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- Herausgeber: John Wiley & Sons
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A comprehensive text that offers a review of the delivery of food active compounds through emulsion-based systems Emulsion-based Systems for Delivery of Food Active Compounds is a comprehensive recourse that reviews the principles of emulsion-based systems formation, examines their characterization and explores their effective application as carriers for delivery of food active ingredients. The text also includes information on emulsion-based systems in regards to digestibility and health and safety challenges for use in food systems. Each chapter reviews specific emulsion-based systems (Pickering, multiple, multilayered, solid lipid nanoparticles, nanostructured lipid carriers and more) and explains their application for delivery of food active compounds used in food systems. In addition, the authors - noted experts in the field - review the biological fate, bioavailability and the health and safety challenges of using emulsion-based systems as carriers for delivery of food active compounds in food systems. This important resource: * Offers a comprehensive text that includes detailed coverage of emulsion-based systems for the delivery of food active compounds * Presents the most recent development in emulsion-based systems that are among the most widely-used delivery systems developed to control the release of food active compounds * Includes a guide for industrial applications for example food and drug delivery is a key concern for the food and pharmaceutical industries Emulsion-based Systems for Delivery of Food Active Compounds is designed for food scientists as well as those working in the food, nutraceutical and pharmaceutical and beverage industries. The text offers a comprehensive review of the essential elements of emulsion-based systems for delivery of food active compounds.
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Table of Contents
Cover
Title Page
Preface
About the Editors
Shahin Roohinejad
Ralf Greiner
Indrawati Oey
Jingyuan Wen
List of Contributors
1 Conventional Emulsions
1.1 Introduction
1.2 Conventional Emulsions Formation and Stability
1.3 Composition of Conventional Emulsions for Food Applications
1.4 Characterization of Conventional Emulsions
1.5 Conventional Emulsions as Carriers for Delivery of Food Active Compounds
1.6
In Vitro
/
In Vivo
Digestion of Conventional Emulsions
1.7 Bioaccessibility and Bioavailability of Food Active Ingredients
1.8 Conclusion and Future Directions
References
2 Pickering Emulsions
2.1 Introduction
2.2 Formation and Stability of Pickering Emulsions
2.3 Pickering Emulsions for Food Applications
2.4 Characterization of Pickering Emulsions
2.5 Pickering Emulsions as Carriers for Active Compounds
2.6
In Vitro
and
In Vivo
Digestion of Pickering Emulsions
2.7 Pickering Emulsions: Rules and Regulations for Health and Safety
2.8 Conclusions and Future Directions
Acknowledgments
References
3 Multiple Emulsions
3.1 Introduction
3.2 Preparation of Multiple Emulsions and Stability
3.3 Multiple Emulsion Compositions for Food Application
3.4 Characterization of Multiple Emulsions
3.5 Multiple Emulsions as Carriers for Delivery of Food Active Compounds
3.6
In Vitro
Digestion of Multiple Emulsions
3.7 Conclusions and Future Focus
Acknowledgment
References
4 Multilayered Emulsions
4.1 Introduction
4.2 Multilayered Emulsion Formation and Stability
4.3 Multilayered Emulsion Compositions for Food Application
4.4 Methods for Characterization of Multilayered Emulsions
4.5 Multilayered Emulsions as Carriers for Delivery of Food Active Compounds
4.6 Oxidative Stability and Digestibility of Multilayered Emulsions
4.7 Conclusions and Future Directions
Acknowledgment
References
5 Solid Lipid Nanoparticles
5.1 Introduction
5.2 Composition and Structure
5.3 Preparation Methods
5.4 Characterization
5.5 Administration Routes of SLNs
5.6 Application of SLN for Delivery of Food Active Compounds
5.7 Safety of SLNs in the Food Industry
5.8 Conclusions and Future Focus
References
6 Nanostructured Lipid Carriers
6.1 Introduction
6.2 Materials Used in NLCs
6.3 Nanostructured Lipid Carrier Fabrication Techniques
6.4 Characterization of NLC
6.6 Conclusions
References
7 Filled Hydrogel Particles
7.1 Introduction
7.2 Filled Hydrogel Particle Formation and Stability
7.3 Filled Hydrogel Particle Compositions for Food Application
7.4 Characterization of Filled Hydrogel Particles
7.5 Filled Hydrogel Particles as Carriers for Delivery of Food Active Compounds
7.6
In Vitro/In Vivo
Digestion of Filled Hydrogel Particles
7.7 Biological Fate, Bioavailability, Health, and Safety
7.8 Conclusion
References
8 Nanoemulsions
8.1 Introduction
8.2 Formation of Nanoemulsions
8.3 Nanoemulsion Composition for Food Application
8.4 Characterization of Nanoemulsions
8.5 Nanoemulsions as Carriers for Food Active Compounds
8.6
In Vitro and In Vivo
Digestion of Nanoemulsions
8.7 Conclusions and Future Focus
References
9 Microemulsions
9.1 Introduction
9.2 Overview of Microemulsion Formation
9.3 Microemulsion Composition for Food Application
9.4 Microemulsions as Carriers of Bioactives and Antibacterial Components
9.5 Food Microemulsions: Health and Safety
9.6
In Vitro
Digestion of Microemulsions
9.7 Conclusions and Future Focus
References
10 Liposomes and Niosomes
10.1 Introduction
10.2 Liposome and Niosome Fabrication
10.3 Liposome and Niosome Composition for Food Applications
10.4 Characterization of Liposomes and Niosomes
10.5 Liposomes and Niosomes as Carriers to Deliver Food Active Compounds
10.6
In Vitro/In Vivo
Digestion of Liposomes and Niosomes
10.7 Biological Fate of Liposomes and Niosomes: Bioavailability, Health, and Safety
10.8 Conclusion and Future Directions
References
Index
End User License Agreement
List of Tables
Chapter 01
Table 1.1 Comparison of emulsifier (protein, polysaccharide, phospholipid, and small molecular surfactant) properties that may be utilized in the food industry (McClements, 2015; McClements and Gumus, 2016).
Chapter 02
Table 2.1 Overview of studies on
in vitro
and
in vivo
digestion of Pickering emulsions
Chapter 03
Table 3.1 W/O/W multiple emulsions for encapsulation of food active ingredients.
Chapter 04
Table 4.1 Hydrophilic‐lipophilic balance (HLB), acceptable daily intake (ADI) and solubility of selected food‐grade ionic small molecule emulsifiers used to produce multilayer m‐o/w emulsions.
Table 4.2 Application of multilayer emulsions in the food industry.
Chapter 05
Table 5.1 Ingredients used for production of solid lipid nanoparticles.
Table 5.2 Advantages and disadvantages of production techniques.
Table 5.3 Examples of bioactive compounds encapsulated by solid lipid nanoparticles.
Chapter 07
Table 7.1 Different biopolymers and their key features.
Table 7.2 Different preparation methods and associated parameters affecting particle size of filled hydrogel particles.
Chapter 08
Table 8.1 Classification of emulsions based on their droplet size.
Table 8.2 Recent studies on nanoemulsions prepared using a high‐pressure valve homogenizer.
Table 8.3 Recent studies on nanoemulsions prepared using a microfluidizer.
Table 8.4 Recent studies on nanoemulsions prepared using an ultrasonic homogenizer.
