Copyright is owned by the Author of the thesis. Permission is given for a copy to be downloaded by an individual for the purpose of research and private study only. The thesis may not be reproduced elsewhere without the permission of the Author. Studies on formation, oxidative stability and plausible applications of food-grade ‘droplet-stabilised’ oil-in-water emulsions A thesis presented in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Food Technology at Riddet Institute, Massey University Palmerston North, New Zealand. Okubanjo Samantha Sewuese 2019 Abstract I Abstract This research was aimed at studying the structural characteristics, chemical stability and plausible functional applications of droplet-stabilised oil-in-water emulsions (DSEs). DSEs consist of oil-in-water droplets (the core) stabilised by submicron protein-stabilised oil droplets (the shell). The first objective was to increase our understanding of their structural properties and processing factors that contribute to DSE formation using food grade ingredients. To achieve this objective, milk protein concentrate (MPC) was chosen as the emulsifier. Four MPCs with different levels of calcium were used. The surface lipid (20 %) consisted of either a low (olive oil), medium (palmolein oil) or high (trimyristin) melting surface lipid. The core lipid (20 %) consisted of either a triglyceride (soybean oil) or pure fatty acid (linoleic acid). Protein- stabilised shell emulsions were processed either via the microfluidizer (170 MPa) or two-stage homogeniser (1st stage-20 MPa; 2nd stage-4 MPa). Results of the study showed that aggregated structure of protein emulsifier, shell droplet concentration, surface and core lipid types influenced the formation and structural properties of DSEs. The second objective focused on investigating the chemical stability of DSEs by evaluating their stability to oxidation and ability of its interfacial structure to protect polyunsaturated lipids incorporated within from oxidation. To achieve this objective, oxidative stability of high linoleic acid oil (safflower oil) stabilised by protein-coated low (olive oil), medium (palmolein oil) and high (trimyristin) melting lipid droplets was evaluated and compared with composition-matched conventional protein-stabilised safflower oil-in-water emulsion as well as a Abstract II conventional protein-stabilised safflower oil-in-water emulsion (reference emulsions). Influence of physical state of high melting lipid droplets on oxidative stability of droplet-stabilised safflower oil emulsion was also evaluated. High linoleic acid (72.54% of total fatty acids) safflower oil (20%) was used because of its high susceptibility to oxidation. Olive oil (low acidity), palmolein oil and trimyristin were chosen because of their low susceptibility to oxidation. The study showed that safflower oil oxidation in DSEs was reduced by about 40-55% in comparison to conventional emulsions. High melting surface lipid DSEs provided better protection for safflower oil than low and medium melting surface lipid DSEs. The third objective aimed at improving our understanding of the influence of antioxidant’s location in emulsions on antioxidant performance. The study was also focused on exploring a plausible functional application of DSEs by incorporating a hydrophobic antioxidant in shell droplets (at the interface) of DSEs rather than in the interior of the core unsaturated lipid. To achieve this objective, butylated hydroxyanisole (BHA) a common commercially used synthetic hydrophobic antioxidant was chosen. BHA was incorporated either in shell droplets or core droplets of DSEs. The ability of BHA to counteract oxidation when incorporated in low (olive oil) and high melting (trimyristin) shell droplets of DSEs was evaluated and compared with BHA’s anti-oxidation performance when incorporated directly in core droplets (safflower oil) stabilised by low (olive oil) and high melting (trimyristin) shell droplets without BHA. Results of the study indicate that ability of BHA-in-shell DSEs to counteract oxidation of core safflower oil better than BHA-in-core DSEs is influenced by BHA’s concentration and transfer mechanism to reaction sites. Abstract III The fourth and final objective was aimed at investigating mobility of a hydrophobic antioxidant incorporated at the interface of DSEs to establish their location after emulsification. The study focused on determining if a hydrophobic antioxidant incorporated in shell droplets remained localised within or migrated overtime to core droplets. The study also investigated the use of two techniques (saturated transfer difference (STD)-nuclear magnetic resonance and confocal Raman microscopy) to determine partitioning of antioxidants in DSE. To achieve this objective, confocal Raman spectroscopy technique was employed to probe antioxidant location without phase separation or destruction of DSE structure. Beta-carotene was chosen for the study for its excellent Raman scattering property. Beta-carotene was incorporated either in shell droplets (olive oil and trimyristin) or core droplets (safflower oil) of DSEs. Location and mobility of beta- carotene was evaluated after three days production. Beta-carotene migration from low (olive) and high melting shell droplets to core safflower oil was minimal. The present study provides processing conditions and structural characteristics required to form food-grade DSEs. The study confirms and establishes the potential of DSEs to effectively protect oxidation-sensitive lipophilic bioactives incorporated within from degradation and confirms the viability of concurrent incorporation of two different bioactives in DSEs emulsions by locating one bioactive in shell droplets and the second within the core. Acknowledgements IV Acknowledgements I am extremely grateful to my main supervisor Dr Simon Loveday for his guidance, patience and encouragement during the entire period of my research. I am profoundly grateful to him for constantly pushing me in the right direction and ensuring I did not lose focus of the main objectives. I am very grateful to my co-supervisors Associate Professor Aiqian Ye, Professor Peter Wilde, and Distinguished Professor Harjinder Singh for their support, guidance and for sharing their vast knowledge and expertise with me. I enjoyed my discussions with each one of them and the expert insights they always provided. A special thank you to Professor Peter Wilde for his committed supervision, contributions, and guidance miles away from New Zealand. I am very grateful to him for attending all my meetings via video conference and for very prompt responses to my emails and questions. A huge thank you to Niki Minards for training on confocal and DIC microscopy and for answering all my questions which ensured I got very clear images. I am also grateful to Sushila Pillai of Victoria University, Wellington for training on how to use Victoria University’s confocal microscope when the confocal at Massey University was unavailable. A huge thanks to Mr. Chris Hall, Dr. Peter Zhu, Ms. Janiene Gilliland and Ms. Maggie Zou for equipment trainings, assistance, advice, and co-operation in my experimental work. I wish to thank Ms. Janine Gilliland and Ms. Maggie Zou specifically for helping with chemicals, reagents and consumable orders and purchases. Acknowledgements V I will like to particularly acknowledge and thank the following for their contribution to this research work  Associate Professor Mark Waterland for his expertise on Raman spectroscopy and for the training on using python to analyse the Raman data and introducing me to Anaconda. I am equally grateful to him for checking the python codes and going over the data analysis.  Sam Brooke also for his expertise on Raman spectroscopy and especially for accompanying me to Auckland university of Technology to collect the Confocal Raman data.  Nic Mostert for writing the python codes used for Confocal Raman data analysis.  Rob Ward for his expertise on microfluidics and for providing the microfluidic channel used for Confocal Raman spectroscopy. Also, for designing and producing the microfluidic channel holder.  Pat Edward for his expertise on Nuclear Magnetic Resonance (NMR) and for data collection and analysis.  Dr. Simon Loveday for oxidation data integration. I will like to acknowledge Massey University and Riddet Institute for providing scholarships to financially support my research. I am particularly grateful to Riddet institute for travel grants to present at international conferences in Australia. I also wish to express my gratitude to Riddet institute staff Ms. Ansley Te Hiwi, Ms. Terri Palmer and Mr. John Henley-King for their administrative Acknowledgements VI support throughout the period of my research. A huge thank you to Mr. Matt Levin for providing technology support throughout the period of my research. I am most grateful to my life-group family for the spiritual and emotional support. To my lovely friends Mr. & Dr. Fong and Maria Au-young for their emotional and spiritual support, for all the times my daughter Ana stayed with you while I attended conferences or worked in the laboratory I am extremely grateful, for all the times you had me and my family over for meals I am sincerely thankful, your support has been tremendously amazing and I really do not have the right words to express my gratitude. To my lovely friend Dr. Adeyinka for always opening your home to my daughter and I, I am sincerely grateful for all the times Ana stayed with you so I could attend conferences, work in the laboratory or focus on my research. Your love, support, and encouragement has been fantastic, and I can never thank you enough. I am also grateful to Prof. Adeyinka for the training on R statistical software and for allowing your wife Dr. Adeyinka support me throughout my PhD. To Ms. Oyelere and family I am very grateful for your support and encouragement, for all the times you helped with Ana I am so grateful. To Ms. Rowena Brown and Family, thank you so much for the encouragements and all the time you had my family over for dinner. To Mr. & Mrs. Brian and Shirley Pegler I am sincerely thankful for the emotional and spiritual support, for all the times you took my family out for lunch and for all the gifts for my daughter. I also acknowledge the emotional support of close friends who have made my stay in Palmerston North enjoyable. This journey could never have been possible without the love and support of my family. To my parents Mr. and Mrs. Jonathan Ichaver, your words of Acknowledgements VII encouragement kept me going on this journey and I am forever grateful for the training and love. To my lovely sister Ier Jonathan-Ichaver, you have been an amazing pillar of support, for all the times we spent talking and you kept cheering me on and your words of encouragement I am extremely grateful, I always came off those calls feeling energized and ready to finish what I started, thank you so much for sharing my dream, you are indeed a priceless sister, To my sisters, Ms Doosuur Tor-anyiin and Ms. Kashimana Ichaver thank you for the emotional support and love. To my dear brother Mr. Bem Ichaver thank you for the love and words of encouragement. To my brothers-in-law Prof. Yinka Okubanjo and Mr. Yemi Okubanjo thank you so much for your love, encouragement and support, for all the chats and words of encouragement I am sincerely grateful. Finally, I will like to acknowledge the tremendous love and support of my best friend and husband, without you Mr. Tosin Okubanjo this journey could not have been successful, thank you for sharing and living this dream with me, thank you for supporting me all the way and for the huge sacrifices you have made to ensure I accomplish this dream. To my beautiful and lovely daughter Ms. Ana Okubanjo, you are the most beautiful gift I have ever received and thank you for embarking on this long, bitter-sweet journey with me, your sweet words of encouragement motivated me every day, I looked forward to your hugs and laughter after each difficult day or moment, My darling Ana I dedicate this thesis to you and I hope that it will inspire you in future and help you accomplish your dream of becoming a scientist and inventor. Table of Contents VIII Table of Contents Abstract ............................................................................................................................. I Acknowledgements ......................................................................................................... IV Table of Contents .......................................................................................................... VIII List of Figures ................................................................................................................ XIV List of Tables .................................................................................................................. XX List of Publications ....................................................................................................... XXII CHAPTER 1: INTRODUCTION .................................................................................................... 