Advances in microencapsulation of polyunsaturated fatty acids (PUFAs)-rich plant oils using complex coacervation: A review

Food Hydrocolloids 69 (2017) 369e381

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Food Hydrocolloids journal homepage: www.elsevier.com/locate/foodhyd

Advances in microencapsulation of polyunsaturated fatty acids (PUFAs)-rich plant oils using complex coacervation: A review Yakindra Prasad Timilsena a, b, Bo Wang c, Raju Adhikari a, b, Benu Adhikari a, b, * a

School of Science, RMIT University, Melbourne, VIC 3083, Australia CSIRO Manufacturing, Clayton South, VIC 3169, Australia c Numega Ingredients, Brisbane, QLD 4109, Australia b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 28 October 2016 Received in revised form 9 February 2017 Accepted 2 March 2017 Available online 6 March 2017

Polyunsaturated fatty acids (PUFAs) are essential fatty acids that are abundantly available in marine fish, algae and some plant seeds. In recent years, the demand of plant-based PUFAs is increasing rapidly due to dietary and lifestyle requirements. The toxicity and environmental sustainability concerns associated with fish oils are driving the demand for plant-based PUFAs. PUFAs-rich oils rapidly undergo deterioration due to oxidation in the course of processing, transportation and storage. They require adequate protection and are microencapsulated before their application or incorporation in food products. Complex coacervation-based microencapsulation technology is increasingly being used in food and pharmaceutical industries due to high encapsulation efficiency, high payload and mild processing conditions associated with this technology. This article provides a comprehensive overview of the relevant scientific literature on complex coacervation with special emphasis on the production of plant protein-plant polysaccharide (gum) complex coacervates. Microencapsulation of PUFAs-rich oils in plant proteingum complex coacervates and resultant oxidative stability of PUFAs is highlighted. Finally, the release and digestion of encapsulated PUFAs in the human physiological conditions are discussed. Coencapsulation of various functional ingredients together with PUFAs derived from plants in plant protein-gum complex coacervate is identified as important future research direction. © 2017 Elsevier Ltd. All rights reserved.

Keywords: Polyunsaturated fatty acids (PUFAs) Plant oil Microencapsulation Complex coacervation Digestion

Contents 1. 2. 3. 4. 5. 6. 7.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 369 Fatty acids and PUFAs profile and content in plant oils . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 372 Complex coacervation process . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 372 Microencapsulation of PUFAs-rich oil using complex coacervation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373 Recent advances in complex coacervation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 Physiological digestion of encapsulated oils and fats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376 Concluding remarks and future perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 379

1. Introduction

* Corresponding author. School of Science, RMIT University, Melbourne, VIC 3083, Australia. E-mail address: [email protected] (B. Adhikari). http://dx.doi.org/10.1016/j.foodhyd.2017.03.007 0268-005X/© 2017 Elsevier Ltd. All rights reserved.

Epidemiological and clinical studies have unveiled that oils and fats that are rich in polyunsaturated fatty acids (PUFAs) such as omega-3 (u-3) and omega-6 (u-6) can provide health benefits in addition to providing energy and being a carrier of lipid soluble

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nutrients. PUFAs play vital role in maintaining health and wellbeing in humans by minimizing the risk of cardiovascular and neurodegenerative disease, arthritis, diabetes and certain types of cancer (Orsavova, Misurcova, Ambrozova, Vicha, & Mlcek, 2015). Since human body is unable to synthesise PUFAs in required quantity, they are known as the essential fatty acids (EFAs) and have to be supplied to the body through diet or dietary supplements. PUFAs are abundantly present in marine fish, algae and some plant seeds. PUFAs contained in plant oils are short chain fatty acids (SCFAs) such as a-linolenic acid (ALA, C18:3 u
3) and linoleic acid (LA, C18:2 u
6), whereas fish and algal oils contain long chain PUFAs (LC PUFAs) such as eicosapentaenoic acid (EPA, C20:5 u
3), docosapentaenoic acid (DPA, C22:5 u
3), docosahexaenoic acid (DHA, C22:6 u
3) and arachidonic acid (AA, C20:4 u
6). It has been commonly accepted that these LC PUFAs can be synthesised in human body using SCFAs (ALA and LA) as precursors, provided they are present in sufficient quantity. Therefore, ALA and LA are known as true “essential” fatty acids (Mehta, 2006; Rubio-Rodríguez et al., 2010). It is also established that the extent or percentage of conversion of ALA to EPA and DHA and that of LA to AA is small, for example, only 15e25% of ALA is converted into DHA in human body (Burdge & Wootton, 2002; Emken, Adlof, & Gulley, 1994). It is also believed that this slower conversion process does not produce sufficient LC PUFAs to meet the physiological demand in human body and thus EPA and DHA should also be supplied in the diet in order to maintain health and well-being. Due to these reasons, EPA and DHA are also considered as EFAs. Commercially available long chain PUFAs mostly come from fish oils derived from fatty fish such as tuna, sardine, mackerel, herring and trout (Meyer et al., 2003). However, these fish oils possess strong fishy flavour and are not favoured by consumers and cannot be incorporated into vegetarian and vegan diets. Furthermore, current global production of fish is unable to meet the demand of u-3 PUFAs which is of the order of 1.25 million metric tonnes per annum (Betancor et al., 2015). Due to the high demand of PUFAsrich fish oil, marine fish has been harvested excessively, which if not managed correctly, would result in the decline and eventually extinction of these fish species. Furthermore, there is an increasing concern of the presence of methyl mercury and organic pollutants such as polychlorinated dibenzo-p-dioxins (PCDD), polychlorinated dibenzofurans (PCDF) and dioxin-like polychlorinated biphenyls (DL-PCB) in marine fish (Smutna, Kruzikova, Marsalek, Kopriva, & Svobodova, 2008). Hence, many food manufacturers and researchers are interested in developing alternative source of PUFAs in order to address the aforementioned limitations in the inherent characteristics and availability of fish oils. One of the important considerations in selecting ideal source of PUFAs is the ratio of u-6 to u-3 in the oil. This ratio is very important because heavily processed western diets are rich in u-6 PUFAs but deficient in u-3, thus creating a gross imbalance in the intake of u-6 and u-3 (De Silva, Francis, & Tacon, 2011, pp. 1e20). In order to address this issue, consumption of oils containing higher concentration of u-3 is to be increased. As a consequence, demand of foods enriched with u-3 is rapidly increasing. However, incorporation of these u-3 PUFAs in processed foods still remains challenging due to their unstable nature. The chemical structure of PUFAs [Fig. 1 (AeE)] shows they have two or more double bonds in their structure. PUFAs are comprised of bis-allylic methylene groups (eCH]HeCH2eCH]CHe) and all the double bonds are present in the cis-configuration. This type of chemical structural make up renders them inherently unstable and sensitive to oxidation, isomerisation and polymerisation when they come in contact with environmental stressors such as oxygen, moisture, heat, light and metallic ions (Rustan & Drevon, 2005). Oxidation of lipids produces undesirable and harmful volatile