Table 8.5 Recent studies on nanoemulsions produced by high‐pressure homogenization combined with solvent evaporation.
Table 8.6 Recent studies on nanoemulsions prepared using low‐energy methods, such as spontaneous emulsification (SE) and emulsion inversion point (EIP).
Table 8.7 A review of recent studies on nanoemulsions as carriers for delivery of food active compounds.
Table 8.8 Recent studies on changes in the particle size and zeta potential of nanoemulsions during
in vitro
GI digestion and on the lipid digestion rate and bioaccessibility of nanoemulsions in the intestinal tract.
Chapter 09
Table 9.1 Comparison between emulsion, nanoemulsions, and microemulsions.
Table 9.2 List of surfactants commonly used in foods.
Table 9.3 Applications of microemulsions for solubilization of valuable food‐derived components.
Chapter 10
Table 10.1 Advantages and disadvantages of liposomes and niosomes.
Table 10.2 Potential entrapment efficiency and vesicle types formed using different methods.
List of Illustrations
Chapter 01
Figure 1.1 Cream and butter as oil‐in‐water and water‐in‐oil food emulsion.
Figure 1.2 Schematic diagram of emulsion destabilization.
Figure 1.3 Schematic diagram of the physiological conditions in the mouth‐GI tract which can cause significant changes in emulsion structures.
Chapter 02
Figure 2.1 Location of a colloidal solid particle at the oil‐water interface, as determined by the contact angle,
θ
, measured through the aqueous phase.
Figure 2.2 Schematic representation of a Pickering particle adsorbed at the oil‐water interface, with its position defined by the contact angle,
θ
, and surface free energy as described in equation 2.3 (situation a, left). For situation b, the surface free energy is defined in equation 2.5; the difference in free energy between situation a and b is the desorption energy.
Figure 2.3 Plot illustrating the desorption energy of a single colloidal solid particle with contact angle 10°, 50° or 90° as function of particle radius, for oil‐water interfacial tension of 20 mN/m and 40 mN/m. The dashed line represents thermal energy at 298 K (1
k
B
T
).
Figure 2.4 Interfacial deformation and associated capillary attractive force between solid particles at an oil‐water interface.
Figure 2.5 Schematic representation of particles at the surface of two neighboring emulsion droplets: (a) two dense particle monolayers separating two adjoining droplets; (b) single particles adsorbed at two distinct adjoining droplets; (c) low‐density network of aggregated Pickering particles.
Figure 2.6 Schematic illustration of Janus particles with several shapes and clear Janus boundaries: (a) spherical‐shaped particles showing the attractive capillary interaction arising from undulations in the Janus boundary, (b) dumbbell shaped, (c) ellipsoid shaped, and (d) cylinder shaped.
Figure 2.7 Size of objects (mainly of biological origin) on a nanometer scale to represent the size of Pickering stabilizers with respect to the definition of nanomaterials.
Chapter 03
Figure 3.1 A schematic representation of the multiple emulsions, their types and formation from basic emulsions.
Figure 3.2 Type A, type B, and type C double emulsions (from left to right).
Figure 3.3 Two‐step emulsification for preparation of stable W/O/W multiple emulsions.
Chapter 04
Figure 4.1 Multilayer emulsions are produced by a multistep procedure: (i) primary emulsion: oil and aqueous phases are homogenized together in the presence of a charged water‐soluble emulsifier; (ii) secondary emulsion: an oppositely charged polyelectrolyte is added to coat the droplets; (iii) tertiary emulsion (multilayer emulsions): sequential polyelectrolyte adsorption steps can be carried out.
Figure 4.2 Influence of pH, number, and type of layers on gravitational separation of multilayer emulsions. Primary: casein; secondary: casein‐pectin; tertiary: casein‐pectin‐chitosan; quaternary: casein‐pectin‐chitosan‐pectin.
Chapter 05
Figure 5.1 Structural models of solid lipid nanoparticles. (a) Food active‐enriched core model. (b) Food active‐enriched shell model. (c) Solid solution model.
Chapter 06
Figure 6.1 Schematic presentation of SLNs showing a burst release from polymorphism. The lipid matrix consists of similar components which will lead to gelation and flocculation after storage.
Figure 6.2 Schematic presentation of nanoparticulate structures of SLNs and NLCs.
Figure 6.3 Algorithm showing the procedures of NLC fabrication using hot and cold homogenization techniques.
Chapter 07
Figure 7.1 A filled hydrogel particle, detailing the different phases.
Figure 7.2 A schematic representation of the common methods for creating filled hydrogel particles. Data from Matalanis et al. (2010); McClements (2010); McClements (2012); Sung et al. (2015); Zeeb et al. (2015); Zhang et al. (2015a).
Chapter 08
Figure 8.1 Schematic illustration of mechanical devices used to produce nanoemulsions using high energy methods.
Chapter 09
Figure 9.1 Schematic of surfactant structure encountered self‐association structures in water, oil, or a mixture.
Figure 9.2 Hypothetical phase diagram of a system containing oil, water, and surfactant.
Figure 9.3 Mixture of oil, water, and surfactant as defined by Winsor.
Figure 9.4 Schematic of interaction energies in the interfacial area of a microemulsion system (oil‐surfactant‐water).
Chapter 10
Figure 10.1 Diagram of a liposome/niosome vesicle detailing different regions and components.
Figure 10.2 Factors to be considered in the selection of a liposomal/niosomal encapsulation method (Dua et al., 2012; Mozafari et al., 2008).
Figure 10.3 Schematic representation of the thin film hydration method detailing the required steps.
Figure 10.4 Schematic representation of the ether injection method.
Figure 10.5 Schematic representation of vesicle size reduction utilizing the bath sonication and probe sonicator methods.
Figure 10.6 Schematic representation of vesicle size reduction utilizing the extrusion technique.
Figure 10.7 Chemical structure of phosphatidyl choline.
Figure 10.8 Structure of non‐ionic surfactants. (a) Span 20; (b) Span 50; (c) Span 80.
Figure 10.9 Chemical structure of cholesterol.
Figure 10.10 Chemical structure of diacetyl phosphate.
Guide
Cover
Table of Contents
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Emulsion‐based Systems for Delivery of Food Active Compounds
Formation, Application, Health and Safety
Edited by
Shahin Roohinejad
Department of Food Technology and Bioprocess Engineering, Max Rubner‐Institut, Karlsruhe, Germany
Ralf Greiner
Department of Food Technology and Bioprocess Engineering, Max Rubner‐Institut, Karlsruhe, Germany
Indrawati Oey
Department of Food Science, University of Otago, Dunedin, New Zealand
Riddet Institute, Palmerston North, New Zealand
Jingyuan Wen
School of Pharmacy, University of Auckland, Auckland, New Zealand
This edition first published 2018© 2018 John Wiley & Sons Ltd
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Preface
Emulsion‐based delivery systems are mainly designed to encapsulate, deliver, and control the release, digestion, and absorption of hydrophobic and hydrophilic food active ingredients including small (e.g. volatiles), medium (e.g. food bioactives such as omega‐3 fatty acids, conjugated linoleic acid, butyrate, phytosterols, carotenoids, antioxidants, vitamins), and large molecular weight compounds (e.g. enzymes) in the food and pharmaceutical industries. Their functionality can be tailored by controlling their chemical composition and physical properties. This has led to the development of different emulsion systems, including conventional emulsions, Pickering emulsions, multiple emulsions, multilayered emulsions, solid lipid nanoparticles, nanostructured lipid carriers, filled hydrogel particles, nanoemulsions, microemulsions, liposomal emulsions, and niosomes. Each of these systems has its own advantages and disadvantages for controlling the absorption and release of food active compounds.
