson, CO) and Giant Eagle brand distilled white vinegar containing 5% acetic acid (Pittsburg, PA) were used to dissolve the chitosan powder. One hundred percent natural vitamin E with mixed tocopherols (400 IU per softgel) was purchased from Whole Foods Market (365™ Everyday Value, Austin, TX). Glycerol USP Kosher 99.7% (Chemical direct, online, USA) and the soy lecithin (Solec® 100L, The Solae Company, St. Lousis, MO) were added as a plasticizer and an emulsifier, respectively. Differential scanning calorimeter (DSC) stainless steel pans, pan crimper/sealer, O-rings, and thermal gravimetric analysis (TGA) platinum pans were purchased from Perkin Elmer Instruments LLC (Shelton, CT). 4.3.2. Edible films formation Edible film solutions were prepared at 22 ± 1°C by dispersing the chitosan powder (1.0, 1.5 and 2% w/w) in distilled water. Lactic acid (1%, w/w) was added to the suspension and stirred (300 rpm) for 3 minutes to dissolve the chitosan powder. Glycerol 116 0.5% (w/w) was added to the solution and mixed for 3 minutes. It was used as a plasticizer to provide flexibility to the films. A 2.0% lecithin solution was then added to the chitosan mixture to produce a 0.1% (w/w) blend as the total dry weight of lecithin in the final films. Lecithin acted as an emulsifier that provided hydrophilic and hydrophobic interactions. The hydrophobic interactions were designed to help the film provide a good attachment to the food surfaces, especially to the high-fat products. The final slurry samples were completed by the addition of vitamin E purchased from Whole Food Market. Two levels of vitamin E (250 and 500 mg) were added in the chitosan slurries by two different steps: (1) vitamin E added before lecithin (VE first); and (2) vitamin E mixed with lecithin and then added to the chitosan solutions (VE mixed). The vitamin E fortified chitosan solutions were settled for one hour prior to further analyses (viscosity and drying rate measurement). Edible films were prepared by casting the edible slurry into 10 inch radius Teflon plates. Aliquots of 33.0± 0.5 ml, 31.0± 0.5 ml, and 29.0± 0.5 ml of 1.0%, 1.5%, and 2.0% of chitosan slurries were cast into the plates in order to obtain films of uniform thicknesses (50.0 ± 2.5 µm). This was oven dried at 45 ± 2 °C for 2 hours, then the dried films peeled off from the plate surfaces. The thickness was controlled by the percentage of solids in the solution. The thicknesses were measured using a 2804S-10 Agd Dialindicator Mitutoyo Guage (Aurora, IL) and reported in mil (1/1000 inch). 117 Scheme 4.1. The flow chart of chitosan-vitamin E slurry formation 1.0, 1.5, 2.0% Chitosan powder Dissolved in 1% (w/w) lactic acid solution Mixed for 3 min Plasticizer added (0.5% w/w glycerol) Mixed for 3 min Added 250/500 mg Vitamin E 250/500 mg Vitamin E blended with Mixed for 3 min 2.0 g (5% w/w) lecithin solution Added 2g (5% w/w) lecithin solution then added into the chitosan slurries Mixed for 3 min Final chitosan-vitamin E slurry 33± 0.5 ml 31± 0.5 ml 9.0± 0.5 ml 1.0% chitosan solution 1.5% chitosan solution 2.0% chitosan solution Cast into Teflon plates 118 4.3.4. Viscosity measurement Viscosity of the edible slurry was measured using a Brookfield DV-E Viscometer (Brookfield, MA) fitted with a LV3 spindle at 12 rpm. Each edible slurry sample (with a minimum volume of 200 ml) was tested at 20.0 ± 0.5°C for 5 minutes. The viscosity was recorded every 30 seconds, and the mean viscosity was the average of triplicates. 4.3.5. Functional groups characterization 4.3.5.1. Attenuated Total Reflection Fourier Transform Infrared spectroscopy (ATR-FTIR) analysis ATR-FTIR was used to characterize the functional groups of the films. A Varian 3100 Infrared Spectroscopy (Varian Inc., Palo Alto CA) with diamond subject at 5 Hz, 4 resolution and 64 scans was used to analysis the samples. All spectra were collected in the frequency range of 4000-700 cm-1 from four testing points. 4.3.5.2. Multivariate analysis The spectra from the ATR-FTIR analyses were evaluated using a Pirouette® Multivariate Analysis Software version 4.0, InfoMetrix, Inc., (Woodville, WA). Soft independent modeling of class analogy (SIMCA) was used to describe and classify the difference between the different edible slurries. The discriminating power was used to compare differences in wavenumbers versus intensity of each sample. Partial least square regression (PLSR) was obtained by converting the absorbance spectra to the secondary derivative spectra. The values of the PLSR showed the linear combination of the variables (Hassel and others 2011). 119 4.3.6. Thermal properties 4.3.6.1. Differential scanning calorimetric (DSC) analysis The thermal stability of the films was evaluated using a 2920 Modulated DSC with Universal Analysis software package v.3.9a (TA Instrument Corp. New Castle, Delaware). The film samples (11± 0.2 mg) were placed into stainless steel pans and sealed with O-rings. The samples were initially cooled from 25°C (at 20°C/ min cooling rate) to -20°C with 30 ml/ min nitrogen purging. After cooling, the samples were heated to 200°C and then cooled to -20°C at the same condition. Thermal parameters such as glass transition temperature (Tg), enthalpy (ΔH), maximum denaturation temperature (Tm), and onset of the crystallization temperature (To) of the film samples, corresponding to the endothermic peak areas on the DSC thermogram were determined by integrating the temperature vs. heat flow curve (Mathew and others 2006). Analysis of the thermograms were done by using the Universal Software that is a part of the equipment software. 4.3.6.2. Thermogravimetric analysis (TGA) This test was done using a Hi-Res Modulated TGA 2950 Thermogravimetric Analyzer with Universal Analysis software package v.3.9a (TA Instrument Corp. New Castle, Delaware). For this test, film samples (11 ± 0.5 mg) were placed in the platinum pans flushed with 90mm Hg of nitrogen, and scanned from 25 to 600ºC at a 30ºC/ min heating rate. Weight loss of the tested samples between 70-100°C was considered as the total moisture in the samples. 120 4.3.7. Statistical analysis This was conducted using a Minitab Software 15 Microsoft Version (Minitab Inc., State College, PA) for both one-way ANOVA with Post Hoc test- Tukey and Student’s t- test to determine the significant effect of chitosan, vitamin E concentrations and the blending processes on the properties of the films with a 95% confidence interval. ANOVA analysis provided the overall comparison between treatments, and the Post Hoc test-Tukey indicated significant differences (p< 0.05) between individual treatments. The statistical analysis was used to: (1) compare chitosan concentrations versus 2 levels of vitamin E; (3) evaluate chitosan, vitamin E and the blending process on their viscosity, temperature of phase change, and the weight loss in TGA. Student’s t-test was used to compare the viscosity, temperature of phase change, and the weight loss in TGA within two levels of vitamin E and two blending processes. Two replicates were used in each treatment. 