AVOCADO BYPRODUCT EXTRACT: POSSIBLE USE AS ANTIOXIDANT COATING ON FLEXIBLE PACKAGING By Jin Zhang A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Packaging¾Doctor of Philosophy 2019 ABSTRACT AVOCADO BYPRODUCT EXTRACT: POSSIBLE USE AS ANTIOXIDANT COATING ON FLEXIBLE PACKAGING By Jin Zhang Food oxidation, as a serious concern of food deterioration, can induce food waste. This spoilage process results from free radical propagation in food molecules and introduces nutrient loss, off-flavor, off-odor and even toxicity issues. Food packaging with an antioxidant coating layer can effectively stabilize free radicals in food products. Avocado byproducts (peel and seed), as reliable and economical sources of natural antioxidants, are rich in phenolic compounds, a predominant group of antioxidants. These food wastes can be utilized for the development of antioxidant packaging. To date, there is no information available in the literature about the applications of avocado byproducts in the packaging field. The purpose of this research was to extract phenolic compounds from avocado byproducts and to use the crude extracts for the development of an antioxidant coating for three types of packaging films commonly used for food products, i.e., PP, PET and LDPE. To achieve this goal, 70% aqueous ethanol and 70% aqueous acetone were first used to recover phenolic compounds from avocado byproducts. An unconventional extraction procedure was employed to maximize extracted phenolic content within a limited time span. To polymerize the phenolic extracts on the polymer films, a non-metal contact dip coater was developed for this research. Alkaline saline (pH = 7.8) and laccase assist (pH = 5) coating methods were applied. Based on SEM observations, the coating layer was evenly distributed on the substrates with a thickness of 37.75 ± 0.30 nm; no polymerized clumps were noticed at a high level of resolution. AgNO3, DPPH•, and ABTS•+ assays were three approaches employed for evaluating the antioxidant efficacy of the phenolic coating in food simulants (95%, 50% and 10% aqueous ethanol). The AgNO3 allowed visual inspection for the existence of phenolic content in the coating layer. The experimental results of DPPH• and ABTS•+ assays showed that the alkaline saline coating technique, an inexpensive approach, could generate a phenolic coating layer with greater antioxidant effectiveness than the laccase assist coating method. Bio-based coating layers with different substrates, extract concentrations, extract ratios (i.e., Wpeel extract/Wseed extract) and coated films with different storage times were tested to analyze antioxidant variation; however, no statistically significant differences were found. While stabilizing free radicals in food simulants, phenolic compounds in the coating layer did not depend on a migration or surface release process for free radical elimination. Instead, they remained in the coating layer. Presumably, there was more than one layer of phenolic compounds polymerized on the substrates. After donating hydrogen atoms to quench free radicals, phenolic compounds at the surface layer of the antioxidant coating abstracted hydrogen atoms from their adjacent phenolic compounds in an inner layer of the antioxidant coating to continuously serve as antioxidants. It was also noticed that different temperature environments did not impact the stability of the coating layer. All these experimental outcomes implied a promising potential of this bio-based antioxidant coating in future commercial use. v Copyright by JIN ZHANG 2019 I dedicate this dissertation to my maternal grandmother v ACKNOWLEDGEMENT “You can’t put a limit on anything. The more you dream, the farther you get.” – Michael Phelps. Pursuing a Ph.D. is a long and tough journey. During the past years, my life was fully filled with tears and joys, ups and downs. Finally, I’m here, standing at the end of the journey. If I were given a second chance, I would still join in the Ph.D. program in the School of Packaging, MSU. First and foremost, I would like to express my deepest appreciation to my major professor, Dr. Susan Selke. Finding a major professor to guide my Ph.D. research is easy, but finding a knowledgeable and experienced major professor with a big heart is truly hard. I feel extremely lucky that I found one and she accepted me as her student. Since the very first day of my program, Dr. Selke never ever stopped caring about me, guiding me to gain professional knowledge and experience, and helping me thrive. I can’t imagine where I would be today without her kind guidance. I’ve already regarded her as a close family member rather than a major professor. My sincere and genuine thanks go to my committee members, Dr. Rafael Auras, Dr. Maria Rubino and Dr. Herlinda Soto Valdez for generously sharing their professional knowledge, experience and valuable recommendations to improve my research. A million thanks to Dr. Arthur Jones at the MSU Mass Spectrometry Facility for kindly providing technical support to analyze the avocado byproduct extracts for us. vi I would like to extend my special thanks to the managers of local Maru Sushi & Grill restaurants. Among all the local restaurants I contacted, Maru was the only one replied to my request, and supplied me fresh avocado byproducts without any hesitation. Without their timely help, I probably would need much longer time to source research materials and complete my experiments. From the bottom of my heart, I’m sincerely grateful for obtaining endless support from my lovely friends (not in any particular order), Hayati, Pom, KK, 286, Unikern, Zuimeili, Guanzi, Doudou and Jinwei. They unconditionally embraced my countless flaws, brought color into my world, saved me from tough situations, and always showed up whenever I requested help. Last but not least, my enormous love goes to my dear parents, Yayan Liu and Yang Zhang, for being my parents and dedicating their whole lives to support me chasing my dream. Because of them, I’m motivated to keep moving forward. Jin Zhang vii TABLE OF CONTENTS LIST OF TABLES……………………………...………………………………………...…….xii LIST OF FIGURES……………………………………….…………………………..………. xvi KEY TO SYMBOLS AND ABBREVIATIONS……………………………..……………….xxi CHAPTER ONE: Introduction and Objectives……………………………………………….....1 CHAPTER TWO: Literature Review………………………………….………..…………….....5 2.1 Food oxidation………………………………………………..…………….……………..…..5 2.1.1 Protein oxidation………………………………………….………………..……………6 2.1.2 Lipid oxidation……………………………………...……………...……..……..………7 2.1.2.1 Autoxidation………………………………………………….……..……..……7 2.1.2.2 Photooxidation………………………………………….……………..…..…… .8 2.1.2.3 Enzymatic oxidation …………………………………………..……….….……8 2.2 Environmental factors influencing food oxidation………………..….…..………..……..……9 2.2.1 Oxygen………………………………………………..……………....………..……..…9 2.2.2 Temperature……..………………………………….……..……..…………...……..… 10 2.2.3 Light exposure…..………………………………………..……..….....……...…….…..11 2.3 Antioxidants……………………………………………………..…………………..……….11 2.3.1 Stoichiometric antioxidants…..…………………………………………..……..…..… 12 2.3.2 Catalytic antioxidants……………………………………..……....………………...… 12 2.4 Mechanisms of antioxidant activity…………………………………..……………………....13 2.4.1 Chain-breaking electron donors (CB-D)……………… ……………………...…….….13 2.4.1.1 Single electron donation………………………..…………...…………....…….13 2.4.1.2 Hydrogen abstraction…………………………………..……………...…….....14 2.4.2 Chain-breaking electron acceptors (CB-A)………………… ………..…….…..…..…..15 2.5 Synthetic antioxidants and natural antioxidants………………………...……….……….…..16 2.5.1 Synthetic antioxidants…………………………………………………...……………..16 2.5.2 Natural antioxidants……………………………………………………...………….....17 2.5.2.1 Natural sources of antioxidants……………………………………………..….18 2.5.2.2 Crude extraction methods for natural antioxidants……………………….…….21 2.6 Most common assays for antioxidant evaluations……………….……………………….......22 2.6.1 ORAC assay……………………………………….…………………..…………….…22 2.6.2 FRAP assay…………………………….....………...……………….…………………23 2.6.3 TEAC assay………………………………….....………...……………………….……23 2.6.4 DPPH• and ABTS•+ assays…………...………………….......……..……………….…24 2.7 Antioxidant applications in the food industry……………………………………….….….…25 viii 2.7.1 Food additives……………………………………………………………................….25 2.7.2 Antioxidant active packaging…………………………..……….……………………...26 2.7.2.1 Active packaging……………………………….………...…………….……....26 2.7.2.2 Antioxidant active packaging………………………...…………………...……28 2.8 Avocado…………………………………………………………………………...…………33 2.8.1 Avocado and its byproducts……………………………………………...….….…....…33 2.8.2 Additional value of avocado byproducts…………………………………..……..…..…35 2.8.3 Applications of avocado byproducts in the packaging field……………….…..…..…....40 2.9 Phenolic compounds………………………………………………………………….……....40 2.9.1 Applications of natural phenolic compounds in the packaging field…………...….....…41 CHAPTER THREE: Crude Extraction of Phenolic Compounds from Avocado Byproducts..…44 3.1 Introduction ………………………………………...……….…...…………………..............44 3.2 Materials and methods……………………………………………………………….…...…..46 3.2.1 Byproducts preparation……………………………...…………………………………46 3.2.2 Extraction of phenolic compounds…………………………………………….….……47 3.2.2.1 Materials…………………………………………………………………..……47 3.2.2.2 Methods……………………………………………………………………...…47 3.3 Phenolic compounds in the crude extracts of Hass avocado byproducts…………...…………48 3.3.1 Materials………………………………………………………………………....……..48 3.3.2 Methods………………………………………………………………...………………48 3.3.3 Phenolic compound identification and quantification results…………………….…….49 CHAPTER FOUR: Non-metal Contact Coating Process……………….…………....................55 4.1 Introduction ………………………………….........................................................................55 4.2 Materials……………………………………………………………………….……………..61 4.2.1 Chemicals……………...........……………………………………….............................61 4.2.2 Sample films…………….......…………………………………….................................61 4.2.3 Coating device…………………………………………...…..........................................62 4.3 Methods……..............………………………………………………………………..............66 4.3.1 Laccase assist coating (sodium acetate) ……..………….……………...........................66 4.3.2 Alkaline saline coating (bicine) …….......………………...….………….......................66 4.3.3 Sample preparation for scanning electron microscope (SEM) observation……......……67 4.3.3.1 Sample preparation for surface observation…………………………..….……..68 4.3.3.2 Sample preparation for cross-sectional observation and thickness measurement…………………………………………………………………………...68 4.4 Antioxidant coating layer under SEM …………………………..……………………………71 CHAPTER FIVE: Evaluation of Antioxidant Activity of Coated Polymer Films Part I: Screening Tests…………………………………………...………………………….……79 5.1 Introduction……………………………………………………………….….………………79 ix 5.1.1 AgNO3…………………………………………………………...……………………..80 5.1.1.1 Introduction…...…...….………………………….………….......……………..80 5.1.1.2 Materials………………………………………………………………....……..82 5.1.1.3 Methods……………………………………………………………...…...….…83 5.1.1.4 Results and discussion…………………………………………………….……83 5.1.2 DPPH• assay………………………………………………………………..…………..….86 5.1.2.1 Introduction………………………………………………………………..……….86 5.1.2.2 Materials…………………………………………………………………..….….…90 5.1.2.3 Methods….…………………………………………………………….………...…90 5.1.2.4 Results and discussion……………………………………………..………….……91 5.1.2.4.1 Coating solution vs. antioxidant efficacy…………………..…………......91 5.1.2.4.2 Plastic substrates vs. antioxidant efficacy……………………..……….…94 5.1.2.4.3 Concentration of phenolic extract vs. antioxidant efficacy…………....….96 5.1.3 ABTS•+ assay…………………………………………………………………...…….……98 5.1.3.1 Introduction……………………………………………………………...…………98 5.1.3.2 Materials………………………………………………...…………...……...…….102 5.1.3.3 Methods…………………………………………………….…………..…..……..103 5.1.3.4 Results and discussion………………………………………………...…………..104 5.1.3.4.1 Antioxidant efficacy of the phenolic extracts from avocado byproducts………………………………………………………………………...104 5.1.3.4.2 Coating solutions vs. antioxidant efficacy……………………………....106 5.1.3.4.3 Plastic substrates vs. antioxidant efficacy………………….…………....108 Part II: Further Tests for Statistical Analysis…………………………………….…………...…111 5.2 Introduction……………………………………………………………………..……..……111 5.2.1 Plastic substrates vs. antioxidant efficacy…………………………………..……...….114 5.2.2 Film storage time vs. antioxidant efficacy……………………….……………………122 5.2.3 Extract ratio vs. antioxidant efficacy…………………………….…………...……….129 CHAPTER SIX: Mode of Antioxidant Activity of Coated Polymer Films………………….....140 6.1 Introduction………………………….……………………….………………………..……140 6.2 Antioxidative recoverability test……………………...…………………………………….142 6.2.1 Materials…………………….………….…………….………………………....…….142 6.2.2 Methods…………….…………………………...….………………………...……….142 6.2.3 Results and discussion………….………………….……..…………………...………142 6.3 Phenolic compound releasing experiments…………….…….……………………….……..152 6.3.1 Screening test for released phenolic compound…………….…...….…………………152 6.3.1.1 Materials…………….……………………….….………..……………...……152 6.3.1.2 Methods…………………….………………………….……………..……….152 6.3.1.3 Results and discussion……………….……….……………….…..…..………152 6.3.2 Migration test…………………….……………….……………………...……..…….156 6.3.2.1 Materials…………………………………………………….……….…….….156 x 6.3.2.2 Methods……………….…………………….……………………...………....156 6.3.2.3 Results and discussion…...…………….……..…………….…...………….…158 CHAPTER SEVEN: Conclusions and Future Work……………………………….………….170 7.1 Crude extraction from avocado byproducts……………………………………….……..….170 7.2 Non-metal contact coating process………………………………………………….…...….170 7.3 Evaluation of antioxidant efficacy of coated polymer films………………………...……….172 7.4 Mode of antioxidant activity of coated polymer films …………………………...…..….…..174 7.5 Future work……………………….………………….………………….……...…………..176 APPENDICES…………………………….………………….………………………………..178 Appendix A Antioxidant efficiency of coated polymer films (screening test results)…………...179 Appendix B Antioxidant efficiency of coated polymer films (further tests for statistical analysis)………………………………………………………………………………………...187 Appendix C Mode of antioxidant activity of coated polymer films……..………………………211 REFERENCES………….…………………………………......................................................229 xi LIST OF TABLES Table 2.1 Examples of natural antioxidants extracted from food byproducts………...…………..20 Table 2.2 Examples of prevalent active packaging in the food industry…………………...……..27 Table 2.3 Selective examples of antioxidant packaging research for food products since 2010….29 Table 2.4 Primary phenolic compounds and their contents in Hass avocado byproducts as reported in literature (Kosińska et al. 2012; Pahua-Ramos et al. 2012; Tremocoldi et al. 2018)…………...36 Table 2.5 Examples of applications of natural phenolic compounds in food packaging……….…43 Table 3.1 Phenolic compounds identified and quantified from the crude extracts of Hass avocado byproducts……….……………………………………………………………………………….51 Table 5.1 ET50 values estimated by the modified dose-response and Boltzmann sigmoidal models for the plastic substrates vs. antioxidant efficacy test. ABTS•+ assay was applied to evaluate polymer films coated with the avocado seed extract.............................................……………....117 Table 5.2 ET50 values estimated by the modified dose-response and Boltzmann sigmoidal models for the plastic substrates vs. antioxidant efficacy test. ABTS•+ assay was applied to evaluate polymer films coated with the avocado peel extract…………………………………………….119 Table 5.3 Antioxidant efficiency of coated PET, LDPE and PP films at different time intervals. The sample films were coated with acetonic seed extract in alkaline saline (pH = 7.8) solution. ABTS•+ assay was selected as the evaluation method…………………………………………..120 Table 5.4 Antioxidant efficiency of coated PET, LDPE and PP films at different time intervals. The sample films were coated with acetonic peel extract in alkaline saline (pH = 7.8) solution. ABTS•+ assay was selected as the evaluation method……………………………………..……121 Table 5.5 ET50 values estimated by the modified dose-response and Boltzmann sigmoidal models for the film storage time vs. antioxidant efficacy test. ABTS•+ assay was applied to evaluate polymer films coated with the avocado seed extract……………………………...………..……125 Table 5.6 ET50 values estimated by the modified dose-response and Boltzmann sigmoidal models for the film storage time vs. antioxidant efficacy test. ABTS•+ assay was applied to evaluate polymer films coated with the avocado peel extract…………………………………………….127 xii Table 5.7 Antioxidative efficacy of coated PP films with different storage times. The sample films were coated with acetonic peel and seed extracts in alkaline saline (pH = 7.8) solution. ABTS•+ radicals in 95% aqueous ethanol was used as the evaluation solution…………………...………128 Table 5.8 ET50 values estimated by the modified dose-response and Boltzmann sigmoidal models for the extract ratio vs. antioxidant efficacy. ABTS•+ assay was applied to evaluate coated PP films…………………………………………………………………………………………….133 Table 5.9 Reaction time comparisons for PP films coated with the phenolic extracts from avocado byproducts at different ratios. ABTS•+ assay was applied for the free radical reduction test…………………………………………………………………………………………..…..133 Table 5.10 Comparisons for the antioxidative efficiency of PP films coated with the phenolic extracts from avocado byproducts at different ratios. The sample films were coated in alkaline saline (pH = 7.8), and tested in ABTS•+ solution diluted with 95% aqueous ethanol……………134 Table 5.11 ET50 values estimated by the modified dose-response and Boltzmann sigmoidal models for the extract ratio vs. antioxidant efficacy. ABTS•+ assay was applied to evaluate coated LDPE films…………………………………………………………………………………………….137 Table 5.12 Reaction time comparisons for LDPE films coated with the phenolic extracts from avocado byproducts at different ratios. ABTS•+ assay was applied for the free radical reduction test………………………………………………………………………………………..……..137 Table 5.13 Comparisons for the antioxidative efficiency of LDPE films coated with the phenolic extracts from avocado byproducts at different ratios. The sample films were coated in alkaline saline (pH = 7.8), and tested in ABTS•+ solution diluted with 95% aqueous ethanol…………....138 Table 6.1 ET50 values estimated by the modified dose-response and Boltzmann sigmoidal models for the antioxidative recoverability test (ABTS•+ assay)………………………………………..149 Table 6.2 Pairwise comparisons for antioxidant efficiency of the coated PP films used for the antioxidative restorability test………………………………………………………..…………150 Table 6.3 Absorbance wavelengths suggested for the detection of different phenolic compounds by UV-Vis spectrophotometry………………………………………………………………….161 Table 6.4 Estimated concentrations of phenolic compounds released from the antioxidant coating layer during the migration test…………………………………………………………………..168 xiii Table A-1 Experimental data of the coating solution vs. antioxidant efficacy test (DPPH• assay)……… ……………………………………………………………………………….…..179 Table A-2 Experimental data of the plastic substrates vs. antioxidant efficacy test (DPPH• assay)…………… …………………………………………………………………………..….180 Table A-3 Experimental data of the concentration of phenolic extract vs. antioxidant efficacy test (DPPH• assay)…… ………………………………………………………………….…………181 Table A-4 Experimental data of the antioxidant efficacy of the phenolic extracts from avocado byproducts test (ABTS•+ assay)……… …………………………………………………..…….182 Table A-5 Experimental data of the coating solutions vs. antioxidant efficacy test (ABTS•+ assay)……… ………………………………………………………………………………..….184 Table A-6 Experimental data of the plastic substrates vs. antioxidant efficacy test (ABTS•+ assay)………… ………………………………………………………...………………………186 Table B-1 Fitted regression equations estimated by the dose-response and Boltzmann sigmoidal models for the plastic substrates vs. antioxidant efficacy test……………………………...……193 Table B-2 Experimental data of the plastic substrates vs. antioxidant efficacy test………….….194 Table B-3 Fitted regression equations estimated by the dose-response and Boltzmann sigmoidal models for the film storage time vs. antioxidant efficacy test……………………………….…..200 Table B-4 Experimental data of the film storage time vs. antioxidant efficacy test……………..201 Table B-5 Fitted regression equations estimated by the dose-response and Boltzmann sigmoidal models for the extract ratio vs. antioxidant efficacy test………………………………….……..208 Table B-6 Experimental data of the extract ratio vs. antioxidant efficacy test………………..…209 Table C-1 Fitted regression equations estimated by the dose-response and Boltzmann sigmoidal models for the antioxidative recoverability test……………………………………………...….216 Table C-2 Experimental data of the antioxidative recoverability test…………………………..217 xiv Table C-3 Phenolic compound size referenced for the mode of activity analysis of the antioxidant coating…………………………………………………………………………………………..228 xv LIST OF FIGURES Figure 2.1 Chemical structure of primary phenolic compounds in Hass avocado byproducts…...37 Figure 3.1 Phenolic compounds identification and quantification results for the crude extracts of Hass avocado byproducts………………………………………………………………….……..54 Figure 4.1 Chemical structure of a DOPA molecule……………………………………………..56 Figure 4.2 Chemical structures of a catechol and a gallol side chain……………………………..57 Figure 4.3 Schematic of interfacial activities between catechol-contained compounds and foreign substrates. R represents the remainder of a catechol-containing molecule, M represents a metal atom in a substrate, X+ represents a cation in a substrate.………………….………………..…….58 Figure 4.4 Schematic of the non-metal contact coating device with horizontal movement…….64 Figure 4.5 Schematic of the non-metal contact coating device with vertical movement………....65 Figure 4.6 SEM sample preparation for surface observation of a coated polymer film……….….68 Figure 4.7 SEM sample preparation for cross-sectional observation of a coated polymer film…..70 Figure 4.8 Topside view of the phenolic coating on a PP film……………………………………73 Figure 4.9 Topside view of the phenolic coating on a LDPE film……………………..…………73 Figure 4.10 Topside view of the phenolic coating on a PET film……………………...…………74 Figure 4.11 Cross-sectional view of the phenolic coating on a PP film……………….……….…74 Figure 4.12 Cross-sectional view of the phenolic coating on a LDPE film………………..……..75 Figure 4.13 Cross-sectional view of the phenolic coating on a PET film………………...…..…..75 xvi Figure 4.14 Cross-sectional view of the phenolic coating on a glass coverslip……….……...…..76 Figure 4.15 Schematic of covalent linkage between catechol-contained phenolic compounds and a LDPE or PP film surface. R represents the remainder of a catechol-containing phenolic compound…………………………………………………………………………………...……77 Figure 4.16 Schematic of p - surface interaction between a catechol-contained phenolic compound and a PET film surface…………………………………………………………………..