This book covers the principles of emulsion‐based systems formation, their characterization and application as carriers for delivery of food active ingredients as well as their digestibility and health and safety challenges for use in food systems. In each chapter, the formation of a specific emulsion‐based system and its application for delivery of food active compounds used in food systems are discussed. Additionally, the biological fate, bioavailability, and health and safety challenges of using emulsion‐based systems as carriers for delivery of food active compounds in food systems are reviewed.
This book is designed to assist food scientists as well as those working in the food, nutraceutical, pharmaceutical, and beverage industries. The topics covered in this book are suitable for teaching in courses such as food chemistry, food biochemistry, sensory science, new product development, and food processing.
We gratefully acknowledge the contribution of all colleagues from around the world and the professional assistance provided by the staff of Wiley.
Shahin Roohinejad, Ralf Greiner, Indrawati Oey, and Jingyuan Wen
About the Editors
Shahin Roohinejad
Dr Roohinejad obtained his BSc in 2000 from the Islamic Azad University, Iran, in the field of food science and technology. He completed his MSc in 2009 in food biotechnology at the University of Putra Malaysia (UPM). In July 2011, he received a full doctoral scholarship award from the Department of Food Science at the University of Otago in New Zealand, and he graduated in December 2014. In June 2015, Dr Roohinejad received a Georg Forster Research Fellowship award granted by the Alexander von Humboldt Foundation to pursue his postdoctoral research at the Department of Food Technology and Bioprocess Engineering, Max Rubner‐Institut (MRI), The German Federal Research Institut of Nutrition and Food. Currently, he is a postdoctoral research associate at the Food Science and Nutrition Department, University of Minnesota, USA. He is a professional member of the Institute of Food Technologists (IFT), graduate member of the New Zealand Institute of Food Science and Technology (NZIFST), associate newsletter editor of the IFT Non‐thermal Processing Division (NPD), a member of the IFT Press Advisory Group and GHI (Global Harmonization Initiative) Ambassador in Germany. In the last 10 years, he has worked on different food areas such as emulsion‐based systems, emerging food processing, nanotechnology, and functional foods. Dr Roohinejad’s research activities have resulted in more than 70 original papers in peer‐reviewed journals, book chapters, abstracts, and short papers in congress proceedings.
Ralf Greiner
Dr Greiner joined the Federal Research Centre for Nutrition, Karlsruhe, Germany, in 1990 as a PhD student after graduating in chemistry at the University of Stuttgart. In the early stages of his career as Deputy Head of the Centre for Molecular Biology, he was mainly engaged in research relating to genetically modified food and enzymes for food processing, with phytases being the center of his interests. In 2007, he held a position as Visiting Professor for Biochemistry and Molecular Biology, Department of Bioprocess Engineering, Federal University of Paraná, Curitiba, Brazil, working on solid‐state fermentation and fungal enzyme production. In 2008, he returned to Karlsruhe where he became Head of the Department of Food Technology and Bioprocess Engineering of the Max Rubner‐Institut (MRI). His research covers the studying and modeling of conventional and new processing technologies, as well as food nanotechnology, but phytases are still the main focus of his interests. Dr Greiner is a representative of MRI in several international and national associations on food technology, food control, and food nanotechnology. In 2012, he accepted a position as an Honorary Assistant Professor in the School of Biological Sciences of the University of Hong Kong. His research activities have resulted in approximately 120 original papers in peer‐reviewed journals, 37 book chapters and 260 abstracts or short papers in congress proceedings. In addition, Dr Greiner is Editor of Food Control.
Indrawati Oey
Professor Oey is Head of the Food Science Department at the University of Otago, Dunedin, New Zealand, and Principal Investigator at the Riddet Institute, Palmerston North, New Zealand. The University of Otago awarded her a full professorial chair as Professor of Food Science in 2009. She graduated with a BSc in Agricultural Technology (Food Science and Nutrition) from Bogor Agricultural University, Indonesia, and earned her MSc in Postharvest Food Processing and Preservation and PhD in Applied Biological Sciences from Katholieke Universiteit Leuven (KU Leuven), Belgium. She was a postdoctoral fellow at the Research Foundation‐Flanders at KU Leuven, Belgium (2000–2009). She was the chair of Training and Career Development for the European Union‐funded NovelQ Integrated Project (2006–2008). She is a Fellow of the New Zealand Institute of Food Science and Technology (NZIFST), a professional member of the Institute of Food Technologists (IFT) and NZIFST, served as Member‐at‐Large for the Executive Committee Board of Non‐thermal Processing Division – Institute of Food Technologists (2012–2015), and as secretary of the NZIFST Otago/Southland branch (2010–2015). She received the George Stewart Award from the Institute of Food Technologists (United States) in 2006. In the last decade, she has been actively involved in building international collaboration in the area of novel food processing, nanotechnology, functional foods, and food innovation. Professor Oey’s research activities have resulted in more than 200 original papers in peer‐reviewed journals, book chapters, abstracts, and short papers in congress proceedings.
Jingyuan Wen
Professor Wen has over 20 years’ research experience in pharmacology, nutraceuticals, pharmaceuticals, drug discovery from natural products, novel drug formulation, and delivery system design. She received her Bachelor of Medicine in 1986 and her Master’s degree in pharmacology in 1991 at the School of Pharmacy, Fudan University, China. She was awarded a PhD in Pharmaceutical Science in 2003 from the School of Pharmacy, University of Otago, New Zealand. She worked as a post‐doc researcher at Neuren Pharmaceuticals Ltd, New Zealand. She was appointed as a lecturer/senior lecturer at the School of Pharmacy, University of Auckland, in 2005. In 2015, she was promoted to Associate Professor, recognizing excellence in teaching and research. She was also appointed as a research theme leader in drug delivery in 2013. Under her supervision, 12 PhD students, 12 Master’s students, and 160 research dissertation and international exchange students have completed their degree. To date, she has published over 200 peer‐reviewed articles, patents, book chapters, and conference abstracts in drug delivery, natural products, and biological science. Professor Wen is a committee member of the Controlled Release Society (CRS), NZ local chapter. She is also an abstracts reviewer for the CRS (from 2007 to present) and was chair of the Oral Drug Delivery Award Committee of the CRS (2007 to 2012). She has established contacts with national and international commercial companies and has been involved in joint projects and consultations for national and international pharmaceutical and healthcare companies since 2005.