121 4.4. Results and Discussion 4.4.1. Viscosity of vitamin E fortified edible slurries In this study, viscosity was used to determine changes in rheological properties of the slurry made with chitosan, vitamin E, and lecithin. Figure 4.1 shows the viscosities of 1.0-2.0% chitosan blended with 0 (control), 250 and 500 mg vitamin E additions. Two different blending processes were used in this experiment: (1) vitamin E added before lecithin (VE first); and (2) vitamin E mixed with lecithin (VE mixed) and then added to the slurry. Results showed that the viscosity increased when vitamin E was added into the system in all chitosan concentrations. Figure 4.1a shows the viscosity of the 1.0- 2.0% chitosan with 250 mg vitamin E addition. Viscosity of the 1.0% chitosan slurries significantly (p< 0.05) increased from 87± 12.3 (control) to 143± 4.9 cp (VE first) and 135± 5.2 cp (VE mixed). In the 1.5% chitosan, the viscosity significantly (p< 0.05) changed from 108± 9.5 cp (control) to 375± 5.2 (VE first) and to 491± 6.2 cp (VE mixed). In the 2.0% chitosan, the viscosity significantly (p< 0.05) increased from 827± 9.8 cp to 957± 11.6 (VE first), and 1200± 20 cp (VE mixed). Figure 4.1b shows the viscosity of the 1.0- 2.0% chitosan with 500 mg vitamin E addition. The viscosity significantly (p< 0.05) increased from 87± 12.3 (control) to 133± 4.9 cp (VE first) and to 142± 7.2 cp (VE mixed) in the 1.0% chitosan slurries. Also, the viscosity significantly (p< 0.05) increased from 108± 9.5 cp (control) to 373± 7.5 (VE first) and to 473± 4.9 cp (VE mixed) in the 1.5% chitosan; and 827± 9.8 cp to 957± 11.6 (VE first), and to 1200± 20 cp (VE mixed) in the 2.0% chitosan, respectively. 122 250mg Vit E 1400 control 1200 VE first 1000 VE mixed 800 600 iscosity (cp) V 400 200 0 1% 1.5% 2.0% Chitosan (a) 500mg Vit E 1200 control 1000 VE first VE mixed 800 600 iscosity (cp) V 400 200 0 1% 1.5% 2.0% Chitosan (b) Figure 4.1. The viscosity of chitosan slurries with (a) 250 mg; (b) 500 mg vitamin E addition. 123 Viscosity analysis is a rapid and simple method to monitor the thoroughness of the raw material mixing process. It represents the level of solids, degree of hydrolysis, and emulsification of the solutions, which can impact the quality of the edible films produced (Rossman 2009). The range of viscosity is an important parameter that influences the manufacturing process for edible films in the food industries. Higher viscosities tend to maintain the stability of the solution, keeping insoluble ingredients in suspension and preserving the homogenity of emulsions (Rossman 2009). However, the viscosity cannot be too high since it will affect the mixing, pumping, filtering, spreading, and transferring of the slurry to the casting line. The addition of vitamin E had a significant effect on increasing the viscosity of all samples. The amine groups from the chitosan molecule provide polycationic electrolytes when dissolved in lactic acid solution (Il’ina and Varlamov 2004; Park and Zhao 2004). These electrolytes are capable of binding with other hydrophilic compounds, as is the case with the hydroxyl group from vitamin E. Figure 4.2 explains how that the hydroxyl groups from vitamin E bonded with the amine or the hydroxyl groups from the chitosan (Figure 4.2). Vitamin E supplies a small amount of hydroxyl bonds, which provides sites for attachment of emulsifier such as lecithin that could be used to improve the incorporation of vitamin E into the chitosan. To compare the lecithin blending processes, the results showed that VE first had a lower viscosity than the VE mixed. Lecithin provided both hydrophobic and hydrophilic interactions that combined with vitamin E on its hydrophobic sites, and then bonded with chitosan by its hydrophilic sites. This allowed a large amount of vitamin E-lecithin mixture to be incorporated into the chitosan polymeric structure and be entrapped into the 124 network. As an explanation for this result, lecithin had a lower probability of bonding to both vitamin E and chitosan when the vitamin E was added before the lecithin. This is so because of the electrolyte properties. Lecithin introduced into the slurry after adding the vitamin E had a lower chance of binding with the chitosan polymers but instead is more prone to bind with vitamin E. In this case, the chitosan/vitamin E/lecithin network system was not as coordinate as the VE mixed-chitosan. !"# Scheme 4.2. The possible molecular binding sites between vitamin E and chitosan. 125 4.4.2. Attenuated Total Reflection Fourier Transform Infrared spectroscopy (ATR- FTIR) characterizations Soft independent modeling of class analogy (SIMCA) from the multivariate data analysis was used to classify the differences between vitamin E and chitosan concentrations by providing a discriminating power and the intensity of the ATR-FTIR reading (Koca and others 2007; Hassel and Rodriguez-Saona 2011). Figure 4.3a shows the SIMCA analysis of 1.0% chitosan films with or without vitamin E addition. Two levels of vitamin E (250 and 500 mg) were used with two blending processes (VE first, and VE mixed), and the discriminating power is shown in Figure 4.4b. Figure 4.4 and Figure 4.5 show the 1.5 and 2.0% chitosan blended with two levels (250 mg and 500 mg) of vitamin E by the VE first and the VE mixed processes. The SIMCA analysis clustered ATR-FTIR data into five groups, and these are demonstrated in Figure 4.3a. The interclass distance of the 1.0% chitosan samples was greater than 3.0, indicating that additional vitamin E versus the blending processes had significant (p< 0.05) influence the intensity on the presence of the functional groups in the ATR-FTIR spectra. The discriminating power generated from the ATR-FTIR spectral data were used to interpret the difference between chitosan and vitamin E treatments. Wavenumbers 874 cm-1 and 1755 cm-1 had the greatest intensity (Figure 4.3b), and this is correspond to the presence of phenyl rings and C=O stretching, respectively. The presence of phenyl ring at 874 cm-1 was related to vitamin E since it was the only ingredient that contained this functional group. However, the 870-890 cm-1 regions were only found in the 1.0% chitosan discriminating spectra (Figure 4.3b). As the chitosan concentrations increased, functional groups such as CH, CH2, CH3, amine 126 and hydroxyl groups also increased. It is believe that the ATR-FTIR responses were higher in intensity to polar bonds were less sensitive to the non-polar bonds in vitamin E. Hassel and Rodriguez-Saona (2011) also indicated that ATR-FTIR analysis is not sutable for differentiating vitamin E when it is in a complicated food matrix. The interaction between chitosan, vitamin E and the blending process became more complicated in the 1.5% chitosan slurry. Compared to the control, the 250 mg VE mixed did not show significant difference (interclass distance< 3) in the SIMCA analysis (Figure 4.4a). The most significant peak in the discriminating power was at 1367 cm-1 (CH bending), which may have been contributed by the polysaccharide, vitamin E, and lecithin (Figure 4.4b). In the 2.0% chitosan samples, five clusters had been differentiated from the SIMCA analysis with interclass distance> 3 (Figure 4.5a). The discriminating -1 power showed characteristic bands associated with wavenumber 1153cm (CH2, CH3 -1 -1 -1 zigzag), 1373 cm (CH3 bonds), 1435 cm (CH2, CH3 bend), 1740 cm (CH stretch) and 2900-3200 cm-1 (CH). Gonçalves (2010) and Marines and Vincente (2010) reported that 2900-3200 cm-1 bands are associated with vitamin E and its symmetric and asymmetric stretching of the CH2 and CH3 functional groups. 