………..78 Figure 5.1 Color changes of polymer films coated with phenolic compounds from avocado byproduct extract. The coated films were reacted with silver nitrate solution for at least 24 hours…………………………………………………………………………...…………………85 Figure 5.2 Single electron donation reaction between a DPPH• radical and a phenolic compound……………………………………………………….………………………...……...89 Figure 5.3 Antioxidant activity of LDPE films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The DPPH• solvent was diluted with 50% aqueous ethanol. Points a and b are estimates of the ET50 of LDPE (alkaline) and LDPE (laccase) respectively…………………..93 Figure 5.4 Antioxidant activity of LDPE films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The DPPH• solvent was diluted with 95% aqueous ethanol. Points a and b are estimates of the ET50 of LDPE (alkaline) and LDPE (laccase) respectively…………………..93 Figure 5.5 Antioxidant activity of PET films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The DPPH• solvent was diluted with 50% aqueous ethanol. Points a and b are estimates of the ET50 of LDPE (alkaline) and LDPE (laccase) respectively……….…...…..…94 Figure 5.6 Antioxidant activity of PET, LDPE and PP films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The DPPH• solvent was diluted with 50% aqueous ethanol. Point a is the estimate of ET50 of LDPE (alkaline), PET (alkaline) and PP (alkaline). Points b and b' are estimates of the ET50 of PET (laccase) and LDPE (laccase) respectively……………….….95 Figure 5.7 Antioxidant activity of PP films coated in solutions with different extract concentrations. The DPPH• solvent was diluted with 50% aqueous ethanol. The films were coated xvii in alkaline saline (pH = 7.8) solution. Points a and b are estimates of the ET50 of PP films coated in solutions with 25mg/mL and 12.5mg/mL avocado seed extract…………………………….…97 Figure 5.8 Oxidation reaction of ABTS with potassium persulfate to generate ABTS•+ radicals…………………………………………………………………….………………..…..101 Figure 5.9 Hydrogen abstraction reaction of ABTS•+ with a phenolic compound……..….…....102 Figure 5.10 Antioxidant activity of acetonic phenolic extracts from avocado byproducts. Points a and b are the estimated ET50 of the peel and seed extract respectively. The ABTS•+ solvent was diluted with 95% aqueous ethanol………………………………………………………..……..105 Figure 5.11 Antioxidant activity of PP films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The ABTS•+ solvent was diluted with 50% aqueous ethanol. Points a and b are estimated ET50 of PP (alkaline) and PP (laccase) respectively……………………………....107 Figure 5.12 Antioxidant activity of PP, PET and LDPE films coated in alkaline saline (pH = 7.8) solutions. The ABTS•+ and DPPH• solvents were diluted with 50% aqueous ethanol. Point a is the estimated ET50 of PP (ABTS) and PET (ABTS), and point b is the estimated ET50 of LDPE (DPPH), PET (DPPH) and PP (DPPH)… ……………………………………………………....108 Figure 5.13 Antioxidant activity of PET and PP films coated with ethanolic seed extract. The films were coated in alkaline saline (pH = 7.8) solution. The ABTS•+ solvent was diluted with 10% aqueous ethanol. Points a and b are the estimated ET50 of PET and PP respectively…….…109 Figure 5.14 Antioxidant activity of PP, PET and LDPE coated with acetonic seed extract. The films were coated in alkaline saline (pH = 7.8) solution. The ABTS•+ solvent was diluted with 95% aqueous ethanol. Points a, b and c are the estimated ET50 of PP, LDPE and PET respectively……………………………………………………………………………………..116 Figure 5.15 Antioxidant activity of PP, PET and LDPE coated with acetonic peel extract. The films were coated in alkaline saline (pH = 7.8) solution. The ABTS•+ solvent was diluted with 95% aqueous ethanol. Points a, b and c are the estimated ET50 of PP, PET and LDPE respectively……………………………………………………………………………….…….118 xviii Figure 5.16 Antioxidant activity of coated PP films with different storage times. The films were coated in alkaline saline (pH = 7.8) solution with acetonic seed extract. The ABTS•+ solvent was diluted with 95% aqueous ethanol. Points a and b are the estimated ET50 of PP (< 36 hours) and PP (7 days) respectively………………………………………………………………..……….124 Figure 5.17 Antioxidant activity of coated PP films with different storage times. The films were coated in alkaline saline (pH = 7.8) solution with acetonic peel extract. The ABTS•+ solvent was diluted with 95% aqueous ethanol. Points a and b are the estimated ET50 of PP (< 36 hours) and PP (7 days) respectively…………………………..…………….………………………………126 Figure 5.18 Antioxidant activity of PP films coated with the phenolic extracts from avocado byproducts at different ratios. The films were coated in alkaline saline (pH = 7.8) solution. The ABTS•+ solvent was diluted with 95% aqueous ethanol. Points a, b and c are the estimated ET50 of PP (100% peel), PP (100% seed) and PP (50% seed + 50% peel) respectively…………….....131 Figure 5.19 Antioxidant activity of LDPE films coated with the phenolic extracts from avocado byproducts at different ratios. The films were coated in alkaline saline (pH = 7.8) solution. The ABTS•+ solvent was diluted with 95% aqueous ethanol. Points a, b and c are the estimated ET50 of LDPE (50% seed + 50% peel), LDPE (100% peel) and LDPE (100% seed) respectively...….135 Figure 6.1 Reaction times (ETTotal) of the antioxidative restorability test. The alkaline saline method was applied to coat PP films with the acetonic peel extract. ABTS•+ stock solution was diluted with 95% aqueous ethanol………………………………………………………………146 Figure 6.2 ABTS•+ reduction process of the antioxidative restorability test. The alkaline saline method was applied to coat PP films with the acetonic peel extract. ABTS•+ stock solution was diluted with 95% aqueous ethanol. Points a, b, c, d and e are the estimated ET50 of trial 1, trial 2, trial 4, trial 3 and trial 5 respectively…………………………………………………………….147 Figure 6.3 Antioxidant efficiency of the after-reaction solutions collected from the antioxidative restorability analysis. ABTS•+ assay was used for the evaluation……………………….………154 Figure 6.4 Cross-sectional SEM image of a coated PP film (alkaline saline coating method) after its initial antioxidative reaction with ABTS•+ radicals……………………………………….…155 Figure 6.5 Two-sided contact migration cell for phenolic compound releasing test….……...…158 xix Figure 6.6 Overlapped UV-Vis spectrum of the crude phenolic extract from avocado peels in 95% aqueous ethanol at various concentrations………………………………..………………..……160 Figure 6.7 Calibration curves for the phenolic compound migration analysis. The wavelength selected for the phenolic compound absorbance measurement was around 280 nm……...……162 Figure 6.8 Schematic of antioxidative restorability of the phenolic coating on a plastic substrate…………………………………………………………………………………...……169 Figure B-1 Goodness of fit results estimated by the dose-response and Boltzmann sigmoidal models for the plastic substrates vs. antioxidant efficacy test……………………………...……187 Figure B-2 Goodness of fit results estimated by the dose-response and Boltzmann sigmoidal models for the film storage time vs. antioxidant efficacy test……………………………….…..196 Figure B-3 Goodness of fit results estimated by the dose-response and Boltzmann sigmoidal models for the extract ratio vs. antioxidant efficacy test…………………………………..…….202 Figure C-1 Goodness of fit results estimated by the dose-response and Boltzmann sigmoidal models for the antioxidative recoverability test…………………………………………………211 Figure C-2 Overlapped UV-Vis spectra of the crude phenolic extract from avocado peels in aqueous ethanol at various concentrations……………………………………………..………..218 Figure C-3 UV-Vis spectra of phenolic compounds released from coated PP films in each migration cell………………………………………………………………….………………..219 xx KEY TO SYMBOLS AND ABBREVIATIONS AAPH 2,2’-azobis(2-amidino-propane) dihydrochloride A1 A2 !′# ABAP ABTS AgNPs The minimum plateau of a free-radical absorbance reading versus phenolic compound concentration curve The maximum plateau of a free-radical absorbance reading versus phenolic compound concentration curve The maximum plateau of a free radical quantity versus reaction time curve 2,20-azo-bis (2-amidino-propane)hydrochloride 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) Silver nanoparticles ANOVA Analysis of variance ANS Ar : H Ar : O• Food additives and nutrient sources added to food An antioxidant A phenoxide ion Ar : OH A phenolic compound Ar• !$%&'()*+,- A new free radical generated by the hydrogen atom transferring process of the antioxidant, Ar : H The absorbance reading of DPPH• or ABTS•+ working solutions used for antioxidant test of coated films at time 23 xxi !$%&'()*+,4 BHA BHT The absorbance reading of DPPH• or ABTS•+ working solutions used for antioxidant test of coated films at time 25, 6=1,2…2 Butylated hydroxyanisole Butylated hydroxytoluene DI water Deionized water DOPA DPPH DW dx EC50 EFSA EVOH ET50 FDA FRAP Gallol HAT 3,4-Dihydroxyphenylalanine 2,2-diphenyl-1-picrylhydrazyl Dry weight Time constant The effective concentration required for antioxidants to reduce 50% free radicals in the working solution. European Food Safety Authority Ethylene vinyl alcohol It refers to the effective time required for the phenolic coating layer to reduce 50% free radicals in a working solution. Food and Drug Administration Ferric reducing antioxidant power 1,2,3-trihydroxyphenyl Hydrogen atom transfer Hillslope The slope or steepness of a free-radical absorbance reading versus phenolic compound concentration curve xxii HS IC Headspace The inhibition capacity of the antioxidant layer. This value is normalized. It varies from 0% to 100% K2S2O8 Potassium persulfate LDPE LSD Mfps •OH ORAC OS PE PES PET PG PLA Low density polyethylene Least significant different Mussel foot proteins Hydroxyl radical Oxygen radical absorbance capacity Oxidative stress Polyethylene Polyethersulfone Polyethylene terephthalate Propyl gallate Polylactic acid Polyphenols Polymerized phenolic compounds PP R• R2 Biaxially oriented Polypropylene An alkyl radical Squared correlation coefficient xxiii ROO : H ROO• ROS SEM Slope t t3 TAC TEAC Trolox USDA UV-Vis UHT =>?@A =BCACDE F F3 y A stabilized radical after the hydrogen abstraction reaction An oxygen-derived peroxyl radical of hydrocarbon substrates Reactive oxygen species Scanning electron microscope The steepness of an inhibition capacity curve Logarithm of reaction time LogET50 Total antioxidant capacity Trolox equivalent antioxidant capacity 6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid, a water- soluble analog of vitamin E United States Department of Agriculture Ultraviolet-Visible Ultra-high temperature Volume of aqueous ABTS solution Volume of aqueous potassium persulfate solution Logarithm of phenolic compound concentration The median value of F or LogEC50 Absorbance readings of a free radical working solution in a UV-Vis spectrophotometer xxiv G′ Normalized absorbance readings of a free radical working solution in a UV-Vis spectrophotometer. It varies from 0% to 100% xxv CHAPTER ONE: Introduction and Objectives Every year, around 1.3 billion tons of food produced for human consumption are either lost or wasted globally. These food wastes and losses account for roughly one-third of the total production (Gustavsson, Cederberg, and Sonesson 2011). Many factors, such as farming technology, food processing problems, and fluctuations of storage environments, are considered as the causes of this issue. Among all the causal factors, food oxidation is a serious concern. This deterioration process in both aqueous and lipid phases of food products results in the development of nutrient loss, off-flavor, off-odor, color change, and even toxicity of food products. The propagation of free radicals in food molecules is the main reason for the deterioration process. By continuously attacking healthy food cells, free radicals can aggravate their deleterious impact on food products. Under this circumstance, food products will not maintain their acceptable quality until their end consumption. Food manufacturers will thus increase their economic cost for food waste management. Applying antioxidants to food products is an effective way to retard or prevent oxidation activity and extend product shelf life. While contacting with oxidants in food molecules, antioxidants can serve as chain-breaking electron donors (CB-D). They can donate single electrons or (and) donate hydrogens to deactivate free radicals and thus slow down or stop free 1 radical propagation. In recent years, the applications of natural antioxidants in active food packaging have drawn remarkable interest from the food industry. Compared with synthetic antioxidants, natural antioxidants are stable, efficient, and more environmentally friendly. Manufacturers normally arouse less concerns if natural antioxidants are employed to protect their food products. Instead of obtaining natural antioxidants from vegetables, herbs, or fruit pulps, extracting the phytochemicals from food byproducts is a more economical alternative. Based on study results from the fields of food safety and food science, the antioxidant content of some fruit byproducts could be up to 27-fold higher than that of the fruit pulp. Avocado byproducts are examples of this case. The seed and peel are rich in phenolic compounds, a predominant group of natural antioxidants. Researchers have noticed the potent antioxidant capability of the phenolic extract from different varieties of avocado seeds and peels. To date, no potential health risk or toxic issues of avocado byproducts have been reported. However, there is very limited information available in the literature regarding the applications of avocado byproducts in the food packaging field. Therefore, the purpose of this research was to extract phenolic compounds from avocado byproducts (seed and peel) and use the extracts to develop an effective antioxidant coating for 2 three types of packaging films commonly used for food products, i.e., polypropylene (PP), polyethylene terephthalate (PET) and low density polyethylene (LDPE). For this research, the goals were to: (1) Develop a method to extract crude phenolic compounds from avocado byproducts (peel and seed). (2) Develop a non-metal contact coating device. (3) Coat the phenolic compounds on PP, PET and LDPE films by using alkaline and acidic solutions. (4) Test and quantify antioxidant activity of the coated polymer films. (5) Understand the mode of antioxidant activity of the phenolic coating. In the next chapter of this document, detailed information is provided to explain food oxidation activity, the reaction mechanisms, types, sources, and the applications of antioxidants in the food industry. In addition to avocado byproducts, phenolic compounds and their current applications in the food packaging field are discussed. Chapter three describes the experimental method used in this research for the extraction of phenolic compounds from avocado byproducts. Chapter four describes the development of a non-metal contact coating device together with two different coating solutions. To test the efficacy of the antioxidant coating by using the two different solutions, chapter five presents the analysis and evaluation results of the antioxidant 3 activities of the coated PP, PET and LDPE films tested in silver nitrate (AgNO3), DPPH• (2,2- diphenyl-1-picrylhydrazyl), and ABTS•+ (2,2'-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid)) assays. Chapter six elaborates the mode of antioxidant activity of the biochemical coating layer. In the last chapter, current achievements and further improvement plans are discussed. 4 CHAPTER TWO: Literature Review 2.1 Food oxidation Food oxidation refers to the chemical reaction between food molecules and oxygen. This process happens in both aqueous phases and lipid phases of food products (Skibsted 2010). It not only results in nutrient loss, unhealthy compounds, undesired color change, and unpleasant flavor, but also, it creates free radicals inside food. These free radicals, as detrimental agents, are classified as reactive oxygen species (ROS). They are highly reactive molecules with an unpaired electron. They can attack most biological molecules at the site of its formation to obtain another electron for stabilization, initiate the propagation of free radical chain reactions, and damage healthy food molecules such as lipids and proteins (Betteridge 2000). As a consequence of both the initial oxidation and the subsequent cascade of reactions, the lifespan of food products can be significantly shortened (Cheeseman and Slater 1993; Lobo et al. 2010; Phaniendra, Jestadi, and Periyasamy 2015; Wasowicz et al. 2004). Food oxidation is always of great concern. It can occur during manufacture, handling, transportation, storage and preparation processes (Soladoye et al. 2015). It expedites the food deterioration activity, resulting in a soaring number of food products becoming of unacceptable 5 quality before consumption. Food manufacturers, grocery stores, and consumers, under this circumstance, have to throw away the oxidized food. This contributes to another global issue, i.e., food waste. In addition, oxidized food products can cause further oxidation reactions during digestion phases of human body, generating other oxidation substances with toxic potential (Van Hecke et al. 2015; Vicente et al. 2012). Via blood distribution and intestinal consumption, the harmful substances may pose risks to internal organs (Estévez and Luna 2017). Under normal conditions, the self-protection system inside the human body is able to scavenge oxidized products so that the quantities of these harmful compounds are controllable. Once this balance is disturbed oxidative stress (OS) will occur (Estévez et al. 2017). Unmanageable development of OS in crucial molecules, for instance proteins, lipids, and DNA, is normally associated with diseases such as inflammatory bowel diseases, fibrotic degeneration of the liver and kidney, Alzheimer and cataractogenesis (Berlett and Stadtman 1997; Keshavarzian et al. 2003; Li et al. 2014). 2.1.1 Protein oxidation Protein oxidation prevails in muscle foods. Because of the large and complex structure of protein molecules, free radicals like superoxide anions can easily locate a spot on the molecules to attack, leading to significant modifications of protein conformation, polymerization, 6 precipitation, etc. (Lund and Baron 2010). This results in food quality degradation; for example meat products with less tenderness (Suman et al. 2014), sharp off-flavors and nutrient loss in dairy products can occur (Citta et al. 2017; Scheidegger et al. 2010). 2.1.2 Lipid oxidation Lipid oxidation can be found in edible oils, nuts, fatty meat and fish products. At the early stage of lipid oxidation, hydroperoxides can be formed. These vulnerable compounds can be further oxidized and decomposed into acids, alcohols and aldehydes, which are well-known as contributors to the development of nutrient loss, off-odor and off-flavor including undesired rancid taste of food products (St Angelo 1996; Wasowicz et al. 2004). Compared with the undesired changes caused by protein oxidation, lipid oxidation normally leads to more noticeable modifications (Lund and Baron 2010). Lipid oxidation is categorized into three classes, namely autoxidation, photooxidation, and enzymatic oxidation. 2.1.2.1 Autoxidation Inside the lipid content of food products, oxygen can serve as the trigger of free radical reactions and a substrate for free radical propagation (Porter 1987). This spontaneous degradation process is known as autoxidation. Hydroperoxides are the primary oxidation products of autoxidation. It can further yield other volatile and non-volatile products causing 7 spoilage and rancidity of food products (Paquette, Kupranycz, and van de Voort 1985; Schultz 1962). 2.1.2.2 Photooxidation As indicated by the reaction name, this oxidative degradation is triggered by the presence of light, and acts in two ways based on Type I and Type II mechanisms. When a singlet state food photosensitizer like chlorophyll contacts with light, it can be excited by absorbing the light energy. Because of intersystem crossing steps, the excited singlet state photosensitizer can evolve to an excited triplet state photosensitizer. If the excited triplet state photosensitizer reacts with triplet oxygen, singlet oxygen can be generated. This process is called a Type II mechanism. If the excited triplet state photosensitizer abstracts an electron or a hydrogen atom from a substance, radicals can be generated. This process is called a Type I mechanism (Lee 2002; Turro 1985). 2.1.2.3 Enzymatic oxidation Enzyme catalysts such as cyclooxygenase and lipoxygenase can increase the rate of the oxidation process in the lipid content of food products. They catalyze polyunsaturated fatty acids to generate unsaturated fatty acid hydroperoxides (Henry et al. 2002; Tripathi and Mishra 2016). The occurrence of this reaction creates a pathway for the introduction of toxic compounds in food products (Kubow 1992). 8 One of the greatest concerns caused by enzymatic oxidation is food product browning. It starts from the oxidation of phenolic compounds in food products, reacts with polyphenol oxidase and other proteins, and finally generates brown pigments, melanosis, on the product surface. This visual deterioration process normally makes food products, especially freshly cut food products, unacceptable (Gonçalves and Oliveira 2016; Jeon, Kim, and Chang 2013; Nirmal et al. 2015). 2.2 Environmental factors influencing food oxidation Food oxidation is a complex process. Individual environmental factors do not influence food deterioration reactions separately. Rather, in real applications, the oxidation activity is subject to multiple impacts concurrently. Its occurrence and reaction rates are influenced by but not limited to the following environmental factors: 2.2.1 Oxygen Food oxidation relies on the involvement of oxygen. For lipid oxidation, oxygen paves the way for fatty acid decomposition to develop food rancidity. For protein oxidation, oxygen involved chemical reactions induce amino acid modification, resulting in protein fragmentation or protein-protein cross-linkage. Storing food products in a low oxygen environment has been recognized as an effective method to slow down food oxidation. When lowering the oxygen concentration of the surrounding environment from 21% to 0.5%, researchers noticed significant 9 decrease in oxygen diffusion in emulsions of fatty substances causing decelerated oxidation activity (Marcuse and Fredriksson 1968). Theoretically, increasing environmental oxygen concentration could ease food oxidation activity. More oxidation products should be detected from food products. However, this was not always the case. Researchers found that within the first 4 days of the experiment, there was no significant difference among the food oxidation product, oxymyoglobin content, of minced beef surrounded by 40, 60, and 80% oxygen in the headspace (O’Grady et al. 2000). 