List of Contributors
Alireza AbbaspourradDepartment of Food ScienceCornell UniversityIthaca, USA
Murad Al GailaniSchool of PharmacyUniversity of AucklandAuckland, New Zealand
Alaa El‐Din A. BekhitDepartment of Food ScienceUniversity of OtagoDunedin, New Zealand
Claire C. Berton‐CarabinLaboratory of Food Process EngineeringWageningen University and ResearchWageningen, The Netherlands
Guanyu ChenSchool of PharmacyUniversity of AucklandAuckland, New Zealand
Shuo ChenSchool of PharmacyUniversity of AucklandAuckland, New Zealand
Meinou N. CorstensLaboratory of Food Process EngineeringWageningen University and ResearchWageningen, The Netherlands
Maral Seidi DamyehDepartment of Food Science and TechnologySchool of Agriculture, Shiraz UniversityShiraz, Iran
David W. EverettRiddet Institute, Palmerston NorthNew ZealandDairy Innovation InstituteCalifornia Polytechnic State UniversitySan Luis Obispo, California, USA
Hadi Hashemi GahruieDepartment of Food Science and TechnologySchool of AgricultureShiraz UniversityShiraz, Iran
Kelvin K.T. GohMassey Institute of Food Science and Technology, College of SciencesMassey UniversityAuckland, New Zealand
Ralf GreinerDepartment of Food Technology and Bioprocess EngineeringMax Rubner‐InstitutFederal Research Institute of Nutrition and FoodKarlsruhe, Germany
Seyedeh Sara HashemiBurn and Wound Healing Research CenterShiraz University of Medical SciencesShiraz, Iran
Kacie K.H.Y. HoLaboratory of Food Process EngineeringWageningen University and ResearchWageningen, The NetherlandsPlants for Human Health InstituteNorth Carolina State UniversityKannapolis, NC, USA
Mohamed KoubaaEcole Supérieure de Chimie Organique et Minérale,Compiègne, France
Sung Je LeeMassey Institute of Food Science and Technology, College of SciencesMassey UniversityAuckland, New Zealand
Mehrdad NiakousariDepartment of Food Science and TechnologySchool of Agriculture, Shiraz UniversityShiraz, Iran
Nooshin NikmaramDepartment of Food Science and TechnologyIslamic Azad University of SabzevarSabzevar, Iran
Indrawati OeyDepartment of Food ScienceUniversity of Otago,Dunedin, New ZealandRiddet Institute, Palmerston North New Zealand
Alireza RafatiDivision of Pharmacology & Pharmaceutical ChemistrySarvestan Branch, Islamic Azad UniversitySarvestan, Iran
Ali RashidinejadRiddet Institute, Centre of Research ExcellenceMassey UniversityPalmerston North, New Zealand
Shahin RoohinejadDepartment of Food Technology and Bioprocess EngineeringMax Rubner‐InstitutFederal Research Institute of Nutrition and FoodKarlsruhe, Germany
Anja SchröderLaboratory of Food Process EngineeringWageningen University and ResearchWageningen, The Netherlands
Karin SchroënLaboratory of Food Process EngineeringWageningen University and ResearchWageningen, The Netherlands
Pankaj SharmaFonterra Co‐operative Group LimitedPalmerston North, New Zealand
Anges TeoMassey Institute of Food Science and Technology, College of SciencesMassey UniversityAuckland, New Zealand
Jingyuan WenSchool of PharmacyUniversity of AucklandAuckland, New Zealand
Marie WongMassey Institute of Food Science and Technology, College of SciencesMassey UniversityAuckland, New Zealand
Naibo YinSchool of PharmacyUniversity of AucklandAuckland, New Zealand
Quan YuanMassey Institute of Food Science and Technology, College of SciencesMassey UniversityAuckland, New Zealand
1Conventional Emulsions
Mehrdad Niakousari1, Maral Seidi Damyeh1, Hadi Hashemi Gahruie1,2, Alaa El‐Din A. Bekhit3, Ralf Greiner4, and Shahin Roohinejad4
1 Department of Food Science and Technology, School of Agriculture, Shiraz University, Shiraz, Iran
2 Biomolecular Engineering Laboratory, Department of Food Science and Technology, School of Agriculture, Shiraz University, Shiraz, Iran
3 Department of Food Science, University of Otago, Dunedin, New Zealand
4 Department of Food Technology and Bioprocess Engineering, Max Rubner‐Institut, Federal Research Institute of Nutrition and Food, Karlsruhe, Germany
1.1 Introduction
Food active ingredients are widely used in the food industry to improve the nutritional and physicochemical properties and prolong the shelf‐life of food products. Incorporation of food active ingredients within foods has its own challenges, such as poor chemical stability, low bioavailability, and low water solubility. Thus, various studies have been conducted to develop effective systems for the delivery of these compounds (Cummings and Overduin, 2007; Maljaars, et al., 2009).
A variety of food active compounds can be incorporated into foods via specifically designed delivery systems with the aim of achieving a certain level of protection and reaching a specific targeted site, i.e. to control their release at specific locations within the gastrointestinal tract (e.g. mouth, stomach, small intestine, or colon) (Kosaraju, 2005). Delivery systems should be able to protect these compounds from physical and/or chemical degradation during processing, handling, and storage, and deliver them to the required site in the gastrointestinal tract without adverse effects on the appearance, stability, texture, or flavor of the food products (McClements, 2010; McClements et al., 2009). Conventional emulsions are considered the most important systems since they are the most widely used in the food industry (Augustin and Sanguansri, 2017).
Conventional emulsions are defined as emulsions having a particle size over 100 nm and are mostly produced using high‐energy techniques. Homogenizers are usually used to facilitate the conversion of two immiscible liquids into an emulsion with the aid of an emulsifier. High‐speed mixers, microchannel homogenizers, high‐pressure valve homogenizers, microfluidizers, colloid mills, membrane and ultrasonic homogenizers are some of the important high‐energy systems used at the industrial and research scales.
This chapter will focus on the formation, characterization, and recent advances in the application of conventional emulsions as a delivery system of valuable food ingredients. The stability of these systems under in vitro/in vivo conditions is also discussed.
1.2 Conventional Emulsions Formation and Stability
1.2.1 Formation
There are two types of naturally occurring emulsions that are found in food systems: typical low‐volume fraction emulsions such as milk and sauces, and a high‐volume fraction such as butter and margarine (Moschakis, 2013). Numerous food emulsions are prepared by combining raw materials, some of which are not found conjointly in nature. For example, a salad dressing product is manufactured by combining water, milk protein, soybean oil, apple vinegar, and seaweed polysaccharides. Each component of the formulation will influence the physical (intermolecular or interdroplet forces, phase separations) and chemical (formation of covalent bonds) properties of the product (Friberg et al., 2004).
Depending on the nature of the starting materials and method used to create an emulsion, the process may involve a single or a number of consecutive steps. In order to convert two separate oil and water phases into an emulsion, different functional ingredients should be dispersed first into the phase in which they are most soluble. For instance, lipid‐soluble compounds (e.g. oil‐soluble vitamins, antioxidants, and pigments) are mixed first in the oil phase, while water‐soluble compounds (e.g. proteins, polysaccharides, phenolic compounds, water‐soluble pigments, and vitamins) are mixed first in water phase (McClements and Li, 2010). However, sometimes it is more convenient to mix powdered functional ingredients directly into a mixture of oil and water. In order to prevent clumping during subsequent mixing and homogenization processes, the intensity and duration of the mixing process need to be optimized (Kinsella and Whitehead, 1989).
The presence of crystalline materials in the lipid phase prevents the formation of a stable emulsion. Therefore, a preheating step to melt fats prior to homogenization is required. Excessive heating, however, may initiate/promote the oxidation of polyunsaturated lipids, which in turn adversely affects the product quality (Ochomogo and Monsalve‐Gonzalez, 2009). Some parameters such as optimum conditions of ingredient mixing, solvent type, and operation temperature are important in the production of a stable functional emulsion.