127 250 mg VE mixed PC2 250 mg VE first control PC1 PC3 500mg VE mixed 500 mg VE first (a) •! SIMCA analysis • ! C1%on chitosantrol: 1.0% with ch i1%tosa lacticn film acid,s wit h0.5%out v glycerolitamin E and 0.1% lecithin VE first: vitamin E added before lecithin VE mixed: vitamin E mixed with lecithin then added to the chitosan slurries (b) Figure 4.3. FTIR analysis of 1.0% chitosan vitamin E fortified films (a) SIMCA analysis; (b) discriminating power. 128 The cross-validated regression (PLSR) based on ATR-FTIR spectral information showed good correlation (r> 0.98) between functional groups and the viscosity of the slurries (Figure 4.6a). Examination of the PLSR loading spectra (Figure 4.6b) indicated that the 900- 1800 cm-1 region was associated with viscosity changes. CH and OH bonds of the films were demonstrated at the 900-920 and 1215 cm-1 regions as reported by Tanka and others (2001). The frequencies at 1124 cm-1, 1373 cm-1, 1460-1490 cm-1, and -1 1740-1770 cm are attributed to the C-OH stretch, CH2, CH3 zigzag, C-H bending, and C=O, respectively. This conclusion is supported from research published by De Nardo and others (2009). Marines and Vicente (2010) also stated that the methyl (CH3) symmetric bending occurs in a broad band between 1215- 1500 cm-1. These functional groups described above may be attributed to chitosan, the solvents, vitamin E, and lecithin. Since ATR-FTIR only indicated the hydrophilic bonds within the samples; therefore, the relationship between vitamin E and the viscosity may not have been fully discovered by these IR spectra. To summarize, the presence of vitamin E in the chitosan film had been successfully indicated by ATR-FTIR at the 827 and 2900-3200 cm-1bands. Hydrophilic bonds such as CH, OH, and C=O significantly affected the viscosity of the edible slurries. However, ATR-FTIR spectra showed no direct evidence that additional vitamin E (above a given threshold) influenced the viscosity of the chitosan slurries. 129 PC2 250 mg VE mixed 500 mg VE mixed 250 mg VE first 500 mg VE first PC1 PC3 control (a) Contr•o! l: SIMCA1.5% ch ianalysistosan fi lms without vitamin E •! 1.5% chitosan with 1% lactic acid, 0.5% glycerol and 0.1% lecithin VE first: vitamin E added before lecithin VE mixed: vitamin E mixed with lecithin then added to the chitosan slurries (b) Figure 4.4. FTIR analysis of 1.5% chitosan vitamin E fortified films (a) SIMCA analysis; (b) discriminating power. 130 PC2 250 mg VE first 500 mg VE first 500 mg VE mixed PC1 PC3 250 mg VE mixed control (a) Control: 2.0% chitosan films without vitamin E •! SIMCA analysis V•E! fi2.0%rst: v chitosanitamin Ewith ad d1%ed lactic befo racid,e le c0.5%ithin glycerol and 0.1% lecithin VE mixed: vitamin E mixed with lecithin then added to the chitosan slurries (b) Figure 4.5. FTIR analysis of 2.0% chitosan vitamin E fortified films (a) SIMCA analysis; (b) discriminating power. 131 (a) (b) Figure 4.6. PLSR analysis of chitosan vitamin E fortified films versus slurry viscosity (a) linear relationship; and (b) loading plot. 132 4.4.3. Thermal analysis of the vitamin E fortified edible films 4.4.3.1. Differential scanning calorimetric (DSC) analysis The differential scanning calorimetric (DSC) curves in Figure 4.7 displayed the thermally-induced endothermic transitions of vitamin E fortified chitosan edible films from 30°C to 100°C. The major endothermic peaks in the DSC curves of the film sample were observed at 60- 75°C (Figure 4.7). Figure 4.7a represents the DSC thermograms of chitosan films (1.0- 2.0%) without vitamin E additions. Figure 4.7 b (1.0% chitosan), Figure 4.7c (1.5% chitosan), and Figure 4.7d (2.0% chitosan) show the chitosan films with vitamin E addition. Results indicated that similar endothermic curves were obtained for all chitosan concentrations. A distinct endothermic peak at 64.8± 0.5 to 73.3± 0.9°C was observed in all samples. However, no significant (p> 0.05) differences in the temperatures were found between all the samples. These endothermic peaks (at 64.8± 0.5 to 73.3± 0.9°C) relate to water evaporation associated with hydrophilic groups from the films (Dhanikula and Panchangnula 2004; El-Hefian and others (2010). The intensity of the peaks decreased as the concentration of chitosan increased, especially in the high vitamin E concentration samples. Martins and others (2012) reported that hydrophobic bonds provided by the vitamin E tend to push the hydrophilic solvents out from the film during drying. In this case, films made with vitamin E would have less water content. No glass transition temperature (Tg) could be identified in the DSC thermograms for all vitamin E incorporated chitosan films. Don and others (2005) reported that it was hard to determine the Tg of chitosan films due to its rigid structure and tight inter- and intra-hydrogen bonding associated with the chitosan chemical structure. On the other 133 hand, Gonçalves and others (2011) illustrated that small amounts of vitamin E addition did not change the Tg in vitamin E enriched polylactic acid (PLA). However, more research is needed to understand the interaction between vitamin E, chitosan, and solvents on phase changes during DSC analyses. 134 (#. 333333333333333333"#(B3CDE26F1GHHHHHHH 333333333333333333"#$B3CDE26F1GHHH3HHH 333333333333333333.#(B3CDE26F1GHHHHH3I (#( !"#$%&' ,(#. !"#+"&' /01234567389:;< ,(#* !(#)*&' ,(#- )( %( JK63L? =0>?0@12A@038&'< LGEM0@F153N*#!O3=O3PGF2@A>0G2F (a) .#) 333333333333333333'6B2@65CCCCCCC 333333333333333333),.3>;3DE@F2CCC3CCC 333333333333333333),.3>;3>EG0HCCCCC3I 333333333333333333,..3>;3DE@F2CCC3C3C 333333333333333333,..3>;3>EG0HC3C3C3C .#. !"#$%&' -.#) !"#$!&' /01234567389:;< !(#")&' -.#$ !(#*+&' !"#*,&' -.#! +. $. ,. !. *. ". (. %.. JG63K? =0>?0@12A@038&'< KBEL0@F153M$#*N3=N3OBF2@A>0B2F (b) Figure 4.7. DSC thermograms of (a) 1.0- 2.0% chitosan films (control); and (b) 1.0% cihtosan with 250 mg and 500 mg vitamin E addition. *Control: films without vitamin E addition *First: vitamin E added before lecithin *Mixed: vitamin E mixed with lecithin 135 )#' 111111111111111111&4B0@43CCCCCCC 111111111111111111'*)1>91DE@F0CCC1CCC 111111111111111111'*)1>91>EG.HCCCCC1I 111111111111111111*))1>91DE@F0CCC1C1C 111111111111111111*))1>91>EG.HC1C1C1C )#) !"#$"%& ,)#' !'#()%& -./01234516789: ($#*$%& ,)#+ (+#!!%& !'#)(%& ,)#( ;) +) *) () !) <) $) ")) JG41K? =.>?.@/[email protected]%&: KBEL.@F/31M+#!N1=N1OBF0@A>.B0F (c) +#- 333333333333333333'6B2@65CCCCCCC 333333333333333333.$+3>;3DE@F2C3C3C3C 333333333333333333.$+3>;3>EG0HCCCCC3I 333333333333333333$++3>;3DE@F2CCC3C3C 333333333333333333$++3>;3>EG0HCCC3CCC +#. !"#$%&' +#+ ()#$)&' ,+#. /01234567389:;< (*#+)&' ()#%%&' ,+#- (+#"!&' ,+#! *+ -+ $+ !