2.2.2 Temperature Heat provides energy to molecules, and elevated temperatures provide more energy. As the Arrhenius equation implies, molecules with higher energy collide with each other more frequently. Frequent collision leads to higher kinetic energy, which meets the requirement for the activation energy of chemical reactions. The activation energy determines the rate of occurrence of chemical reactions. Therefore, increasing temperature could promote food oxidation reactions. It has been reported that ultra-high temperature (UHT) treatment for dairy product processing could accumulate protein oxidation products such as oxidized amino acid residues causing cross- linked protein species, which could constitute major food allergens (Fenaille et al. 2005). Also, high cooking temperature for meat products could increase the formation of protein carbonyls (Roldan et al. 2014). 10 2.2.3 Light exposure Light waves catalyze food oxidation. They influence food oxidation from two aspects: light density and light exposure time. Short light waves with high energy can ease food photooxidation activity (Bekbölet 1990). It was reported that the hydroperoxide content in ice cream, generated by food photooxidation process, could bring about the development of an off- flavor issue (Shiota et al. 2004). Color stability of fresh meat could be disturbed by light-induced myoglobin oxidation (Cooper et al. 2017). After exposing milk products to light, nutrients like vitamin A and riboflavin were degraded and off-flavor was noticed due to the formation of aldehydes in the fat content and the degradation of sulfur-containing amino acids (Brothersen et al. 2016). Light exposure also facilitates oxygen consumption for food oxidation. For cream powders with 35 weeks of storage time, researchers evaluated the product oxidation by measuring the remaining oxygen concentration in the packaging headspace (HS) at each time interval. Compared with the cream powders stored in the dark, the HS oxygen concentration of the cream powders kept in light was significantly lower (Andersson and Lingnert 1998). 2.3 Antioxidants Antioxidants are chemical compounds which react with free radicals to slow down or inhibit the oxidation activity in food products and thus to extend product lifetimes (Choe and 11 Min 2009). This defense reaction in food molecules minimizes the negative impact of free radicals on food quality without changing food taste and odor. 2.3.1 Stoichiometric antioxidants Antioxidants can be either stoichiometric or catalytic reagents. As stoichiometric reagents, antioxidants sacrifice themselves to stabilize free radicals, and thus slow down the product deterioration process. This consumption activity is permanent. When free radicals propagate at a very high rate, stoichiometric antioxidants may not make desired contributions to slow down or stop this detrimental propagation because the total amount of antioxidants is limited (Haber and Gross 2015; Scott 1989). 2.3.2 Catalytic antioxidants Catalytic antioxidants act differently. They induce antioxidant activity, repeatedly being involved in the free radical stabilization reaction, but they are not consumed. This process is considered as ROS (reactive oxygen species) detoxification activity without self-sacrifice (Golden and Patel 2008). On that account, catalytic antioxidants, even at a low concentration, are still able to present their potent antioxidant ability to inhibit the damaging impacts caused by free radical production (Franck et al. 2013). 12 2.4 Mechanisms of antioxidant activity To stabilize free radicals, antioxidants primarily act as kinetic chain-breaking agents. These agents are generally classified into two categories, namely, chain-breaking electron donors (CB-D) and chain-breaking electron acceptors (CB-A). 2.4.1 Chain-breaking electron donors (CB-D) Scheme 1.1 represents the mechanism of the CB-D reaction. The left part of the equation explains the electron donation process of an antioxidant to a free radical; the right part of the reaction demonstrates the hydrogen abstraction activity of a free radical, Where + e- + H+ ROO• ROO- ROO : H (Scheme 1.1) ROO• is an oxygen-derived peroxyl radical of hydrocarbon substrates ROO : H is a stabilized radical after the hydrogen abstraction reaction 2.4.1.1 Single electron donation For the reaction of single electron donation, an antioxidant stabilizes a free radical by donating an electron to the free radical which otherwise could attack and damage healthy molecules. Similar to the hydrogen abstraction reaction, the antioxidant becomes a new free radical after reacting with this free radical. The ionization potential of the antioxidant determines 13 the occurrence of this reaction. It is also positively correlated to the degree of difficulty of the free radical stabilization process. In other words, the lower the ionization potential of the antioxidant, the easier the single electron donation will be initiated (Ashby 1988; Bendary et al. 2013). 2.4.1.2 Hydrogen abstraction For the reaction of hydrogen abstraction, antioxidants are used to eliminate peroxidation processes participated in by oxygen-derived peroxyl radicals of hydrocarbon substrates, ROO•. During the elimination process, peroxyl radicals abstract hydrogen atoms from antioxidants (Scheme 1.2) to transfer to a more stabilized and less reactive state (Luzhkov 2005; Morello, Shahidi, and Ho 2002). The hydrogen bond dissociation enthalpy influences this antioxidation reaction. It is positively correlated to the degree of difficulty of this free radical stabilization process. In other words, the lower the hydrogen bond dissociation enthalpy in antioxidants, the easier the hydrogen abstraction reaction will take place (Mader, Davidson, and Mayer 2007). Where ROO• + Ar : H = ROO : H + Ar• (Scheme 1.2) ROO• is an oxygen-derived peroxyl radical of hydrocarbon substrates Ar : H is an antioxidant 14 ROO : H is a stabilized radical after the hydrogen abstraction reaction Ar• is a new free radical generated by the hydrogen atom transferring process of the antioxidant, Ar : H Both the hydrogen abstraction and single electron donation activities are prevalent reactions occurring between antioxidants and free radicals (Morello et al. 2002). They, in most cases, take place concurrently. It is hard to differentiate and identify one reaction from another (Liang and Kitts 2014). In both reactions, the new free radicals are more stable and less reactive compared with the free radicals the antioxidants neutralized, and thus remarkably reduce the risk to unharmed food molecules (Barzegar 2012; Lü et al. 2010; Zhuravlev et al. 2016). 2.4.2 Chain-breaking electron acceptors (CB-A) Scheme 1.3 represents the mechanism of CB-A reaction. Unlike the CB-D reaction, the - e- free radical in this reaction loses an electron. It basically oxidizes alkyl radicals into non radical products. This reaction efficiency relies on oxygen deficiency of the reaction condition (Al- Malaika et al. 2017). - e- R• Non radical product (Scheme 1.3) O2 deficient R• is an alkyl radical Where 15 2.5 Synthetic antioxidants and natural antioxidants There are two types of antioxidants widely used in the food industry, i.e., synthetic antioxidants and natural antioxidants. 2.5.1 Synthetic antioxidants Synthetic antioxidants are chemical compounds produced by chemical processes. Butylated hydroxyanisole (BHA) and butylated hydroxytoluene (BHT) are commonly used synthetic antioxidants to prevent oxidation activity of fats. They are phenolic compounds, and can be applied for butter, snacks, and processed meat products, for example, pork sausage and beef patties. However, it was reported that BHA and BHT have the potential of carcinogenicity (Olsen et al. 1986; Sasaki et al. 2002), and this potential risk to human health has driven researchers to develop different experimental methods to further evaluate the properties of these two synthetic antioxidants (S.-H. Jeong et al. 2005; Vandghanooni et al. 2013). Propyl gallate (PG), as another type of commonly used synthetic antioxidant, can also be found in the food industry. This phenolic compound is widely applied to meat products, frozen meals, edible oils and soup mixes. Despite the fact that this synthetic compound serves as an effective antioxidant to prevent food rancidity, researchers are unveiling its adverse effects on human health. It has been noticed that the side effects of PG can be skin allergy (García- Melgares et al. 2007), stomach damage, and kidney problems (EFSA Panel on Food Additives 16 and Nutrient Sources added to Food (ANS), 2014). In the United States, the Food and Drug Administration (FDA) allows the application of PG in the food industry under strict regulations (21 C.F.R. § 582. 3660 (2018)), In European countries, PG is permitted for very limited categories of food products (Annex II of Regulation (EC) No 1333/2008). 2.5.2 Natural antioxidants Natural antioxidants are phytochemicals. They are natural compounds mainly derived from vegetables, fruits, spices, herbs, and tea leaves. Normally, natural antioxidants are obtained by aqueous extraction processes, safer and more environment-friendly methods compared with the chemical processing methods for obtaining synthetic antioxidants. Considering their antioxidative efficiency, and stability while reacting with oxidants in food products, during recent years, natural antioxidants have drawn increasing attention from the market for food quality protection (Caleja et al. 2017; Carocho et al. 2014; Carocho and Ferreira 2013). There are three main groups of natural antioxidants: vitamins, carotenoids and phenolic compounds. In order to understand the antioxidative activity of these three categories of antioxidants, researchers evaluated their efficiency in different cultivars. Based on the experimental results of their free radical scavenging ability, phenolic compounds comparatively had greater effect on oxidation prevention (Gil et al. 2002). 17 Natural ascorbic acid, also known as Vitamin C, is a powerful natural antioxidant (Carocho et al. 2014). It is obtainable in fruits and vegetables. Due to its hydrophilic characteristics, this antioxidant is capable of reacting with free radicals in both lipid and aqueous contents to stabilize food products such as fish, vegetable oils, milk and beverages (Cort, 1982). Since natural ascorbic acid is also served as a nutrient supplement, it has been approved by the FDA as an antioxidant preservative (21 C.F.R. § 145. 110 (2018)). Lycopene is a well-known natural antioxidant in the category of carotenoids. It is also used as a natural pigment on account of its red color. Tomatoes, guavas, watermelon, and pink grapefruit are rich in lycopene. The food industry extracts this antioxidant from vegetables and fruits and then applies it to beverages, dairy products and sauces to extend product shelf life. Different from BHA and BHT, there is no research revealing that lycopene has the potential of carcinogenicity, nor toxicity (Bánhegyi 2005). On the contrary, researchers presented promising data to indicate that lycopene, other than serving as a strong antioxidant for food products, can be recommended as a dietary supplement as well to improve human health (Paetau et al. 1998; Riccioni et al. 2008; Wood et al. 2008). 2.5.2.1 Natural sources of antioxidants In addition to vegetables, fruit pulps, spices and herbs, food byproducts from the industry, for instance husks, peels and seeds, also contain antioxidant phytochemicals. Instead of 18 directly throwing them away to landfills causing environmental impact, an additional value of these non-edible parts can be achieved if they are utilized as a source of natural antioxidants. Compared with vegetables and fruit pulp, using food byproducts is a more economical option. Moreover, experimental results have shown that the antioxidant content of some fruit byproducts could be up to 27-fold higher than that of the fruit pulp (Goulas and Manganaris 2012; Guo et al. 2003; Someya, Yoshiki, and Okubo 2002). The types of natural antioxidants extracted from food byproducts are not limited to a small range. Phenolic compounds, carotenoids and vitamins can all be found in food byproducts (Selvamuthukumaran and Shi 2017). Among them, phenolic compounds with high antioxidant potency are predominant (Moure et al. 2001). Due to the aforementioned facts, increasing numbers of researchers have started using food byproducts as attractive sources for natural antioxidant experiments (Table 2.1). 19 Table 2.1 Examples of natural antioxidants extracted from food byproducts. Antioxidant Source Types of Antioxidants Pomegranate peel and seed Phenolic compounds Start fruit residue Grape waste Citrus peel Mango peel powder Avocado seeds and peels Coffee spent grounds and silverskin Tomato peel Orange, lemon and grapefruit wastes References Singh et al. (2002) Shui et al. (2006) Lafka et al. (2007) Xu et al. (2008) Ajila et al. (2010) Kosińska et al. (2012) Jiménez-Zamora et al. (2015) Phenolic compounds Phenolic compounds Phenolic compounds Phenolic compounds and carotenoids Phenolic compounds Phenolic compounds Lycopene and phenolic compounds Elbadrawy et al. (2016) Vitamin C and phenolic compounds Sir Elkhatim et al. (2018) 20 2.5.2.2 Crude extraction methods for natural antioxidants To utilize antioxidants from natural sources, crude extraction is the first step. This process is primarily influenced by solvent type, concentration and polarity, extraction duration and temperature, pH value of the extraction solution, and solubility of natural antioxidants in the extraction solvent. A wide range of crude extraction solvents have been reported for natural antioxidants. For water-soluble antioxidants, for instance ascorbic acid, phenolic compounds, flavonoids and glutathione, pure or aqueous solvents with high to medium polarity were used for the extraction. Common solvents for water-soluble antioxidants are water, ethanol, methanol and acetone (Abu et al. 2017; Boeing et al. 2014). For lipid-soluble antioxidants, for example vitamin A, vitamin E, and carotenoids, organic solvents with medium to low polarity, such as ethanol, ether and benzene, were reported for the extraction (Ghasemzadeh et al. 2015; Traber and Atkinson 2007). Crude extraction procedures can be categorized into two groups, i.e., conventional and unconventional. Hot water bath, maceration and Soxhlet extraction method are classified as conventional extraction approaches. They are regarded as time-consuming and costly means with low yield of efficiency (Selvamuthukumaran and Shi 2017; Zhang, Lin, and Ye 2018). In contrast, unconventional extraction procedures like ultrasound, microwave, pressurized, pulsed 21 electric field and enzyme hydrolysis are proposed due to their economic, environmental friendly and high efficiency attributes (Hidalgo and Almajano 2017; Xu et al. 2017). 2.6 Most common assays for antioxidant evaluations To evaluate free radical elimination ability of antioxidants, researchers have developed various experimental methods. The free radical decreasing process can be quantified by measuring a free radical absorbance peak with a UV-Vis spectrophotometer at a particular wavelength. For some of the testing approaches, the free radical elimination process can also be visualized by observing color change of the free radical working solutions. 2.6.1 ORAC assay ORAC stands for oxygen radical absorbance capacity. This assay is utilized for testing the antioxidant efficiency of hydrophilic antioxidants (Huang et al. 2002). In ORAC testing, natural antioxidants slow down the degradation rate of fluorescent molecules (normally fluorescein) due to the attack from a free radical generator, such as AAPH ((2,2’-azobis(2-amidino-propane) dihydrochloride). To reduce free radical attack, natural antioxidants transfer their hydrogen atoms (HAT) to stabilize free radicals. This transfer efficiency is quantified by creating decay curves of fluorescent molecules, compared with standard decay curves, Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid, a 22 water-soluble analog of vitamin E) curves, and expressed as Trolox equivalence (Kohri et al. 2009; Roy et al. 2010). 2.6.2 FRAP assay FRAP stands for ferric reducing antioxidant power. It determines the antioxidant ability of phytochemicals by measuring the reduction volume of ferric ions. At low pH value (3.6), ferric ions (Fe3+) in working solution will be gradually transformed to ferrous ions (Fe2+). The transformation process can be visualized by a blue color development of the working solution, and quantified by measuring the absorbance peak at 593 nm with a UV-Vis spectrophotometer (Benzie and Strain 1996). The FRAP assay is easy to prepare. It does not require a complicated detection method to evaluate the rapid transformation process. However, the low pH value of the working solution does not represent physiological conditions (López-Alarcón and Denicola 2013). 2.6.3 TEAC assay Trolox equivalent antioxidant capacity (TEAC) assay was developed for total antioxidant capacity (TAC) determination (Miller et al. 1993). This method was later modified by Van den Berg et al. (1999). According to Van de Berg et al., fresh radical anions of ABTS (2,2'-azino-bis(3- ethylbenzothiazoline-6-sulphonic acid)) working solution, ABTS•-, need to be prepared every 23 time for the antioxidant testing. It can be obtained by mixing ABAP (2,20-azo-bis (2-amidino- propane)hydrochloride) with ABTS•2- stock solution. After adding natural antioxidants to the working solution, the antioxidant efficiency can be analyzed by measuring the mixture at 734 nm with a UV-Vis spectrophotometer for 6 minutes. The testing result is expressed as Trolox equivalents by comparing it with Trolox standard graphs. To minimize environmental interference, light exposure should be avoided throughout the test. 2.6.4 DPPH• and ABTS•+ assays DPPH• (2,2-diphenyl-1-picrylhydrazyl) and ABTS•+ are stable free radicals. Their working solutions are purple and blue-green in color, respectively. To conduct the antioxidant test, ABTS•+ needs to be prepared by oxidizing ABTS with K2S2O8, whereas DPPH• can be directly purchased. During the test, antioxidants stabilize these radicals resulting in free radical reduction. This radical concentration decreasing process can be measured with a UV-Vis spectrophotometer at 517 nm (DPPH•) or 734 nm (ABTS•+). For this research, DPPH• and ABTS•+ assays were selected to evaluate the antioxidant efficacy of the phenolic coating layer on PP, PET and LDPE films. More discussions regarding these testing methods are included in chapter five. 24 2.7 Antioxidant applications in the food industry Synthetic and natural antioxidants have two main applications in the food industry: either directly introduced to food products as additives or applied to food packaging to maintain product smell, color, taste, and texture until final consumption. 2.7.1 Food additives Food additives can be introduced to food products at any step of their processing and production procedures. Manufacturers, traditionally, add BHA, BHT and PG into edible oils to ensure product quality (Raikos 2017), and ascorbic acid and tocopherols into milk products to prevent milk protein oxidation (L.H. Skibsted 2010). The introduced volume of different additives is under strict control by regulatory authorities. Food products in European countries with antioxidant additives must comply with European Food Safety Authority (EFSA) regulations. For food products manufactured and consumed in the United States, they must follow food safety regulations issued by the Food and Drug Administration of the United States of America (FDA) and United States Department of Agriculture (USDA). The allowed content of each antioxidant is not invariable. Based on researchers’ investigation, study and experimental results, the authorities inspect and re-evaluate potential safety risks of each antioxidant and revise the corresponding regulations regularly. 25 2.7.2 Antioxidant active packaging When antioxidants were employed in the packaging field, researchers developed antioxidant packaging. Antioxidant packaging is classified as active packaging. It is widespread in the food, pharmaceutical and cosmetic industries due to their potent ability to prolong product shelf life. 2.7.2.1 Active packaging In order to obtain desired surrounding conditions for a product, active packaging applies different technologies to adjust the inside environment of a sealed packaging system. Depending on the type of the packaged product, the applied technologies may vary. During transportation and delivery, even faced with the fluctuations of temperature and relative humidity, the packaged product, under the effective protection from its active packaging, can still be prevented from spoilage, off-flavor, and off-odor issues, and finally present its high quality to end consumers. Thanks to active packaging, manufacturers, especially food manufacturers, each year significantly reduce their expenses for handling products with unacceptable quality. The environment thus can suffer less impact. The following table includes active packaging types that are prevailing in the food industry (Coles, McDowell, and Kirwan 2003; Ozdemir and Floros 2004; Patel 2018; Prasad and Kochhar 2014; Yildirim et al. 2018). 