As mentioned earlier, homogenization is used to convert two immiscible liquids into an emulsion with the aid of an emulsifier. Depending on the initial concentration of the two liquid phases, two different types of emulsions can be obtained in the presence of an emulsifier; at high oil and low oil concentrations, water in oil (w/o) and oil in water (o/w) emulsions will be formed, respectively (Figure 1.1). The balance between two opposing physical processes as well as the droplet disruption and coalescence will have a huge impact on the size of the droplets produced by the homogenization process (Schubert et al., 2003; Walstra, 2003).
Figure 1.1 Cream and butter as oil‐in‐water and water‐in‐oil food emulsion.
1.2.2 Stability
Emulsion stability is defined as the ability of an emulsion to resist changes to its properties over time. Stable emulsions have high resistance against changes in their properties and vice versa. Various physical and chemical processes may cause instability of an emulsion. As shown in Figure 1.2, several mechanisms such as creaming, sedimentation, flocculation, coalescence, partial coalescence, phase inversion, and Ostwald ripening can contribute to physical instability of an emulsion (Walstra, 2003). Oxidation and hydrolysis are the most common reasons for chemical instability of emulsions (McClements and Decker, 2000). A clear understanding of each mechanism, the interplay of relationships between oxidation and hydrolysis, and the processing and storage factors that influence them should be established in order to achieve more stable products.
Figure 1.2 Schematic diagram of emulsion destabilization.
Although several studies have tried to elucidate the general principles governing the stability of emulsions in order to predict their behavior under different processing conditions, the compositional and structural complexities of some emulsions do not allow accurate prediction of stability during storage (Goff and Hartel, 2003). Analytical techniques are normally used to monitor changes in emulsion properties over time. Environmental conditions (e.g. temperature and humidity of storage, mechanical agitation, and storage time) as well as composition and microstructure of an emulsion have major effects on the rate of breakdown and the mechanism by which this breakdown occurs.
1.3 Composition of Conventional Emulsions for Food Applications
The aqueous phase (W) of conventional emulsions can solubilize different amounts of water‐soluble ingredients such as acids, minerals, bases, preservatives, flavors, vitamins, surfactants, sugars, polysaccharides, and proteins. The partitioning, solubility, conformation, volatility, and chemical reactivity of some of these food components are measured by their interactions with water.
Lipids play an important part in beverage emulsions as flavor carriers or as the source of flavor themselves, such as essential oils (Tan, 2004). The choice of a suitable lipid in a formulation will have a huge impact on the nutritional, physicochemical, and sensory properties of conventional emulsions. In the formulation of a conventional emulsion, lipid acts as a solvent for hydrophobic food active ingredients such as oil‐soluble vitamins and antioxidants, preservatives, and essential oils.
The term “emulsifier” describes any surface active component that is able to interact with oil and water phases and create an oil‐water interface and protect emulsions from destabilization through aggregation, flocculation, and coalescence. In the food industry, low molecular weight surfactants, amphiphilic biopolymers, and surface active particulates are the emulsifiers most commonly used to form and stabilize emulsions (Table 1.1). An ideal emulsifier must be rapidly adsorbed to the oil‐water interface, eliminates the interfacial tension, and prevents the occurrence of droplet coalescence during homogenization through the creation of an interfacial membrane.
Table 1.1 Comparison of emulsifier (protein, polysaccharide, phospholipid, and small molecular surfactant) properties that may be utilized in the food industry (McClements, 2015; McClements and Gumus, 2016).
Source: McClements and Gumus (2016). Reproduced with permission of Elsevier.
Emulsifier
Molecular properties
Emulsion properties
Phospholipids
Lecithin
Surface active because of polar head group (phosphate moiety) and non‐polar (two fatty acids) tail group
Can form fairly small droplets at low levels using high‐pressure homogenization. Unstable under acidic conditions (pH <3), and at high ionic strength. May break down at high temperatures
Lysolecithin
Surface active because of polar head group (phosphate moiety) and non‐polar (one fatty acid) tail group
Small molecule surfactants
Quillija saponins
Surface active because they contain both hydrophilic (e.g. sugars) and hydrophobic (e.g. phenolics) regions
Can form small droplets at low levels using high‐pressure homogenization. Emulsions unstable at highly acidic conditions (pH <3), and at high ionic strengths. Stable to heating
Proteins
Whey protein
Mıxture of globular proteins from milkMW ≈ 18 kDa; pI ≈ 5, T
m
≈ 80 °C
Unstable at pH near pI, at high ionic strength, and at temperatures > T
m
. Stable at pH below or above pI, at low ionic strength, and at temperatures below T
m
Beta‐lactoglobulin
Globular protein from whey proteinMW ≈ 18.4 kDa; pI ≈ 5.4; T
m
≈ 83 °C
Unstable at pH near pI, at high ionic strength, and at temperatures > T
m
. Stable at pH below or above pI, at low ionic strength, and at temperatures below T
m
Alpha‐lactalbumin
Globular protein from whey proteinMW ≈ 14.2 kDa; pI ≈ 4.4; T
m
≈ 83 °C
Bovine serum albumin
Globular protein from whey proteinMW ≈ 66.3 kDa; pI ≈ 5.1; T
m
≈ 75 °C
Lactoferrin
Globular glycoprotein from whey proteinMW ≈ 80 kDa; pI ≈ 8; T
m
≈ 60 °C and 85 °C
Caseinates
Mixtures of flexible proteins from milkMW ≈ 24 kDa; pI ≈ 5
Unstable at pH near pI, and at high ionic strength. Stable at pH below or above pI, at low ionic strength, and to heating
Alpha‐s‐casein
Flexible protein from milkMW ≈ 23.6 kDa; pI ≈ 5.1
Beta‐casein
Flexible protein from milkMW ≈ 24.0 kDa; pI ≈ 5.5
Egg proteins
Mixture of globular proteins from egg white or yolk
Unstable at pH near pI, at high ionic strength, and at temperatures > T
m
. Stable at pH below or above pI, at low ionic strength, and at temperatures below T
m
Ovalbumin
Globular protein from egg white MW ≈ 45 kDa; pI ≈ 4.5; T
m
≈ 80 °C
Lysozyme
Globular protein from egg white MW ≈ 14.3 kDa; pI ≈ 11.3; T
m
≈ 72 °C
Legume proteins (soy, pea, lentil, chickpea, fava bean, etc.)
Mixture of globular proteins from legumes with variable molecular weightspI ≈ 4.3–5.0; T
m
≈ 82–90 °C
Unstable at pH near pI, at high ionic strength, and at temperatures > T
m
. Stable at pH below or above pI, at low ionic strength, and at temperatures below T
m
Gelatin
Fairly hydrophilic flexible protein from animal sources (collagen). Variable molecular weight depending on processing conditionspI ≈ 5 (Type B) or 8 (Type A); T
m
≈ 10–30 °C
Often not very surface active due to high hydrophilic character. Some types of gelatin can be used successfully as emulsifiers
Polysaccharides
Gum arabic
Branched glycoproteinMW ≈ 1000 kDa; pKa ≈ 3.5
Requires a high surfactant‐to‐oil ratio, but forms droplets stable to a wide range of pH, ionic strength, and temperature
Beet pectin
Branched anionic hydrophilic polysaccharide with hydrophobic ferulic acid groups
Citrus pectin
Branched anionic hydrophilic polysaccharide with hydrophobic protein groups attached
MW, molecular weight; pI, isoelectric point; Tm, melting temperature.