1 1.1 GENERAL INTRODUCTION ....................................................................................................... 1 CHAPTER 2: LITERATURE REVIEW ......................................................................................... 8 2.1 CLASSES OF EMULSIONS, PREPARATION METHODS AND EMULSIFIERS ....................................... 8 2.1.1 Emulsion preparation .................................................................................................. 9 2.1.2 Micro-emulsions and nano-emulsions ......................................................................... 9 2.1.3 Double Emulsions ..................................................................................................... 10 2.1.4 Pickering emulsions .................................................................................................. 12 2.1.5 Multi-layered emulsions ............................................................................................. 13 2.1.6 Emulsifier types ......................................................................................................... 14 2.2 STRUCTURE OF FOOD EMULSIONS ........................................................................................ 16 2.2.1 Physical stability ........................................................................................................ 16 2.2.2 Chemical Instability ................................................................................................... 24 2.3 LIPID OXIDATION .................................................................................................................. 24 2.3.1 Auto-oxidation............................................................................................................ 25 2.3.2 Enzymatic oxidation .................................................................................................. 28 2.3.3 Photo-oxidation ......................................................................................................... 28 2.4 STRUCTURAL FACTORS AFFECTING THE OXIDATIVE STABILITY OF EMULSIONS ......................... 31 2.5 ANTIOXIDANTS .................................................................................................................... 35 2.5.1 Primary antioxidants .................................................................................................. 35 2.5.2 Secondary Antioxidants ............................................................................................. 36 Table of Contents IX 2.5.3 The Polar Paradox .................................................................................................... 38 2.5.4 Antioxidant location ................................................................................................... 42 2.6 EMULSIONS AS DELIVERY SYSTEMS FOR BIOACTIVES ............................................................. 44 2.6.1 Influence of droplet size on encapsulated bioactives ................................................ 47 2.6.2 Influence of emulsion composition and interfacial structure on encapsulated bioactives ............................................................................................................................ 48 2.7 PICKERING EMULSIONS IN FOOD ........................................................................................... 51 2.8 SUMMARY AND CONCLUSIONS ............................................................................................. 54 CHAPTER 3: MATERIALS AND METHODS ............................................................................. 56 3.1 MATERIALS ......................................................................................................................... 56 3.1.1 Water ......................................................................................................................... 56 3.1.2 Milk Protein concentrate ............................................................................................ 56 3.1.3 Surface lipids ............................................................................................................. 59 3.1.4 Core Lipids ................................................................................................................ 59 3.1.5 Antioxidants ............................................................................................................... 60 3.1.6 Reagents/Chemicals ................................................................................................. 61 3.2 EQUIPMENT......................................................................................................................... 62 3.2.1 Waterbath .................................................................................................................. 62 3.2.2 pH Meter .................................................................................................................... 62 3.2.3 Centrifuge .................................................................................................................. 62 3.2.4 Spectrophotometer .................................................................................................... 62 3.2.5 Gas Chromatograph .................................................................................................. 63 3.2.6 Homogenisers, microfluidizer and mixers ................................................................. 63 3.3 METHODS ........................................................................................................................... 66 3.3.1 Shell emulsion preparation ........................................................................................ 66 3.3.2 Droplet-stabilised emulsions ..................................................................................... 67 3.3.3 Particle size characterisation..................................................................................... 67 3.3.4 Confocal microscopy ................................................................................................. 67 3.3.5 Differential Interference Contrast (DIC) Microscopy ................................................. 68 3.3.6 Accelerated Lipid Oxidation ....................................................................................... 69 Table of Contents X 3.3.7 Preparation of Ferrous ion ......................................................................................... 70 3.3.8 Lipid Oxidation Measurements .................................................................................. 70 3.3.9 Sample extraction ...................................................................................................... 70 3.3.10 Determination of Conjugated Dienes ...................................................................... 70 3.3.11 Determination of Lipid Hydroperoxides ................................................................... 72 3.3.12 Determination of Volatile Secondary Oxidation Product (Hexanal) ........................ 73 3.4 STATISTICAL ANALYSIS ........................................................................................................ 74 3.5 SOFTWARE PACKAGES ........................................................................................................ 74 CHAPTER 4: FORMATION AND PROPERTIES OF DROPLET-STABILISED FOOD GRADE OIL-IN-WATER EMULSIONS..................................................................................................... 75 4.1 ABSTRACT .......................................................................................................................... 75 4.2 INTRODUCTION .................................................................................................................... 79 4.3 MATERIALS AND METHODS .................................................................................................. 81 4.3.1 Droplet stabilised emulsions (DSEs) ......................................................................... 82 4.4 RESULTS AND DISCUSSION .................................................................................................. 85 4.4.1 Formation of droplet-stabilised soybean oil-in-water emulsions with high protein MPC ............................................................................................................................................ 85 4.4.2 Effect of shell emulsion concentration on formation of droplet-stabilised emulsions (high protein MPC) ............................................................................................................. 89 4.4.3 Effect of low protein MPC on formation of droplet-stabilised emulsions ................... 93 4.4.4 Effect of calcium-depleted MPC on the formation of droplet-stabilised emulsions ... 97 4.4.5 Effect of microfluidization on formation of droplet-stabilised emulsions with high protein and calcium-depleted MPC .................................................................................. 101 4.4.6 Effect of core lipid type on formation of droplet-stabilised emulsions ..................... 106 4.4.7 Effect of surface lipid type and state on formation of droplet-stabilised emulsions 111 4.5 CONCLUSIONS .................................................................................................................. 120 CHAPTER 5: EFFECT OF DROPLET-STABILISED OIL-IN-WATER EMULSIONS ON OXIDATIVE STABILITY OF UNSATURATED LIPID INCORPORATED WITHIN .................. 122 5.1 ABSTRACT ........................................................................................................................ 123 Table of Contents XI 5.2 INTRODUCTION .................................................................................................................. 125 5.3 MATERIALS AND METHODS ................................................................................................. 128 5.3.1 Materials .................................................................................................................. 128 5.3.2 Droplet-stabilised emulsion preparation .................................................................. 128 5.3.3 Accelerated Lipid Oxidation ..................................................................................... 131 5.3.4 Lipid Oxidation Measurements ................................................................................ 131 5.3.5 Lipid oxidation measurements................................................................................. 131 5.3.6 Statistical Analysis ................................................................................................... 