26 Table 2.2 Examples of prevalent active packaging in the food industry. Mechanisms Use oxygen absorbing sachets inside packaging to reduce excessive oxygen in packaging headspace Use ethylene absorbing sachets inside packaging to reduce excessive ethylene in packaging headspace Packaging Type Oxygen scavenger Ethylene scavenger Carbon dioxide controlled Odor removal Packaging Application Cheese spread, cakes, chocolate, vegetable juice, etc. Climacteric fruits and vegetables, etc. Fresh fish and meat products, etc. Wine, peanut butter, etc. Mushroom, tomato, strawberries, etc. Use carbon dioxide releasing or absorbing sachets inside packaging to regulate gas composition in packaging headspace In order to remove odoriferous byproducts in packaging headspace, a. Use sachets filled with active carbon, b. Incorporate antioxidants or other odor removers in packaging substrates, or c. Apply an antioxidant coating onto packaging substrates In order to maintain the product quality of moisture sensitive products, use a. Desiccant sachets, b. Packaging systems with humidity regulators, or c. Packaging materials with selective gas permeability Humidity-controlled 27 Table 2.2 (cont’d) Temperature-controlled Antimicrobial Antioxidant Seafood, sake, coffee, tea, ready meals, etc. Fresh and processed meat, fresh seafood, dairy products, etc. Seed, nuts, edible oil, fried products, fresh vegetables and fruits, etc. In order to maintain the product quality of temperature sensitive products, Use a. Self-cooling cans b. Special insulating materials c. Dry ice packs d. Self-heating packaging In order to prevent microbial growth in food products, a. Incorporate antimicrobial agents in packaging substrates, or b. Apply an antimicrobial coating onto packaging substrates In order to slow down food deterioration process and prevent food rancidity, a. Incorporate antioxidants in packaging substrates, or b. Apply an antioxidant coating onto packaging substrates 2.7.2.2 Antioxidant active packaging Compared with directly adding antioxidants in food products, incorporating antioxidants in packaging substrates or applying antioxidant coatings onto packaging substrates may use a smaller amount of antioxidants. When a packaging substrate starts releasing antioxidants to its food product, or an antioxidant coating is contacting with its packaged food product, the 28 antioxidant activity is initiated. To retard the deterioration process of the food products, free radicals in the food cells are scavenged by antioxidants to stop further deleterious chemical reactions. The potency of the antioxidant activity depends on various aspects, including the release rate of the antioxidants in the packaging substrate, the density of the antioxidant coating, the mechanisms of the antioxidants reacting with different types of free radicals, the water content of the food products, the lipid content of the food products, and environmental factors. Table 2.3 lists some examples of recent research on antioxidant packaging that can be applied for food products. Table 2.3 Selective examples of antioxidant packaging research for food products since 2010. Packaging Technology Tested Food Products Experimental Results Packaging Substrate Poly (lactic acid) film EVOH film Antioxidant(s) a-Tocopherol Natural flavonoids (quercetin and catechin) Twin screw extrusion (Antioxidant incorporation) Twin screw extrusion (Antioxidant incorporation) Soybean oil Fried peanuts and sunflower oil Reference Manzanarez- López et al. (2011) López de- Dicastillo et al. (2011) Retarded the oxidation process of soybean oil The flavonoids effectively reduced the radical oxidative species presented in both food products 29 Table 2.3 (cont’d) Chitosan film Green tea extract Pork sausages Solvent casting (Antioxidant incorporation) LDPE film LDPE film PET/PE/ EVOH/PE film Tocopherol mixture Natural phenolic compounds from brewery residual and rosemary extract Natural oregano essential oil or solid green tea extract Salmon (Salmo salar) Twin screw extrusion (Antioxidant incorporation) Beef Antioxidant coating Antioxidant coating Foal meat 30 The green tea extract enhanced the antioxidant and antimicrobial properties of the chitosan film Effectively conserved the salmon samples for long-term storage The coated film enhanced the oxidative stability of the meat product Siripatrawan et al. (2012) Barbosa- Pereira et al. (2013) Barbosa- Pereira et al. (2014) Lorenzo et al. (2014) The antioxidant coating film containing oregano essential oil presented better protection to retard oxidation, retained product color and odor Hauser et al. (2016) Battisti et al. (2017) (Battisti et al. 2017) The antioxidant and antimicrobial effectiveness of the coated film could last for 60 days. Plus, food off- flavor and off-odor issues were not noticed After four days of storage, the meat product had a lower microbial population, better oxidation stability, and presented desired red color Table 2.3 (cont’d) LDPE film Murta leaf extract in a methylcellulose layer Antioxidant coating Milk chocolate Raw paper sheets Citric acid in gelatin Beef Antioxidant coating 31 Wrona et al. (2017) Navikaite- Snipaitiene et al. (2018) Table 2.3 (cont’d) PE film Green tea extract Antioxidant encapsulation in extruded PE film Minced pork meat Oriented PP film Clove essential oil or eugenol Antioxidant coating Beef The antioxidant packaging extended the meat product shelf life for 3 days and preserved the red color of the pork meat During the 14 days of storage time, the antioxidant coating layer reduced the formation of oxidative compounds and protected the color of the meat product 32 Table 2.3 (cont’d) Bio-based (Polylactic acid) emitting sachets Eugenol, carvacrol, and trans- anethole Inserted the sachets into cellulose and PP pillow Fresh-cut iceberg lettuce PET/LDPE film Sage leaf or bay leaf extracts Added the antioxidants in the film adhesive Fried potatoes Eugenol and trans-anethole reduced discoloration of the lettuce, preserved sensory quality of the lettuce, and introduced formation and accumulation of phenolic compounds inside the packaging The antioxidant film effectively retarded the food oxidation process and decreased oxidative products Wieczyńska et al. (2018) Oudjedi et al. (2019) 2.8 Avocado 2.8.1 Avocado and its byproducts Avocado (Persea americana Mill.) is one of the most common fruits in the United States, originating from Central Mexico. Among the hundreds of varieties of avocados, Fuerte and Hass are the most common varieties in the food market. Because of the high content of bioactive 33 compounds (including vitamin C, vitamin E, carotenoids and phenolic compounds (Antasionas, Riyanto, and Rohman 2017)) in its fruit pulp, avocado has been recognized as a functional fruit providing great benefits for human health. Based on clinical study results, consuming avocados aided the healthy aging process, benefited cardiovascular health, and helped cholesterol level management in the human body (Dreher and Davenport 2013). Nowadays, avocado, a highly nutritious fruit, has become widely known around the world. Its production has spread from Mexico to the United States, Australia, South Africa, and Spain (Rodríguez-Carpena et al. 2011). From 1994 to 2004, the global production volume of avocado dramatically soared from 4.6 billion pounds to 6.8 billion pounds. The United States is ranked as the second largest avocado producer in the world, following Mexico. The fruit growers can be found in California, Florida and Hawaii. Every year, California alone is estimated to produce 400 million pounds of avocados (Dreistadt 2007). These huge numbers imply a large quantity of avocado waste could be generated every year, including spoiled avocado, and avocado seed and peel left by human consumption and the food processing industry. Thus, handling the fruit waste requires substantial costs for food waste management. If additional value of the food waste could be added to avocado, the food industry would show further interest in this crop. This would also bring economic benefit to the food growers and manufacturers. 34 2.8.2 Additional value of avocado byproducts Avocado seed and peel are ideal sources of natural phenolic compounds, which belong to the major group of natural antioxidants. According to study results from food safety and food science fields, the peels and seeds of Fuerte and Hass avocados contain significantly higher phenolic contents than the avocado pulps, and the phenolic extract from these avocado byproducts presented robust antioxidant capability to inhibit oxidation reactions. (Rodríguez- Carpena et al. 2011; Soong and Barlow 2004; Torres, Mau-Lastovicka, and Rezaaiyan 1987; Wang, Bostic, and Gu 2010). Avocado seed and peel are rich in a mixture of phenolic compounds. The prevalent natural phenolic compounds, such as catechin, epicatechin gallate, procyanidin, chlorogenic acid, protocatechuic acid, syringic acid, rutin, and quercetin, all exist in avocado seed and peel (Kosińska et al. 2012; Pahua-Ramos et al. 2012; Tremocoldi et al. 2018). Table 2.4 and Figure 2.1 present primary phenolic compounds and their contents in Hass avocado byproducts. The phenolic content was determined based on dry weight (DW). In addition to the antioxidant property, avocado byproducts also have anti-inflammatory and analgesic properties (Kristanti et al. 2017; Tremocoldi et al. 2018). These great potentials have aroused interest from the pharmaceutical industry. Researchers can use them as reliable sources to prevent inflammatory diseases. 35 Table 2.4 Primary phenolic compounds and their contents in Hass avocado byproducts as reported in literature (Kosińska et al. 2012; Pahua-Ramos et al. 2012; Tremocoldi et al. 2018). Content (µg/g DW) Peel 148.80 ± 5.95 40.21 ± 0.24 135.40 ± 7.44 81.80 ± 5.95 55.10 ± 4.46 31.20 ± 4.46 23.80 ± 2.98 N/A N/A N/A Seed 152.80 ± 14.60 10.27 ± 0.08 ( with epicatechin gallate) N/A N/A N/A N/A N/A 128.18 ± 0.01 107.42 ± 0.04 89.30 ± 9.73 Compound Name Type of Byproduct Catechin Epicatechin Procyanidin dimer B (I) 5-O-caffeoylquinic acid Procyanidin dimer B (II) Quercetin 3-O-galactoside Quercetin-3-O-rutinoside (rutin) Protocatechuic acid Kaempferide Procyanidin trimer A (II) Peel & Seed Peel & Seed Peel Peel Peel Peel Peel Seed Seed Seed 36 Table 2.4 (cont’d) Procyanidin trimer A (I) 3-O-Caffeoylquinic acid Vanillic acid Seed Seed Seed N/A N/A N/A 81.70 ± 6.49 57.50 ± 6.49 28.67 ± 0.001 Catechin Epicatechin 5-O-caffeoylquinic acid Procyanidin dimer B (I) Figure 2.1 Chemical structure of primary phenolic compounds in Hass avocado byproducts. 37 Figure 2.1 (cont’d) Procyanidin dimer B (II) Quercetin 3-O-galactoside Quercetin-3-O-rutinoside (rutin) Epicatechin gallate 38 Figure 2.1 (cont’d) Protocatechuic Acid 3-O-Caffeoylquinic acid 39 Kaempferide Vanillic acid 2.8.3 Applications of avocado byproducts in the packaging field To date, there is no information available in the literature discussing applications of avocado byproducts in the packaging field. However, in 2012, a Mexican company named Biofase claimed that they developed a biodegradable plastic by using avocado waste (Anon 2012). According to the chemical engineer and founder of Biofase, around 30,000 tons of avocado seeds are discarded by the Mexican industry each month. In most cases, these avocado seeds were directly burned at landfill sites. The biomaterial offered an environmentally friendly solution to improve the situation. No further information was provided to show that this technology had been put into production. Experimental data in terms of the chemical and physical properties of the biomaterial were missing. 2.9 Phenolic compounds Phenolic compounds refer to chemical compounds with one or more hydroxyl groups attached to a carbon atom of an aromatic ring. A phenolic compound should have at least one aromatic ring. There are more than 8000 types of phenolic compounds with different chemical structures that have been identified (Martínez-Valverde, Periago, and Ros 2000). They vary from a single-aromatic ringed chemical with low molecular weight to a complicated polyphenol with multiple aromatic rings. Based on the number of aromatic rings and the carbon atom arrangement, phenolic compounds can be grouped into 9 categories. They are phenolic acids, 40 acetophenones, phenylacetic acid, hydroxycinnamic acids, coumarins, naphthoquinones, xanthones, stilbenes, and flavonoids (Crozier, Jaganath, and Clifford 2007). Phenolic compounds utilize their redox properties to serve as antioxidants. While reacting with oxidative chemicals, they can donate electrons, transfer hydrogens, and chelate metal ions such as iron and copper (Bendary et al. 2013; Estévez et al. 2008; Zhuravlev et al. 2016). These different reaction mechanisms allow phenolic compounds to exhibit strong antioxidant capability. 2.9.1 Applications of natural phenolic compounds in the packaging field Because of the remarkable interest in the applications of natural antioxidants from the food industry, natural phenolic compounds, as the predominant phytochemicals, have been evaluated by a variety of packaging researchers. Based on the experimental results, the biochemicals exhibit robust antioxidant capability to scavenge free radicals in tested samples, slow down or prevent the browning process of food products, and maintain food taste and texture. In view of the stability and solubility of natural phenolic compounds, researchers recommended natural phenolic compounds as trustworthy biochemicals to protect food products from oxidation and rancidity. Natural phenolic compounds are easy to obtain. Extracts from many herbs, spices, vegetables, and fruits contain phenolic compounds. Määttä-Riihinen et al. (2004) identified and 41 quantified the phenolic compounds in berries. Rusak et al. (2008) and Yang et al. (2012) evaluated the phenolic contents of different types of tea plants. Roby et al. (2013) tested the total phenolic compounds in thyme, sage, and marjoram. Cheng et al. (2013) examined the antioxidant efficacy of phenolic compounds in red and yellow onions. There is a broad range of food products that can be protected by natural phenolic compounds. The phytochemicals can be utilized for red meat, fish, dairy products, beverages, edible oils, etc. (L. H. Skibsted 2010). Table 2.5 lists several examples of the applications of natural phenolic compounds in food packaging. 42 Table 2.5 Examples of applications of natural phenolic compounds in food packaging. Phenolic Sources Grapefruit extract Barley husk extract Green tea extract Grape seed extract Oregano extract Packaging Type A bio-based film layer containing the antioxidants Antioxidant coating on PE film Packaging film containing antioxidants Modified atmosphere packaging Antioxidant coating on PP film Food Application Pork loins Atlantic salmon (Salmo salar L.) All type of foods, from aqueous to fatty products Pork patties Beefsteak Reference Hong et al. (2009) Pereira de Abreu et al. (2010) López de Dicastillo et al. (2011) Kumar et al. (2015) Djenane et al. (2016) 43 CHAPTER THREE: Crude Extraction of Phenolic Compounds from Avocado Byproducts 3.1 Introduction For the crude extraction of phenolic compounds from food or food byproducts, researchers commonly utilize methanol, ethanol, acetone, and ethyl acetate as the solvents. Both pure and aqueous mixtures (DI water and solvent mixture) can be used. Normally, 70% aqueous mixtures are used for extraction to recover phenolic compounds with desired antioxidant efficacy from food or food byproducts. Solvent polarity is one key factor that determines the concentration and antioxidant efficacy of phenolic extracts from food or food byproducts (Antasionas et al. 2017; Naczk and Shahidi 2006). In addition, the solubility of extracted phenolic compounds in solvents also affects the phenolic content recovered from food or food byproducts (Alothman, Bhat, and Karim 2009). While extracting phenolic compounds from food or food byproducts, temperature is another important aspect that needs to be considered. Moderate heat could assist the extraction, and thus increase phenolic content. However, when the temperature is elevated to a certain level, decomposition of phenolic compounds may be introduced, causing the reverse effect (Liyana- Pathirana and Shahidi 2005; Tan, Tan, and Ho 2013). 44 The experimental conditions for phenolic extraction from different food or food byproducts are not universal. In addition to solvent polarity, the solubility and polarity of extracted phenolic compounds and extraction time are other critical factors that need to be considered. Based on the literature, extraction time varied based on the types of food and food byproducts. Alothman et al. (2009) used 3 hours to obtain phenolic compounds from honey pineapple (Ananas comosus Merr.), banana (Musa paradasiaca) and guava (Psidium guajava L.). Lafka et al. (2007) changed the extraction time from 30 minutes to 24 hours to obtain phenolic content from winery wastes. Ajila et al. (2010) used 15 minutes to complete the extraction process. To extract phenolic compounds from avocado byproducts, Folasade et al. (2016) used 48 hours to allow the solvent to completely react with avocado seed powder. Calderón-Oliver et al. (2016) shortened the time to 30 minutes to extract phenolic contents from avocado seed and peel powders. Due to the acceptability for human consumption and the solubility of a wide range of phenolic compounds, ethanol was selected as one solvent for the phenolic extraction from avocado byproducts. Moreover, to understand the influence of different solvents with different polarities on the antioxidant effectiveness of coated films, acetone, as a most efficient solvent for phenolic extraction reported by Alothman et al. (2009) and Folasade et al. (2016), was also 45 selected for this research. Trial-and-error was used to determine the byproduct quantity, extraction time, and solvent temperature for the extraction process of this research. 3.2 Materials and methods 3.2.1 Byproducts preparation Raw avocado seed powder was purchased from Addicted 2 Healthy Nutritional SuperFoods, LLC. The powder was stored under refrigeration at 4 °C until use. Fresh Hass avocado (product of Mexico) peels were supplied by local sushi restaurants. They were first cleaned using tap water to remove dust particles and other contaminants on the peel surface, and air dried at room temperature for 12 hours. To completely remove the moisture content, the avocado peels were then put in a freeze dryer for another 12 hours. Afterwards, the peels were ground into small particles using an Eberbach E3300 mill (Belleville, MI, U.S.A.) with a number 40 mesh sieve. The peel particles passing through the sieve were around 0.42 mm in diameter. Finally, the ground peels were stored in plastic jars with screw-on lids under refrigeration at 4 °C until use. In order to control the relative humidity surrounding the ground peels, the plastic jars were placed inside Ziploc bags together with desiccant sachets. 46 3.2.2 Extraction of phenolic compounds 3.2.2.1 Materials Ethanol (200 proof) was purchased from VWR International (Radnor, PA, U.S.A.). Acetone and 250 mL SigmaÒ filter systems were purchased from Sigma-Aldrich Corporation (St. Louis, MO, U.S.A.). The acetone and ethanol were of reagent grade. 3.2.2.2 Methods Aqueous solvent (either 70% ethanol or 70% acetone/DI water) was prewarmed at 40 °C. One-gram avocado byproduct powder (either seed or peel powder) was then added into 10 mL of the aqueous solvent. Afterwards, the powder-solvent mixture was mixed using a Vortex mixer for 30 minutes, and a Fisher ScientificÒ ultrasonic cleaner (Pittsburgh, PA, U.S.A., model: FS30D) for another 45 minutes. Next, an Eppendorf centrifuge (Hamburg, Germany, model: 5804 R) at 3000 rpm was used for 10 minutes at room temperature to collect supernatant (crude phenolic extract) from the powder-solvent mixture, and the supernatant was filtered using a SigmaÒ filter system with a 0.22 µm pore size polyethersulfone (PES) membrane. Each gram of avocado byproduct powder was extracted twice, and the supernatants were combined. Finally, solvent contained in the filtered supernatant was evaporated using an IKAÒ RV10 rotary evaporator (Staufen im Breisgau, Germany) at 30 °C, and the phenolic extract was stored in a dark bottle under refrigeration at 4 °C until use. 47 3.3 Phenolic compounds in the crude extracts of Hass avocado byproducts 3.3.1 Materials The acetonic peel and seed extracts obtained from section 3.2.2 were evaluated for phenolic compound identification and quantification. Mass spectrometry standards used for the phenolic compound quantification analysis were purchased from Sigma-Aldrich Corporation (St. Louis, MO, U.S.A.). All the standards were of reagent grade. 3.3.2 Methods This experiment was performed by the Mass Spectrometry Research Technology Support Facility at Michigan State University. Phenolic compounds in both peel and seed extracts were identified and quantified using liquid chromatography/mass spectrometry (LC/MS) on a Waters Xevo® G2-XS QTof (Manchester, UK) interfaced to a Waters I-class Acquity solvent delivery system. Compounds were separated on a Waters HSS-T3 column (2.1 x 100 mm, held at 40˚C) using a gradient based on 10 mM ammonium formate adjusted to pH 2.8 using formic acid (Solvent A) and acetonitrile (Solvent B) at a total flow rate of 0.40 mL/minute. Solvent gradient was as follows (%A/%B): initial: (98/2); hold at (98/2) until 4.0 minutes, linear gradient to (90/10) at 8.0 minutes, (75/25) at 20 minutes, (5/95) at 32 minutes, with a hold until 37 minutes, followed by return to initial conditions. Mass spectra were acquired using electrospray ionization in negative-ion mode, with 48 a capillary voltage of -2.0 KV. Centroided mass spectra were acquired over m/z 100-1500 at 0.3 seconds/spectrum. Data were acquired using MSE (alternating low- and high-energy collision conditions, collision potential was ramped from 20-80 V during high energy spectrum acquisition). Argon was used as collision gas at a manifold pressure of 1.2 x 10-1 mbar. Leucine enkephalin was used as lock mass with real-time correction of ion masses. The Avocado byproduct extracts were diluted in Milli-Q water at the ratio of 1:10 before analysis, and injections of 10 µL were made for each sample using a Waters 2777 autosampler (Milford, MA, U.S.A.). Two cocktails of external standards were analyzed to generate calibration curves. Post-acquisition data analysis was performed using Waters QuanLynx software, integrating peak areas for [M-H] - ions. Quantitative analyses for those analytes for which authentic standards were not available, e.g. catechin isomers, were quantified using the assumption that response factors were the same as the standard with greatest structural similarity (A. Jones, personal communication, Nov. 14, 2019). 3.3.3 Phenolic compound identification and quantification results As shown in Table 3.1 and Figure 3.1, there were 28 types of phenolic compounds identified from the avocado peel and seed extracts. The identification results, in general, agreed with the mass spectral analyses reported by Kosińska et al. (2012), Pahua-Ramos et al. (2012) and Tremocoldi et al. (2018). Based on the literature, there were only 2 out of 13 types of 49 phenolic compounds could be identified from both avocado peel and seed extracts, namely, catechin and epicatechin. However, the Mass Spectrometry Facility reported that 22 out of 28 types of phenolic compounds could be found in both byproduct extracts utilized for this research. The remaining 6 types of phytochemicals were presented in only one of the extracts. According to the quantification result of each phenolic compound in the mass spectra (Figure 3.1), procyanidin dimer B (I), catechin isomer 2, procyanidin trimer 2, 5-O-caffeoylquinic acid and procyanidin trimer 1 were ranked as the top five phenolic compounds in the avocado peel extract, and the avocado seed extract was rich in oxidized procyanidin trimer 2, followed by catechin isomer 2 and oxidized procyanidin trimer 1. 