Texture modifiers are divided into thickening agents and gelling agents, depending on the molecular origin of their functional characteristics. These ingredients are added to the formulation of conventional emulsions to modify the texture of the continuous phase. In practice, there is often no clear distinction between thickening and gelling agents due to the ability of thickening agents to form gels when they are added at high concentrations as well as the ability of gelling agents to increase the viscosity of aqueous solutions without forming gels when they are used at low concentrations. A particular biopolymer can act as either a thickening or a gelling agent under particular conditions, for example temperatures, pH, or ionic strengths. Texture modifiers in conventional emulsions are used to provide the product with appropriate textural and mouthfeel characteristics, and to enhance emulsion stability by decreasing the rate at which particulates diffuse (McClements, 2005).
1.4 Characterization of Conventional Emulsions
Various analytical techniques are used to determine the physicochemical and sensory properties of conventional emulsions, such as stability, texture, flavor, and appearance. Some of the methods which are used to characterize the conventional emulsions are described below.
1.4.1 Testing Emulsifier Effectiveness
The effectiveness of an emulsifier for a specific food application depends on the minimum concentration needed to prepare a stable emulsion and to prevent droplets from aggregation and consequently destabilization of the emulsion during storage. In order to assess emulsifier efficiency, two simple empirical procedures (e.g. emulsifying capacity (EC) and emulsion stability index (ESI)) and several sophisticated methods (e.g. surface activity, saturation surface pressures, excess surface concentration, critical micelle concentrations (CMC), adsorption kinetics, and interfacial rheology) are used (McClements, 2007).
1.4.2 Dispersed Phase Volume Fraction
The dispersed phase volume fraction of a conventional emulsion can be simply determined by measuring its density (Pal, 1994). Densities of the dispersed (ρ1) and continuous (ρ2) phases are used to determine the density of emulsion (ρe) using the following equation: ρe = φρ1 + (1 − φ) ρ2 (φ = (ρe−ρ1)/(ρ2−ρ1)). With knowledge of the density of oil (~900 kg m−3), aqueous (1000 kg m−3), and dispersed phases, the dispersed phase volume fraction can be calculated. The density of emulsion is simply measured to 0.2 kg m−3 and the dispersed phase volume fraction is 0.002 or 0.2% of the volume. Increasing the oil content leads to a reduction in emulsion density (due to the lower density of liquid oil compared to water) (Pal, 1994).
The proximate analysis method is applied to determine the concentration of the dispersed phase in a conventional emulsion (Nielsen, 2003). Solvent extraction techniques such as continuous, semi‐continuous, and discontinuous phase of extraction can be used for measuring the fat content in an emulsion (Min and Boff, 2003). If the sample can be dried, then it can be ground and mixed with a solvent to extract fat. After the lipid extraction step, the solvent is separated from the sample, evaporated, and the remaining fat is weighed. A non‐solvent extraction technique can also be used to measure the fat content of an o/w emulsion in dairy products using the Babcock and Gerber methods.
Electrical conductivity (ε) of the dispersed phase of a conventional emulsion is used to determine its volume fraction (Asami, 1995). Water has a much higher electrical conductivity than oil, so the greater the oil content of an emulsion, the less the electrical conductivity (Clausse, 1983).
1.4.3 Measurement of Droplet Size Distribution and Microstructure
Several analytical instruments, such as photon correlation spectroscopy (PCS) and Doppler shift spectroscopy (DSS), were primarily designed to measure the particle size of emulsions. Dynamic light scattering (DLS) is one of the most popular techniques used to effectively determine particle size and distribution of emulsions. There is no particle‐particle interaction in a diluted emulsion and the size of the droplets can be determined by the Stokes–Einstein equation.
Static light scattering is another rapid and reproducible technique that has been used to measure the particle size of emulsions, varying from 0.05 to 2000 µm. In this method, a beam of light passes through an emulsion and is scattered by the droplets in a well‐defined manner (Hiemenz and Rajagopalan, 1997). Some instruments are used only to measure the light intensity, especially non‐scattered light by passing it directly through the emulsion, while others determine the light intensity which has been scattered by the emulsion droplets.
Nuclear magnetic resonance (NMR) investigates the interaction between hydrogen atom nuclei and electromagnetic waves. NMR can be used for the measurement of particle size distribution in conventional emulsions based on limited diffusion of molecules within the droplets (Balinov et al., 2004). The hydrogen atoms in the sample, which is located in a static magnetic field gradient, are excited by exposure to radiofrequency pulses. These hydrogen atoms will emit signals that are detectable by NMR. The amplitude of emitted signals is used to examine molecular movements. Sizes in the range of 0.2–100 µm can be determined by this method (Dickinson and McClements, 1995).
In the ultrasonic spectrometry techniques, the size of droplets with radii between 10 nm and 1000 µm can be measured from the interactions between ultrasonic waves and emulsions (Coupland and McClements, 2004). Compared to other traditional technologies, this method has some advantages due to its ability to analyze concentrated and optically opaque emulsions without any sample preparation.
The neutron scattering method determines the particle size, size distribution of particles, and the thickness of the interfacial layer in a conventional emulsion on the basis of the interaction between emulsion and neutron beam (Reynolds et al., 2000). Two specific features of this method are low scattering of emulsions that make it possible to use concentrated emulsion without dilution, and the ability to manipulate the contrast between different ingredients in heterogeneous foods to differentiate between their neutron scatterings. Thus, particular structural features can be detected in the emulsion. The main limitation of this method is the need for a nuclear reactor to create the neutron beam (Hone et al., 2002).
A combination of electricity and sound waves, known as electroacoustics, is used for the measurements of zeta potential and particle size distribution of emulsions (Dukhin et al., 2000). Particle sizes in the range of 0.1–10 µm can be measured by electroacoustics. Coupled with ultrasound spectrometry, this technique can determine the particle size in a larger range of 10 nm to 1000 µm (Hsu and Nacu, 2003). Particle size of high concentration emulsions and a wide variety of w/o and o/w conventional emulsions can be measured using this technique (Djerdjev et al., 2003; Kong et al., 2003; O’Brien et al., 2003). The main disadvantages of this method are the presence of charged droplets, a significant difference between the density of droplets and continuous phase, and a defined viscosity of continuous phase in each frequency.
In the dielectric spectroscopy technique, the conventional emulsion is exposed to a wide range of electromagnetic frequencies and its electric permittivity measured, which can be converted to particle size distribution and droplet size (Sjoblom et al., 1996). Only emulsions containing charged particles can be analyzed using this method. In addition to particle size distribution, zeta potential can also be determined. Dielectric spectroscopy is suitable to measure particle size distribution of opaque and concentrated emulsions without dilution.