132 5.4 RESULTS .......................................................................................................................... 133 5.4.1 Formation of droplet-stabilised and control emulsions ............................................ 133 5.4.2 Impact of droplet-stabilised emulsion structure on oxidative stability of core safflower oil ...................................................................................................................................... 137 5.4.3 Effect of surface lipid type on oxidative stability of droplet-stabilised safflower oil emulsion ........................................................................................................................... 148 5.4.4 Impact of surface lipid state on oxidative stability of droplet-stabilised emulsion ... 149 5.5 DISCUSSION ...................................................................................................................... 152 CHAPTER 6: IMPACT OF ANTIOXIDANT LOCATION IN DROPLET-STABILISED OIL-IN- WATER EMULSIONS ON OXIDATIVE STABILITY OF UNSATURATED LIPID INCORPORATED WITHIN ....................................................................................................... 160 6.1 ABSTRACT ........................................................................................................................ 160 6.2 INTRODUCTION .................................................................................................................. 162 6.3 MATERIALS AND METHODS ................................................................................................. 164 6.3.1 Antioxidant-loaded droplet-stabilised emulsions ..................................................... 164 6.3.2 Accelerated Lipid Oxidation ..................................................................................... 168 6.3.3 Lipid Oxidation Measurements ................................................................................ 168 6.4 RESULTS .......................................................................................................................... 175 6.4.1 Particle size and microscopic structure of droplet-stabilised oil-in-water emulsions .......................................................................................................................................... 175 6.4.2 Impact of incorporating BHA (50 ppm) in shell droplets versus core droplets of droplet-stabilised emulsions on oxidative stability ............................................................ 181 Table of Contents XII 6.4.3 Impact of incorporating BHA (500 ppm) in shell droplets versus core droplets of droplet-stabilised emulsions on oxidative stability ............................................................ 186 6.5 DISCUSSION ...................................................................................................................... 191 CHAPTER 7: PROBING THE LOCATION OF ANTIOXIDANTS INCORPORATED IN DROPLET-STABILISED OIL-IN-WATER EMULSIONS ......................................................... 202 7.1 ABSTRACT ........................................................................................................................ 202 7.2 INTRODUCTION .................................................................................................................. 205 7.3 MATERIALS AND METHODS ................................................................................................. 207 7.3.1 Saturation Transfer Nuclear Magnetic Resonance ................................................. 207 7.3.2 Confocal Raman Spectroscopy ............................................................................... 209 7.3.3 Raman Data Analysis .............................................................................................. 210 7.4 RESULTS .......................................................................................................................... 218 7.4.1 Particle size of antioxidant loaded droplet-stabilised emulsions ............................. 218 7.4.2 Saturated transfer difference (STD) ........................................................................ 221 7.4.3 Confocal Raman Microscopy (microscope slides) .................................................. 224 7.4.4 Confocal Raman Microscopy (Microfluidic channel) ............................................... 229 7.5 DISCUSSION ...................................................................................................................... 243 CHAPTER 8: FINAL PERSPECTIVES AND RECOMMENDATIONS .................................... 247 8.1 SUMMARY ......................................................................................................................... 247 8.2 FORMATION OF FOOD-GRADE DROPLET-STABILISED OIL-IN-WATER EMULSIONS ..................... 248 8.3 LIPID OXIDATION IN OIL-IN-WATER EMULSIONS ..................................................................... 251 8.4 LOCATION-BASED ANTIOXIDANT PERFORMANCE IN EMULSIONS ............................................. 254 8.5 MOBILITY OF ANTIOXIDANTS INCORPORATED IN EMULSIONS ................................................. 257 8.6 OUTCOMES AND APPLICATIONS .......................................................................................... 259 8.7 RECOMMENDATIONS FOR FUTURE RESEARCH ..................................................................... 262 References............................................................................................................... 266 Appendix A (chapter 4) ........................................................................................ 286 A.1 Exploring uniform interfacial droplet stabilisation with MPC shell emulsions processed via the microfluidizer .............................................. 286 Table of Contents XIII A.2 Crystal morphology in Trimyristin droplet-stabilised emulsions ..... 291 Appendix B (chapter 5) ..................................................................................... 296 B.1 Accelerated lipid oxidation conditions (Dark Vs Light) ..................... 296 B.2 Spontaneous adsorption of shell droplets .......................................... 298 Appendix C (chapter 6) ..................................................................................... 299 C.1 Evolution of hexanal in DSEs with BHA (500 ppm) ............................ 299 Appendix D (chapter 7) ..................................................................................... 300 D.1 Raman spectroscopy of Olive DSEs with BHA (Microscope slides) 300 List of Figures XIV List of Figures FIGURE 1.1: SCHEMATIC DIAGRAM OF ‘DROPLET-STABILISED’ EMULSION ............................................. 3 FIGURE 1.2 THESIS STRUCTURE ...................................................................................................... 7 FIGURE 2.1 SCHEMATIC DIAGRAM OF SIMPLE (O/W & W/O). ADAPTED FROM CHUNG AND MCCLEMENTS (2015) ............................................................................................................. 8 FIGURE 2.2: SCHEMATIC DIAGRAM OF DOUBLE EMULSIONS (W/O/W & O/W/W). ADAPTED FROM MARTÍNEZ-PALOU ET AL. (2011) ............................................................................................ 11 FIGURE 2.3: SCHEMATIC DIAGRAM OF PICKERING EMULSION. ADAPTED FROM CHEVALIER AND BOLZINGER (2013) ............................................................................................................... 12 FIGURE 2.4: SCHEMATIC DIAGRAM OF MULTI-LAYERED EMULSION AND PREPARATION STEPS. ADAPTED FROM MCCLEMENTS AND LI (2010) ....................................................................................... 14 FIGURE 2.5 EMULSION INSTABILITY SCHEMATICS. ............................................................................ 18 FIGURE 2.6 BRIDGING AND DEPLETION FLOCCULATION SCHEMATICS ................................................ 20 FIGURE 2.7 INITIATION OF LIPID OXIDATION PATHWAYS- A) INITIATION BY HYDROGEN ABSTRACTION; B) INITIATION BY FREE-RADICAL ATTACK ON A DOUBLE BOND; C) INITIATION BY SINGLET OXYGEN. ADAPTED FROM KANNER ET AL. (1987). ................................................................................. 25 FIGURE 2.8: OXIDATION PATHWAY OF LINOLEIC ACID. ADAPTED FROM WHEATLEY (2000) ................. 30 FIGURE 2.9: SCHEMATIC DIAGRAM OF THE POLAR PARADOX THEORY: PARTITIONING OF ANTIOXIDANTS IN BULK OILS (A & B) AND OIL-IN-WATER EMULSIONS (C). ADAPTED WITH PERMISSION FROM FEREIDOON SHAHIDI AND ZHONG (2011), COPYRIGHT (2018). ............................................... 39 FIGURE 3.1 SUMMARY OF EXPERIMENTAL SET-UP .......................................................................... 57 FIGURE 3.2 MICROFLUIDIZER (MICROFLUIDICS M-110P) ................................................................. 64 FIGURE 3.3 TWO-STAGE HOMOGENISER (RANNIE 2.5H) ................................................................. 64 FIGURE 3.4 TWO-STAGE HOMOGENISER (FBF ITALIA) ..................................................................... 65 FIGURE 3.5 HAND HELD MIXER (LABSERV D-130) ........................................................................... 65 FIGURE 3.6: ACCELERATED LIPID OXIDATION SET-UP WITH TWO FLUORESCENT LAMPS- A) FLUORESCENT LAMPS; B) EMULSION SAMPLES IN PLATES AND VIALS EXPOSED TO FLUORESCENT LAMPS .................................................................................................................................. 69 FIGURE 4.1 FLOW CHART OF DROPLET-STABILISED OIL-IN-WATER EMULSION PRODUCTION ................ 84 List of Figures XV FIGURE 4.2 PARTICLE SIZE DISTRIBUTION OF SHELL EMULSIONS (A) AND DROPLET-STABILISED SOYBEAN OIL-IN-WATER EMULSIONS (B) MADE WITH MPC 1 AND 2 .......................................... 87 FIGURE 4.3 CONFOCAL MICROSCOPY IMAGES OF DROPLET-STABILISED EMULSIONS MADE WITH MPC 1 AND 2 SHELL EMULSIONS PROCESSED VIA THE TWO-STAGE HOMOGENISER (B & D = ZOOM). ..... 88 FIGURE 4.4: PARTICLE SIZE DISTRIBUTION AND CONFOCAL MICROSCOPY IMAGES OF DROPLET- STABILISED EMULSIONS PROCESSED WITH MPC 1 AND VARYING CONCENTRATIONS OF SHELL EMULSION ............................................................................................................................. 92 FIGURE 4.5: PARTICLE SIZE DISTRIBUTION (A & B) AND CONFOCAL MICROSCOPY IMAGES (C, D, E & F) OF DROPLET-STABILISED EMULSIONS PROCESSED WITH MPC 1 AND 3. .................................... 96 FIGURE 4.6: PARTICLE SIZE DISTRIBUTION OF SHELL (A) AND DROPLET-STABILISED (B) EMULSIONS MADE WITH MPC 4 SHELL EMULSION PROCESSED VIA THE TWO-STAGE HOMOGENISER ............. 99 FIGURE 4.7: CONFOCAL MICROSCOPY IMAGES OF DROPLET-STABILISED EMULSIONS MADE WITH MPC 4 SHELL EMULSIONS PROCESSED VIA THE TWO-STAGE HOMOGENISER ...................................... 100 FIGURE 4.8 PARTICLE SIZE DISTRIBUTION OF SHELL (A) AND DROPLET-STABILISED (B) EMULSIONS MADE WITH MPC 1 AND 4 SHELL EMULSION PROCESSED VIA THE MICROFLUIDIZER ................. 104 FIGURE 4.9 CONFOCAL MICROSCOPY IMAGES OF DROPLET-STABILISED EMULSIONS MADE WITH MPC 1, 2 AND 4 SHELL EMULSIONS PROCESSED VIA THE MICROFLUIDIZER .......................................... 105 FIGURE 4.10: PARTICLE SIZE DISTRIBUTION OF DROPLET-STABILISED SOYBEAN OIL AND LINOLEIC ACID CORE EMULSIONS PREPARED WITH MPC 1 AT 10% (A) AND 16% (B) SHELL EMULSION CONCENTRATIONS............................................................................................................... 109 FIGURE 4.11: CONFOCAL MICROSCOPY IMAGES OF DROPLET-STABILISED LINOLEIC ACID CORE EMULSIONS MADE WITH MPC 1 (A & B= 10% SHELL EMULSION; C, D, E, F, G & H= 16% SHELL EMULSION) ......................................................................................................................... 110 FIGURE 4.12 PARTICLE SIZE DISTRIBUTION OF SHELL (A) AND DROPLET STABILISED EMULSIONS (B) COMPRISING SURFACE LIPIDS OF VARYING MELTING TEMPERATURE ....................................... 