50 Table 3.1 Phenolic compounds identified and quantified from the crude extracts of Hass avocado byproducts. Content (µg/mL) Seed 48.3 273.0 75.9 8.2 16.0 66.4 4.9 40.8 13.5 33.2 Compound Name Type of Byproduct Catechin isomer 1 Catechin isomer 2 Procyanidin dimer B (I) Procyanidin dimer B (II) Procyanidin trimer 1 Procyanidin trimer 2 Procyanidin trimer 3 Alternate procyanidin dimer 1 Alternate procyanidin dimer 2 Alternate procyanidin dimer 3 Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed 51 Peel 10.3 1101.3 1516.0 88.5 181.0 950 71.9 3.7 0.47 25.6 Table 3.1 (cont’d) Oxidized procyanidin trimer 1 Oxidized procyanidin trimer 2 Protocatechuic acid 3-O-caffeoyl quinic acid 4-O-caffeoyl quinic acid 5-O-caffeoyl quinic acid 5-O-p-coumaroyl quinic acid Quercetin 3-O- galactoside Quercetin 3-O-glucoside Quercetin-3-O- rutinoside (rutin) Quercetin rhamnoside hexoside 2 Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed Peel & Seed 52 5.9 30.6 1.5 0.16 4.2 425.8 0.46 3.5 1.3 16.9 1.0 256.0 288.0 4.9 5.7 139 7.0 8.1 0.32 0.38 0.19 0.36 79.7 N/A N/A 25.1 37.0 13.3 31.3 Table 3.1 (cont’d) Quinic acid Kaempferol Kaempferol dihexoside Caffeoyl quinic acid isomer 4 Catechin isomer 3 3-O-p-coumaroyl quinic acid 4-O-p-coumaroyl quinic acid Peel & Seed 130.0 0.01 3.7 N/A N/A N/A N/A Peel Peel Seed Seed Seed Seed 53 Figure 3.1 Phenolic compounds identification and quantification results for the crude extracts of Hass avocado byproducts. 54 54 CHAPTER FOUR: Non-metal Contact Coating Process 4.1 Introduction Phenolic compounds can be coated onto material surfaces via an oxidative polymerization reaction (Roman, Decker, and Goddard 2016). Due to the significant similarities of the chemical structures between phenolic compounds and mussel foot proteins (Mfps), phenolic compounds, after polymerization, present adhesive properties (Sileika et al. 2013). Mfps are widely known because of their durable and strong adhesive properties (Kord Forooshani and Lee 2017). These bio-based proteins enable marine mussels to stick to foreign surfaces in wet, dry and salty environments (Lee, Lee, and Messersmith 2007; Waite 1987; Zhao et al. 2006). It is believed that 3,4-Dihydroxyphenylalanine (DOPA), as an amino acid in Mfps, plays a vital role in the adhesive actions. A DOPA molecule contains a catechol side chain (see Figure 4.1). This adhesive moiety can be involved in a) redox activities, b) metal chelating reactions, c) cross-linking actions, and d) interfacial activities (covalent and noncovalent) to allow Mfps to tightly bind to all types of materials under water, such as glass, plastics, and metal oxides (d’Ischia and Ruíz-Molina 2017; Lu et al. 2013; Mian and Khan 2017). After the adhesion on foreign surfaces, marine mussels are almost motionless. It is hard to remove them. 55 According to the literature, the tenacity (a size independent detachment force) of a California mussel (Mytilus Californianus) in the perpendicular direction was up to 300 N and in the parallel direction was 180 N (Mian and Khan 2017). Catechol (adhesive moiety) Figure 4.1 Chemical structure of a DOPA molecule. Phenolic compounds contain catechol and/or gallol (1,2,3-trihydroxyphenyl, see Figure 4.2 and Figure 2.1 for examples on phenolic compounds) functional parts. Similar to DOPA, these side chains in the antioxidant molecules have interfacial binding properties and thus serve as strong and versatile adhesive moieties attaching phenolic compounds to various foreign surfaces. The foreign surfaces can be either organic or inorganic, such as metals, ceramics, plastics, glass, and biological materials (Barrett, Sileika, and Messersmith 2014; Forooshani, Meng, and Lee 2017; Sileika et al. 2013; Zhan et al. 2017). 56 (cid:5)(cid:3)(cid:1) (cid:5)(cid:3)(cid:1) (cid:5)(cid:3)(cid:1) (cid:5)(cid:3)(cid:1) (cid:5)(cid:3)(cid:1) Catechol side chain Gallol side chain Figure 4.2 Chemical structures of a catechol and a gallol side chain. Catechol has strong adaptability to foreign surfaces. Based on the catechol content in phenolic compounds, the nature of substrates, and the pH value of surrounding environment, catechol-contained compounds interact with foreign surfaces in four ways, i.e., a) hydrogen bonding, b) coordination (monodentate, bidentate, and chelating bidentate), c) p - surface interaction (p - p and p - cation interaction) and d) covalent bonding via Michael-type addition (see Figure 4.3, Andersen, Chen, and Birkedal 2019; Saiz-Poseu et al. 2019). The same interfacial activities were reported between gallol-containing compounds and foreign surfaces. In addition to bidentate coordination, tridentate coordination was noticed. Compared with bidentate coordination, tridentate interfacial activity resulted in stronger binding strength (Zhan et al. 2017). 57 (cid:6)(cid:1) (cid:3)(cid:5)(cid:1) (cid:5)(cid:3)(cid:1) Substrate a) Hydrogen bonding (cid:6)(cid:1) (cid:6)(cid:1) (cid:5)(cid:1) (cid:5)(cid:3)(cid:1) (cid:5)(cid:1) (cid:5)(cid:1) Substrate Substrate (cid:6)(cid:1) (cid:5)(cid:1) (cid:5)(cid:1) (cid:4)(cid:1) Metal/metal oxide b) Monodentate Bidentate Chelating bidentate Figure 4.3 Schematic of interfacial activities between catechol-contained compounds and foreign substrates. R represents the remainder of a catechol-containing molecule, M represents a metal atom in a substrate, X+ represents a cation in a substrate. 58 Figure 4.3 (cont’d) (cid:7)(cid:2)(cid:1) c) p - p interaction p - cation interaction (cid:3)(cid:5)(cid:1) (cid:6)(cid:1) (cid:5)(cid:3)(cid:1) Substrate d) Covalent bonding (via Michael-type addition) There are two mechanisms suggested in the literature as coating methods for phenolic compounds: laccase assisted enzymatic polymerization (Jeon et al. 2010, 2013) and alkaline saline assisted oxidative polymerization (Geißler et al. 2016; Sileika et al. 2013). For both methods, oxygen involvement and moderate mechanical agitation are required. These two approaches utilize coating solutions with different pH values and different catalysts to 59 polymerize phenolic compounds onto flexible films. For the laccase assisted coating method, researchers proposed adding laccase obtained from Trametes versicolor into an acid buffer solution (pH = 5) to expedite the phenolic compound polymerization process, whereas the alkaline saline assisted method used an alkaline buffer solution (pH = 7.8) to complete the phenolic compound polymerization process. In the presence of sodium chloride, the coating efficiency of the alkaline saline method could be enhanced. Considering the requirement of enzymes for the laccase assisted approach, the coating cost is much higher than that of the alkaline saline method. Roller coating is a conventional process to apply a coating layer onto a flexible plastic substrate. Basically, it utilizes a metal roll to fully contact with the surface of a substrate, and thus physically transfer a coating layer from the roll to the substrate. The thickness of the coating layer and the coating speed are adjustable. Due to the high transfer efficiency and minimal labor requirement, roller coating is widely accepted. It is an ideal method for stable coating solutions that do not react with metals. In addition to free radical scavenging and electron-donating abilities, polymerized phenolic compounds (polyphenols) are metal chelators (Chew et al. 2008). They can be utilized to chelate ferrous, ferric, and cupric ions in different environments with various pH values. The binding abilities of the phytochemicals to different metals depend on their phenolic structure and 60 the location of their hydroxyl groups (Senevirathne et al. 2006; Thompson, Williams, and Elliot 1976). To avoid the metal chelating reaction happening during the coating process, conventional roller coating process was not considered as a feasible method for this research. A non-metal contact coating system with an agitation feature was developed. 4.2 Materials 4.2.1 Chemicals Sodium chloride, sodium hydroxide, bicine, and glacial acetic acid were purchased from VWR International (Radnor, PA, U.S.A.). Sodium acetate and laccase from Trametes versicolor were purchased from Sigma-Aldrich Corporation (St. Louis, MO, U.S.A.). All the chemicals were of reagent grade. 4.2.2 Sample films LDPE film was selected from the polymer films available in the School of Packaging (East Lansing, MI); the manufacturer was unknown. Biaxially oriented PP and oriented PET film were supplied by Dow Chemical Company (Midland, MI, U.S.A.). By using Dyne test pens, the surface energy of the PP and PET films were determined to be 30 and 34 dyne/cm, and the surface energy of the LDPE film was lower than 30 dyne/cm. The thickness of the LDPE, PP and PET films were 38.80 ± 0.45 µm, 20.00 ± 1.00 µm, and 14.20 ± 61 1.30 µm respectively, determined by averaging five measurements using a TMI digital micrometer (Ronkonkoma, NY, U.S.A., model number: 49-70-01-001). 4.2.3 Coating device Figures 4.4 and 4.5 present the schematics of the non-metal contact coating device developed for this research. It is a dip coating system. Basically, it utilizes a slider-crank mechanism to convert rotary motion to linear motion. There are two ways of linear motion designed for the coating process, i.e., horizontal and vertical movements. A sample film can be either horizontally moved inside the coating solution by using the connecting bar attached to the rotation bar and slider (Figure 4.4), or, it can be vertically moved if the connecting bar is removed (Figure 4.5). Both linear movements allow the involvements of oxygen from the atmosphere and moderate agitation during the coating process to meet the polymerization requirements. The movement’s frequency is determined by the speed of the electric motor underneath the rotation bar. In other words, the dip coating speed is a controllable variable. The diameters of different beakers used for coating are not the same and limit the moving range of a sample film inside the coating solution if horizontal movement is required. The connecting bar is designed to attach to different fitting holes on the rotation bar based on beaker 62 size to make sure the sample film movement is within the range, allowing different sizes of vessels to be used. For this research, the vertical movement design was employed based on the preliminary results of coating tests. To meet the agitation requirement for the coating process, in addition to using the sample holder to stir the coating solution while dip coating a sample film, the hot plate stirrer placed under the beaker can also agitate the solution while controlling the coating temperature. 63 A. Front View B. Top View 1 2 3 4 5 6 7 8 9 10 11 Hot plate stirrer Guides A Power switch Fitting holes Rotation bar Connecting bar Slider Beaker Electrical energy Electric motor Sample holder Figure 4.4 Schematic of the non-metal contact coating device with horizontal movement. 64 A. Front View B. Top View 1 2 3 4 5 6 7 8 9 10 11 12 Hot plate stirrer Guides A Power switch Fitting holes Rotation bar Electrical energy Electric motor Sample holder Slider Guide B Fixed guide Beaker Figure 4.5 Schematic of the non-metal contact coating device with vertical movement. 65 4.3 Methods 4.3.1 Laccase assist coating (sodium acetate) Glacial acetic acid was used to adjust the pH value of 0.1 M sodium acetate buffer to 5. Then, the crude phenolic extract (100% peel extract, 100% seed extract, or 50% peel extract and 50% seed extract) was dissolved into the buffer solution at a concentration of 25 mg/mL, followed by adding 1 mg/mL laccase. Afterwards, a 4.5 ´ 2 cm sample film was rinsed with DI water, purged by pure nitrogen flow, and attached to the sample holder of the coating device for 24-hour dip coating at 23 °C. To meet the mild agitation requirement of the coating process, the dipping speed of the coating device (the speed of the electronic motor) was 25 rpm, and the coating solution stirring speed of the hot plate (Scilogex LLC, Rocky Hill, CT, U.S.A., model: MS-H280-Pro) was set at 350 rpm. Finally, the coated sample film was detached from the sample holder, rinsed with DI water, and purged by pure nitrogen flow again to remove any residue on the film surface. 4.3.2 Alkaline saline coating (bicine) This coating method was based on the experimental design proposed by Geißler et al. 2016. In essence, the coating buffer was made by mixing 0.1 M bicine (end concentration) with 0.6 M NaCl (end concentration). The pH value of the buffer solution was then adjusted to 7.8 using NaOH. To make the final coating solution, the crude phenolic extract (100% peel extract, 66 100% seed extract, or 50% peel extract and 50% seed extract) was added into the buffer solution at a concentration of 25 mg/mL. Afterwards, a 4.5 ´ 2 cm sample film was rinsed with DI water, purged by pure nitrogen flow, and attached to the sample holder of the coating device for 24- hour dip coating at 23 °C. To meet the mild agitation requirement of the coating process, the dipping speed of the coating device (the speed of the electronic motor) was 25 rpm, and the coating solution stirring speed of the hot plate was set at 350 rpm. Finally, the coated sample film was detached from the sample holder, rinsed with DI water, and purged by pure nitrogen flow again to remove any residue on the film surface. 4.3.3 Sample preparation for scanning electron microscope (SEM) observation To understand the distribution of the antioxidant coating on the substrates, coated LDPE, PP and PET films were used for surface and cross-sectional SEM observations, and a coated glass coverslip (diameter: 12 mm, # 1) was also used for coating thickness measurement. All the substrates were coated with the acetonic avocado peel extract by following the alkaline saline coating method stated in section 4.3.2. The reason for choosing this coating method will be discussed in chapter 5. 67 4.3.3.1 Sample preparation for surface observation At room temperature, a coated polymer film was cut into 1 ´ 1 cm samples with a utility knife, and then attached to an aluminum SEM specimen stub (diameter: 25 mm) using clear epoxy adhesive (see Figure 4.6). Coated PP film SEM specimen stub Figure 4.6 SEM sample preparation for surface observation of a coated polymer film. 4.3.3.2 Sample preparation for cross-sectional observation and thickness measurement To minimize deformation of the antioxidant coating caused by the sample preparation process, liquid nitrogen was used to temporarily freeze the coated substrates and a SEM specimen stub. As shown in Figure 4.7, a 25 mm stub was first placed into liquid nitrogen for 2 ~ 3 minutes; then, a coated polymer film was placed into liquid nitrogen for around 30 seconds. After that, the SEM stub and coated PP film were removed from the liquid nitrogen. A new razor blade was used to immediately cut the temporarily frozen film into small samples on the 68 temporarily frozen SEM stub. Finally, the sample films were vertically attached onto a 12.5 mm SEM specimen stub by using clear epoxy adhesive. For a clear cross-sectional view under SEM, a new razor blade was used for each cut. The coated glass coverslip was treated in the same way. Rather than using a razor blade, the frozen coverslip was manually fractured into small pieces after removing it from the liquid nitrogen. After curing for 15 hours the epoxy adhesive, all the SEM samples were added a thin layer of iridium as the conductive coating. The SEM photos for both surface and cross-sectional observations were obtained by placing the coated films and glass coverslips into the chamber of a JEOL JSM 7500F (JEOL Ltd., Tokyo, Japan), and visualized under an accelerating voltage of 5 kV. 69 Step 1: Step 2: Step 3: Step 4: Liquid nitrogen A SEM specimen stub A coated polymer film New razor blade The coated polymer film from step 2 (temporarily frozen) The SEM specimen stub from step 2 (temporarily frozen) A new SEM specimen stub The coated polymer films from step 3 A new SEM specimen stub Figure 4.7 SEM sample preparation for cross-sectional observation of a coated polymer film. 70 4.4 Antioxidant coating layer under SEM Figures 4.8, 4.9 and 4.10 illustrate the surface SEM images of the antioxidant coating on a PP, LDPE and PET film, respectively. The cracked texture was caused by the conductive coating layer, iridium, at a high level of resolution. As reflected in the images, each coating surface presented in almost the same grayscale. In addition, no charge-contrast spots, air bubbles, gaps, pinholes and fisheyes were noticed. These phenomena implied that the phenolic compounds from the avocado peel extract did not form small clumps on the film surface during the alkaline saline coating process; instead, they were uniformly distributed. This finding was later supported by the cross-sectional images of the coated polymer films (Figures 4.11, 4.12 and 4.13). While observing the surface characteristic of the coated LDPE film, the coating layer actively interacted with the electron beam of SEM, resulting in wrinkles on the film surface within 5 seconds. A better preparation method should be developed for surface observation of the coated LDPE film to avoid sample deformation. While observing the antioxidant coating from its cross-sectional view, the SEM images did not exhibit a sharp and clear distinction between the substrates and the coating layer. Possibly, the razor blade, when cutting the coated films during the SEM sample preparation process, compressed the coating layer and thus caused deformation of the coating on its cross- sectional surface. Based on Figures 4.11, 4.12 and 4.13, the deformation caused the edge of the phenolic coating to stretch on its substrate. In this circumstance, measuring the coating thickness 71 was challenging. Therefore, a glass coverslip was used to replace the polymer films as the coating substrate. Under SEM, a clear edge line was found between the coating layer and the glass coverslip (Figure 4.14). The phenolic compounds were evenly polymerized on the glass coverslip, which indicates a great potential of using other types of materials as the substrate coated with the avocado peel extract by employing alkaline saline coating method. By averaging the coating thickness at 4 different locations, the coating thickness was determined to be 37.75 ± 0.30 nm. It was noticed that the SEM sample preparation method for the coated glass coverslip resulted in deformation at some spots of the coating layer. At high magnification (35000x), clumps were observed at some spots of the coating layer surface. This deformation might be introduced by the SEM sample preparation process. After removal from liquid nitrogen, the coated glass coverslip became very brittle. Manually fracturing the sample, in this case, might result in deformation at some spots of the coating layer. Therefore, a better preparation method should be developed for thickness measurement of future samples to avoid sample deformation. 72 Figure 4.8 Topside view of the phenolic coating on a PP film. Figure 4.9 Topside view of the phenolic coating on a LDPE film. 73 Figure 4.10 Topside view of the phenolic coating on a PET film. A ntioxidant coating Figure 4.11 Cross-sectional view of the phenolic coating on a PP film. 74 A ntioxidant coating Figure 4.12 Cross-sectional view of the phenolic coating on a LDPE film. i n g c o a t i o x i d a n t A n t Figure 4.13 Cross-sectional view of the phenolic coating on a PET film. 75 g xi d a n t c o ati n n ti o A Figure 4.14 Cross-sectional view of the phenolic coating on a glass coverslip. To uniformly attach phenolic compounds from the avocado byproduct extracts onto nonpolar LDPE and PP films, and PET film with low surface energy (34 dyne/cm), two types of interfacial reactions might be initiated during the coating process, namely, covalent bonding via Michael-type addition and p - surface interaction. For the LDPE and PP films, covalent linkage (Michael-type addition) could occur between catechol functional group(s) of the phenolic compounds and the film surfaces. In the presence of oxygen, the adhesive moiety of phenolic compounds could be firstly oxidized to quinone form. After that, carbons in the polymeric chains of LDPE or PP might serve as nucleophiles to react with the oxidized adhesive moiety in the wet environment. In this way, the 76 phenolic compounds could finally attach to the film surfaces, see Figure 4.15. Similar mechanisms were reported by Saiz-Poseu et al. (2019) and Yang, Stuart, and Kamperman (2014). For the PET film, the interfacial reaction might not be limited to covalent linkage only. Considering the existence of aromatic rings in the polymeric chain of PET, p - surface interaction (Figure 4.16) might also be initiated during the coating process to allow the phenolic compounds to anchor on the film surface. (cid:6)(cid:1) (cid:6)(cid:1) Catechol oxidation (cid:6)(cid:1) (cid:3)(cid:5)(cid:1) (cid:5)(cid:3)(cid:1) LDPE or PP Wet environment (cid:3)(cid:5)(cid:1) (cid:5)(cid:3)(cid:1) (cid:5)(cid:1) (cid:5)(cid:1) quinone form (cid:6)(cid:1) (cid:3)(cid:5)(cid:1) (cid:5)(cid:3)(cid:1) Figure 4.15 Schematic of covalent linkage between catechol-contained phenolic compounds and a LDPE or PP film surface. R represents the remainder of a catechol-containing phenolic compound. 77 Figure 4.16 Schematic of p - surface interaction between a catechol-contained phenolic compound and a PET film surface. 78 CHAPTER FIVE: Evaluation of Antioxidant Activity of Coated Polymer Films Part I: Screening Tests 5.1 Introduction Before determining the potential applications of plastic substrates coated with the phenolic compounds extracted from avocado byproducts, three types of screening tests were employed to evaluate the antioxidant efficacy of the phenolic coating. By applying the laccase assist coating (section 4.3.1) and alkaline saline assist coating (section 4.3.2) methods, the phenolic content from avocado byproducts was attached onto PP, PET and LDPE films for inspection. Both the ethanolic and acetonic extracts from avocado peel and seed (section 3.2.2) were utilized. At the first stage, silver nitrate (AgNO3) solution was used for visually inspecting the existence of the phenolic coating. In contact with this solution, the phenolic coating acted as a reducing agent to convert Ag+ to Ag0, and excited surface plasmon resonance. The excitation, in this way, changed the coating to a dark yellow or brown color indicating the presence of phenolic content. At the second stage, a DPPH• assay was used. The phenolic coating served as an electron donor to stabilize the free radicals reducing the concentration of DPPH• in the working solution. 79 The potential influences of the coating environment, the concentration of phenolic extract in the coating solution, and different plastic substrates on the antioxidant effectiveness of the coating layer were investigated. An ABTS•+ assay, as the third evaluation method, was employed to confirm the experimental results obtained from the previous stages. In this reaction, the phenolic coating transferred hydrogen atoms to ABTS•+ to decrease the radicals in the working solution. The potential influences of the coating environment and different plastic substrates on the antioxidant efficacy of the coating layer were studied. In addition, the avocado peel and seed extracts were directly evaluated to understand their antioxidant potency to quench ABTS•+ radicals in the working solution. For both the DPPH• and ABTS•+ assays, food simulants (95%, 50%, and 10% aqueous ethanol) were used to analyze the antioxidant performance of the phenolic coating from avocado byproducts. 