A number of microscopic techniques, such as conventional optical microscopy, laser scanning confocal microscopy, electron microscopy (transmission electron microscopy (TEM), scanning electron microscopy (SEM)), and atomic force microscopy, can be used to study the structure, dimensions, and organization of the components of a conventional emulsion (Kirby et al., 1995; Morris et al., 1999). According to Aguilera and Stanley (1990), any type of microscopy that is going to be used in examining the structure of small objects must have three qualities: resolution, magnification, and contrast.
1.4.4 Droplet crystallinity
Nuclear magnetic resonance has good potential for measurement of the solid content in conventional emulsions (Dickinson and McClements, 1995). Despite the high initial cost of the NMR apparatus, rapid analysis of emulsions without any preparation of concentrated or optically opaque samples has resulted in it replacing the time‐consuming dilatometry method. When an emulsion is exposed to a radiofrequency pulse, the change of some of the hydrogen nuclei into an excited state leads to the generation of a detectable NMR signal. The basic parameters of this signal include frequency, amplitude, and decay time. Analyzing these parameters provides a wide range of information about the solid content of the material.
Dilatometry is a useful technique to detect the crystallinity of dispersed and continuous phases of conventional emulsions based on density changes that occur during melting or crystallization of materials (Phipps, 1964). In principle, the dilatometer measures the decrease and increase in density of a material when it melts or crystallizes, respectively. Compared to the liquid state of a material, the density of solid state is usually greater due to the more efficient packing of the molecules.
Differential scanning calorimetry (DSC) and differential thermal analysis (DTA) can be used to monitor the melting and crystallization behavior of conventional emulsion droplets (Palanuwech and Coupland, 2003). Since the materials have a tendency to release heat during crystallization and absorb heat during melting, these methods have been used to measure the release or absorption of heat by a sample when it is exposed to a controlled temperature change process.
Ultrasound is another method which can be applied to detect phase transitions in a conventional emulsion (Coupland and McClements, 2004). The melting or crystallization process can be altered significantly by the ultrasonic characteristics of a material. Since the velocity of ultrasonic waves in a solid medium is higher than in a liquid one, cooling of an emulsion from initial temperature of liquid droplets to the temperature where droplets start to crystallize causes a sharp increase in sound velocity. Conversely, increasing the emulsion temperature leads to a sharp decrease in sound velocity, due to the droplet melting process. Generally, supercooling effects cause droplet crystallization to occur at much lower temperatures than the melting point of the bulk oil (Dickinson et al., 1992).
1.4.5 Droplet Charge
Different instruments are used based on electroacoustics for determination of the concentration, size, and zeta potential of particles in emulsion systems (Kong et al., 2001a–c). Electroacoustic properties analysis can be conducted in two different ways:
colloid vibration potential (CVP), in which an acoustic signal is sent to a sample and the resulting electric signal obtained by the oscillating particles is detected
electrosonic amplitude (ESA), in which an electric signal is applied to a sample and the resulting acoustic signal formed by the oscillating particles is registered.
In the particle electrophoresis technique, the conventional emulsion is located in a cell and a static electrical field is transmitted through it via two electrodes. This results in charged particles being absorbed by the electrode on the side having an opposite charge (Hunter, 1993). The movement of droplets can be detected using different analytical methods. For instance, optical microscopy or static light scattering can be used to monitor the movements of large particles (>1 µm), while for detection of the movement of smaller particles, ultramicroscope, DLS, or static light scattering is normally used.
The relationship between droplet interaction and the physicochemical properties of a conventional emulsion is an interesting area in emulsion research. Unfortunately, there are no commercially available devices to study the charge, magnitude, and range of droplet interactions. However, some experimental techniques have been introduced to predict and control the fundamental interactions and to provide information about how these forces can influence the stability of an emulsion. For instance, in order to investigate the attractive forces between flocculated emulsion particles, plotting apparent viscosity as a function of shear stress by applying a suitable mathematical model can be used (Berli et al., 2002; Quemada and Berli, 2002).
Evaluation of emulsion stability is also a good tool to assess the magnitude of certain types of dispersed particle interactions. For example, it is possible to obtain indirect information about the strength of attractive interactions by determining the minimum salt concentration needed to improve flocculation of droplets in an emulsion stabilized electrostatically (Montagne et al., 2003).
The magnetic chaining approach directly monitors the force‐separation distance profile between colloidal particles. When an emulsion‐based paramagnetic colloid is exposed to an external magnetic field, monodispersed particles spontaneously align into linear chains due to magnetic dipole induction (Dimitrova and Leal‐Calderon, 1999). The equilibrium between repulsive and attractive magnetic forces gives rise to the separation of droplets within chains, which can be measured using light scattering techniques. In this method, changing the strength of the applied external magnetic field can control the magnitude of the magnetic forces.
Atomic force microscopy has permitted the probing of droplet‐droplet interactions by directly measuring force versus distance profiles for a particle attached to the end of a cantilever and a particle immobilized on a solid surface (Gunning et al., 2004).
1.5 Conventional Emulsions as Carriers for Delivery of Food Active Compounds
Conventional emulsions have been considered as stable delivery systems especially for lipophilic components that can be solubilized in the hydrophobic area of the oil droplet in an o/w emulsion (Iwamoto et al., 1991; O’Mullane et al., 1987). A hydrophobic or hydrophilic bioactive component is dispersed in the oil or aqueous phase, respectively, prior to homogenization and then a stabilized emulsion system will be created to provide protection to the bioactives in food products (Fustier et al., 2010).
Encapsulation and delivery of lipophilic compounds in o/w conventional emulsions have numerous advantages, including relative ease of preparation and low cost (McClements et al., 2009), high physical and chemical stability of the component by designing the oil‐water interface (Mao et al., 2013), ability to produce different rheological properties (Genovese et al., 2007), and versatility of production in wet state (Ru et al., 2010) or as solid powders, which facilitates their transportation and storage (Kumari et al., 2011).
However, conventional emulsions as delivery systems have some disadvantages, such as susceptibility to environmental stresses (e.g. heating, extreme pH, chilling, and high ionic strength) that may lead to physical and chemical instability under these conditions. Examples of chemical instabilities include oxidation and hydrolysis, and examples of physical instabilities include coalescence, flocculation, Ostwald ripening, and creaming (Dickinson, 2010; McClements and Decker, 2000). Applying chelating agents and antioxidants such as tocopherols results in sequestering heavy metals and consequently reduces the oxidation of lipophilic bioactive components (Hu et al., 2004; Ribeiro and Shubert, 2003). Maillard reaction products created by heat treatment of aqueous protein‐carbohydrate mixtures were reported by Augustin et al. (2006) to protect oxidation‐sensitive compounds such as fish oil. Protein (e.g. beta‐lactoglobulin), carbohydrates (in the amorphous state), and gums (e.g. gum arabic, guar gum) as encapsulating matrices may be able to stabilize emulsions by acting as an oxygen barrier around the emulsion oil droplets. Moreover, milk protein, as an emulsifier system, can be used to protect polyunsaturated lipids from oxidation (Fustier et al., 2010). The protection afforded by the emulsifiers and release of the encapsulated component may result in small droplets (approximately µm) and small interfacial layers (approximately nm) due to very short time scales for molecular diffusion (McClements et al., 2007, 2009).