116 FIGURE 4.13 CONFOCAL MICROSCOPY IMAGES OF DROPLET STABILISED EMULSIONS COMPRISING SURFACE LIPIDS OF VARYING MELTING TEMPERATURE ........................................................... 117 FIGURE 4.14 CONFOCAL MICROSCOPY IMAGES OF DROPLET STABILISED EMULSIONS WITH TRIMYRISTIN SURFACE LIPID- NT= FINAL HOMOGENISATION ABOVE 56°C AND NT2 = FINAL HOMOGENISATION BELOW 56°C. ..................................................................................................................... 118 List of Figures XVI FIGURE 4.15 CONFOCAL MICROSCOPY IMAGES OF MIXED SAFFLOWER OIL AND TRIMYRISTIN- A & B (MIXTURE HEATED TO 65°C) AND SAFFLOWER OIL/TRIMYRISTIN COARSE EMULSION HOMOGENISED BELOW MELTING TEMPERATURE- C, D & E). IMAGE B & D TAKEN IN DIC MODE. ..................... 119 FIGURE 5.1 FLOW CHART OF DROPLET-STABILISED OIL-IN-WATER EMULSION PRODUCTION .............. 130 FIGURE 5.2: CONFOCAL MICROSCOPY IMAGES OF CONTROL AND ‘DROPLET-STABILISED’ EMULSIONS ......................................................................................................................................... .136 FIGURE 5.3 EVOLUTION OF PRIMARY (CONJUGATED DIENE-A, B & C AND HYDROPEROXIDES- D, E & F) AND SECONDARY (HEXANAL- G, H & I) OXIDATION PRODUCTS IN CORE SAFFLOWER OIL IN ‘DROPLET-STABILISED’ AND CONTROL EMULSIONS EXPOSED TO FLUORESCENT LAMP IN THE PRESENCE OF FERROUS IRON (100ΜM). .............................................................................. 140 FIGURE 5.4 CONFOCAL MICROSCOPY IMAGES OF CONTROL EMULSIONS FRESHLY PREPARED (DAY 0) AND AFTER ACCELERATED OXIDATION OF CORE SAFFLOWER OIL (DAY 5 AND 7). ..................... 144 FIGURE 5.5 CONFOCAL MICROSCOPY IMAGES OF DROPLET STABILISED EMULSIONS FRESHLY PREPARED (DAY 0) AND AFTER ACCELERATED OXIDATION OF CORE SAFFLOWER OIL (DAY 5 AND 7). ........ 146 FIGURE 5.6 PARTICLE SIZE DISTRIBUTION OF ‘DROPLET STABILISED’ EMULSION AND CONTROL EMULSIONS FRESHLY PREPARED (A, B & C) AND DURING ACCELERATED OXIDATION-DAY 5 (D, E & F) AND DAY 7 (G, H & I). ..................................................................................................... 147 FIGURE 5.7 CONFOCAL (TOP) AND DIFFERENTIAL INTERFERENCE CONTRAST (BOTTOM) MICROSCOPY IMAGES OF DROPLETS STABILISED EMULSIONS WITH TRIMYRISTIN SURFACE LIPID; NT-PROCESSED ABOVE MELTING TEMPERATURE; NT2-PROCESSED BELOW MELTING TEMPERATURE - ARROWS SHOW DIFFERENT CRYSTAL MORPHOLOGY FOR NT2 ............................................................. 150 FIGURE 5.8 SCHEMATIC DIAGRAM OF OXIDATION IN CONVENTIONAL AND DROPLET-STABILISED EMULSIONS. A)- CONVENTIONAL EMULSION; B)- DROPLET-STABILISED EMULSION WITH LOW OR MEDIUM MELTING SURFACE LIPID; C)- DROPLET-STABILISED EMULSION DROPLET WITH HIGH MELTING SURFACE LIPID. ..................................................................................................... 151 FIGURE 6.1 FLOW CHART OF SURFACE-LOADED BHA DROPLET-STABILISED EMULSION FORMATION… .......................................................................................................................................... 166 FIGURE 6.2 FLOW CHART OF CORE-LOADED BHA DROPLET-STABILISED EMULSION FORMATION ...... 167 FIGURE 6.3 CONFOCAL MICROSCOPY IMAGES OF DROPLET-STABILISED EMULSIONS WITHOUT BHA AND WITH BHA (50 PPM) IN SHELL DROPLETS OR CORE DROPLETS ............................................... 177 List of Figures XVII FIGURE 6.4 CONFOCAL MICROSCOPY IMAGES OF DROPLET-STABILISED EMULSIONS WITHOUT BHA AND WITH BHA (500 PPM) IN SHELL DROPLETS OR CORE DROPLETS ............................................. 179 FIGURE 6.5 EVOLUTION OF CONJUGATED DIENES, LIPID HYDROPEROXIDES AND HEXANAL IN DROPLET- STABILISED SAFFLOWER OIL-IN-WATER EMULSIONS WITHOUT BHA AND WITH BHA (50 PPM). .. 183 FIGURE 6.6 PARTICLE SIZE DISTRIBUTION OF DROPLET-STABILISED SAFFLOWER OIL-IN-WATER EMULSIONS WITHOUT BHA AND WITH BHA (50 PPM) FRESHLY PREPARED AND DURING ACCELERATED OXIDATION (DAY 5 AND 7) .............................................................................. 184 FIGURE 6.7 EVOLUTION OF CONJUGATED DIENES, LIPID HYDROPEROXIDES AND HEXANAL IN DROPLET- STABILISED SAFFLOWER OIL-IN-WATER EMULSIONS WITHOUT BHA AND WITH BHA (500 PPM) . 188 FIGURE 6.8 PARTICLE SIZE DISTRIBUTION OF DROPLET-STABILISED SAFFLOWER OIL-IN-WATER EMULSIONS WITHOUT BHA AND WITH BHA (500 PPM) FRESHLY PREPARED AND DURING ACCELERATED OXIDATION (DAY 5 AND 7) .............................................................................. 189 FIGURE 6.9 POSSIBLE PATHWAYS OF LIPID OXIDATION IN DROPLET-STABILISED EMULSIONS EXPOSED TO LIGHT IN THE PRESENCE OF FERROUS IONS (500 µM) ....................................................... 192 FIGURE 6.10 SCHEMATIC ILLUSTRATION OF POSSIBLE BHA TRANSFER PATHWAY IN DROPLET- STABILISED EMULSIONS- A= BHA-IN-SHELL DROPLETS; B = BHA-IN-CORE DROPLETS. RED ARROWS INDICATE SHELL DROPLET SPONTANEOUS ADSORPTION AND BLACK ARROWS INDICATE BHA TRANSFER AFTER SHELL DROPLET ADSORPTION. .......................................................... 200 FIGURE 6.11 SCHEMATIC ILLUSTRATION OF POSSIBLE BHA TRANSFER PATHWAY IN DROPLET- STABILISED EMULSIONS- A= BHA-IN-SHELL DROPLETS; B = BHA-IN-CORE DROPLETS ............ 201 FIGURE 7.1 FLOW CHART OF SURFACE-LOADED ANTIOXIDANT DROPLET-STABILISED EMULSION PRODUCTION ...................................................................................................................... 211 FIGURE 7.2 FLOW CHART OF CORE-LOADED ANTIOXIDANT DROPLET-STABILISED EMULSION PRODUCTION ...................................................................................................................... 212 FIGURE 7.3: SCHEMATIC DIAGRAM OF MICROFLUIDIC CHANNEL CHIP USED TO ISOLATE DSE DROPLETS FOR CONFOCAL RAMAN MICROSCOPY .................................................................................. 213 FIGURE 7.4: PARTICLE SIZE DISTRIBUTION OF COCONUT OIL DSES WITH BHA IN SHELL AND CORE USED FOR SATURATED TRANSFER DIFFERENCE EXPERIMENT .......................................................... 219 FIGURE 7.5: PARTICLE SIZE DISTRIBUTION OF OLIVE (A) AND TRIMYRISTIN (B) DSES WITH BETA- CAROTENE INCORPORATED IN SHELL AND CORE .................................................................... 220 List of Figures XVIII FIGURE 7.6: STD FULL SPECTRAL WIDTH OF COCONUT OIL WITH BHA; SAFFLOWER OIL; AND DROPLET- STABILISED EMULSIONS WITH BHA INCORPORATED IN SHELL; & IN CORE. ............................... 222 FIGURE 7.7: STD EXPANDED AROMATIC BHA PEAKS IN COCONUT OIL; DSE WITH BHA IN SHELL AND DSE WITH BHA IN CORE. BLUE SPECTRA = REFERENCE STD SPECTRA RECORDED FIRST; RED SPECTRA = STD SPECTRA WITH IRRADIATED SAFFLOWER OIL PEAK ....................................... 223 FIGURE 7.8: RAMAN SPECTRA OF DSES WITH AND WITHOUT ANTIOXIDANTS (BHA & BETA-CAROTENE) .......................................................................................................................................... 225 FIGURE 7.9 OLIVE OIL DSE (WITHOUT BETA-CAROTENE) WITH 150 POINTS (TOP) & 265 POINTS (BOTTOM) (A & C) BRIGHT FIELD REFLECTANCE MODE IMAGES, (B & D) PEAK INTEGRAL DISTRIBUTION (RED TRACE= BETA-CAROTENE; BLUE TRACE= CH BAND; GREENT TRACE= OLIVE OIL). ................................................................................................................................... 226 FIGURE 7.10 OLIVE DSE (BETA-IN-SHELL) WITH 122 POINTS (TOP); & 250 POINTS (BOTTOM). (A & C) BRIGHT FIELD REFLECTANCE MODE IMAGE, (B & D) PEAK INTEGRAL DISTRIBUTION (RED TRACE= BETA-CAROTENE; BLUE TRACE= CH BAND; GREENT TRACE= OLIVE OIL). ................................ 227 FIGURE 7.11 OLIVE DSE (BETA-IN-CORE) WITH 158 POINTS (TOP) & 115 POINTS (BOTTOM). (A & C) BRIGHT FIELD REFLECTANCE MODE IMAGE, (B & D) PEAK INTEGRAL DISTRIBUTION (RED TRACE= BETA-CAROTENE; BLUE TRACE= CH BAND; GREENT TRACE= OLIVE OIL). ................................ 228 FIGURE 7.12 RAMAN SPECTRA OF BETA-CAROTENE, OLIVE OIL, SAFFLOWER OIL, AND TRIMYRISTIN 233 FIGURE 7.13 CONFOCAL RAMAN MICROSCOPY IMAGES AND INTENSITY PROFILE OF OLIVE OIL CONTROL EMULSIONS (SHELL GENTLY STIRRED-IN) - A, B, & C= BETA-CAROTENE, SAFFLOWER OIL AND CH BAND RAMAN CHANNELS RESPECTIVELY; D= LINE SCAN LOCATION ACROSS DROPLETS; E= INTENSITY PROFILES FOR SELECTED RAMAN CHANNELS ACROSS LINE SCAN SHOWN IN D ........ 234 FIGURE 7.14 CONFOCAL RAMAN MICROSCOPY IMAGES AND INTENSITY PROFILE OF OLIVE OIL DSES WITH BETA-CAROTENE-IN-SHELL DROPLETS- A, B, & C= BETA-CAROTENE, SAFFLOWER OIL AND CH BAND RAMAN CHANNELS RESPECTIVELY; D= LINE SCAN LOCATION ACROSS DROPLETS; E= INTENSITY PROFILES FOR SELECTED RAMAN CHANNELS ACROSS LINE SCAN SHOWN IN D ........ 235 FIGURE 7.15 CONFOCAL RAMAN MICROSCOPY IMAGES AND INTENSITY PROFILE OF OLIVE OIL DSES WITH BETA-CAROTENE-IN-CORE DROPLETS- A, B, & C= BETA-CAROTENE, SAFFLOWER OIL AND CH BAND RAMAN CHANNELS RESPECTIVELY; D= LINE SCAN LOCATION ACROSS DROPLETS; E= INTENSITY PROFILES FOR SELECTED RAMAN CHANNELS ACROSS LINE SCAN SHOWN IN D ........ 236 List of Figures XIX FIGURE 7.16 CONFOCAL RAMAN MICROSCOPY IMAGES AND INTENSITY PROFILE OF TRIMYRISTIN CONTROL EMULSION (SHELL GENTLY STIRRED-IN) - A, B, & C= BETA-CAROTENE, SAFFLOWER OIL AND CH BAND RAMAN CHANNELS RESPECTIVELY; D= LINE SCAN LOCATION ACROSS DROPLETS; E= INTENSITY PROFILES FOR SELECTED RAMAN CHANNELS ACROSS LINE SCAN SHOWN IN D .. 237 FIGURE 7.17 CONFOCAL RAMAN MICROSCOPY IMAGES AND INTENSITY PROFILE OF TRIMYRISTIN DSES WITH BETA-CAROTENE-IN-SHELL DROPLETS- A, B, & C= BETA-CAROTENE, SAFFLOWER OIL AND CH BAND RAMAN CHANNELS RESPECTIVELY; D= LINE SCAN LOCATION ACROSS DROPLETS; E= INTENSITY PROFILES FOR SELECTED RAMAN CHANNELS ACROSS LINE SCAN SHOWN IN D ........ 238 FIGURE 7.18 CONFOCAL RAMAN MICROSCOPY IMAGES AND INTENSITY PROFILE OF TRIMYRISTIN DSES WITH BETA-CAROTENE-IN-CORE DROPLETS- A, B, & C= BETA-CAROTENE, SAFFLOWER OIL AND CH BAND RAMAN CHANNELS RESPECTIVELY; D= LINE SCAN LOCATION ACROSS DROPLETS; E= INTENSITY PROFILES FOR SELECTED RAMAN CHANNELS ACROSS LINE SCAN SHOWN IN D ........ 239 FIGURE 7.19: CONFOCAL RAMAN MICROSCOPY IMAGES AND INTENSITY PROFILE OF OLIVE DSES WITH BETA-CAROTENE-IN-SHELL DROPLETS SHOWING EVIDENCE OF POSSIBLE MIGRATION TO CORE – A & B= BETA-CAROTENE AND SAFFLOWER OIL RAMAN CHANNELS RESPECTIVELY; C= LINE SCAN LOCATION ACROSS DROPLETS; D= INTENSITY PROFILES FOR SELECTED RAMAN CHANNELS ACROSS LINE SCAN SHOWN IN C .......................................................................................... 240 FIGURE 7.20 CONFOCAL RAMAN MICROSCOPY IMAGES AND INTENSITY PROFILE OF TRIMYRISTIN DSES WITH BETA-CAROTENE-IN-SHELL DROPLETS SHOWING EVIDENCE OF POSSIBLE MIGRATION TO CORE- A & B= BETA-CAROTENE AND SAFFLOWER OIL RAMAN CHANNELS RESPECTIVELY; C= LINE SCAN LOCATION ACROSS DROPLETS; D= INTENSITY PROFILES FOR SELECTED RAMAN CHANNELS ACROSS LINE SCAN SHOWN IN C. ......................................................................................... 241 FIGURE 7.21: CONFOCAL RAMAN MICROSCOPY IMAGES OF DSES SCANNED AT HIGH POWER (15 MW) SHOWING EVIDENCE OF POSSIBLE PHOTO-BLEACHING OF BETA-CAROTENE. A & C= BETA- CAROTENE RAMAN CHANNELS, B & D= SAFFLOWER OIL RAMAN CHANNELS. ........................... 242 FIGURE 8.1: SUMMARY OF STUDY OUTCOMES ............................................................................... 261 List of Tables XX List of Tables TABLE 2.1: FOOD EMULSIFIERS ..................................................................................................... 15 TABLE 2.2: FACTORS INFLUENCING THE PHYSICAL STABILITY OF EMULSIONS .................................... 23 TABLE 2.3: SUMMARY OF RESEARCH ON IMPACT OF INTERFACIAL LOCATION OF ANTIOXIDANTS ON OXIDATIVE STABILITY ............................................................................................................. 43 TABLE 2.4: BIOACTIVE COMPOUNDS AND THEIR POTENTIAL HEALTH BENEFITS .................................. 45 TABLE 3.1 COMPOSITION OF MILK PROTEIN CONCENTRATE (MPC) ................................................. 58 TABLE 3.2: REAGENTS AND CHEMICALS USED, SUPPLIERS AND GRADE ............................................ 