+ (+ %+ "+ )++ JG63K? =0>?0@12A@038&'< KBEL0@F153M-#(N3=N3OBF2@A>0B2F (d) Figure 4.7. DSC thermograms of (c) 1.5%; and (d) 2.0% cihtosan with 250 mg and 500 mg vitamin E addition. *Control: films without vitamin E addition *First: vitamin E added before lecithin *Mixed: vitamin E mixed with lecithin 136 4.4.3.2. Thermogravimetric analysis (TGA) Thermogravimetric analysis (TGA) is used to understand the correlation between temperature and a sample’s weight changes. In Figure 4.8, the smooth decreasing lines indicate the percent weight changes percent (%; Y-1) and sharp peaks indicate the derivative of the weight percentage per degree (%/°C; Y-2) versus temperature (X). Similar weight change curves were obtained for all samples. Figure 4.8a shows the 1.0- 2.0% chitosan films without vitamin E addition. The films with vitamin E addition are shown in Figure 4.8b (1.0% chitosan), Figure 4.8c (1.5% chitosan), and Figure 4.8d (2.0% chitosan). Three major peaks were located at 65-75°C, 140-210°C, and 270-430°C in all samples. A weight loss at 430-450°C became visible when the chitosan and vitamin E concentrations increased. Significant (p< 0.05) differences were found in the weight changes at 430-450°C when comparing the 1.5% and the 2.0% chitosan with 500 mg VE first and 500 mg VE mixed with the controls. The first peak from the left (65-75°C) was caused by water evaporation. Martins and others (2012) supported this assumption in the study they reported. The second peak from the left (140- 210°C) corresponded to chitosan decomposition as reported by El- Hefian and others (2010). The weight change at 140- 210°C (Figure 4.8a) was significantly (p< 0.05) higher in the 1.0% chitosan due to the instability of chitosan networks. This occurred because chitosan may have hydrolyzed in the solvents and then formed a network with its amine and hydrogen bonds. Increasing the chitosan concentrations increased the strength of the networks as supported in the report from El- Hefian and others (2010). Also, Martins and others (2012) illustrated that weight loss at the 200°C related to the chemical absorption of water by hydrogen bonds and the 137 elimination reaction of NH3. Increasing chitosan concentration, the chem-sorption of water decreased because of a lowering of the amorphous regions within the polymeric matrix. Moreover, the strength of the polycationic interactions increased with increasing chitosan concentrations and this had stronger solid-solid interactions instead of solid- water interactions (Martins and others 2012). The highest weight loss occurred at 270- 320°C (the third peak), which was the thermal degradation temperature of chitosan (Don and others 2005). On the contrary, the 2.0% chitosan showed 5.0% more weight loss when compared with the 1.0% chitosan at the thermal degradation temperature. This may have been caused by the larger presence of chitosan molecules in the 2.0% chitosan films. 138 #$! !"% ++++++++++++++++++#"!1+4?+4@A/(@BCCCCCCC ++++++++++++++++++#"91+4?+4@A/(@BC+C+C+C ++++++++++++++++++$"!1+4?+4@A/(@BCCC+CCC #!! !"$ 8! ,')-./+015 &'()*"+,')-./+012345 !"# 7! 6! !"! ! #!! $!! %!! 6!! 9!! 7!! :';<'(=/>('+0345 DA)*'(E=B+F6"GH+:H+IAE/(>;'A/E (a) (#$ //////////////////'@A3,@BCCCCCCC //////////////////):(/<1/D-,E3CCC/CCC //////////////////):(/<1/<-F+GCC/CC/C //////////////////:((/<1/D-,E3CCC/C/C %(( //////////////////:((/<1/<-F+GC/C/C/C (#) 9( 0+-123/457 *+,-.#/0+-123/456&'7 (#% 8( !!"#$%&' !( (#( ( %(( )(( $(( !(( :(( 8(( ;+<=+,>3?,+/4&'7 HA-.+,E>B/I!#JK/;K/LAE3,?<+A3E (b) Figure 4.8. TGA thermograms of (a) 1.0- 2.0% chitosan films (control); and (b) 1.0% cihtosan with 250 mg and 500 mg vitamin E addition *Control: films without vitamin E addition *First: vitamin E added before lecithin *Mixed: vitamin E mixed with lecithin 139 +#( 111111111111111111&?@5.?ABBBBBBB 111111111111111111$)+1;31C/.D5BBB1BBB 111111111111111111$)+1;31;/E-FBBBBB1G 111111111111111111)++1;31C/.D5BBB1B1B "++ 111111111111111111)++1;31;/E-FB1B1B1B +#$ '+ !!"#$$%& 2-/3451679 ,-./0#12-/3451678%&9 +#" *+ !($#)'%& !!'#*!%& !!'#"(%& !+ +#+ + "++ $++ (++ !++ )++ *++ :-;<-.=5>.-16%&9 H@/0-.D=A1I!#JK1:K1L@D5.>;-@5D (c) $#( 000000000000000000'?@4-?ABBBBBBB 000000000000000000*9$0;20C.-D4BBB0BBB 000000000000000000*9$0;20;.E,FBBBBB0G 0000000000000000009$$0;20C.-D4BBB0B0B )$$ 0000000000000000009$$0;20;.E,FB0B0B0B $#* "$ !!)#!*&' 1,.2340568 !!"#("&' +,-./#01,.2340567&'8 $#) %$ !!"#$%&' !!(#!%&' !$ $#$ $ )$$ *$$ ($$ !$$ 9$$ %$$ :,;<,-=4>-,05&'8 H@./,-D=A0I!#JK0:K0L@D4->;,@4D (d) Figure 4.8. TGA thermograms of (c) 1.5%; and (d) 2.0% cihtosan with 250 mg and 500 mg vitamin E addition. *Control: films without vitamin E addition *First: vitamin E added before lecithin *Mixed: vitamin E mixed with lecithin 140 A small peak at 430-450°C was seen in the 1.0% chitosan with 500 mg VE first (Figure 4.8b). This was associated with the decomposition of the phenyl ring from vitamin E. This assumption is supported in reports published by Martins and others (2012). More than 2.88% weight loss were demonstrated at 430-450°C in 1.5 chitosan with 250 mg VE first; and 2.0% chitosan with 500 mg VE mixed (Figure 4.9). Both treatments had the highest overall vitamin E concentration within the films (Table 3.2). The TGA thermograms showed that the 1.5% chitosan with 250 mg VE first and the 2.0% chitosan with 500 mg VE mixed had the sharpest derivative weight percentage per degree curves (%/°C) within the groups. This indicated that the incorporation of vitamin E within the chitosan polymeric structure was well uniformed. Also, incorporated vitamin E tightly bonded with chitosan, which made it difficult to be removed when the energy of the system was too low (low temperature). This result also related to the HPLC analysis in Table 3.2, where more vitamin E incorporated within the films showed sharper and more intense peaks in the TGA thermograms. Contrarily, higher concentrations of vitamin E on the film surface showed a wider decomposition temperature range in the TGA thermograms. As could be seen, the 1.5% and 2.0% chitosan 500 mg VE first demonstrated the broadest derivative weight percentage per degree curves (%/°C). The vitamin E on the film surface (unbonded with chitosan polymers) was relatively unstable when compared with the incorporated vitamin E within the chitosan films. Therefore, less energy was needed to evaporate those unbonded vitamin E from the system. For example, vitamin E decomposition started at 375°C in the 2.0% chitosan 500 mg VE first (Figure 4.9a), which was 75°C less than in the 2.0% chitosan 500 mg VE mixed. 141 (#"+ (#"+ 222222222222222222@AB6/ACDDDDDDD !!"#$$%& 222222222222222222$+(2<42E0/F6D2D2D2D 222222222222222222$+(2<42<0G.HDDDDD2I 250 mg first 222222222222222222+((2<42E0/F6DDD2D2D 222222222222222222+((2<42<0G.HDDD2DDD (#"( (#"( !*$#+)%& !!)#,!%& 500 mg first 500 mg mixed -./01#23.04562789%&: !!'#()%& (#(+ 250 mg mixed (#(+ (#(( (#(( *+( !(( !+( +(( ++( ;.<=./>6?/.27%&: JB01./F>C2K!#LM2;M2NBF6/?<.B6F (a) $#)+ $#)+ 111111111111111111'?@5.?ABBBBBBB 111111111111111111*+$1;31C/.D5BBB1BBB !!)#!*&' 111111111111111111*+$1;31;/E-FBBBBB1G 111111111111111111+$$1;31C/.D5BBB1B1B 500 mg mixed 111111111111111111+$$1;31;/E-FB1B1B1B !!"#("&' 500 mg first $#)$ $#)$ !!"#$%&' 250 mg first ,-./0#12-/3451678&'9 $#$+ !!(#!%&' $#$+ 250 mg mixed $#$$ $#$$ (+$ !$$ !+$ +$$ ++$ :-;<-.=5>.-16&'9 H@/0-.D=A1I!#JK1:K1L@D5.