5.1.1 AgNO3 5.1.1.1 Introduction In the nanotechnology field, food and food byproduct extracts are utilized as reliable sources to synthesize silver nanoparticles (AgNPs). Compared with the chemical and physical synthesis methods, this biological technique provides an environment-friendly alternative to 80 easily obtain silver nanoparticles in a more economical way (Ibrahim 2015; Mohanpuria, Rana, and Yadav 2008). Silver nanoparticles are non-toxic and inorganic. They can serve as great antibacterial agents to kill around 650 disease-causing organisms in the body (S. H. Jeong, Yeo, and Yi 2005), as excellent sterilizers and UV-protectors on cotton fabric (Rai et al. 2014), and as a spectrally selective coating for solar energy absorption (Mohanpuria et al. 2008). The potential applications of this nanoparticle have been extensively studied in recent years. To synthesize AgNPs, food and food byproduct extracts need to react with aqueous silver nitrate solution. During this reaction, the extracts are used as reducing agents to convert Ag+ to Ag0, and thus cause metal deposition (Jeeva, Thiyagarajan, Elangovan, Geetha, & Venkatachalam, 2014). Other than metal deposition, the reduction process activates the excitation of surface plasmon resonance, resulting in color change of the solution (Mulvaney 1996; Sosa, Noguez, and Barrera 2003). Normally, a dark yellow or brown color can be observed indicating the existence of biosynthesized silver nanoparticles. There is no universal condition that works for all the biosynthesis reactions of silver nanoparticles. The silver ion reduction potential is mainly determined by reaction time, solution temperature, the concentrations of aqueous silver nitrate solution, and the volume of natural extracts. According to the literature, the reaction time used for the synthesis of AgNPs varied 81 from 5 minutes to 72 hours; the solution temperature varied from room temperature to 100 °C; the range of aqueous silver nitrate concentration was from 0.25 mM to 5.0 mM, and from 0.25 mL to 3 mL of natural extracts were used to reduce silver ions (Ibrahim 2015; Jeeva et al. 2014; Jeon et al. 2013; Sileika et al. 2013). In the natural extracts employed for the synthesis of silver nanoparticles, phenolic compounds were evaluated as effective ingredients. By utilizing phenolic compounds extracted from rice husk, Satureja intermedia C.A. Mey, and Ananas comosus, Liu et al. (2018), Firoozi et al. (2016), and Ahmad et al. (2012) reported the formation of AgNPs from the silver ion reduction test. This indicated the possibility of utilizing aqueous silver nitrate solution to inspect for the presence of phenolic compounds. Hence, aqueous silver nitrate solution was employed, for this research, to visually inspect for the presence of the phenolic compounds, which were extracted from avocado byproducts, in the coating layer of the sample films (based on color change of the coated films). Trial-and-error was used to determine the reaction time, solution temperature and concentration of the aqueous silver nitrate solution. 5.1.1.2 Materials Silver nitrate powder was purchased from VWR International (Radnor, PA, U.S.A.). This chemical was of reagent grade. 82 Ethanolic seed extract was utilized to coat PP, PET, and LDPE films by applying the laccase assist coating (section 4.3.1) and alkaline saline assist coating (section 4.3.2) methods. 5.1.1.3 Methods PP, PET and LDPE films coated with the phenolic layer were immersed into aqueous silver nitrate solution (100 mM) for at least 24 hours at room temperature. After that, the sample films were removed from the silver nitrate solution, rinsed with DI water, and purged by pure nitrogen flow. For comparison purposes, uncoated PP, PET and LDPE films were used as control samples and treated in the exact same way. Both the coated and uncoated films were photographed for evaluation. 5.1.1.4 Results and discussion After immersing the coated polymer films into silver nitrate solution, a slight color change was observed on the sample films after 24 ~ 48 hours. Starting from a light-yellow color, the coated sample films gradually changed to a brown or greyish brown color as time increased. This implied the deposition of silver nanoparticles on the surfaces of sample films, and the excitation of surface plasmon resonance resulted from the silver ion reduction process. For each coated sample film, the color evolution process did not start at the same time. Generally, the polymer films coated in the alkaline saline solution initiated the color change first. After 24-hour immersion, a noticeable light-yellow color was perceived on the surface of sample 83 films coated in the alkaline solution. On the other hand, it took at least 48 hours for the sample films coated in the laccase assist solution to start changing their color. At the end of the reaction, a strong color contrast was observed between the films coated in the alkaline solution and the laccase assist solution (Figure 5.1). Dark yellowish or greyish brown colors showed on alkaline solution coated sample films, while a light brown color presented on laccase solution coated sample films. This might imply that the alkaline saline solution, which is a comparatively low cost coating solution, could provide more potent phenolic coating on the sample films. Other evaluation methods were required to further support this statement. 84 Experimental Condition Control group Coated in laccase assist solution (pH = 5) Coated in alkaline saline solution (pH = 7.8) Film PP PET LDPE Figure 5.1 Color changes of polymer films coated with phenolic compounds from avocado byproduct extract. The coated films were reacted with silver nitrate solution for at least 24 hours. 85 5.1.2 DPPH• assay 5.1.2.1 Introduction DPPH is a chromogen. This organic chemical is normally utilized to detect the antioxidant efficacy of phenolic compounds. In order to evaluate the antioxidant property, aqueous DPPH• solution needs to be prepared by mixing DPPH powder into a solvent. When DPPH powder completely dissolves in the solvent, a purple color solution can be obtained indicating the formation of DPPH•. Then, phenolic compounds can be added into the DPPH• working solution to initiate the antioxidant activity. During this process, phenolic compounds locate DPPH• radicals in the solution, donate single electrons and/or hydrogen atoms to DPPH• radicals (Figure 5.2), and thus decrease the concentration of DPPH•. At the same time, the DPPH• solution gradually transforms to a yellow color, implying the end of the reaction. The color-change speed depends on the concentration of the phenolic compounds in the DPPH• working solution. The decrease in DPPH• concentration can be detected in a UV-Vis spectrophotometer at 517 nm and quantified by Equation 5.1. DPPH• inhibition capacity (%)= Abs56789:;<−Abs56789:;> Abs56789:;< ×100 (Equation 5.1) 86 Where BCDEFGHIJK< = The absorbance reading of DPPH• working solutions used for antioxidant BCDEFGHIJKM = The absorbance reading of DPPH• working solutions used for antioxidant test of coated films at time tL test of coated films at time tN, i=1,2…t To acquire an accurate understanding about the DPPH• inhibition ability of various phenolic compounds, researchers normally introduce and calculate EC50. EC50 refers to the effective concentration required for antioxidants to reduce 50% of the free radicals in a working solution. In order to determine this value, the DPPH• assay needs to be performed to establish a correlation between the total quantity of DPPH• radicals in the working solution (the UV-Vis absorbance readings of a DPPH• working solution) and the concentration of phenolic compound(s) added in the DPPH• working solution. Once the correlation is confirmed, EC50 can be determined by fitting the experimental results in different mathematical models, including logistic, Boltzmann sigmoidal and dose-response models (Suriyatem et al. 2017). For this research, coated sample films with the same surface area were tested for the evaluation of antioxidant efficiency. The goal was to understand the correlation between the total quantity of DPPH• radicals in the working solution and the effectiveness of the phenolic coating layer in eliminating DPPH• radicals. In other words, rather than the phenolic compound 87 concentrations studied in other research, the reaction time used for DPPH• elimination in this research was the parameter of interest. Therefore, ET50 was introduced, referring to the effective time required for the phenolic coating layer to reduce 50% of the free radicals in a working solution. The DPPH• assay is easy to prepare. It is an effective method to study the efficacy of phenolic compounds stabilizing free radicals. However, DPPH is insoluble in water (Stasko et al. 2007), the entire reaction process is time consuming, and it is expensive. These non-negligible factors limit its applications for various antioxidants in different environments. 88 DPPH• H • DPPH-H Where Ar : OH is a phenolic compound Ar : O• is a phenolic radical Figure 5.2 Single electron donation reaction between a DPPH• radical and a phenolic compound. 89 5.1.2.2 Materials DPPH powder was purchased from Sigma-Aldrich Corporation (St. Louis, MO, U.S.A.). Ethanol (200 proof) was purchased from VWR International (Radnor, PA, U.S.A.). These chemicals were of reagent grade. Ethanolic seed extract was utilized to coat PP, PET, and LDPE films by applying the laccase assist coating (section 4.3.1) and alkaline saline assist coating (section 4.3.2) methods. 5.1.2.3 Methods To make the DPPH• working solution, DPPH powder was dissolved in 95% aqueous ethanol (fatty food simulants) and 50% aqueous ethanol (simulating milk and products with high alcohol content) to obtain a concentration of 0.1 mM. Then, a 2 ´ 1.5 cm sample film (PP, PET, or LDPE) with the phenolic coating layer was immersed into 1 mL of DPPH• working solution to initiate the antioxidant activity at room temperature. At each predetermined time interval, the immersed sample film was temporarily removed from the working solution, and the absorbance reading of the DPPH• working solution was taken at 517 nm in a Shimadzu UV-Vis spectrophotometer (Kyoto, Japan, model: UV- 1800). After the measurement, the sample film was placed back in the working solution to allow further reaction. Light exposure was avoided during the entire electron donation reaction. The DPPH• working solution was freshly made every time before the analysis. 90 For comparison purposes, uncoated sample films immersed in DPPH• working solution were used as control samples and treated in the exact same way. 5.1.2.4 Results and discussion 5.1.2.4.1 Coating solution vs. antioxidant efficacy Sample films coated by the two different solutions did not behave in the same way to reduce DPPH• radicals in the working solutions. As shown in Figures 5.3 and 5.4, the free radical reduction process of LDPE films coated in the alkaline saline solution (pH = 7.8) was rapid. For both DPPH• tests, the alkaline solution coated LDPE films stabilized around 86% of the free radicals within 5 hours. To eliminate the same amount of DPPH• radicals, the laccase solution coated LDPE film in 50% aqueous ethanol required 9 hours. After 48 hours, the laccase solution coated LDPE film in 95% aqueous ethanol reduced only 60% of the free radicals in the working solution. Based on the ET50 values of the coated LDPE films in both 50% and 95% aqueous ethanol, the alkaline saline coating method was, again, suggested as a more effective approach than the laccase assist coating method. In 50% aqueous ethanol, the coated sample film used around 30 minutes to eliminate 50% of DPPH• radicals, which is at least 6 times faster than the laccase solution coated LDPE film. In 95% aqueous ethanol, the time difference increased to 93 times. 91 To further confirm the effectiveness of the alkaline saline solution, coated PET films were tested in 50% aqueous ethanol. As shown in Figure 5.5, the phenolic coating layer generated by the alkaline solution still displayed potent antioxidant ability. Similar to the alkaline solution coated LDPE films, the alkaline solution coated PET film presented a faster DPPH• reduction process. The ET50 of the laccase solution coated PET film was around 2.5 hours, which was 6.8 times longer than that of the alkaline solution coated PET film. Based on the above analysis, the alkaline saline solution resulted in a more active and efficient coating to stabilize DPPH• radicals in 50% aqueous ethanol (milk and high alcohol content food simulant) and 95% aqueous ethanol (fatty food simulant). This result agreed with the indications from the previous silver nitrate test. Since DPPH powder is insoluble in water, another assay was desired to understand whether the same experimental outcome could be obtained in a water-based food simulant (10% aqueous ethanol). 92 Figure 5.3 Antioxidant activity of LDPE films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The DPPH• solvent was diluted with 50% aqueous ethanol. Points a and b are estimates of the ET50 of LDPE (alkaline) and LDPE (laccase) respectively. Figure 5.4 Antioxidant activity of LDPE films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The DPPH• solvent was diluted with 95% aqueous ethanol. Points a and b are estimates of the ET50 of LDPE (alkaline) and LDPE (laccase) respectively. 93 Figure 5.5 Antioxidant activity of PET films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The DPPH• solvent was diluted with 50% aqueous ethanol. Points a and b are estimates of the ET50 of LDPE (alkaline) and LDPE (laccase) respectively. 5.1.2.4.2 Plastic substrates vs. antioxidant efficacy It is worth mentioning that there was no apparent time difference between the estimated ET50 of alkaline solution coated PP, LDPE and PET films in 50% aqueous ethanol (see Figure 5.6). These coated sample films exhibited similar antioxidant behavior. Within the first hour, all the alkaline solution coated sample films eliminated 80% of the DPPH• radicals in the working solution. 94 A small time difference was noticed between the estimated ET50 of laccase solution coated LDPE and PET films. The ET50 of laccase solution coated PET film was estimated at 2.5 hours, which was 0.8 hour faster than the LDPE film coated in the same type of solution. These experimental outcomes suggested that the polymer substrates did not exert a major influence on the antioxidant efficacy of the phenolic coating. Figure 5.6 Antioxidant activity of PET, LDPE and PP films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The DPPH• solvent was diluted with 50% aqueous ethanol. Point a is the estimate of ET50 of LDPE (alkaline), PET (alkaline) and PP (alkaline). Points b and b' are estimates of the ET50 of PET (laccase) and LDPE (laccase) respectively. 95 5.1.2.4.3 Concentration of phenolic extract vs. antioxidant efficacy In order to understand the correlation between the concentration of phenolic extract in the coating solution and the antioxidant efficacy of the coating layer, the alkaline solution coated PP films were tested. For the purpose of comparison, 12.5 mg/mL and 25 mg/mL ethanolic avocado seed extract were used in the coating process. As shown in Figure 5.7, no significant time difference was observed between the free radical inhibition process of the two samples. The alkaline solution with doubled seed extract appeared to slightly reduce the estimated ET50 but the difference was small. Within 1 hour, both coated films reduced DPPH• concentration in the working solution by at least 70%. Based on this finding, the excess phenolic content in the coating solution might not attach to the film surface during the coating process. Alternatively, provided the coating duration and phenolic content in the coating solution were sufficient, the antioxidant efficiency of the coating layer might be determined by substrate surface area rather than the amount of phenolic content in the coating solution. Further evaluation is desired to understand the minimum coating duration and phenolic concentration in the coating solution to obtain an adequate coating layer on different surface areas. 96 Figure 5.7 Antioxidant activity of PP films coated in solutions with different extract concentrations. The DPPH• solvent was diluted with 50% aqueous ethanol. The films were coated in alkaline saline (pH = 7.8) solution. Points a and b are estimates of the ET50 of PP films coated in solutions with 25mg/mL and 12.5mg/mL avocado seed extract. In conclusion, the DPPH• assay was employed at the secondary experimental stage to evaluate potential influences of the coating environment, plastic substrates and the concentration of phenolic extract in the coating solution on the antioxidant effectiveness of the phenolic coating layer. As was indicated by the silver nitrate test, the sample films coated in alkaline saline solution (pH = 7.8) presented greater antioxidant effectiveness in 50% aqueous ethanol 97 (milk and high alcohol content food simulant) and 95% aqueous ethanol (fatty food simulant). While testing the impact of plastic substrates and phenolic concentration in the coating solution on the free radical reduction process, significant variation was not observed. It seemed that the antioxidant efficiency of the coating layer was not linearly correlated to the concentration of phenolic extract in the coating solution. As long as enough phenolic content could be provided for substrate surface attachment, excess phenolic content in the coating solution did not appear to create a more potent coating layer. Further experiments are desired to understand the minimum coating duration and phenolic concentration in the coating solution to obtain an effective coating layer on various surface areas. In addition, another assay is desired to understand the antioxidant efficiency of the coating layer in water-based food simulant (10% aqueous ethanol) as DPPH powder is insoluble in water. 5.1.3 ABTS•+ assay 5.1.3.1 Introduction ABTS is another type of chromogen. It is a common chemical used for understanding the dynamic process of the antioxidant activity of phenolic compounds. In order to evaluate the antioxidant property, aqueous ABTS solution needs to be oxidized by potassium persulfate (K2S2O8) first to obtain ABTS•+ solution (Figure 5.8). During 98 the oxidation process, the solution gradually changes its color to dark blue-green indicating the formation of ABTS•+. After that, a small portion of diluted ABTS•+ solution is used to evaluate the antioxidant ability of phenolic compounds by mixing it with the phytochemicals (Figure 5.9). During this process, phenolic compounds locate ABTS•+ radicals in the solution, transfer their hydrogen atoms to ABTS•+ radicals (hydrogen abstraction of antioxidants), and thus decrease the concentration of ABTS•+ in the working solution. Under this circumstance, the ABTS•+ working solution gradually loses its blue-green color. The color-change speed depends on the concentration of the phenolic compounds mixed with the diluted ABTS•+ solution. The decrease of ABTS•+ concentration can be detected in a UV-Vis spectrophotometer at 734 nm and quantified by Equation 5.2. ABTS•U inhibition capacity (%)= Abs56789:;<−Abs56789:;> Abs56789:;< ×100 (Equation 5.2) Where BCDEFGHIJK< = The absorbance reading of ABTS•+ working solutions used for antioxidant BCDEFGHIJKM = The absorbance reading of ABTS•+ working solutions used for antioxidant test of coated films at time tL test reacted with coated films at time tN, i=1,2…t 99 ABTS•+ radicals react with antioxidants rapidly. Unlike DPPH•, this free radical can be dissolved in both aqueous and organic solvents, and the pH value of the working solution does not negatively impact the existence of ABTS•+ (Shalaby 2013). All these aforementioned facts enable ABTS•+ to be widely utilized as an efficient agent to evaluate the antioxidant effectiveness of phenolic compounds. For this test, coated sample films with the same surface area were tested for the evaluation of antioxidant efficiency. The aim was to understand the correlation between the total quantity of ABTS•+ radicals in the working solution and the effective time the phenolic coating layer used for ABTS•+ elimination. To acquire an accurate understanding about the ABTS•+ inhibition ability of the phenolic coating layer, and conduct valid comparisons among the ABTS•+ tests, ET50 was introduced again. This term refers to the effective time required for the phenolic coating layer to reduce 50% of the free radicals in a working solution. 100 ABTS ABTS•+ Figure 5.8 Oxidation reaction of ABTS with potassium persulfate to generate ABTS•+ radicals. 101 ABTS•+ ABTSH+ Figure 5.9 Hydrogen abstraction reaction of ABTS•+ with a phenolic compound. 5.1.3.2 Materials ABTS and potassium persulfate powders were purchased from Sigma-Aldrich Corporation (St. Louis, MO, U.S.A.). All the chemicals were of reagent grade. Before the experiment, ABTS powder was stored under refrigeration at 4 °C and protected from light until use. 102 Both ethanolic and acetonic extracts of avocado seed and peel were utilized to coat PP, PET, and LDPE films by applying the laccase assist coating (section 4.3.1) and alkaline saline assist coating (section 4.3.2) methods. 