61 TABLE 4.1 PROTEIN, CALCIUM AND SUGAR CONTENTS OF MPCS ..................................................... 82 TABLE 4.2 EMULSION FORMULATIONS ......................................................................................... 111 TABLE 5.1: AVERAGE PARTICLE SIZE OF DROPLET-STABILISED AND CONTROL EMULSIONS ............... 135 TABLE 5.2: OXIDATION PARAMETERS DERIVED FROM FITTING EQUATIONS 2, 3 AND 4 TO OXIDATION DATA. VALUES IN BRACKETS ARE STANDARD ERRORS. .......................................................... 141 TABLE 5.3: AREA UNDER THE CURVE (AUC) VALUES FOR CONJUGATED DIENES (CD), LIPID HYDROPEROXIDES (LHP) AND HEXANAL, INTEGRATED OVER THE WHOLE OXIDATION TIME COURSE. .......................................................................................................................................... 142 TABLE 6.1 FORMULATION OF ANTIOXIDANT (BHA) LOADED DROPLET-STABILISED EMULSIONS WITH LOW BHA LEVEL (50 PPM) IN SHELL AND CORE DROPLETS USED FOR OXIDATION STUDY ................. 169 TABLE 6.2 FORMULATION OF ANTIOXIDANT (BHA) LOADED DROPLET-STABILISED EMULSIONS WITH HIGH BHA LEVEL (500 PPM) IN SHELL AND CORE DROPLETS USED FOR OXIDATION STUDY ....... 170 TABLE 6.3: EMULSION FORMULATIONS FOR LOW BHA LEVEL (50 PPM). ......................................... 171 TABLE 6.4: EMULSION FORMULATIONS FOR HIGH BHA LEVEL (500 PPM). ....................................... 173 TABLE 6.5 AVERAGE PARTICLE SIZE OF DROPLET-STABILISED EMULSIONS WITH AND WITHOUT BHA 180 TABLE 6.6: PARTICLE SIZE CHANGES OF DROPLET-STABILISED SAFFLOWER OIL-IN-WATER EMULSIONS WITHOUT BHA AND WITH BHA (50 PPM) FRESHLY PREPARED AND DURING ACCELERATED OXIDATION (DAY 5 AND 7) .................................................................................................... 185 TABLE 6.7: PARTICLE SIZE CHANGES OF DROPLET-STABILISED SAFFLOWER OIL-IN-WATER EMULSIONS WITHOUT BHA AND WITH BHA (500 PPM) FRESHLY PREPARED AND DURING ACCELERATED OXIDATION (DAY 5 AND 7) .................................................................................................... 190 List of Tables XXI TABLE 7.1 FORMULATION OF ANTIOXIDANT (BHA) LOADED DROPLET-STABILISED EMULSIONS WITH BHA IN SHELL AND CORE DROPLETS USED FOR NMR SATURATED TRANSFER DIFFERENCE STUDY .......................................................................................................................................... 214 TABLE 7.2 FORMULATION OF ANTIOXIDANT (BETA-CAROTENE) LOADED DROPLET-STABILISED EMULSIONS WITH BETA-CAROTENE-IN-SHELL AND CORE DROPLETS USED FOR CONFOCAL RAMAN MICROSCOPY STUDY ........................................................................................................... 215 TABLE 7.3: BHA LOADED DROPLET-STABILISED EMULSIONS (DSE) ............................................... 216 TABLE 7.4: BETA-CAROTENE LOADED DROPLET-STABILISED EMULSIONS (DSE) ............................. 217 List of Publications & Presentations XXII List of Publications Okubanjo, S. S., Loveday, S. M., Ye, A., Wilde, P. J., & Singh, H. (2019). Droplet-Stabilized Oil-in-Water Emulsions Protect Unsaturated Lipids from Oxidation. Journal of Agricultural and Food Chemistry, 67(9), 2626-2636. Conference Presentations Okubanjo, S. S., Loveday, S. M., Ye, A., Wilde, P. J., & Singh, H. (2019). Droplet-stabilised emulsions: colloidal and oxidative stability advantages. Presented at the 9th Australian Colloid and Interface Symposium. Hobart, Tasmania. Okubanjo, S. S., Loveday, S. M., Ye, A., Wilde, P. J., & Singh, H. (2017). Droplet-stabilized emulsions: formulation and processing effects on structure. Presented at the 7th international symposium on delivery of functionality in complex food systems. Auckland, New Zealand. Okubanjo, S. S., Loveday, S. M., Ye, A., Wilde, P. J., & Singh, H. (2017). Anti-oxidative properties of droplet-stabilized emulsions. Presented at the 4th international food structures, digestion and health conference. Sydney, Australia. Chapter 1: Introduction 1 Chapter 1: Introduction 1.1 General Introduction Over the years, efforts are been made to provide food solutions that deliver health benefits beyond basic nutrition. This concept birthed the functional food industry. The functional food centre in Dallas recently proposed a unique definition of functional foods as ‘’natural or processed foods containing known or unknown bioactive compounds which if present in effective, non-toxic amounts provide a clinically proven and recorded health benefit for the prevention and treatment of chronic diseases’’ (Martirosyan & Singh, 2015). Food bioactives are defined as naturally occurring compounds in the food chain which are capable of conferring a health benefit (Biesalski et al., 2009). The biological activities of many bioactive compounds have been researched and documented, but these compounds are prone to degradations that result in a loss in their bioactive functionality. Conventional, micro/nano and multi-layered emulsions can improve the stability, solubility and bioavailability of lipid-soluble bioactives. Over the years, a lot of research has been focused on the stability of food emulsions (C. Berton, Ropers, Bertrand, Viau, & Genot, 2012; Mezdour, Lepine, Erazo-Majewicz, Ducept, & Michon, 2008; Phan, Le, Van de Walle, Van der Meeren, & Dewettinck, 2016; Singh, Tamehana, Hemar, & Munro, 2003), and their applications (Aditya et al., 2015; Piorkowski & McClements, 2013). Chapter 1: Introduction 2 The use of emulsions as carriers of active ingredients in the food industry is also on the rise. Increased protection, stability and efficient delivery of various lipophilic, hydrophilic and amphiphilic bioactives such as vitamin E, ß-carotene, ascorbic acid, resveratrol, omega 3 fatty acids etc. within emulsion-based carrier systems have been explored for applications in the food industry (Hategekirnana, Masamba, Ma, & Zhong, 2015; Hemar, Cheng, Oliver, Sanguansri, & Augustin, 2010; Tan & Nakajima, 2005; Tokle, Mao, & McClements, 2013; L. Wang et al., 2016; Yang & McClements, 2013; Yi, Li, Zhong, & Yokoyama, 2015). Recently, a novel emulsion structure made up of oil droplets stabilised by protein- stabilised nano-droplets was reported (Ye, Zhu, & Singh, 2013). This novel emulsion which is referred to as a ‘droplet-stabilised’ emulsion consists of ‘core lipid’ stabilised by protein-coated smaller lipid droplets that are referred to as ‘shell droplets’. The structure of this novel emulsion has the potential to protect and enhance the delivery and bioavailability of bioactive compounds, thus maximising their health benefits. Chapter 1: Introduction 3 Figure 1.1: Schematic diagram of ‘droplet-stabilised’ emulsion These functionalities if successfully explored, studied, and understood will positively contribute to the growth of the functional food industry; therefore, it is necessary to fully understand the structure, stability and plausible applications of this novel emulsion. This research sought to answer the following questions: 1. How do the emulsion composition and emulsification conditions of droplet- stabilised emulsions affect their formation and properties? 2. How does the unique interfacial structure and composition of droplet- stabilised emulsions affect their stability to oxidation in comparison to conventional emulsion? Chapter 1: Introduction 4 3. How does incorporating antioxidants in the shell droplets of droplet- stabilised emulsions rather than in the core lipid affect their stability to oxidation? 4. Can droplet-stabilised emulsions be used for concurrent encapsulation of lipophilic bioactives by incorporating one bioactive in the shell droplets and another in the core droplets? To achieve these objectives, four studies were conducted. These studies are organized into chapters in the form of manuscripts. Chapter 2 is a detailed literature review carried out to establish current knowledge on the subject matter. Chapter 3 describes the materials and methods used in this research as well as some of the challenges experienced with some methods and how they were overcome. Chapter 4 (research question 1) details the structural characteristics of droplet- stabilised emulsions as affected by varying the type of emulsifier (milk protein concentrate), homogenisation conditions, surface lipid and core lipid. The aims of the study were to explore the production of droplet-stabilised emulsions with food-grade ingredients and examine the effect of formulation and processing conditions on formation and structural characteristics of DSEs. The study tested the hypothesis that the ‘formation and structural characteristics of DSEs emulsions could be greatly influenced by its composition (interfacial and core) Chapter 1: Introduction 5 and processing conditions’. The study provided critical and insightful knowledge about the structural characteristics of DSEs. Chapter 5 (research question 2) details the impact of ‘droplet-stabilised emulsions’ structure and interfacial composition on oxidative stability of polyunsaturated rich oil incorporated within. The aim was to examine the influence of the emulsions’ interfacial structure and varied interfacial composition on the oxidative stability of PUFA rich oil. The study also investigated the effect of physical state of surface lipid on oxidative stability. The study tested the hypothesis that ‘droplet-stabilised emulsions could provide greater oxidative stability than conventional emulsions and core droplets stabilised by high melting surface lipid may provide greater oxidative stability than low melting surface lipid’. The study confirmed the potential oxidation resistance of droplet-stabilised emulsions. Chapter 6 (research question 3) describes the effect of locating antioxidants at the interface of droplet-stabilised emulsions on oxidative stability of PUFA rich oil incorporated within in comparison with locating antioxidants in the core PUFA rich oil. The aim was to explore the possibility of incorporating antioxidants in the surface lipid located at the interface of DSEs rather than the current practice of incorporating directly in PUFA oil. The study tested the hypothesis that locating antioxidant at the interface of DSEs could provide greater oxidative stability than locating antioxidant directly in the core unsaturated lipid. The study revealed that location-based antioxidant performance in DSEs was concentration dependent. Chapter 7 (research question 4) describes new methods used to probe the location of antioxidants incorporated in DSEs and monitor the antioxidant’s Chapter 1: Introduction 6 mobility. The objective was to determine if antioxidant incorporated in the surface lipid would migrate or diffuse into the core lipid thus exploring the possibility of using DSEs for concurrent encapsulation of bioactives. The study revealed that antioxidant mobility from shell to core was minimal and the antioxidant remained mostly localized at the interface. Chapter 8 summarizes the outcomes of this research and details the impact on applicable subject areas such as development of structured oil-in-water delivery systems, bioactive compound encapsulation and protection, oxidation of polyunsaturated rich oils, location-based antioxidant performance in oil-in-water emulsion systems, and functional food applications of emulsions. The chapter also highlights future research prospects based on this research. Chapter 1: Introduction 7 Figure 1.2 Thesis structure Chapter 2- Literature Review Chapter 3- Materials and Methods Chapter 4- Formation and properties of droplet- stabilised oil-in-water emulsion: Structure characteristics of droplet-stabilised oil- in-water emulsion as affected by  Homogenization equipment  Emulsifier type  Surface and core lipid types Chapter 5- Droplet-stabilised oil-in-water emulsions protect unsaturated lipids from oxidation  Oxidative stability of unsaturated lipid incorporated in droplet-stabilised oil-in-water emulsion