>;-@5D (b) Figure 4.9. TGA thermograms at 350-550°C of (a) 1.5%; and (b) 2.0% cihtosan with 250 mg and 500 mg vitamin E addition. *Control: films without vitamin E addition *First: vitamin E added before lecithin *Mixed: vitamin E mixed with lecithin 142 4.5. Conclusion The amount of vitamin E and the types of blending process significantly affected the viscosity of the chitosan edible slurries. Vitamin E-lecithin mixture had the greatest impact on improving its incorporation into the chitosan slurries. ATR-FTIR identified the functional groups related to vitamin E at 827 and 2900-3200 cm-1. PLSR generated from the ATR-FTIR showed a linear correlation between the functional groups and the viscosity (r>0.98). The DSC thermograms did not change after vitamin E was introduced into the system. However, vitamin E fortified chitosan films demonstrated a significant weight loss at 430-450°C in the TGA analysis. This was associated with vitamin E decomposition. This present work suggested that the 2.0% chitosan blended with the 500 mg VE mixed had the most efficient incorporated activity. 4.6. References Bjorsvik HR, Martens H. Data analysis: calibration of NIR instruments by PLS regression. 1992. In handbook of near-infrared analysis. Burns D, Ciurczak E. Dekker. New York, NY. P 159-180. Darmadji P, Izumimoto M. 1994. Effect of chitosan in meat preservation. Meat Sci. 38: 243–254. De Nardo, T., Shiroma-Kian, C., Halim, Y., Francis, D., Rodriguez-Saona, L. 2009. Rapid and simultaneous determination of lycopene and b-carotene contents in tomato juice by infrared spectroscopy. J Agric Food Chem. 57:1105–1112. Dhanikula AB, Panchangnula R. 2004. Development and characterization of biodegradable chitosan films for local delivery of paclitaxel. The AAPS J 6:(3) 1-12. 143 Don T-M, King C-F, Chiu W-Y, Peng C-A. 2005. Preparation and characterization of chitosan-g-poly (vinyl alcohol)/ poly (vinyl alcohol) blends used for the evaluation of blood-cotacting compatibility. Carbohyd Polym. 63: 331-339. El-Hefian EA, Elgannoudi ES, Mainal A. 2010. Characterization of chitosan in acetic acid: Rheological and thermal studies. Turk J Chem. 24: 47-56. Gonçalves CMB, Tomé LC, Coutinho JAP. Marrucho IM. 2011. Addition of α- tocopherol on poly(lactic acid): thermal, mechanical, and sorption properties. J Appl Polym Sci. 119: 2468-2475. Halim Y, Schwartz S, Francis D, Baldauf N, Rodriguez-Saona L. 2006. Direct determination of lycopene content in tomatoes (Lycopersicon esculentum) by attenuated total reflectance infrared spectroscopy and multivariate analysis. J AOAC Int. 89: 1257-1262. Hassel S, Rodriguez-Saona LE. 2011. Application of a handheld infrared sensor for monitoring the mineral fortification in whole grain cornmneal. Food Anal Methods. 5: 571-578. Il’ina AV. Varlamov VP. 2004. Hydrolysis of chitosan in lactic acid. Appl Biochem Microbio+. 40: 354-358. Janjarasskul T and Krochta JM. 2010. Edible Packaging Materials. Annu. Rev. Food Sci. Technol. 1: 415-448. Jayakumar R, Nwe NT, Tokura S, Tamura H. 2007. Sulfated chitin and chitosan as novel biomaterials. Int J Bio Macromolecules. 40: 175–181. Jayakumar R, Reis RL, Mano JF. 2006. Chemistry and Applications of Phosphorylated Chitin and Chitosan. e-Polymers 035. 144 Jongrittiporn S, Kungsuwan A, Rakshit SK. 2001. A study on the preservation of fishballs using chitosan. In European Conference on Advanced Technology of Safe and High Quality Foods-EUROCAFT, 5–7 December 2001, Berlin. Koca N, Rodriguez-Saona L, Harper W, Alvarez V. 2007. Application of fourier transform infrared spectroscopy for monitoring short-chain fatty acids in Swiss cheese. J Dairy Sci. 90: 2596-2603. Martins JT and Vicente AA. Incorporation of α-tocopherol into chitosan films: effects on film properties. Semana de Engenharia 2010. Mathew S, Brahmakumar M, Abraham TE. 2006. Microstructureal imaging and characterization of the mechanical, chemical, thermal, and swelling properties of starch-chitosan blend films. Biopolymers. 82: 176-187. Park S-I, Zhao Y. 2004. Incorporation of high concentration of mineral or vitamin into chitosan-based films. J Agric Food Chem. 52: 1933-1939. Rossman JM. 2009. Edible films and coatings for food applications. In: Embuscado ME, Huber KC. Edible films and coatings for food applications. Springer Science Business Media, LLC, New York, NY. p367-390. Tanaka T, Nagao S, Ogawa H. 2001. Attenuated total reflection fourier transform infrared (ATR-FTIR) spectroscopy of functional groups of humic acid dissolving in aqueous solution. Anal Sci 1081-1084. Wang GH. 1992. Inhibition and inactivation of five species of foodborne pathogens by chitosan. J Food Protect. 55(11), 916–919. 145 Wessling C, Nielsen T, Leufven A. 2000. The influence of α-tocopherol concentration on the stability of ilnoleic acid and the properties of low-density polyethylene. Packag Technol Sci. 13: 19-28. 146 CHAPTER 5 CONCLUSION This study demonstrated that chitosan films made with acetic acid-ethanol and vitamin E could be used for selected food processing applications. Chitosan dissolved in the selected solvents and at an optimal concentration for film formation had promising potential for commercial-scale production as an edible film. This study also demonstrated that chitosan can be used as a polymeric matrix for incorporation of micronutrients to enhance the functional properties of edible films. Such films could be used for food wraps, surface coating, microwavable food covers, and packaging of ready- to-eat products. The properties of the chitosan slurries were affected by the solvents and the micronutrient interactions by bonding with the amine groups and the hydrogen bonds on the chitosan molecule. These interactions directly affected the solubility of the chitosan films. In addition, hydrophilic and hydrophobic interactions within the polymer influenced the viscosity and the drying rate of the edible slurries. ATR-FTIR with multivariate data analysis was used to determine the biochemical compositions and the presence of functional groups in the film formulations. The micronutrient incorporation was achieved through adequate used of the surfactant and the appropriate processing 147 conditions. However, addition of the micronutrient and the surfactant significantly effected the films properties. Further studies are needed to understand the solvent residuals and micronutrient interactions within the chitosan polymeric matrix and their effect on the film properties. In addition, sensory analysis should be included to evaluate potential applications of these developed edible films. Additional research developments should focus on improvements to the technology transformation from a lab-scale to a commercial-scale process using continuous-casting equipments. 148 LIST OF REFERENCES André Guinier X-ray diffraction in crystals, imperfect crystals, and amorphous bodies. 1994. Dover Publications, Inc, NY p.3 Badawy MEI and Rebea EI. 2009. Potential of the biopolymer chitosan with different molecular weights to control postharvest gray mold of tomato fruit. Postharvest Bio Tech. 51: 110-117. Bajpai SK, Chand N, Chaurasia V. 2009. Investigation of water vaporpermeability and antimicrobial property of zinc oxide nanoparticles-loaded chitosan-based edible film. J of Appl Polym Sci. 115: 674-683. Balasubramaniam VM, Chinnan MS, Mallikarjunan P, Philips RD. 2007. The effect of edible film oil uptaking and moisture retention of deep-fried poultry product. J Food Process Eng. 20: 17-20. Baldwin EA. 2007. Surface treatments and edible coatings in food preservation. In Handbook of Food Preservation, ed. MS Rahman, 21:478–508. Boca Raton, FL: CRC Press. Baldwin EA, Nisperos MO, Baker RA. 1995. Edible coatings for lightly processed fruits and vegetables. Hortscience. 