5.1.3.3 Methods ABTS•+ stock solution was made by mixing 7 mM aqueous ABTS solution with 2.45 mM aqueous potassium persulfate solution. The ratio of VWXYZ to V[\Z\]^ was 1:1. The mixture was then stored in a dark bottle and dark room at room temperature for 16 hours to allow completion of the oxidation reaction between ABTS and potassium persulfate. To make the ABTS•+ working solution, 1 mL ABTS•+ stock solution was diluted with around 60 mL 95% aqueous ethanol (fatty food simulant), 50% aqueous ethanol (simulant of high alcohol containing products and milk), or 10% aqueous ethanol (simulant of water-based food) to obtain an absorbance of 0.7 ± 0.05 at 734 nm in a Shimadzu UV-Vis spectrophotometer (Kyoto, Japan, model: UV-1800). After that, a 2 ´ 1.5 cm sample film (PP, PET, or LDPE) with the phenolic coating layer was immersed into 1 mL of ABTS•+ working solution to initiate the antioxidant activity at room temperature. Every 6 minutes, the immersed sample film was temporarily removed from the working solution, and the absorbance reading of the working solution was taken at 734 nm in the UV-Vis spectrophotometer. After the measurement, the sample film was placed back in the 103 working solution to allow further reaction. Light exposure was avoided during the entire hydrogen abstraction reaction. The ABTS•+ working solution was freshly made every time before the analysis. For comparison purposes, uncoated sample films immersed in ABTS•+ working solution were used as control samples and treated in the exact same way. 5.1.3.4 Results and discussion 5.1.3.4.1 Antioxidant efficacy of the phenolic extracts from avocado byproducts In order to understand the antioxidant efficacy of the phenolic extracts from avocado byproducts, one drop (around 0.83 µl) of the acetonic peel or seed extract was directly added into 1mL ABTS•+ working solution (diluted with 95% aqueous ethanol) for evaluation. This screening test was conducted before evaluating the coated sample films. though _`abc•d Figure 5.10 presents the free radical reduction process of the peel and seed extracts. Even _efKghiK≈1200, both peel and seed extracts exhibited potent antioxidant ability stabilizing 50% of ABTS•+ radicals within 1 minute. Compared with the seed extract, the peel extract was more powerful. The ET50 of the peel extract was 9.5 seconds, which was only 19% of the time the seed extract used to reach its ET50 point. In addition, at the end of the test (3 minutes), the peel extract had stabilized 100% of the free radicals in the working solution, whereas 40% of ABTS•+ radicals still remained in the other working solution. This experimental 104 outcome agreed with the phenolic quantification and identification results discussed in section 3.3. Based on the analysis result of the mass spectra (Table 3.1 and Figure 3.1), there were 22 out of 28 types of phenolic compounds available in both avocado peel and seed extracts. Among these phytochemicals, the concentrations of catechin isomer 2, procyanidin dimer B (I), 5-O- caffeoylquinic acid, procyanidin trimer 2 and procyanidin trimer 1 in the peel extract were significantly greater than that in the seed extract. These high content phenolic compounds enabled the peel extract to eliminate more ABTS•+ radicals in the working solution at a faster rate. Figure 5.10 Antioxidant activity of acetonic phenolic extracts from avocado byproducts. Points a and b are the estimated ET50 of the peel and seed extract respectively. The ABTS•+ solvent was diluted with 95% aqueous ethanol. 105 5.1.3.4.2 Coating solutions vs. antioxidant efficacy In order to confirm that the alkaline saline solution could provide a more powerful phenolic coating than the laccase assist solution, ethanolic seed extract was utilized again for this screening test. As shown in Figure 5.11, the ABTS•+ evaluation result further confirmed that the alkaline solution coated sample film was more potent than the laccase solution coated polymer substrate. Within 18 minutes, the PP film coated by the alkaline solution completely eliminated ABTS•+ radicals in the working solution. During the same time period, the laccase solution coated PP film only reduced 30% of ABTS•+ radicals. After 90 minutes, 16% of ABTS•+ radicals still remained in the working solution used for testing the laccase solution coated PP film. The free radical reduction process of ABTS•+ assay was rapid. Different from the long reaction time of DPPH• assay, the effective duration of ABTS•+ assay could be controlled within 2 hours. In comparing the free radical reduction process of alkaline solution coated polymer films (as shown in Figure 5.12), significant time difference was noticed. The coated films could stabilize ABTS•+ radicals in 30 minutes, whereas the DPPH• evaluation time was extended to 5 hours. After 5 hours, around 15% of DPPH• radicals still remained in the working solutions. 106 Figure 5.11 Antioxidant activity of PP films coated in laccase assist (pH = 5) and alkaline saline (pH = 7.8) solutions. The ABTS•+ solvent was diluted with 50% aqueous ethanol. Points a and b are estimated ET50 of PP (alkaline) and PP (laccase) respectively. 107 Figure 5.12 Antioxidant activity of PP, PET and LDPE films coated in alkaline saline (pH = 7.8) solutions. The ABTS•+ and DPPH• solvents were diluted with 50% aqueous ethanol. Point a is the estimated ET50 of PP (ABTS) and PET (ABTS), and point b is the estimated ET50 of LDPE (DPPH), PET (DPPH) and PP (DPPH). 5.1.3.4.3 Plastic substrates vs. antioxidant efficacy To examine the influence of plastic substrates on the antioxidant efficacy of the phenolic coating, ethanolic seed extract was used to coat PP and PET films in alkaline saline solution, and then immersed in 10% aqueous ethanol (water-based food simulant) containing ABTS•+ radicals for a quick preliminary test. Based on Figure 5.13, it took nearly equal time for both films to 108 quench 50% of ABTS•+ radicals in the solution. There was only a 0.4 minute gap between the estimated ET50 of the coated PET and PP films. This experimental result further supported the conclusion that polymer substrates did not act as a major factor influencing the antioxidant efficacy of the phenolic coating. Figure 5.13 Antioxidant activity of PET and PP films coated with ethanolic seed extract. The films were coated in alkaline saline (pH = 7.8) solution. The ABTS•+ solvent was diluted with 10% aqueous ethanol. Points a and b are the estimated ET50 of PET and PP respectively. 109 To sum up, the ABTS•+ assay is a rapid testing method compared with the DPPH• assay. It was employed at the third experimental stage to evaluate the antioxidant efficacy of the avocado peel and seed extracts, and the potential influences of the coating environment and plastic substrates on the antioxidant effectiveness of the phenolic coating layer. In addition, the avocado peel and seed extracts were directly evaluated to understand their antioxidant potency to quench free radicals. According to the analysis results, the high content phenolic compounds enabled the peel extract to eliminate more ABTS•+ radicals in the working solution at a faster rate. Like what was implied from the silver nitrate test and DPPH• assay, the sample films coated in alkaline saline solution presented greater antioxidant effectiveness than the laccase solution coated polymer substrates. Polymer substrates did not play a crucial role in influencing the antioxidant efficacy of the phenolic coating. 110 5.2 Introduction Part II: Further Evaluation with Statistical Analysis In this part, coated PP, PET and LDPE films were tested in triplicate in order to a) further verify the influence of plastic substrates, b) analyze the potential effect of film storage time, and kleem efKghiK kneeo efKghiK in the coating solution on the antioxidant efficiency of the c) evaluate the impact of phenolic coating. Considering the fast reaction rate, the solubility in both aqueous and organic solvents, and the stability of the working solution in different pH environments, the ABTS•+ assay (section 5.1.3) was selected to evaluate coated PP, PET and LDPE films in 95% aqueous ethanol (fatty food simulant). Due to the potent coating efficacy evaluated in the previous silver nitrate, DPPH• and ABTS•+ tests, the alkaline saline method (section 4.3.2) was employed to coat acetonic seed and peel extracts on the plastic substrates. As discussed in section 5.1.2.1, researchers could fit their experimental results to different mathematical models to evaluate the antioxidant efficiencies of various phenolic compounds, including the following logistic, Boltzmann sigmoidal and dose-response models (Suriyatem et al. 2017)(cid:2)(cid:1) (cid:1) (cid:1) 111 sU(ff<)tMmmnmule+Bw Logistic model p= WqrW\ sUJxH(fyf 0.001%), the absorbance peak gradually rose at around 280 nm and eventually formed a sharp peak (see Figure C-2 in Appendix C for the overlapped UV-Vis spectra of the peel extract in 50% and 10% aqueous ethanol). This peak absorbance wavelength agreed with the experimental results of other research regarding total phenolic compound detection in wine (Aleixandre-Tudo and Toit 2018; Ribereau-gayon 1974), plants (Engida et al. 2015; Owades, Rubin, and Brenner 158 1958), coffee and teas (Cohen 2000; Meireles et al. 2012). Although other absorbance wavelengths were also suggested, using 280 nm for phenolic compound determination is still widely accepted. For a particular phenolic compound, the absorbance wavelength may vary around 280 nm. Table 6.3 includes the suggested absorbance wavelengths for the detection of different phenolic compounds by UV-Vis spectrophotometry. These compounds are the primary antioxidants in the avocado byproduct extracts used for this research. Considering the crude extracts used for this research were mixtures of phenolic compounds with other ingredients, the UV-Vis absorbance peak might slightly shift during the migration test. The selected wavelength range for the phenolic compound releasing experiment was 270 ~300 nm, and the concentration of released phenolic compounds from the antioxidant coating could be estimated by using the calibration curves in Figure 6.7. 159 Figure 6.6 Overlapped UV-Vis spectrum of the crude phenolic extract from avocado peels in 95% aqueous ethanol at various concentrations. 160 Table 6.3 Absorbance wavelengths suggested for the detection of different phenolic compounds by UV-Vis spectrophotometry. Absorbance Wavelength ~ 288 nm 275 nm Compound Name Catechin 5-O-caffeoylquinic acid 217 nm with shoulder at 240 nm 324 nm with shoulder at 296 nm Belay (2010) ~296 nm, 326 nm References Aleixandre et al. (2018) Bark et al. (2011) Peres et al. (2013) Wang et al. (2017) Aleixandre et al. (2018) Abualhasan et al. (2017) Procyanidin 280 nm Quercetin-3-O-rutinoside (rutin) ~ 263 nm 260nm, 360 nm 161 A. Avocado peel extract in 95% aqueous ethanol Figure 6.7 Calibration curves for the phenolic compound migration analysis. The wavelength selected for the phenolic compound absorbance measurement was around 280 nm. 162 Figure 6.7 (cont’d) B. Avocado peel extract in 50% aqueous ethanol C. Avocado peel extract in 10% aqueous ethanol 163 Table 6.4 presents migration results of the phenolic compound releasing experiment (see Figure C-3 in Appendix C for the UV-Vis spectra of the migration results). The ‘initial time’ noted in the table refers to the starting time point for obtaining detectable released phenolic compounds in each migration cell. The concentration of released phenolic compounds in each migration cell was quantified using the calibration curves. Based on the experimental results, the coated films started releasing phenolic compounds into each migration cell at different time points. Within the first 30 minutes, released phenolic compounds could be detected from 6 out of 18 migration cells. At 1 hour, the number of samples with detectable compounds increased from 6 to 10. Noticeable phenolic content was observed from two more migration cells within 24 hours. Five coated sample films did not release detectable phenolic compounds until the end of migration test. As for the concentrations of released phenolic compounds, they did not increase as the testing time increased. Instead, they either stabilized at 0.01% during the entire testing process or ranged from 0% to 0.04%. At both experimental temperatures, the concentrations of released phenolic compounds in 10% ethanol varied within a comparatively wider range. Still, concentration stayed at a very low level. No food simulant containing more than 0.04% released phenolic compounds was found. 164 The aforementioned two facts verified the speculation of the previous screening test. During those 168 hours, only a small number of phenolic compounds were released from the antioxidant coating layer. It is highly possible that this limited number of compounds resulted from surface residues left by the coating process. That being said, the phenolic coating layer did not rely on a surface releasing or migration process to stop or slow down free radical reactions. For the sample films in the same food simulants, different experimental temperatures did not have any significant influence on the release of phenolic compounds. As shown in Table 6.4, estimated concentrations of the migration cells at 4 °C were either the same or similar to that of the migration cells with the same food simulant at 40 °C. This experimental result is further evidence that the phenolic compounds from the avocado byproducts were stabilized on the substrate. They did not depend on a migration or surface releasing process for the free radical quenching process. Since a) at least the majority of the polymerized phenolic compounds remained inside of the coating layer during the free radical reduction process, b) the coating layer could restore its antioxidant ability after quenching the ABTS•+ radicals in the working solution, and c) the diameters of the primary phenolic compounds identified from the avocado byproduct extracts, such as catechin, protocatechuic acid and rutin, were around 1 nm (see Table C-3 in Appendix C), which were much smaller than the estimated coating thickness (37.75 ± 0.30 nm, see section 165 4.4), the antioxidant coating layer can be presumed to have more than one layer of phenolic compounds polymerized on the polymer substrates. After donating hydrogen atoms to stabilize DPPH• or ABTS•+ radicals, phenolic compounds at the surface layer of the antioxidant coating become phenolic radicals (Ar : O• , see Figure 6.8). In order to continuously serve as antioxidants, the surface phenolic radicals may abstract hydrogen atoms from their adjacent phenolic compounds (Ar’ : OH) in an inner layer of the antioxidant coating to restore their antioxidant ability. The longer the elapsed time between two consecutive DPPH• or ABTS•+ stabilization reactions, the more Ar : O• at the coating surface could regain their antioxidant ability, and Ar’ : OH, in this case, turn to new phenolic radicals, Ar’ : O•. It is worth noting that not all the surface phenolic radicals are able to regain their antioxidant ability. Also, the O-H bond dissociation enthalpy of Ar’ : OH may influence this hydrogen abstraction reaction. The lower the O-H bond dissociation enthalpy in the Ar’ : OH, the more readily the hydrogen abstraction reaction will take place. To continue the hydrogen abstraction reaction between Ar : O• and Ar’ : OH every time after the DPPH• or ABTS•+ stabilization reaction, Ar’ : O• needs to abstract hydrogen atoms from their adjacent phenolic compounds, Ar’’ : OH, to transfer the hydrogen atoms to Ar : O• at the surface layer. As the number of the DPPH• or ABTS•+ stabilization reaction increases, the hydrogen abstraction reaction will take place in a much deeper layer of the antioxidant coating to 166 transfer hydrogens, causing longer time for the surface phenolic radical, Ar : O•, to regain their antioxidant ability. Since hydrogen atoms in the phenolic coating layer would be gradually consumed, there would be less and less Ar : O• that could recover their antioxidant ability. Eventually, the phenolic coating layer will lose its antioxidant property when the last hydrogen atom is used up for the surface phenolic radicals to regain antioxidant ability. 167 Table 6.4 Estimated concentrations of phenolic compounds released from the antioxidant coating layer during the migration test. 95% ethanol 50% ethanol 10% ethanol Cell 1 0.5 Cell 2 Cell 3 0.5 1 Cell 1 168 Cell 2 Cell 3 168 168 Cell 1 24 Cell 2 Cell 3 0.5 1 Initial Time (hours) Concentration (%) 0.02 ~ 0.04 0.01 S 0.01 ~ 0.02 ND ND ND 0 ~0.04 0.02~0.04 0.02~0.04 Initial Time (hours) 0.5 1 0.5 168 0.5 168 1 48 24 0 ~ 0.01 0.02 ~ 0.04 0.01 ~ 0.02 ND 0 ~ 0.02 ND 0.02 ~ 0.04 0.02 ~ 0.04 0.02 ~ 0.04 Concentration (%) 4 ◦C 40 ◦C Initial time refers the starting time point for obtaining detectable released phenolic compounds The superscript S refers to the concentration of released phenolic compounds stabilized at a particular value during the entire migration test ND refers to a non-detectable concentration of released phenolic compounds during the entire migration test 168 168 Ar : OH is a phenolic compound inside the antioxidant coating Ar’ : OH is a phenolic compound close to Ar : OH inside the antioxidant coating Ar’’ : OH is a phenolic compound close to Ar’ : OH inside the antioxidant coating Ar’’’ : OH is a phenolic compound close to the plastic substrate Ar : O• is a phenolic radical of Ar : OH Ar’ : O• is a phenolic radical of Ar’ : OH Ar’’ : O• is a phenolic radical of Ar’’ : OH Ar’’’ : O• is a phenolic radical of Ar’’’ : OH Figure 6.8 Schematic of antioxidative restorability of the phenolic coating on a plastic substrate. 169 CHAPTER SEVEN: Conclusions and Future Work 7.1 Crude extraction from avocado byproducts Both avocado peel and seed powders were utilized for this research to obtain crude phenolic extracts. In order to maximize phenolic content recovered from the byproduct powders, 70% ethanol and 70% acetone, as the most common solvents, were selected. The selections were based on acceptability for human consumption, solvent polarity and the solubility of the extracted phenolic compounds. In addition, moderate heat treatment was used to further promote the extraction process. Within a limited time period, a Vortex mixer and ultrasonic bath, as unconventional crude extraction procedures, were utilized to allow the solvents to completely mix with avocado byproduct powders and thus increase the concentration of extracted phenolic compounds in the aqueous solvents. 7.2 Non-metal contact coating process The coating process was a polymerization reaction. It required the involvement of oxygen and moderate mechanical agitation. More importantly, a metal contact coating technique, conventional roller coating process for example, could not be employed due to the metal 170 chelating ability of phenolic compounds. Considering no coating equipment was available in the lab to meet the aforementioned requirements, a non-metal contact dip coater was developed. The dip coater applied a slider-crank mechanism to convert rotary motion into two ways of linear motion, i.e., horizontal and vertical movements. Together with a hot plate stirrer, this coating device could be used for beakers with different sizes, and the temperature of coating solution, dip coating and agitation speed were all controllable variables. Two types of coating solutions were utilized for this research, namely, laccase assist (pH = 5) and alkaline saline (pH = 7.8) coating solutions. By mixing with the phenolic extracts at different ratios (100% peel extract, 100% seed extract, or 50% peel extract and 50% seed extract), the coating solutions successfully polymerized the phenolic compounds onto PP, LDPE, and PET films, and SEM showed the coating layer was uniform on the substrates with thickness of approximately 37.75 ± 0.30 nm. It is worth mentioning that for the thickness measurement of the phenolic coating, a glass coverslip was used as the coating substrate. This material was able to present a distinct edge line between itself and the coating layer under SEM to facilitate the measurement. More importantly, even on a different type of substrate, the phenolic coating was still evenly distributed. This experimental result indicated the substrate-independent characteristic of the phenolic coating, 171 and thus implied a broader range of applications of this antioxidant coating on other types of packaging materials. 7.3 Evaluation of antioxidant efficacy of coated polymer films To understand the antioxidant efficacy of the phenolic coating, experimental analysis was separated into two parts. In the screening test, AgNO3, DPPH•, and ABTS•+ assays were employed to analyze coated PP, LDPE and PET films in food simulants based on 3 reaction mechanisms. Both avocado peel and seed extracts was used to coat the polymer films using the laccase assist and alkaline saline approaches stated in chapter four. In addition, the avocado peel and seed extracts were directly tested to understand the greater antioxidant efficiency of the peel extract. The AgNO3 method first provided a visual inspection result to prove the existence of phenolic content in the polymerized coating layer. After immersing coated polymer films into AgNO3 solution, phenolic compounds in the antioxidant coating layer resulted in film color change, synthesis and deposition of silver nanoparticles. This simple testing method is recommended to preliminarily verify phenolic coating. Moreover, the darker color change on the coated polymer films indicated the higher coating efficacy of the alkaline saline method. 172 DPPH• radicals are purple in color. To stabilize these radicals, phenolic compounds donated single electrons and/or hydrogen atoms to gradually transform the free radical solution into a yellow color. ABTS•+ radicals are blue-green in color. To stabilize these radicals, phenolic compounds transferred their hydrogen atoms and thus gradually removed the color in the working solution. Based on the experimental results of DPPH• and ABTS•+ assays, the alkaline saline coating method, as an inexpensive approach, could generate a phenolic coating layer with greater antioxidant effectiveness. The polymer films coated by this approach required less time to reach ET50 in both DPPH• and ABTS•+ assays. Also, the different plastic substrates did not result in any statistically significant difference in terms of the antioxidant efficiency of the phenolic coating. During the coating process, the phenolic compounds acted as a layer of adhesive attached onto the plastic surface. They did not interact with the substrates to cause any variance in the antioxidant ability of the phenolic coating. When the polymer films were coated in solutions with the same type of crude extract but different concentrations, they did not present significant difference in the antioxidant ability to reduce DPPH• radicals. It seemed that the antioxidant efficiency of the coating layer was not linearly correlated to the concentration of phenolic extract in the coating solution. Provided the coating duration and phenolic content in the coating solution were sufficient, the antioxidant 173 efficiency of the coating layer might be determined by substrate surface area rather than the amount of phenolic content in the coating solution. To further evaluate the antioxidant coating, coated PP, PET and LDPE films were tested in triplicate in the second part of the experimental analysis. The statistical analyses, again, agreed that the different plastic substrates did not result in any statistically significant difference in terms of the antioxidant efficiency of the phenolic coating. Even though no significant variance was found among the overall antioxidant rates of coated films with different storage times, the antioxidant efficiencies of the coated films did vary significantly during some time intervals. Although the avocado peel extract exhibited more potent antioxidant efficiency than the seed extract, no statistically significant difference was found among the polymer substrates !"##$ #&'()*' !