compared to conventional emulsion  Impact of surface lipid state on resistance to oxidation Chapter 6- Impact of antioxidant location on oxidation resistance of droplet-stabilised oil-in-water emulsions  Oxidative stability of unsaturated lipid incorporated in droplet-stabilised oil-in-water emulsion as affected by locating antioxidant at the interface versus the core unsaturated lipid Chapter 8- Perspectives and future research prospects Chapter 7- Probing the location of antioxidants incorporated in droplet-stabilised oil-in-water emulsions  Diffusion or migration of antioxidant incorporated in shell droplets to core droplets Chapter 2: Literature Review 8 Chapter 2: Literature Review 2.1 Classes of emulsions, preparation methods and emulsifiers Two distinct phases are observed in an emulsion; the dispersed and continuous phase. A food system made up of oil droplets dispersed in an aqueous phase is called an oil-in-water (O/W) emulsion while water droplets dispersed in oil phase is referred to as water-in-oil emulsion (W/O). Examples of O/W food emulsions are milk, cream, mayonnaise etc. while W/O food emulsions include margarine, butter and some spreads/sauces. Figure 2.1 Schematic diagram of simple (O/W & W/O). Adapted from Chung and McClements (2015) Generally, emulsions are classified according to the composition of their dispersed and continuous phases however, classification of emulsions have become more complex and are now also classified based on their dispersed phase size (Micro, nano, macro) or structured interfacial phase (Multi-layered, Pickering). Chapter 2: Literature Review 9 2.1.1 Emulsion preparation To create an emulsion, it is usually necessary to apply mechanical energy and decrease the interfacial tension. Generally, emulsions are created by a process called homogenisation whereby the immiscible liquids are mixed by high speed or agitation (McClements, 2005). Spontaneous emulsification is another method of creating an emulsion without application of mechanical energy (Bibette, Leal- Calderon, & Schmitt, 2007; López-Montilla, Herrera-Morales, Pandey, & Shah, 2002). 2.1.2 Micro-emulsions and nano-emulsions Micro-emulsions and nano-emulsions are terms that are sometimes used in literature without clear distinctions however; growing research involving these systems makes it necessary to provide a clear distinction. It is generally accepted that micro-emulsions are thermodynamically stable and have very small droplet sizes (r<50nm) whereas nano-emulsions also have small droplet sizes (r<100nm) but are kinetically stable (J. Rao & McClements, 2011). Anton and Vandamme (2011) in providing clarification on the differences between these systems made reference to the methods by which they are created and their resulting structure and stability. McClements (2012) in an attempt to also distinguish between the two systems highlighted the differences in their preparation, but also noted that it is often difficult to distinguish between the two systems merely by their formation methods. Both writers proposed methods such as particle size distribution that can be used to distinguish between these two systems however; the proposed methods are only indicative and not conclusive. They also emphasized that creation of nano-emulsions usually requires application of high energy. Chapter 2: Literature Review 10 Nano-emulsions are usually prepared by high energy methods using high pressure homogenisers. Low energy and phase inversion temperature methods have also been explored in producing nano-emulsions. T. Tadros, Izquierdo, Esquena, and Solans (2004) compared the formation of nano-emulsions using high pressure homogenisers and phase inversion temperature (PIT) methods. The emulsions produced by the homogenisers were more stable to Ostwald ripening over those made by the PIT method even though it was not clear if the composition of the nano-emulsions made with homogenisers was identical to the PIT produced nano-emulsions. On the other hand, Solans, Izquierdo, Nolla, Azemar, and Garcia-Celma (2005) emphasized that efficient formation of nano- emulsions can be achieved by low energy methods however it was evident that the size of droplets was largely dependent on the structure of the surfactant phase. The ultimate goal in selecting a method of preparing emulsions should be its stability and final application moreover, methods of overcoming high droplet polydispersity associated with oil-in-water nano-emulsions produced by low- energy methods are proposed in literature (Gutiérrez et al., 2008). 2.1.3 Double Emulsions Over the years, more complex emulsions have emerged which consists of multiple emulsions (Figure 2.2) referred to as water-in oil-in water (W/O/W) or oil- in-water-in-oil (O/W/O). In this case, the dispersed phase is dispersed within a larger dispersed phase which is in turn dispersed in the continuous phase (Muschiolik, 2007). Chapter 2: Literature Review 11 Figure 2.2: Schematic diagram of double emulsions (W/O/W & O/W/W). Adapted from Martínez-Palou et al. (2011) The formation of double emulsions typically involves two stages however; other methods of preparation have been explored. Sachio Matsumoto carried out a series of studies on the preparation and stability of double emulsions specifically water-in-oil-in-water (W/O/W) emulsions and their potential applications in foods (Matsumoto, 1983, 1985, 1986; Matsumoto, Koh, & Michiura, 1985). In these studies three main methods of forming double emulsions were discussed; mechanical agitation, phase inversion and the two-stage emulsification process. The formation of W/O/W emulsions by these methods requires that attention is given to the ratio of hydrophobic and hydrophilic emulsifiers employed. It is imperative to optimize the concentration of emulsifiers, as the structure of the emulsions has a tendency to change entirely once the optimal ratio of hydrophilic to hydrophobic emulsifier is exceeded. Double emulsions formed by these methods are highly prone to instability, as the hydrophobic emulsifiers tend to migrate over time to the continuous aqueous phase leading to a breakage in the oil layer adsorbed on the surface of the aqueous phase. Matos, Gutiérrez, Coca, and Pazos (2014) produced double emulsions for encapsulation of trans-resveratrol by mechanical agitation and membrane emulsification. The results showed higher stability with emulsions produced by Chapter 2: Literature Review 12 mechanical agitation over those made by membrane emulsification, but higher release of bioactive was obtained with the latter over the former. These results were not surprising as double emulsions were formed by mechanical agitation where high energy was applied therefore, it is expected that stability with membrane emulsification where low energy was applied will differ. 2.1.4 Pickering emulsions Pickering emulsions (Figure 2.3) are emulsions stabilised by solid particles that are smaller than the emulsion droplets (Berton-Carabin & Schroën, 2015; Chevalier & Bolzinger, 2013). Figure 2.3: Schematic diagram of Pickering emulsion. Adapted from Chevalier and Bolzinger (2013) Pickering (1907) was the first to introduce the concept of emulsions stabilised with solid particles. He emphasized that emulsions stabilised with organic substances/surfactants separate over time or as soon as the emulsifier is destroyed by addition of acid or certain salts and highlighted the benefit of high interfacial adsorption energy of solid particles which are difficult to displace once adsorbed at the interface. Chevalier and Bolzinger (2013) reviewed the possible methods of preparing Pickering emulsions, in most methods, nanoparticles are Chapter 2: Literature Review 13 dispersed in either the oil or aqueous phase depending on the type of emulsion desired. The type of nanoparticle employed depends on the type of emulsion for example hydrophilic particles are dispersed in aqueous phase for O/W emulsions and both hydrophilic and hydrophobic particles are used for double emulsions (W/O/W), in all cases emulsification was achieved by high shearing except for preparation of solid particle stabilised double emulsions in which final emulsification was done by low shearing. The review revealed that Pickering emulsions can easily be used in many applications to replace classical emulsions and provide improved stability. Aveyard, Binks, and Clint (2003) also used similar methods in their study on emulsions stabilised with colloidal particles and established that hydrophilic particles create oil-in-water emulsions while hydrophobic particles create water-in-oil emulsions. 2.1.5 Multi-layered emulsions Multi-layered emulsions (Figure 2.4) were designed as a strategy to improve emulsion stability against droplet aggregation. Multi-layered emulsions are typically made up of several layers of emulsifiers and this is achieved by a layer- by-layer (LBL) electrostatic deposition principle (Beicht, Zeeb, Gibis, Fischer, & Weiss, 2013; Gudipati, Sandra, McClements, & Decker, 2010; Jo, Chun, Kwon, Min, & Choi, 2015). Multi-layered emulsions are typically prepared by a two stage process; first a primary emulsion stabilised by an ionic emulsifier is made by homogenisation secondly, a polymer with an opposite charge is electrostatically deposited onto the charged primary lipid droplets and this can be repeated to achieve the desired number of layers (Aoki, Decker, & McClements, 2005; Guezey & McClements, Chapter 2: Literature Review 14 2006; Guzey & McClements, 2006; Thanasukarn, Pongsawatmanit, & McClements, 2006). Figure 2.4: Schematic diagram of multi-layered emulsion and preparation steps. Adapted from McClements and Li (2010) 2.1.6 Emulsifier types Emulsifiers are surface-active molecules capable of stabilizing emulsions and protecting emulsion droplets from flocculation or coalescence (McClements, 2005). Surfactants are surface-active agents made up of a non-polar (hydrophobic) portion attached to a polar (hydrophilic) portion (T. F. Tadros, 2014). The oil-water interfaces in emulsions are stabilised with surfactants or emulsifiers which are both amphiphilic molecules used to modify the surface tension between two phases (Rosen & Kunjappu, 2012; T. Tadros, 2013). Norn (2014) classified emulsifiers based on their nature or charge, and their hydrophilic-lipophilic balance (HLB). The hydrophilic-lipophilic balance is a classification system introduced by Griffin (1949) which measures the extent to which a surfactant is hydrophilic or lipophilic. On the basis of charge, Norn (2014) classified emulsifiers as non-ionic, cationic, anionic and zwitterionic. The non- Chapter 2: Literature Review 15 ionic emulsifiers are mostly used in foods and include monoglycerides, diglycerides, polysorbates and sorbitans etc. Anionic emulsifiers include lactylates and carboxylates however they are rarely used in foods. The most common zwitterionic emulsifier is lecithin. Table 2.1: Food Emulsifiers Food emulsifiers Common examples Surfactants Mono-diglycerides, polysorbate 60, sodium stearoyl-lactylate, lecithin, sorbitan monostearate Biopolymers Proteins: Milk proteins, meat proteins, egg proteins Polysaccharides: modified starch, gum Arabic, cellulose (HPMC) It is important to note that not only surfactants are used as emulsifiers. Biopolymers which also exhibit good surface activity are used as emulsifiers. The most widely used include proteins and polysaccharides that display amphiphilic properties. Mixtures of polymers are also used to stabilise emulsions (Akhtar & Dickinson, 2007; Dickinson & Galazka, 1991; Einhorn-Stoll, Ulbrich, Sever, & Kunzek, 2005). Chapter 2: Literature Review 16 2.2 Structure of food emulsions The structure of emulsions can be very complex as they are usually not just simple mixtures of lipids and water. Emulsions usually contain polysaccharides, sugars, flavouring agents, texture modifiers, proteins, and surfactant molecules etc., which contribute greatly to the complexity of their structure. These ingredients also interact in different ways with emulsion droplets giving rise to the structure of the product. For example, there is a marked difference in structure when cream dessert is produced with a polysaccharide and when it is produced without a polysaccharide. The physical and chemical stability of an emulsion greatly depends on the structure and composition of the interfacial phase, which also has a significant influence on how effective the emulsion will be in protecting or enhancing delivery of bioactives incorporated within. 