30(1):35–38. Baldwin EA, Nisperos MO, Chen X, Hagenmaier RD. 1996. Improving storage life of cut apple and potato with edible coating. Postharvest Biol. Technol. 9:151–63. 149 Barone JR, Schmidt WF. 2006. Nonfood application of proteinaceous renewable materials. J Chem Education. 83: 1003-1009. Beverly RL, Janes ME, Prinyawiwatkula W, No HK. 2007. Edible chitosan films on ready-to-eat roast beef for the control of Listeria monocytogenes. Food Microbiol. 25: 534-537. Bjorsvik HR, Martens H. 1992. Data analysis: calibration of NIR instruments by PLS regression. In handbook of near-infrared analysis. Burns D, Ciurczak E. Dekker. New York, NY. P 159-180. Bravin B, Perssini D, Sensidoni A. 2006. Development and application of polysaccharide-lipid edible coating to extend shelf-life of dry bakery products. J Food Eng. 76: 280-290. Chai Z, Shang J, Jiang Y, Ren F, Leng X. 2010. Effects of the free and pre-encapsulated calcium ions on the physical properties of whey protein edible film. Inter J of Food Sci Technol. 45: 1532-1538. Chen CS, Liaun WY, Tsai GJ. 1998. Antimicrobial effects of N-sulfonated and N- sulfobenzoyl chitosan and application to oyster preservation. J Food Protec. 61: 1124- 1128. Chien PJ, Sheu F, Yang FH. 2007. Effects of edible chitosan coating on quality and shelf life of sliced mango fruit. J Food Eng. 78: 225–229. Chillo S, Mastromatteo M, Conte A, Gerschenson L, Del Nobile MA. 2008. Influence of glycerol and chitosan on tapioca starch-based edible film properties. J Food Eng. 88: 159-168. 150 Cho SY, Park JW, Batt HP, Thomas RL. 2007. Edible film made from membrane processed soy protein concentrates. LWT Food Sci Technol. 40: 418-423. Coma V. 2008. A review: Bioactive packaging technologies for extended shelf life of meat-based products. Meat Science. 78(1) 90–103. Coma V, Martial-Gros A, Garreau S, Copinet A, Salin F, Deschamps A. 2002. Edible antimicrobial film based on chitosan matrix. J Food Sci 67: 1162-1169. Chrissafis K, Paraskevopoulos KM, Papageorgiou GZ, Bikiaris DN. 2008. Thermal and dynamic mechanical behavior of bionanocomposites: found silica nanoparticles dispersed in poly(vinyl pyrrolidone), chitosan, and poly(vinyl alcohol). J Appl Polym Sci. 110: 1739-1749. Coggins C, Blanchard K, Alvarez F, Brache V, Weisberg E. Kilmarx P, Lacarra M, Massai R, Mishell D, Salvatierra A, Witwatwongwana P, Elias C, Ellertson C.2000. Preliminary safety and acceptability of a carrageenan gel for possible use as a vaginal microbicide. Sex Transm Infect. 76: 480–483. Cuero RG, Osuji G, Washington A. 1991. N-carboxymethyl chitosan inhibition of aflatoxin production: role of zinc. Biotechnol Lett 13(6): 441-444. Cunningham P, Ogale AA, Dawson PL, Acton JC. 2000. Tensile properties of soy protein isolate films produced by a thermal compaction technique. J Food Sci. 65(4): 668– 671. Damonte EB, Matulewicz MC, Cerezo AS. 2004. Sulfated seaweed polysaccharides as antiviral ageants. Curr Med Chem. 11: 2399-1419. Darmadji P, Izumimoto M. 1994. Effect of chitosan in meat preservation. Meat Sci. 38: 243–254. 151 Debeaufort F, Voilley A. 2009. Lipid-based edible films and coatings. Embuscado ME Huber KC eds. Edible Films and Coatings for Food. Springer Science Business Media, LLC, New York, NY. De Moura MR, Aouada FA, Avena-Bustillos RJ. McHugh TH. Krochta JM. Mattoso LHC. 2009. Improved barrier and mechanical properties of novel hydroxypropyl methylcellulose edible films with chitosan/tripolyphosphate nanoparticles. J Food Eng. 92(4): 448–453. De Nardo, T., Shiroma-Kian, C., Halim, Y., Francis, D., Rodriguez-Saona, L. 2009. Rapid and simultaneous determination of lycopene and b-carotene contents in tomato juice by infrared spectroscopy. J Agric Food Chem. 57:1105–1112. Dhanikula AB, Panchangnula R. 2004. Development and characterization of biodegradable chitosan films for local delivery of paclitaxel. The AAPS J 6:(3) 1-12. Don T-M, King C-F, Chiu W-Y, Peng C-A. 2005. Preparation and characterization of chitosan-g-poly (vinyl alcohol)/ poly (vinyl alcohol) blends used for the evaluation of blood-cotacting compatibility. Carbohydr Polym. 63: 331-339. Dragich AM. Krochta JM. 2010. Whey protein solution coating for fat-update reduction in deep fried chicken breast strips. J Food Sci. 75(1): 43-47. Du W-X, Olsen CW, Avena-Bustillos RJ, McHugh TH, Levin CE, Friedman M. 2008. Storage stability and antibacterial activity against Escherichia coli O157:H7 of carvacrol in edible apple films made by two different casting methods. J Agric Food Chem. 56: 3082-3088. El-Hefian EA, Elgannoudi ES, Mainal A. 2010. Characterization of chitosan in acetic acid: Rheological and thermal studies. Turk J Chem. 24: 47-56. 152 Garcia D, Guo R, Bhalla AS. 2000. Growth and properties of BA0.9Sr0.1TiO3 single crystal fibers. Mater Lett. 42: 136-141. Gaontard N, Guilbert S. 1994. Bio-packaging: Technology and properties of edible and/or biodegradable material of agricultural origin in food packaging and preservation. ed., Mathlouthim, London: Blackie Academic & Professional. p159-181. Gasparoux J, Laux D, Ferrandis JY, Attal J, Tordjeman P. 2008. Large frequency bands viscoelastic properties of honey. J Non-Newtonian Fluid Mechanics, 153:46-52. Gennadios, A. and C.L. Weller, 1990. Edible films and coatings from wheat and corn proteins. Food Technol. 44:63-69. Gonçalves CMB, Tomé LC, Coutinho JAP. Marrucho IM. 2011. Addition of α- tocopherol on poly(lactic acid): thermal, mechanical, and sorption properties. J Appl Polymer Sci. 119: 2468-2475. González ME, Alarcón B, Carrasco L. 1987. Polysaccharides as antiviral agents: Antiviral activity of carrageenan. Antimicrob Agents Chemother. 31: 1388-1393. Gómez-Estaca J, López de Lacey A, López-Caballero ME, Gómez-Guillén MC, Montero P. 2010. Biodegradable gelatin–chitosan films incorporated with essential oils as antimicrobial agents for fish preservation. Food Microbiol. 27: 889-896. GSM industries. 2012. http://www.dsm.com/en_US/html/dep/barrier_film_extr_coating.htm Guinier A. 1994. X-ray diffraction in crystals, imperfect, crystals, and amorphous bodies. W.H. Freeman & Company, San Francisco, CA. Halim Y, Schwartz S, Francis D, Baldauf N, Rodriguez-Saona L. 2006. Direct determination of lycopene content in tomatoes (Lycopersicon esculentum) by 153 attenuated total reflectance infrared spectroscopy and multivariate analysis. J AOAC Int. 89: 1257-1262. Hassel S, Rodriguez-Saona LE. 2011. Application of a handheld infrared sensor for monitoring the mineral fortification in whole grain cornmneal. Food Anal Methods. 5: 571-578. Han C, Lederer C, McDaniel M, Zhao Y. 2005. Sensory evaluation of fresh strawberries (Fragaria ananassa) coated with chitosan-based edible coatings. J Food Sci. 70: 173- 178. Han C, Zhao Y, Leonard SW, Traber MG. 2004. Edible coating to improve storability and enhance nutritional value of fresh and frozen strawberries (Fragaria × ananassa) and raspberries (Rubus ideaus). Postharvest Bio and Tech. 33:67-84. Han JH, Hwang HM, Min S, Krochta JM. 2008. Coating of peanuts with edible whey protein film containing alpha-tocopherol and ascorbyl palmitate. J Food Sci. 73: 349- 355. Hauck BW, Huber GR. 1989. Single screw vs twin screw extrusion. Cereal Food World. 