Using emulsifying agents, such as dairy proteins, gives rise to lipid droplets that carry cationic charge and repelling cationic transition metal ions (e.g. Fe2+). In order to sequester the transition metal ions and avoid contact with the lipid phase, a chelating agent (e.g. EDTA) can be added to the aqueous phase. Antioxidants with the ability to partition into the lipid droplet interfaces can be used to prevent oxidation reactions (McClements et al., 2007). Due to the emulsifying and ligand‐binding characteristics of whey protein isolate (WPI), it can be used in an emulsion formulation for simultaneous encapsulation of food active ingredients with various physicochemical properties. Accordingly, binding of bioactive components such as vitamins, polyphenols, and fatty acids to the oil droplets membrane in the o/w emulsion can be stabilized by whey protein (Wang et al., 2016a).
Wang et al. (2016a) demonstrated the possibility of simultaneous encapsulation of alpha‐tocopherol and resveratrol in the oil phase and at the oil‐water interface of o/w emulsions, in which roughly 94% of alpha‐tocopherol and 50% of resveratrol were encapsulated in oil droplets stabilized by 0.01% WPI. Binding to WPI leads to partitioning of amphiphilic resveratrol between the aqueous phase and the oil‐water interface (Wang et al., 2016a). Due to a physical barrier created by WPI, alpha‐tocopherol was protected from decomposition (Ries et al., 2010).
Different bioactive lipophilic components have been encapsulated by conventional emulsions to prevent their oxidation, such as omega‐3 fatty acids (Chee et al., 2007; McClements and Decker, 2000), lycopene (Ribeiro and Shubert, 2003; Ribeiro et al., 2006; Tyssandier et al., 2001), astaxanthin (Ribeiro et al., 2006), lutein (Losso et al., 2005; Santipanichwong and Suphantharika, 2007), beta‐carotene (Santipanichwong and Suphantharika, 2007), plant sterols (Sharma, 2005), and conjugated linoleic acids (Jimenez et al., 2004), and have been incorporated into different food products (e.g. milk, yoghurts, ice cream, and meat patties) (Chee et al., 2007; McClements and Decker, 2000; Sharma, 2005).
Beta‐carotene, a natural colorant and antioxidant, is a compound beneficial to human health through its ability to decrease the risk of cancer, cardiovascular disease, and cataracts (Hou et al., 2014). Encapsulating beta‐carotene in emulsion‐based delivery systems can help in overcoming drawbacks such as poor water solubility and chemical instability (Roohinejad et al., 2014a, 2014b, 2015). Gu et al. (2017) investigated beta‐carotene protection in o/w conventional emulsion using conjugates made from egg white protein and catechin as emulsifiers. Applying the egg white protein and prepared conjugate in the emulsion resulted in droplets with a mean diameter of 0.203 and 0.328 µm, respectively. Unexpected large oil droplets were produced by the conjugate due to its impact on lowering the interfacial tension, but a physically stable emulsion was produced (McClements and Gumus, 2016).
The mean droplet diameter of emulsions stabilized by polyphenol‐protein conjugates is influenced by the type of proteins and polyphenols used in the emulsion (Gu et al., 2017). For example, Wei et al. (2015) reported that droplet size was reduced in emulsions stabilized by epigallocatechin gallate attached to alpha‐lactalbumin or beta‐lactoglobulin compared to the protein alone. This was in contradiction to the findings of Gu et al. (2017) who used egg white protein‐catechin conjugates that resulted in larger droplet size. Interestingly, in emulsion stabilized by egg white protein‐catechin conjugates, the degradation rate of beta‐carotene was significantly less than those emulsions prepared with egg white protein alone (Gu et al., 2017). Moreover, better physical and chemical stability was observed for emulsions stabilized by egg white protein‐catechin conjugates, due to an increase in the thickness of the interfacial layer and strong antioxidant activity of conjugates.
Crystalline bioactive lipid components such as carotenoids should be melted prior to homogenization in order to prevent fouling of homogenizers as well as to ensure that they are below saturation concentration in the carrier oil to prevent emulsion instability (McClements et al., 2007). Lycopene, for example, is crystalline at ambient temperature (melting point 173 °C); dispersing its crystals into carrier oil (e.g. medium chain fatty acid triacylglycerols (MCT)) and heating it until melted (about 140–210 °C) is critical to reach a stable emulsion as a delivery system. Also, the homogenization condition should be controlled to prevent chemical degradation of highly susceptible lycopene by minimizing homogenization time and reducing the oxygen content in the system (Ribeiro and Schubert, 2003). Investigation of emulsified lycopene incorporated into milk, orange juice, and water demonstrated more chemical stability in orange juice than in milk or water, which can be enhanced by adding alpha‐tocopherol as an antioxidant (Ribeiro and Schubert, 2003). Homogenization conditions (i.e. temperature, pressure, and cycles) should be carefully controlled to avoid labile bioactive compounds such as omega‐3 fatty acids and carotenoids being subjected to factors enhancing the degradation rate, including high temperatures, oxygen, light, or transition metals (McClements et al., 2007). The homogenization conditions largely affect the properties of emulsions, including droplet size, stability, and viscosity, and consequently must be considered in the design of emulsions for targeted delivery (Lu et al., 2015; Yuan et al., 2008).
The effect of conventional emulsion components (i.e. Arabic gum, xanthan gum, orange oil) on volatile flavor release from a model orange beverage was investigated by Mirhosseini et al. (2008b). Increasing orange oil content resulted in an increase in the average droplet size (Mirhosseini et al., 2008a), which consequently enhanced flavor release due to an increase in the total oil‐water interfacial surface area. In fact, the transfer rate of the hydrophobic component from oil phase to water phase may be increased as a result of an increase in interfacial surface area (Mirhosseini et al., 2008b). A high content of xanthan gum enhanced the resistance to transfer hydrophobic components due to its distinctive hydrophobic character, and consequently results in a negative impact on flavor release (Mirhosseini et al., 2008b). This reduction in flavor release was caused by adsorption, entrapment, and binding forces as a result of interactions between flavor components and matrix constituents (Shahidi and Zhong, 2011). This phenomenon was also observed by other researchers using other materials or processing conditions, such as enhanced release of aroma components from o/w emulsions containing Tween 20 (van Ruth et al., 2000), and enhanced release of lemon and citrus aromas (Charles et al., 2000) due to an increase in droplet size.
Vitamin E, an oil‐soluble antioxidant, exhibits various health effects, such as reducing cardiovascular disease, diabetes, and cancer, and has consequently gained interest as an ingredient for food products (Sylvester et al., 2011). Partial absorption of vitamin E at intestinal sites reduces its bioavailability. A better absorption was found in the presence of surfactants or emulsions (Julianto et al., 2000; Nacka et al., 2001). Yang and McClements (2013) investigated o/w emulsion delivery of vitamin E using a natural food‐grade surfactant (Q‐Naturale®) and compared its performance to Tween 80. Q‐Naturale was effective at producing emulsions with relatively small droplets, which was found to decrease even more by adding glycerol in the aqueous phase prior to homogenization. Higher percentages of small droplets were produced by the presence of Tween 80 and Q‐Naturale in an oil phase containing low levels (<40%) and high levels (60–80%) of vitamin E acetate, respectively. It was previously reported that a reduction in emulsion droplet size could be achieved by either a decrease in vitamin concentration in the oil phase or an increase in glycerol content in the aqueous phase (Yang and McClements, 2013).