2.2.1 Physical stability Emulsion stability refers to the potential of an emulsion to resist various forces or interactions within the system which could lead to changes in its state, structure or composition over time. When an emulsion is unable to resist changes in its physiochemical properties, it is said to be unstable. Stability of emulsions is sometimes defined in relation to their thermodynamic or kinetic stability. An emulsion is said to be thermodynamically stable when the free energy of the droplets in aqueous phase is lower than the separate phases (oil, water and surfactants) and unstable when the free energy is higher than the separate phases. A kinetically stable emulsion has higher free energy than its Chapter 2: Literature Review 17 component phases, but an energy barrier inhibits droplet coalescence, thus preventing it from reverting to separate phases (McClements, 2012). Emulsions become unstable due to physical and chemical processes that occur within the system. The physical processes responsible for emulsion instability include creaming, sedimentation, flocculation, coalescence, phase inversion, and Ostwald ripening (Figure 2.5) while the chemical processes include oxidation and hydrolysis. It is possible that two or more of these processes will occur in an emulsion at the same time. Chapter 2: Literature Review 18 Figure 2.5 Emulsion instability schematics. Oil and water have different densities hence there is usually the effect of gravitational force which leads to phase separation either by creaming or sedimentation. 1. Creaming: This phenomenon occurs when there is a difference in density between the droplets and the liquid in which they are dispersed Chapter 2: Literature Review 19 such that when the density of the droplets is lower than the liquid, the droplets travel upwards to the surface of the system. This is very common in oil-in-water emulsions because oils usually have lower densities than water, although density differences can be overcome with interfacially adsorbed ‘weighting agents’, i.e. polymers that increase the mass of droplets. 2. Sedimentation: This is because of a higher droplet density over that of the continuous phase. In this case the droplets settle to the bottom of the container. Emulsion droplets are constantly in motion due to kinetic energy (Brownian motion) or gravitational forces (creaming/sedimentation). As droplets move, they tend to collide with each other. Upon collision, two droplets either move apart, flocculate or coalesce. 3. Flocculation: This is when droplets aggregate such that each droplet retains its individual identity. This process can be reversible or irreversible. Flocculation increases emulsion viscosity which may be undesirable in some foods as it leads to higher instability thus reducing the shelf life (Demetriades, Coupland, & McClements, 1997). Bridging flocculation is a type of flocculation that occurs when an electrically charged high molecular weight polymer adsorbs to the surface of two or more droplets through electrostatic interactions, forming bridges (Dickinson, 2003). On the other hand, depletion flocculation occurs due to osmotic effects resulting from the exclusion of non-adsorbing Chapter 2: Literature Review 20 polymers from regions surrounding two droplets, this leads to an increase in the attractive forces between the droplets and eventual flocculation by depletion (McClements, 2004). Figure 2.6 Bridging and depletion flocculation schematics 4. Coalescence: This is a process in which two or more droplets merge together into a single bigger droplet as a result of collision, and it is irreversible. This interaction results in a decrease in the area of oil-water interface. This droplet interaction is not always between the liquid of two droplets; it could also be between a solid fat crystal of one droplet and liquid of the other droplet this type of interaction is referred to as ‘partial coalescence’. Partial coalescence will happen in an emulsion only when there are partially crystalline droplets. This mechanism when controlled can be desirable in the production of some foods such as ice cream where the emulsion has to be cooled thus resulting in development of fat crystals hence when coalescence occurs it improves the texture of the product (Goff, 1997). Bridging flocculation Depletion flocculation Chapter 2: Literature Review 21 5. Ostwald ripening: This is a phenomenon which has been described as occurring in solid and liquid solutions. It is described by Kabalnov (2001) as a process whereby smaller particles dissolve and larger particles grow at the expense of the smaller ones because solubility increases due to small radius of curvature. In emulsions, Taylor (1998) describes Ostwald ripening as the growth of a droplet at the expense of a smaller droplet arising from a difference in radius of curvature of the droplets which is inversely related to surface pressure of the dispersed phase. He states that as the smaller droplets dissolve the average radius of the droplets increases with time. 6. Phase inversion: This is when a system inversion occurs such that for example an oil-in-water emulsion changes to a water-in-oil emulsion or the opposite way. Phase inversion is usually caused by changes in the composition or environment of an emulsion. Phase inversion in food emulsions is reported to be induced by surfactants caused by a change in molecular geometry or induced by fat-crystallization caused by partial coalescence (McClements, 2005). This type of instability has a negative impact on the sensory properties of foods but is also very important in the production of foods like butter and margarine. The stability of an emulsion has important consequences on the digestion and functionality of the emulsion or its components. As the utilization of emulsions as delivery systems of bioactive compounds expands, the effect of stability on their ability to perform efficiently cannot be overemphasized. Chapter 2: Literature Review 22 Golding and Wooster (2010) pointed out that the stability of emulsions in the digestive tract plays a role in lipid digestion and gastric emptying rate. For example, emulsions that have undergone phase separation under gastric conditions have faster gastric emptying which will subsequently affect satiety signals (Marciani et al., 2007). The structure of emulsions can change very easily during storage, processing and digestion. These changes may result in any one of the unstable forms discussed above. According to Yao, Xiao, and McClements (2014), structure of micro-emulsions may be altered as they encounter bile salts or minerals while indigestible nano-emulsions are more likely to keep the core material intact as they pass through the gastric environment. Chapter 2: Literature Review 23 Table 2.2: Factors influencing the physical stability of emulsions Emulsion Instability Driving forces Control measures References Gravitational separation (creaming and sedimentation)  Difference in density between droplets and surrounding liquid.  Increased droplet poly-dispersity.  Decrease droplet size  Increase droplet concentrat ion and viscosity of continuous phase McClements (2005) Flocculation  Droplet collisions due to Brownian motion, gravitational separation and shearing.  Retard or decrease collision frequency.  Prevent unnecessary agitation.  Increase viscosity of continuous phase.  Increase repulsive forces (e.g. electrostatic, steric) between droplets over attractive. McClements and Weiss (2005) Coalescence  Film rupture and thinning.  Emulsifier type.  Processing and storage conditions.  Prevent interfacial membrane rupture.  Increase repulsive forces. McClements (2005) Ostwald ripening  Increased solubility of molecules in droplet in continuous phase.  Increased diffusion of molecules in droplet to aqueous phase.  Prevent solubility of dispersed phase in continuous phase.  Decrease interfacial tension.  Minimize droplet polydispersity Walstra (2003); Weers (1998) Phase Inversion  Temperature  Applied mechanical energy.  Surfactant type and composition.  Fat crystallization.  Prevent partial coalescence.  Select suitable emulsifier and concentration. McClements and Weiss (2005) Chapter 2: Literature Review 24 2.2.2 Chemical Instability Emulsions are susceptible to chemical reactions that could negatively impact their sensory and functional properties. These reactions include lipid oxidation and biopolymer hydrolysis. Lipid oxidation will be discussed extensively in the next sections. 2.3 Lipid oxidation Lipid oxidation can be defined as the removal of one or more electrons from lipids either by the addition of oxygen or hydrogen abstraction. It is a complex process that involves the production of free radicals and hydroperoxides, alterations in the double bond arrangement of unsaturated lipids, and final breakdown of the lipid. The rate of lipid oxidation is influenced by catalysts or factors such as light, metals, enzymes and temperature. Therefore, lipids could undergo auto- oxidation, thermal oxidation, enzymatic oxidation and photo-oxidation depending on the environmental conditions to which they are subjected. Mechanism of lipid oxidation in emulsions differs from that in bulk oil because in emulsions, the aqueous phase may contain pro-oxidants which are not usually present in bulk oils and these pro-oxidants may partition within the different phases in emulsions. Oxidation rates in emulsions are reported to be usually faster in emulsions than bulk oils because of the large surface area of interfaces created in emulsions which allows interaction of the aqueous and oil phases (Logan, Nienaber, & Pan, 2013). Chapter 2: Literature Review 25 2.3.1 Auto-oxidation Auto-oxidation as the name implies is a spontaneous reaction. It involves the reaction of atmospheric oxygen with lipids, and unsaturated lipids are very prone to these reactions. Autoxidation and thermal oxidation are chain reactions that involve three stages; initiation, propagation and termination. Kanner, German, Kinsella, and Hultin (1987) reported three possible pathways by which lipid oxidation can be initiated as shown below. Figure 2.7 Initiation of lipid oxidation pathways- a) Initiation by hydrogen abstraction; b) Initiation by free-radical attack on a double bond; C) Initiation by singlet oxygen. Adapted from Kanner et al. (1987). Initiation: Atmospheric oxygen is in the triplet ground state therefore to react with lipid molecules it requires excitation or activation. Three mechanisms by which oxygen can be activated have been identified; 1. Formation of singlet oxygen (Excited state oxygen) 2. Formation of partially reduced or activated oxygen species like hydrogen peroxide, superoxide anion, or hydroxyl radical. 