24: 930-939 Hernandez-Izquierdo VM, Krochta JM. 2008. Thermoplastic processing of proteins for film formation-A review. J Food Sci. 73: 30-39. Hernandez-Izquierdo VM. 2007. Thermal transitions, extrusion, and heat-sealing of whey protein edible films [dissertation]. Davis, Calif.: Univ. of California. p110. Henrique CM, Teófilo RF, Sabino L, Ferreira MMC, Cereda MP. 2007. Classification of cassava starch films by physicochemical properties and water vapor permeability quantification by FT-IR and PLS. J Food Sci. 72 (4): 184-189. 154 Höhne N, Galleguillos C, Blok K, Harnisch J, Phylipsen D. 2003. Evolution of Commitments under the UNFCCC: Involving Newly Industrialized Countries and Developing countries. Research Report 20141255, UBA-FB 000412. Ecofys, Berlin, Germany. Il’ina AV, Varlamov VP. 2003. Hydrolysis of chitosan in lactic acid. Appl Biochem Microbiol. 40 (3): 300-303. ISO 11357-2: Plastics - Differential scanning calorimetry (DSC) - Part 2: Determination of glass transition temperature 1999. Janjarasskul T and Krochta JM. 2010. Edible Packaging Materials. Annu. Rev. Food Sci. Technol. 1: 415-448. Jansson A, Thuvander F. 2004. Influence of thickness on the mechanical properties for starch films. Carbonhydrate Polymers. 56: 499-503. Jayakumar R, Nwe NT, Tokura S, Tamura H. 2007. Sulfated chitin and chitosan as novel biomaterials. Int J Bio Macromolecules. 40: 175–181. Jayakumar R, Prabaharan M, Reis RL, Mano JF. 2005. Graft copolymerized chitosan- present status and applications. Carbohyd Polym 62: 142-158. Jayakumar R, Reis RL, Mano JF. 2006. Chemistry and Applications of Phosphorylated Chitin and Chitosan. e-Polymers 035. Jongrittiporn S, Kungsuwan A, Rakshit SK. 2001. A study on the preservation of fishballs using chitosan. In European Conference on Advanced Technology of Safe and High Quality Foods-EUROCAFT, 5–7 December 2001, Berlin. Karbowiak T, Hervet H, Léger L, Champion D, Debeaufort F, Voilley A. 2006. Effect of plasticizers (water and glycerol) on the diffusion of a small molecule in Iota- 155 carrageenan biopolymer films for edible coating application. Biomacromolecules. 7: 2011-2019. Kester JJ, Fennema OR. 1986. Edible films and coatings: a review. Food Technol. 40 (12): 47-59. Khondka D, Tester RF, Hudson N, Karkalas J, Morrow J. 2007. Rheological behavior of uncross-linked and cross-linked gelatinized waxy maize with pectin gels. Food Hydrocolloids. 21: 1296-1301. Khondkar P, Aidoo KE, Tester RF. 2010. The ffects of emperature, pH, and cations on the rheological properties of the extracellular polysaccharides of Medicinal Species of Genus Tremella Pers. (Heterobasidiomycetes). Inter J Med Mushr. 12:73-83 Koca N, Rodriguez-Saona L, Harper W, Alvarez V. 2007. Application of fourier transform infrared spectroscopy for monitoring short-chain fatty acids in Swiss cheese. J Dairy Sci. 90: 2596-2603. Krochta JM. 2002. Protein as raw materials for films and coatings: definitions, current status, and opportunities. In Protein-Based Films and Coatings, ed., A Gennadios, 1:1–40. New York: CRC Press. Krochta JM, Baldwin EA,Nisperos-Carriedo M. 1994. Edible Coating and Films to Improve Food Quality. Technomic Pub Co., Lancaster, USA. Krochta JM, De Mulder-Johnston C. 1997. Edible and biodegradable polymer films: challenges and opportunities. Food Technol. 51(2):61–73. Ku KJ, Hong YH, song KB. 2008. Mechanical properties of a Gelidium corneum edible film containing catechin and its application in sausage. J Food Chem. 73: 217-221. 156 Lacroix M. 2009. Mechanical and permeability properties of edible films and coatings for food and pharmaceutical application. In: Embuscado ME, Huber KC. Edible films and coatings for food applications. Springer Science Business Media, LLC. New York, NY. p367-390. Larotonda FDS, Hilliou L, Sereno AMC, Gonçalves MP. 2005. Green edible films obtained from starch-domestic carrageenan mixtures. 2nd Mercosur Congress on Chemical Engineering 1-10. Lee SH. 1996. Effect of chitosan on emulsifying capacity of egg yolk. J Korean Soc Food Nutr. 25: 118-122. Li B, Kennedy JF, Peng JL, Yie X, Xie BJ. 2006. Preparation and performance evaluation of glucomannan-chitosan-nisin ternary antimicrobial blend film. Carbohydrate Polym. 65: 488–494. Lodi A and Vodovotz Y. 2008. Physical properties and water state changes during storage in soy bread with and without almond. Food Chem.110:554-561. Lu Y, Wang YJ, Tang X. 2008. Formulation and thermal sterile stability of a less painful intravenous clarithromycin emulsion containing vitamin E. Int J Pharm. 346: 47-56. Macquarrie R. Edible film formulation. 2002. United States Patent Application Publication US 2002/0155200 A1. Malkin AY and Isayev AI 2006. Rheology: Concepts, Methods, and Applications, ChemTec Publishing, Toronto Canada. p9-20. Mark HF, Atlas S. 1997. Introduction to polymer science. Kauiman HS ed., Introduction to polymer science and technology. Wiley Interscience. 157 Martins JT and Vicente AA. Incorporation of α-tocopherol into chitosan films: effects on film properties. Semana de Engenharia 2010. Mathew S, Brahmakumar M, Abraham TE. 2006. Microstructureal imaging and characterization of the mechanical, chemical, thermal, and swelling properties of starch-chitosan blend films. Biopolymers. 82: 176-187. Mauriello G, De Luca E, La Storia A, Villani F, Ercolini D. 2005. Antimicrobial activity of a nisin-activated plastic film for food packaging. Letters in Applied Microbiol. 41: 464–469. McFarland MJ. 2011. Biosolids Engineering, McGraw-Hill Companies, Inc. New York, NJ, B-2. Mei Y, Zhao Y, Yang Y, Furr HC. 2002. Using edible coating to enhance nutritional and sensory qualities of baby carrots. J Food Sci. 67: 1964-1968. Miller KS, Krochta JM. 1997. Oxygen and aroma barrier properties of edible films: A review. Trends Food Sci Tech. 8(7): 228-237. Morr CV, Ha EYW, 1993. Whey protein concentrates and isolate processing andfunctional properties. CRC Cr Rev Food Sci. 33:431-479. No HK, Meyers SP, Prinyawiwatkul W, Xu Z. 2007. Applications of chitosan for improvement of quality and shelf life of foods: a review. J Food Sci 72: 87-100. Ogale AA, Cunningham P, Dawson PL, Acton JC. 2000. Viscoelastic, thermal and microstructural characterization of soy protein isolate films. J Food Sci. 65(4): 672- 679. Park IK, Lee YK, Kim ML, Kim SD. 2002a. Effect of surface treatment with chitosan on shelf-life of baguette. J Chitin Chitosan. 7: 214-218. 158 Park HJ. 1999. Review: Development of advanced edible coatings for fruits. Trends Food Sci Tech. 10: 254-260. Park HJ, Chinnan MS. 1995. Gas and water vapor barrier properties of edible films from protein and cellulosic materials. J Food Eng. 25(4):497–507. Park HJ, Chinnan MS, Shewfelt RL. 1994. Edible corn-zein film coatings to extend storage life of tomatoes. J Food Proc Preserv. 18(4):317–31. Park HJ, Rhim JW, Weller CL, Gennadios A, Hanna M. 2002b. Films and coatings from proteins of limited availability. In Protein-Based Films and Coatings. A Gennadios, 12:305–28. New York: CRC Press. Park HJ, Weller CL, Vergano PJ, Testin RF. 1993. Permeability and mechanical properties of cellulose-based edible films. J. Food Sci. 58(6):1361–70. Park JW, Whiteside WS, Cho SY. 2008. Mechanical and water vapor barrier properties of extruded and heat-pressed gelatin films. LWT-Food Sci Technol. 