+##, #&'()*' = 100% to 0%, 0% to 100%, and coated with the phenolic extracts at different ratios ( 50% to 50% respectively). On the contrary, the coated films presented similar performance in stabilizing ABTS•+ radicals in the working solution. 7.4 Mode of antioxidant activity of coated polymer films While stabilizing free radicals in food simulants, at least the majority of the phenolic compounds remained in the antioxidant coating layer. They did not depend on a migration or surface releasing process to quench free radicals. 174 According to the phenolic releasing experiments, only a small number of phenolic compounds were released into the food simulant. The majority of phenolic compounds remained in the coating layer. Possibly, this small number of migrating compounds was surface residues left by the coating process. The cross-sectional SEM image of the antioxidant coating after the initial antioxidant reaction supported this speculation. Under SEM, the phenolic coating still presented a uniform layer on the substrate; no form change was noticed. It was also worth mentioning that the different migration test temperatures did not impact the stability of the coating layer. At 4 and 40 °C, the phenolic concentrations in the migration cells containing the same type of food simulants did not present any significant differences. This experimental result further suggested that the phenolic compounds from avocado byproducts were stabilized on the substrate. They did not depend on a migration or surface release process for free radical elimination. Presumably, there was more than one layer of phenolic compounds extracted from the avocado byproducts polymerized on the polymer substrates. After donating hydrogen atoms to stabilize free radicals, phenolic compounds at the surface layer of the antioxidant coating became phenolic radicals. In order to continuously serve as antioxidants, the surface phenolic radicals might abstract hydrogen atoms from their adjacent phenolic compounds in an inner layer of the antioxidant coating to restore their antioxidant ability. As the number of free radical stabilization 175 reaction increased, the hydrogen abstraction reaction would take place in a much deeper layer of the antioxidant coating to transfer hydrogens, causing longer time for the surface phenolic radical to regain their antioxidant ability. Eventually, the phenolic coating layer would lose its antioxidant property when the last hydrogen atom was used up for the surface phenolic radicals to regain antioxidant ability. 7.5 Future work For this research, experiments were conducted to obtain crude phenolic extracts from avocado byproducts, polymerize the crude extracts on substrates uniformly, evaluate antioxidant efficiency of the coating layer, and understand the mode of antioxidant activity of the coating layer. Still, improvements and further evaluations are required to thoroughly understand this bio- based antioxidant coating. To obtain the antioxidant coating layer, polymer films were dip coated for 24 hours at room temperature. Although this coating duration was shorter than the experimental time recommended by other researchers, a relatively higher temperature together with a different dipping speed may expedite the coating process without introducing decomposition of phenolic compounds in the coating solution. During each coating process, the dip coater developed for this research could only coat substrates with small surface areas. For the purpose of commercial application, a better coating 176 method or device, which can coat a substrate with large surface area, is desired to enhance the coating efficiency. While measuring the coating thickness under SEM, slight deformation on the coating surface was noticed at high magnification (35000x). This deformation might result from the sample preparation process for the SEM images. After removal from liquid nitrogen, the coated substrate became very brittle. Manually fracturing the sample, in this case, might result in deformation at some spots of the coating layer. A better sample preparation method is needed to avoid any possible deformation. Before applying the antioxidant coating in real applications, knowing the expiration time of the antioxidant coating is a must. Despite the fact that no statistically significant difference was noticed between the antioxidant efficacy of coated polymer films with less than 36 hours and 7 days of storage times, a coated film with longer storage time needs to be tested to further understand the antioxidant efficacy of the phenolic coating layer. To better understand the antioxidant efficacy of the coating layer in real applications, food products, such as meat patties, beverage and dairy products, should be used. Different from food simulants, chemical reactions are more complicated in real food products because of their complex ingredients. If the polymerized coating layer could present its potent antioxidant efficacy to food products, it would raise its promising value in commercial use. 177 APPENDICES 178 Appendix A Antioxidant efficiency of coated polymer films (screening test results) Table A-1 Experimental data of the coating solution vs. antioxidant efficacy test (DPPH• assay). Inhibition Capacity Time (hour) 50% aqueous ethanol 95% aqueous ethanol 50% aqueous ethanol LDPE (laccase) LDPE (alkaline) Negative Control LDPE (laccase) LDPE (alkaline) Negative Control PET (laccase) PET (alkaline) Negative Control 0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 5 6 7 8 9 24 48 0.00% 0.00% 12.66% N/A 0.00% 0.00% 19.78% 85.10% 0.34% 28.86% 85.15% 0.17% 35.73% 84.62% 0.17% 0.00% 3.74% 7.32% 8.93% 9.44% 0.00% 0.00% 0.00% 0.00% 56.31% 1.55% 15.98% N/A 70.87% 2.77% 32.22% 82.50% 76.08% 4.02% 37.45% N/A 79.29% 4.26% 44.94% 80.52% 41.76% 86.37% 0.34% 10.27% 81.77% 4.76% 50.80% N/A 47.07% 85.96% 0.07% 11.12% 83.80% 4.86% 49.67% 81.48% N/A N/A N/A N/A N/A N/A 47.21% N/A 56.28% 88.56% 0.07% 17.54% 85.81% 5.12% 55.03% 82.02% N/A N/A N/A N/A N/A N/A 60.97% N/A 64.15% 89.98% 1.16% 20.35% 86.65% 5.25% 66.65% 82.66% 72.53% 90.28% 2.51% 21.60% 87.13% 5.29% 71.41% 75.96% 90.46% 2.90% 24.18% 86.94% 5.48% 71.41% 82.96% 86.43% N/A N/A N/A N/A N/A N/A 3.55% 27.27% 86.23% 5.56% 4.43% 29.67% 86.29% 6.79% N/A N/A 42.71% 86.17% 7.34% 63.72% N/A 8.57% N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A 0.00% 0.06% 0.12% 0.17% 0.23% 0.30% 0.35% 0.48% 0.58% 0.61% 0.74% 1.05% 2.34% N/A N/A N/A N/A 179 Table A-2 Experimental data of the plastic substrates vs. antioxidant efficacy test (DPPH• assay). Time (hour) 0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 5 6 7 8 9 24 48 Inhibition Capacity 50% aqueous ethanol LDPE (laccase) LDPE (alkaline) Negative Control PET (laccase) PET (alkaline) Negative Control PP (alkaline) Negative Control 0.00% 0.00% 0.00% 0.00% 0.00% 12.66% N/A 0.00% 15.98% N/A 0.00% 0.06% 0.00% 0.00% N/A N/A 19.78% 85.10% 0.34% 32.22% 82.50% 0.12% 84.73% 0.35% 28.86% N/A 0.17% 37.45% N/A 0.17% N/A N/A 35.73% 84.60% 0.17% 44.94% 80.52% 0.23% 86.53% 2.97% 41.76% N/A 0.34% 50.80% N/A 0.30% N/A N/A 47.07% 86.00% 0.07% 49.67% 81.48% 0.35% 86.40% 3.01% N/A N/A N/A 47.21% N/A 0.48% N/A N/A 56.28% 88.60% 0.07% 55.03% 82.02% 0.58% 86.20% 3.25% N/A N/A N/A 60.97% N/A 0.61% N/A N/A 64.15% 90.00% 1.16% 66.65% 82.66% 0.74% 85.17% 4.11% 72.53% 75.96% 82.96% 86.43% N/A N/A N/A N/A N/A N/A N/A N/A 2.51% 71.41% 2.90% 71.41% 3.55% 4.43% N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A 1.05% 2.34% N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A 180 Table A-3 Experimental data of the concentration of phenolic extract vs. antioxidant efficacy test (DPPH• assay). Time (hour) 0 0.5 1 1.5 2 2.5 3 4 5 Inhibition Capacity 50% aqueous ethanol PP (12.5 mg/mL) PP (25 mg/mL) 0.00% 35.61% 73.42% 80.26% 85.04% 87.94% 90.19% 92.76% 93.50% 0.00% N/A 84.73% N/A 86.53% N/A 86.40% 86.20% 85.17% Negative Control 0.00% N/A 0.35% N/A 2.97% N/A 3.01% 3.25% 4.11% 181 Table A-4 Experimental data of the antioxidant efficacy of the phenolic extracts from avocado byproducts test (ABTS•+ assay). Time (second) Inhibition Capacity 95% aqueous ethanol Peel extract Seed extract Negative Control 0 2 8 14 20 26 32 38 44 50 56 62 68 74 80 86 92 98 0.00% 28.96% 43.95% 63.11% 78.67% 86.89% 91.79% 94.52% 96.25% 97.41% 97.98% 98.41% 98.56% 98.70% 98.70% 98.85% 99.14% 98.99% 0.00% 20.48% 26.91% 29.45% 34.83% 38.57% 41.70% 45.44% 48.13% 50.22% 52.17% 53.96% 55.75% 56.80% 57.85% 58.74% 59.49% 60.09% 0.00% 0.15% 0.29% 0.15% 0.00% 0.44% 0.15% 0.15% 1.03% 0.15% 0.29% 0.00% 0.00% 0.15% 0.15% 0.15% 0.15% 0.15% 182 Table A-4 (cont’d) 104 110 116 122 128 134 140 146 152 158 164 170 176 182 98.41% 98.27% 98.41% 98.56% 98.70% 98.70% 98.85% 98.99% 99.14% 99.28% 99.57% 99.71% 99.86% 100.00% 60.54% 60.99% 61.29% 61.58% 61.88% 62.33% 62.48% 62.78% 62.93% 63.08% 63.08% 63.08% 62.93% 63.08% 0.15% 0.44% 0.15% 0.29% 0.00% 0.00% 0.29% 0.29% 0.29% 0.29% 0.29% 0.29% 0.15% 0.00% 183 Table A-5 Experimental data of the coating solutions vs. antioxidant efficacy test (ABTS•+ assay). Time (minute) 0 6 12 18 30 42 54 66 78 90 PP (laccase) 0.00% 15.39% 22.03% 30.13% 37.94% 44.84% 51.39% 63.83% 79.38% 83.68% Inhibition Capacity 50% aqueous ethanol PP (alkaline) 0.00% 63.51% 95.81% 100.00% N/A N/A N/A N/A N/A N/A Negative Control 0.00% 0.58% 1.75% 2.66% 3.41% 4.29% 5.99% 7.06% 7.06% 7.00% A. Antioxidant efficacy of PP films coated with ethanolic seed extract 184 Table A-5 (cont’d) Time (hour) Inhibition Capacity 50% aqueous ethanol DPPH• assay ABTS•+ assay PP Cntrl PET Cntrl LDPE Cntrl PP Cntrl PET Cntrl 0 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.1 0.2 0.3 1 2 3 4 5 N/A N/A N/A N/A N/A N/A 63.51% 0.58% 82.25% 1.49% N/A N/A N/A N/A N/A N/A 95.81% 1.75% 99.43% 2.87% N/A N/A N/A N/A N/A N/A 100.00% 2.66% 100.00% 2.96% 84.73% 0.35% 82.50% 1.14% 85.10% 0.23% 86.53% 2.97% 80.52% 2.01% 84.62% 0.69% 86.40% 3.01% 81.48% 3.53% 85.96% 1.87% 86.20% 3.25% 82.02% 3.66% 88.56% 2.94% 85.17% 4.11% 82.66% 4.41% 89.98% 3.61% N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A N/A B. Antioxidant activity of coated polymer films tested by DPPH• and ABTS•+ assays 185 Table A-6 Experimental data of the plastic substrates vs. antioxidant efficacy test (ABTS•+ assay). Time (minute) 0 6 12 18 24 30 36 PP 0.00% 22.19% 53.94% 74.00% 88.64% 97.64% 99.42% Inhibition Capacity 10% aqueous ethanol Negative Control 0.00% 0.01% 0.40% 1.07% 1.37% 2.06% 2.44% PET 0.00% 28.12% 56.03% 69.89% 78.96% 88.71% 93.50% Negative Control 0.00% 0.12% 0.17% 0.23% 0.30% 0.35% 0.48% 186 Appendix B Antioxidant efficiency of coated polymer films (further tests for statistical analysis) Dose-response model Boltzmann sigmoidal model A. PP films coated with the seed extract Figure B-1 Goodness of fit results estimated by the dose-response and Boltzmann sigmoidal models for the plastic substrates vs. antioxidant efficacy test. 187 Figure B-1 (cont’d) Dose-response model Boltzmann sigmoidal model B. PET films coated with the seed extract 188 Figure B-1 (cont’d) Dose-response model Boltzmann sigmoidal model C. LDPE films coated with the seed extract 189 Figure B-1 (cont’d) Dose-response model Boltzmann sigmoidal model D. PP films coated with the peel extract 190 Figure B-1 (cont’d) Dose-response model Boltzmann sigmoidal model E. PET films coated with the peel extract 191 Figure B-1 (cont’d) Dose-response model Boltzmann sigmoidal model F. LDPE films coated with the peel extract 192 Table B-1 Fitted regression equations estimated by the dose-response and Boltzmann sigmoidal models for the plastic substrates vs. antioxidant efficacy test. Boltzmann Sigmoidal Model − C′E ;<=>?)+C′E A= 1+FGH(7978 − 0.710 A= 9I.LPM)+0.710 1+FGH(79I.KIN −0.680 A= 9I.EIE)+0.680 1+FGH(79I.NLN −0.707 A= 9I.LPT)+0.707 1+FGH(79I.NLL −0.663 A= 9I.EIK)+0.663 1+FGH(79I.KIL −0.660 A= 9I.LTQ)+0.660 1+FGH(79I.KQK −0.686 A= 9I.EEK)+0.686 1+FGH(79I.KNN PP (seed) PET (seed) LDPE (seed) PP (peel) PET (peel) LDPE (peel) Dose-response Model IC (%)= IC (%)= IC (%)= IC (%)= IC (%)= IC (%)= IC (%)= 100 100 100 100 100 100 100 1+10[(7897)×;<=>?] 1+10[(I.KLM97)×E.EKN] 1+10[(I.NEQ97)×E.LKP] 1+10[(I.NLK97)×E.EMP] 1+10[(I.KIQ97)×E.LIK] 1+10[(I.KQN97)×E.TVE] 1+10[(I.KPL97)×L.PMI] 193 Table B-2 Experimental data of the plastic substrates vs. antioxidant efficacy test. Trial Sample 0 min 6 min 12 min 18 min 24 min 30 min 36 min 1 2 PP (seed) 3 Negative Control 1 2 PET (seed) 3 Negative Control 1 2 LDPE (seed) 3 Negative Control 1 2 PP (peel) 3 Negative Control 0.696 0.346 0.174 0.037 0.002 N/A N/A 0.710 0.296 0.057 0.018 0.000 N/A N/A 0.703 0.206 0.125 0.019 0.000 N/A N/A 0.712 0.705 0.708 0.701 0.696 N/A N/A 0.680 0.363 0.169 0.054 0.000 N/A N/A 0.669 0.370 0.188 0.072 0.005 N/A N/A 0.668 0.355 0.172 0.067 0.000 N/A N/A 0.664 0.653 0.653 0.648 0.645 N/A N/A 0.707 0.243 0.021 0.000 0.000 0.000 0.000 0.702 0.459 0.256 0.141 0.087 0.036 0.016 0.690 0.432 0.190 0.066 0.000 0.000 0.000 0.672 0.668 0.672 0.668 0.664 0.663 0.662 0.663 0.267 0.123 0.031 0.000 0.000 N/A 0.657 0.334 0.205 0.096 0.017 0.000 N/A 0.663 0.170 0.041 0.000 0.000 0.000 N/A 0.712 0.705 0.708 0.701 0.696 0.693 N/A 194 Table B-2 (cont’d) 1 2 PET (peel) 3 Negative Control LDPE (peel) 1 2 3 Negative Control 0.656 0.285 0.077 0.000 0.000 N/A N/A 0.656 0.308 0.114 0.024 0.000 N/A N/A 0.660 0.292 0.094 0.009 0.000 N/A N/A 0.664 0.653 0.653 0.648 0.645 N/A N/A 0.677 0.221 0.074 0.000 0.000 0.000 0.000 0.686 0.310 0.129 0.011 0.001 0.000 0.000 0.683 0.480 0.314 0.215 0.129 0.067 0.023 0.672 0.668 0.672 0.668 0.664 0.663 0.662 195 Dose-response model Boltzmann sigmoidal model A. PP (<36 hours) films coated with the seed extract Figure B-2 Goodness of fit results estimated by the dose-response and Boltzmann sigmoidal models for the film storage time vs. antioxidant efficacy test. 196 Figure B-2 (cont’d) Dose-response model Boltzmann sigmoidal model B. PP (7 days) films coated with the seed extract 197 Figure B-2 (cont’d) Dose-response model Boltzmann sigmoidal model C. PP (<36 hours) films coated with the peel extract 198 Figure B-2 (cont’d) Dose-response model Boltzmann sigmoidal model D. PP (7 days) films coated with the peel extract 199 Table B-3 Fitted regression equations estimated by the dose-response and Boltzmann sigmoidal models for the film storage time vs. antioxidant efficacy test. Boltzmann Sigmoidal Model − C′E A= ;<=>?)+C′E 1+FGH(7978 − 0.710 A= 9I.LPM)+0.710 1+FGH(79I.KIN −0.656 A= 9I.ELE)+0.656 1+FGH(79I.NPQ −0.663 A= 9I.EIK)+0.663 1+FGH(79I.KIL −0.659 A= 9I.LTM)+0.659 1+FGH(79I.NVI PP (< 36 hours, seed) PP (7 days, seed) PP (< 36 hours, peel) PP (7 days, peel) Dose-response Model IC (%)= IC (%)= IC (%)= IC (%)= IC (%)= 100 100 100 100 100 1+10[(7897)×;<=>?] 1+10[(I.KLM97)×E.EKN] 1+10[(I.NPV97)×E.IVK] 1+10[(I.KIQ97)×E.LIK] 1+10[(I.NVE97)×E.TKM] 200 Table B-4 Experimental data of the film storage time vs. antioxidant efficacy test. Trial Sample 0 min 6 min 12 min 18 min 24 min 30 min 36 min PP (< 36 hours, seed) Negative Control PP (7 days, seed) Negative Control PP (< 36 hours, peel) Negative Control PP (7 days, peel) 1 2 3 1 2 3 1 2 3 1 2 3 0.696 0.346 0.174 0.037 0.002 N/A N/A 0.710 0.296 0.057 0.018 0.000 N/A N/A 0.703 0.206 0.125 0.019 0.000 N/A N/A 0.712 0.705 0.708 0.701 0.696 N/A N/A 0.654 0.379 0.217 0.081 0.000 0.000 0.000 0.656 0.364 0.201 0.107 0.024 0.000 0.000 0.654 0.441 0.263 0.170 0.082 0.033 0.000 0.712 0.705 0.708 0.701 0.696 0.693 0.689 0.663 0.267 0.123 0.031 0.000 0.000 N/A 0.657 0.334 0.205 0.096 0.017 0.000 N/A 0.663 0.170 0.041 0.000 0.000 0.000 N/A 0.712 0.705 0.708 0.701 0.696 0.693 N/A 0.656 0.431 0.192 0.049 0.008 N/A N/A 0.659 0.372 0.096 0.000 0.000 N/A N/A 0.653 0.381 0.167 0.051 0.000 N/A N/A Negative Control 0.712 0.705 0.708 0.701 0.696 N/A N/A 201 Dose-response model Boltzmann sigmoidal model A. PP (100% seed) Figure B-3 Goodness of fit results estimated by the dose-response and Boltzmann sigmoidal models for the extract ratio vs. antioxidant efficacy test. 202 Figure B-3 (cont’d) Dose-response model Boltzmann sigmoidal model B. PP (100% peel) 203 Figure B-3 (cont’d) Dose-response model Boltzmann sigmoidal model C. PP (50% seed + 50% peel) 204 Figure B-3 (cont’d) Dose-response model Boltzmann sigmoidal model D. LDPE (100% seed) 205 Figure B-3 (cont’d) Dose-response model Boltzmann sigmoidal model E. LDPE (100% peel) 206 Figure B-3 (cont’d) Dose-response model Boltzmann sigmoidal model F. LDPE (50% seed + 50% peel) 207 Table B-5 Fitted regression equations estimated by the dose-response and Boltzmann sigmoidal models for the extract ratio vs. antioxidant efficacy test. PP (100% seed) PP (100% peel) PP (50% seed + 50% peel) LDPE (100% seed) LDPE (100% peel) LDPE (50% seed + 50% peel) Boltzmann Sigmoidal Model − C′E A= ;<=>?)+C′E 1+FGH(7978 − 0.710 A= 9I.LPM)+0.710 1+FGH(79I.KIN −0.663 A= 9I.EIK)+0.663 1+FGH(79I.KIL −0.666 A= 9I.EEI)+0.666 1+FGH(79I.NLT −0.707 A= 9I.LPT)+0.707 1+FGH(79I.NLL −0.686 A= 9I.EEK)+0.686 1+FGH(79I.KNN −0.651 A= 9I.LPV)+0.651 1+FGH(79I.KLL Dose-response Model IC (%)= IC (%)= IC (%)= IC (%)= IC (%)= IC (%)= IC (%)= 100 100 100 100 100 100 100 1+10[(7897)×;<=>?] 1+10[(I.KLM97)×E.EKN] 1+10[(I.KIQ97)×E.LIK] 1+10[(I.NEI97)×L.PPT] 1+10[(I.NLK97)×E.EMP] 1+10[(I.KPL97)×L.PMI] 1+10[(I.KLL97)×E.EEK] 208 Table B-6 Experimental data of the extract ratio vs. antioxidant efficacy test. Trial Sample 0 min 6 min 12 min 18 min 24 min 30 min 36 min PP (100% seed) Negative Control PP (100% peel) Negative Control PP (50% seed + 50% peel) Negative Control LDPE (100% seed) 1 2 3 1 2 3 1 2 3 1 2 3 0.696 0.346 0.174 0.037 0.002 N/A N/A 0.710 0.296 0.057 0.018 0.000 N/A N/A 0.703 0.206 0.125 0.019 0.000 N/A N/A 0.712 0.705 0.708 0.701 0.696 N/A N/A 0.663 0.267 0.123 0.031 0.000 0.000 N/A 0.657 0.334 0.205 0.096 0.017 0.000 N/A 0.663 0.170 0.041 0.000 0.000 0.000 N/A 0.712 0.705 0.708 0.701 0.696 0.693 N/A 0.658 0.330 0.154 0.067 0.019 0.000 N/A 0.666 0.399 0.222 0.121 0.047 0.000 N/A 0.658 0.313 0.178 0.049 0.036 0.007 N/A 0.712 0.705 0.708 0.701 0.696 0.693 N/A 0.707 0.243 0.021 0.000 0.000 0.000 0.000 0.702 0.459 0.256 0.141 0.087 0.036 0.016 0.690 0.432 0.190 0.066 0.000 0.000 0.000 Negative Control 0.672 0.668 0.672 0.668 0.664 0.663 0.662 209 Table B-6 (cont’d) LDPE (100% peel) 1 2 3 0.677 0.221 0.074 0.000 0.000 0.000 0.000 0.686 0.310 0.129 0.011 0.001 0.000 0.000 0.683 0.480 0.314 0.215 0.129 0.067 0.023 Negative Control 0.672 0.668 0.672 0.668 0.664 0.663 0.662 LDPE (50% seed + 50% peel) 1 2 3 0.651 0.315 0.162 0.068 0.023 N/A N/A 0.651 0.234 0.106 0.019 0.000 N/A N/A 0.650 0.235 0.057 0.000 0.000 N/A N/A Negative Control 0.672 0.668 0.672 0.668 0.664 N/A N/A 210 Appendix C Mode of antioxidant activity of coated polymer films Dose-response model Boltzmann sigmoidal model A. Trial 1 Figure C-1 Goodness of fit results estimated by the dose-response and Boltzmann sigmoidal models for the antioxidative recoverability test. 211 Figure C-1 (cont’d) Dose-response model Boltzmann sigmoidal model B. Trial 2 212 Figure C-1 (cont’d) Dose-response model Boltzmann sigmoidal model C. Trial 3 213 Figure C-1 (cont’d) Dose-response model Boltzmann sigmoidal model D. Trial 4 214 Figure C-1 (cont’d) Dose-response model Boltzmann sigmoidal model E. Trial 5 215 Table C-1 Fitted regression equations estimated by the dose-response and Boltzmann sigmoidal models for the antioxidative recoverability test. Boltzmann Sigmoidal Model − C′E ;<=>?)+C′E A= 1+FGH(7978 − 0.748 A= 9I.LET)+0.748 1+FGH(79I.TNI − 0.736 A= 9I.EMI)+0.736 1+FGH(79I.PMT −0.717 A= 9I.ELM)+0.717 1+FGH(79L.LNI −0.751 A= 9I.LEN)+0.751 1+FGH(79L.LLV −0.727 A= 9I.LNQ)+0.727 1+FGH(79L.EVE Trial 1 Trial 2 Trial 3 Trial 4 Trial 5 Dose-response Model IC (%)= IC (%)= IC (%)= IC (%)= IC (%)= IC (%)= 100 100 100 100 100 100 1+10[(7897)×;<=>?] 1+10[(I.TNE97)×M.VLN] 1+10[(I.PMP97)×L.NPM] 1+10[(L.LNI97)×E.IQI] 1+10[(L.LEI97)×M.QQM] 1+10[(L.EVE97)×E.MTM] 216 Table C-2 Experimental data of the antioxidative recoverability test. Absorbance 0 min 6 min 12 min 18 min 24 min 30 min 36 min 42 min 48 min 54 min 0.748 0.252 0.017 0.000 N/A N/A N/A N/A N/A N/A 0.745 0.155 0.000 0.000 N/A N/A N/A N/A N/A N/A 0.748 0.295 0.083 0.002 N/A N/A N/A N/A N/A N/A 0.734 0.397 0.153 0.330 0.000 0.000 0.000 N/A N/A N/A 0.735 0.504 0.293 0.152 0.043 0.004 0.000 N/A N/A N/A 0.736 0.492 0.367 0.188 0.118 0.069 0.020 N/A N/A N/A 0.716 0.608 0.350 0.286 0.226 0.160 0.082 0.032 0.006 0.000 0.717 0.543 0.347 0.275 0.215 0.174 0.114 0.072 0.018 0.000 0.717 0.655 0.532 0.423 0.288 0.185 0.095 0.050 0.009 0.000 0.751 0.682 0.282 0.147 0.042 0.000 N/A N/A N/A N/A 0.736 0.689 0.511 0.346 0.098 0.007 N/A N/A N/A N/A 0.744 0.701 0.452 0.215 0.022 0.000 N/A N/A N/A N/A 0.727 0.632 0.560 0.472 0.387 0.231 0.181 0.128 0.075 0.021 0.726 0.646 0.508 0.391 0.261 0.152 0.052 0.000 0.000 0.000 0.727 0.577 0.410 0.326 0.221 0.151 0.074 0.010 0.000 0.000 Trial Sample 1 2 3 4 5 1 2 3 1 2 3 1 2 3 1 2 3 1 2 3 Negative Control 0.712 0.705 0.708 0.701 0.696 0.693 0.689 0.690 0.692 0.697 217 . s b A . s b A 0.773 0.600 0.400 0.200 0.000 -0.100 190.00 0.773 0.600 0.400 0.200 0.000 -0.100 190.00 350.00 400.00 250.00 300.00 Wavelength ( ) nm. A. The crude phenolic extract in 50% ethanol 250.00 300.00 Wavelength ( ) nm. 350.00 400.00 B. The crude phenolic extract in 10% ethanol Figure C-2 Overlapped UV-Vis spectra of the crude phenolic extract from avocado peels in aqueous ethanol at various concentrations. 218 . s b A 0.773 0.600 0.400 . s b A 0.200 0.000 -0.100 190.00 Sample 1 250.00 300.00 Wavelength ( ) nm. 350.00 400.00 Sample 2 Figure C-3 UV-Vis spectra of phenolic compounds released from coated PP films in each migration cell. 219 Figure C-3 (cont’d) Sample 3 A. Coated PP films in 95% aqueous ethanol at 4°C 0.773 0.600 . s b A 0.400 . s b A 0.200 0.000 -0.100 190.00 250.00 300.00 Wavelength ( ) nm. 350.00 400.00 Sample 1 220 Figure C-3 (cont’d) Sample 2 250.00 300.00 nm. Wavelength ( ) 350.00 400.00 Sample 3 0.773 0.600 0.400 . s b A . s b A 0.200 0.000 -0.100 190.00 B. Coated PP films in 50% aqueous ethanol at 4°C 221 Figure C-3 (cont’d) Sample 1 Sample 2 222 Figure C-3 (cont’d) Sample 3 C. Coated PP films in 10% aqueous ethanol at 4°C . s b A 0.773 0.600 0.400 . s b A 0.200 0.000 -0.100 190.00 250.00 300.00 nm. Wavelength ( ) 350.00 400.00 Sample 1 223 Figure C-3 (cont’d) Sample 2 Sample 3 D. Coated PP films in 95% aqueous ethanol at 40°C 224 Figure C-3 (cont’d) 0.773 0.600 0.400 . s b A . s b A 0.200 0.000 -0.100 190.00 250.00 300.00 nm. Wavelength ( ) 350.00 400.00 Sample 1 Sample 2 225 Figure C-3 (cont’d) . s b A . s b A 0.773 0.600 0.400 0.200 0.000 -0.100 190.00 250.00 300.00 nm. Wavelength ( ) 350.00 400.00 Sample 3 E. Coated PP films in 50% aqueous ethanol at 40°C Sample 1 226 Figure C-3 (cont’d) Sample 2 Sample 3 F. Coated PP films in 10% aqueous ethanol at 40°C 227 Table C-3 Phenolic compound size referenced for the mode of activity analysis of the antioxidant coating. Compound Name Catechin Quercetin-3-O-rutinoside (rutin) Protocatechuic Acid Size (nm) 0.8×1.3 1.27×0.6 0.9×0.68×0.29 References Ariga (2012) Yang et al. (2015) Barahuie et al. 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