3. Formation of active oxygen-iron complexes like ferric-oxygen-ferryl complexes. Chapter 2: Literature Review 26 At initiation, a hydrogen radical is removed from an unsaturated fatty acid (LH) resulting in a free lipid radical (L●). LH L● + H● (Eq. 2-1) Propagation During the propagation stage the free radical (L●) in turn reacts with oxygen to form an unstable lipid peroxyl radical (LOO●), which reacts with lipids by abstracting hydrogen from unsaturated fatty acid to form hydroperoxide (LOOH) and an unstable lipid radical (L●). This lipid radical again reacts with oxygen initiating another free radical resulting in a chain reaction. Propagation gives rise to the chain reaction in oxidation. It involves the following: - Radical coupling with oxygen: L● + 3O2 LOO● (Eq. 2-2) - Atom or group transfer: LOO● + LH LOOH + L● (Eq.2-3) - Fragmentation: LOO● L● + 3O2 (Eq. 2-4) Hydroperoxides are unstable so they continue to degrade and produce radicals. According to Fereidoon Shahidi and Wanasundara (2008) there is an induction period when hydroperoxide formation is low. Termination To terminate the propagation chain reaction, two types of reactions are possible; radical-radical coupling and radical-radical disproportionation. These reactions Chapter 2: Literature Review 27 involve the formation of non-radical products such as ketones, aldehydes etc. (Erickson, 2002). L● + L● LL (Eq. 2-5) L● + LOO● LOOL (Eq. 2-6) LOO● + LOO● LOOL + 3O2 (Eq. 2-7) Lipid oxidation pathways are not the same for every food matrix it depends on the nature of the molecules present and the environmental conditions (Erickson & Sista, 1996). The mechanisms of lipid oxidation differ for different bulk lipids so it will also not be the same for emulsions (Naz, Siddiqi, Sheikh, & Sayeed, 2005). The rate of oxidation of fatty acids increases with an increase in their unsaturation. The rate of oxidation in terms of oxygen uptake in oleic (18:1), linoleic (18:2), and linolenic (18:3) acid is in the order of 1:50:100 with respect to oxygen consumption and 1:12:25 with respect to peroxide production (Hsieh & Kinsella, 1989). Hsieh and Kinsella also reported that the initial rate of linolenic oxidation was twice that of linoleic which is attributed to the number of double bonds present. Triglycerides yield oxidation products that reflect their fatty acid composition. Figure 2.8 shows an example of oxidation reactions of linoleic acid producing hexanal and concentrations of the products (conjugated diene, hydroperoxide and hexanal) shown in the pathway provide a good indication of oxidative deterioration of lipids rich in linoleic acid. Chapter 2: Literature Review 28 Lipids when subjected to high temperatures are highly susceptible to auto- oxidation resulting in rapid formation and decomposition of hydroperoxides. Low temperature treatments does not mean there will be no oxidation, the formation of autoxidation products are just slower in this case (Choe & Min, 2006). 2.3.2 Enzymatic oxidation Lipase enzymes present in lipids breakdown triglycerides into free fatty acids. Lipoxygenase which is an iron containing enzyme found in plants and animals is capable of catalysing (with polyunsaturated fatty acid as a suitable substrate) the reaction between free fatty acid and oxygen thus forming hydroperoxide (LOOH). 2.3.3 Photo-oxidation Photo-oxidation occurs when there is a reaction between light-activated singlet oxygen and unsaturated fatty acids, leading to the formation of hydroperoxides. Singlet oxygen is a high-energy form of oxygen that is highly reactive with organic compounds. It is not a radical compound therefore reacts mostly with non-radical double-bonded compounds. Singlet oxygen is formed from triplet oxygen (ground state atmospheric oxygen) when photosensitizers present in foods absorb energy from light and transfer this energy to triplet oxygen. The photo-sensitization mechanisms are classified as type I and type II (Davidson, 1979; Min & Boff, 2002b). Sensitization occurs when molecules usually called sensitizers (Sen) such as chlorophylls, riboflavin, myoglobin etc present in lipids, get sensitized by light (absorb light) and become excited (1Sen*). The excited sensitizer may also be Chapter 2: Literature Review 29 converted to a triplet sensitizer (3Sen*) by a process called intersystem crossing (Kanofsky, 2016). Sen 1Sen* 3Sen* (Eq. 2-8) Type I Mechanism: The excited sensitizer (3Sen*) directly abstracts hydrogen from a fatty acid and produces a free-radical this mechanism is most likely to occur under low oxygen concentrations. The free radical in turn initiates the free radical chain reaction. 3Sen* + LH ●Sen H + L● (Eq. 2-9) Type II Mechanism: When oxygen is readily available, the excited sensitizer (3Sen*) is short-lived and thus they emit or transfer the energy absorbed to triplet oxygen (3O2) generating singlet oxygen (1O2). Singlet oxygen directly reacts with the double bonds of unsaturated fatty acids and produces conjugated and non- conjugated hydroperoxides (Edwin N. Frankel, 2012). 3Sen* + 3O2 1O2 + Sen (Eq. 2-10) 1O2 + LH LOOH (Eq. 2-11) hv Intersystem crossing Chapter 2: Literature Review 30 Figure 2.8: Oxidation pathway of linoleic acid. Adapted from Wheatley (2000) Chapter 2: Literature Review 31 2.4 Structural factors affecting the oxidative stability of emulsions The oxidative stability of emulsions must be given good attention as it has subsequent consequences on emulsion functionality. S. J. Lee, Choi, Li, Decker, and McClements (2011) evaluated the oxidative stability of nanoemulsions and conventional emulsions stabilised by whey protein isolates. The emulsions were prepared by a two-step homogenisation process and its oxidative stability was monitored over four weeks’ storage at 37°C. The rate of oxidation was higher in nano-emulsions. The authors expected slower oxidation in the nano-emulsion as a consequence of higher protein concentration (some amino acids have antioxidant effects) and attributed the observed faster oxidation to pro-oxidant impurities in the protein. Haahr and Jacobsen (2008) compared oxidative stability of fish oil-in-water emulsions stabilised by tween 80, citrem, lecithin and caseinate. The emulsions were prepared by a two-step process and oxidation monitored over 12 days’ storage in the dark at 20°C. The authors reported slightly higher oxidation rates in citrem and tween emulsions which had smaller droplet sizes over lecithin and caseinate emulsions which had larger droplets however, they emphasized that differences in oxidative stability between these emulsions could not have been solely due to differences in droplet sizes. Similarly, C. Jacobsen et al. (2000) investigated the influence of droplet size on oxidative stability of fish oil-enriched mayonnaise, and reported lower concentrations of volatile oxidation compounds in mayonnaise with large oil droplets and higher concentrations in mayonnaise with smaller droplets during storage at 20°C. The authors attributed the observed Chapter 2: Literature Review 32 faster oxidative deterioration in mayonnaise with smaller droplet sizes to increased interfacial area. An opposite trend of faster oxidation in emulsions with smaller droplet sizes over larger droplet sizes was reported by Nakaya, Ushio, Matsukawa, Shimizu, and Ohshima (2005). They observed lower levels of hydroperoxides for emulsions with smaller droplet sizes (0.8µm) over bigger size (12.8µm) indicating improved oxidative stability with smaller droplets. This improved stability was attributed to a ‘wedge effect’ in smaller droplets which prevents mobility of lipid molecules whereas in larger droplets mobility of lipid molecules is not retarded by this effect. This wedge effect is thought to be imposed by hydrophobic residues of emulsifier thus inducing shorter oxidation chain reactions in small droplets. Walker, Decker, and McClements (2015) also examined the effect of droplet size and surfactant concentration on oxidative stability of fish oil-in-water emulsions prepared by spontaneous emulsification (low energy) and microfluidization (high energy). Additional surfactant was added to portions of the prepared emulsions to produce emulsions with large droplets and high surfactant concentration. Oxidation rate was monitored over 14days storage in the dark at 55°C. The authors expected a higher rate of oxidation in emulsions with smaller particle sizes because of increased surface area of the oil droplets but observed slightly higher hydroperoxide values in low-energy emulsions with added surfactant which had greater particle size, and attributed the observed higher oxidation rate to possible partitioning of reactants into surfactant micelles which altered the lipid oxidation rate. Lethuaut, Métro, and Genot (2002) also investigated the influence of droplet size on oxidative stability of protein stabilised emulsions and reported higher conjugated diene concentrations in emulsions with smaller droplet sizes over Chapter 2: Literature Review 33 emulsions with larger droplet sizes however, the authors also report that the faster oxidation rates in emulsions with smaller sizes may have been partially counteracted by antioxidant activity of adsorbed proteins which led to similar concentrations of conjugated dienes between emulsions with small and large droplet sizes at a certain point. The effect of droplet size on oxidative stability appears to be controversial however from these literatures, it is apparent that the composition of the emulsions and methods of preparation were different even though it is possible that the structures of these emulsions were similar, it shows that the oxidative stability of an emulsion is not only dependent on the droplet size but on other factors such as the composition of the interfacial layer, emulsifier type, oil phase type and transportation mechanisms of reactants. Moreover, Osborn and Akoh (2004) investigated the influence of droplet size of emulsions on oxidative stability and reported that droplet size had no effect on lipid oxidation. Also examined effect of droplet size on oxidative stability of soybean oil and fish oil-in-water emulsions and reported reverse effects of droplet size whereby oxidative stability of fish oil-in-water emulsions increased with decreasing droplet size while the reverse effect was observed in soybean oil-in-water emulsions. The interfacial phase plays an important role in the oxidative stability of emulsions in fact it is believed to be the gateway between pro-oxidants or hydrophobic phase and the lipid phase (C. Berton, Ropers, Viau, & Genot, 2011; Silvestre, Chaiyasit, Brannan, McClements, & Decker, 2000). Hence it is no wonder that structuring the interfacial phase of emulsions have become a popular approach in designing emulsions. Chapter 2: Literature Review 34 Kargar, Spyropoulos, and Norton (2011) monitored the oxidation of emulsions stabilised by tween 20 and sodium caseinate over seven days. Oxidation rates were lower for protein-stabilised emulsions over tween 20-stabilised emulsions and this was attributed to the ability of proteins to form a thicker interfacial layer. The authors also monitored oxidative stability of Pickering emulsions stabilised by silica particles and reported that the thick interfacial layer formed by Pickering particles also reduced lipid oxidation rate in Pickering emulsions compared to emulsions stabilised by tween 20. In this study, the effect of other factors such as oil-phase fraction, pH, and emulsifier concentration in the aqueous phase were assessed and all these seemed to influence the rate of lipid oxidation. It is also interesting to note in Kargar et al’s study that as the concentration of Pickering particles increased, the droplet size decreased leading to a decrease in oxidation. This signifies that contact between the lipid phase and pro-oxidants must have been minimized by the particles at the interface. Some of the factors that affect lipid oxidation in emulsions include; chemical structure of lipids, quality of ingredients, oxygen concentration, interfacial characteristics, and interactions with aqueous phase (McClements & Decker, 2000). All these factors should be taken into consideration in preventing lipid oxidation in emulsions. Flaxseed oil-in-water emulsions stabilised by multi-layered membranes (sodium caseinate and pectin) at pH between 3 and 5 showed better oxidative stability than single layered sodium caseinate stabilised emulsion as evidenced from the levels of hydroperoxides and TBARS c