41(4):692–700. Park S, Zhao Y. 2006. Development and characterization of edible films from cranberry pomace extracts. J Food Sci. 71(2):E95–101. Park S-I, Zhao Y. 2004. Incorporation of high concentration of mineral or vitamin into chitosan-based films. J Agric Food Chem. 52: 1933-1939. Park SK, Hettiarachchy NS, Ju ZY, Gennadios A. 2002c. Formation and properties of soy protein films and coatings. In Protein-Based Films and Coatings, ed. A Gennadios, 4:123–38. New York: CRC Press. Park SY, Lee BI, Jung ST, Park HJ. 2001. Biopolymer composite films based on κ- carrageenan and chitosan. Mater Res Bull. 36: 511-519. 159 Pavlath AE, Orts WJ. 2009. Edible Films: Why, What and How! Edible Films and Coatings for Food and Other Applications. Springer Science Business Media, LLC, New York, NY p1-24. Peressini D, Bravin B, Lapasin R, Rizzotti C, Sensidoni A. 2003, Starch-methylcellulose based edible films: rheological properties of film-forming dispersions. J Food Eng. 9: 25-32. Perkinelmer, 2011. http://www.perkinelmer.com/CMSResources/Images/44- 74844TCH_FTIRATR.pdf Pommet M, Redl A, Guilbert S, Morel M-H. 2005. Intrinsic influence of various plasticizers on functional properties and reactivity of wheat gluten thermoplastic materials. J Cereal Sci. 42:81–91. Pranoto Y, Salokhe VM, Rakshit SK. 2005. Physical and antimicrobial properties of alginate-based edible film incorporated with garlic oil. Food Res Int. 38: 267-272. Porous Material Inc. 2012. Water vapor permeability analyzer. Online source. http://www.pmiapp.com/products/water_vapor_transmission_analyzer.html Prommakool A, Sajjaanantakul T, Janjarasskul T, Krochta JM. 2011. Whey protein-okra polysaccharide fraction blend edible films: tensile properties, water vapor permeability and oxygen permeability. J Sci Food Agric. 92: 263-269. Pujol CA, Scolaro LA, Ciancia M, Matulewicz MC, Cerezo AS, Damonte EB. 2006. Antiviral activity of a carrageenan from Gigartina skottsbergii against intraperitoneal murine Herpes simplex virus infection. Planta Med. 72: 121-125. 160 Qussahlah M, Caillet S, Salmieri S, Saucier L, Lacroix M. 2004. Antimicrobial and antioxidant effects of milk protein-based film containing essential oils for the preservation of whole beef muscle. J Agric Food Chem. 52 (18): 5598-5605. Quintavalla S and Vicini L. 2002. Antimicrobial food packaging in meat industry. Meat Sci. 62: 373–380 Rao MA. 2007. Rheology of Fluid and Semisolid Foods: Principles and Applications 2nd , Springer Science Business Media, LLC, New York, NY Chapter 1. Redl A, Morel MH, Bonicel J, Vergnes B, Guilbert S. 1999. Extrusion of wheat gluten plasticized with glycerol: influence of process conditions on flow behavior, rheological properties, and molecular size distribution. Cereal Chem. 76: 361-370. Robertson GL. 1993. Food packaging. Principles and practice. New York: Mercel Dekker Inc. p 676. Rosenthal AJ 1999. Food Texture: Measurement and Perception, Aspen Publishers, Inc, p. 274. Rossman JM. 2009. Edible films and coatings for food applications. In: Embuscado ME, Huber KC. Edible films and coatings for food applications. Springer Science Business Media, LLC, New York, NY. p367-390. Sabato SF, Ouattara B, Yu H, D’Aprano G, Lacroix M. 2001. Mechanical and barrier properties of cross-linked soy and whey protein based films. J Agric Food Chem. 49: 1397-1403. Sarikus G, Seydim AC. 2006. Antimicrobial activity of whey protein based edible films incorporated with oregano, rosemary, and garlic essential oils. Food Res Int. 39: 639- 644. 161 Saucedo-Pompa S, Rojas-Molina R, Aguilera-Carbó AF, Saenz-Galindo A, De la Garza H. 2009. Edible film based on candelilla wax to improve the shelf life and quality of avocado. Food Res Int. 42: 511-515. Seol KH, Lim DG, Jang A, Jo C, Lee M. 2009. Antimicrobial effect of κ-carrageenan- based edible film containing overtransferrin in fresh chicken breast stored at 5°C. Meat Sci. 83: 479-483. Sepe MP. 1997. Thermal Analysis of Polymers, RAPRA Technology LTD, UK, p.13. Seydim AC. Sarikus G. 2006. Antimicrobial activity of whey protein based edible films incorporated with oregano, rosemary and garlic essential oils. Food Res Int. 39: 639- 644. Siracusa V, Rocculi P, Romani S, Rosa MD. 2008. Biodegradable polymers for food packaging: a review. Trends Food Sci Tech. 19: 634-643. Sorrentino A, Gorrasi G, Vittoria V. 2007. Potential perspective of bio-nanocomposites for food packaging applications. Trends Food Sci Tech. 18: 84-95. Steffe JF. 1996. Rheological Methods in Food Process Engineering 2nd, Freeman Press, East Lansing, MI, p 2. Suppakul P, Miltz J, Sonneveld K, Biggar SW. 2003. Active packaging technologies with an emphasis on antimicrobial packaging and its applications. J of Food Sci 68: 408- 420. Tanaka T, Nagao S, Ogawa H. 2001. Attenuated total reflection fourier transform infrared (ATR-FTIR) spectroscopy of functional groups of humic acid dissolving in aqueous solution. Anal Sci 1081-1084. 162 Tharanathan RN. 2003. Biodegradable films and composite coatings: Past, present and future. Trends Food Sci Tech. 14: 71–78. Troesch B, Eggersdorfer M, Weber P. 2012. 100 Years of Vitamins: Adequate intake in the elderly is still a matter of concern. J Nutr. 142: 979-980. United States Department of Agriculture, National agriculture online library. 2012. http://fnic.nal.usda.gov/dietary-guidance/dietary-reference-intakes Urano S, Yano K, Matsuo M. 1988. Membrane-stabilizing effect of vitamin E: effect of - tocopherol and its model compounds on fluidity liposomes. Biochem Biophys Res Commun. 150: 469-475. US Apple Association. 2012. http://www.usapple.org/apples-n-wax Wang GH. 1992. Inhibition and inactivation of five species of foodborne pathogens by chitosan. J Food Protect. 55(11): 916–919. Wang J, Wang B, Jiang W, Zhao Y. 2007. Quality and shelf life of Mango (Mangifera Indica L. cv. ‘Tainong’) coated by using chitosan and polyphenols. Food Sci Technol Int. 13(4):317–22. Wang Y, Padua GW. 2003. Tensile properties of extruded zein sheets and extrusion blown films. Macromol Mater Eng. 288(11):886–93 . Wessling C, Nielsen T, Leufven A. 2000. The influence of α-tocopherol concentration on the stability of ilnoleic acid and the properties of low-density polyethylene. Packag Technol Sci. 13: 19-28. Williams PA, Phillips GO. 2000. In Handbook of hydrocolloids; Phillips GO, Willians PA. ed., CRC Press, Cambridge, England, pp 1-19. 163 Witvrouw M, DeClercq E. 1997. Sulfated polysaccharides extracted from seaalgae as potential antiviral drugs. Gen Pharmacol. 29: 497-511. Yang L, Paulson AT. 2000. Mechanical properties of water vapour barrier properties of edible gellan films. Food Res Int. 33(7): 563–570. Yoo S and Krochta JM. 2011. Whey protein-polusaccharide blended edible film formation and barrier, tensile, thermal and transparency properties. J Sci Food Agric. 91: 2628-2636. Zacharopoulos VR, Phillips DM. 1997. Vaginal formulations of carrageenan protect mice from herpes simplex virus infection. Clin Diagn Lab Immunol. 4(4):465–468. Zhang J, Mungara P, Jane J. 2001. Mechanical and thermal properties of extruded soy protein sheets. Polymers. 42: 2569–2578. Zhong Q-P, Xia W-S. 2008. Physicochemical properties of edible and preservative films from chitosan/cassava starch/gelatin blend plasticized with glycerol. Food Technol Biotehnol. 46: 262-269. 164