compounds and causes rancidity in foods (German, 1999). Therefore, suitable protection and delivery mechanisms have to be developed in order to prevent the degradation of PUFAs-rich oils during processing, storage and transportation and to facilitate their incorporating in processed foods. Two main strategies are being pursued by food industries to minimise the deterioration of PUFAs-rich oils during processing, transportation and storage. They are: 1) incorporation of natural or synthetic anti-oxidants in oils and fats to minimise oxidation, and 2) microencapsulation of oils and fats to prevent their direct contact with environmental stressors including oxygen, heat, moisture and light. In many cases, both strategies are applied concurrently to achieve synergistic effects. This review will focus only on the second strategy. Microencapsulation is a process during which a core material (e.g. PUFAs-rich oil) is entrapped inside a thin layer of coating material (shell). Protein and polysaccharides including gums, either individually or in complexed form, are commonly used as shell materials for microencapsulation of food ingredients including PUFAs-rich oils. The shell matrix creates a physical barrier between the core and the external environment so that the core is prevented from direct contact of moisture, oxygen, heat, light, acid or alkali. The shell matrix also helps to mask the undesirable odour, taste or colour of the core so that sensory attributes and palatability of the product are either enhanced or at least not compromised. Furthermore, shell can also prevent undesirable interaction between core material and other food components and can enhance their bioavailability (Sanguansri & Augustin, 2011). In recent years, microencapsulation is used to design controlled release formulations in order to achieve targeted and sustained delivery of lipids, increase satiety and help control the body weight (Ubbink & Krüger, 2006; Wilson & Shah, 2007). Thus, microencapsulation enhances the nutritional efficacy of food additives and helps broadening the application of unstable food ingredients in a wider range of food products (Desai & Park, 2005). Microencapsulation has become an important and ubiquitous technology for stabilizing and delivering numerous unstable and lipophilic ingredients in food and pharmaceutical products. The first successful microencapsulation of flavour oil was carried out in gum acacia matrix through spray drying in 1927 (Fanger, 1974). Spray drying is now more frequently and widely used in industry to encapsulate many unstable food ingredients and active pharmaceutical agents (Arslan, Erbas, Tontul, & Topuz, 2015). Several other methods such as freeze drying (Anwar & Kunz, 2011), fluidized bed coating (Dewettinck & Huyghebaert, 1998), centrifugal extrusion (Desai & Park, 2005), inclusion complexation (Song, Bai, Xu, He, & Pan, 2009), complex coacervation (Wang, Adhikari, & Barrow, 2014; Eratte et al., 2015; Timilsena, Wang, Adhikari, & Adhikari, 2016; Timilsena, Adhikari, Barrow, & Adhikari, 2016), ionotropic gelation (Bokkhim, Bansal, Grøndahl, & Bhandari, 2016), liposome entrapment (Mozafari et al., 2008), and electro-spraying (Coghetto et al., 2016) have also been developed in order to encapsulate large array of unstable food and pharmaceutical components. The choice of microencapsulation methods depends on the nature of the core and shell material and intended application of the final product. The design of an effective encapsulation system requires a sound understanding of the physicochemical mechanisms of encapsulation, possible interaction of the active compounds with the shell matrix and the nature of shell matrix (Augustin & Hemar, 2009). Complex coacervation is regarded as one of the simplest, low cost, scalable, reproducible and most effective methods of microencapsulation of hydrophobic substances (Barrow & Shahidi, 2007). This method yields considerably high encapsulation efficiency and low surface oil and provides high oxidative stability of

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371

Fig. 1. Chemical structure of common PUFAs. (A): Linoleic acid (C18:2, u-6), (B): Alpha linolenic acid (C18:3, u-3), (C): Eicosapentaenoic acid (C20:5, u-3), (D): Docosahexaenoic acid (C22:6, u-3) and (E): Arachidonic acid (C20:4, u
6) [Adapted from Holub (2002)].

the core (Gouin, 2004; Wang et al., 2014). In addition, microencapsulation using complex coacervation can be carried out at mild processing conditions without requiring toxic solvents (Gouin, 2004). Complex coacervates formed by the interaction of oppositely charged polymers are believed to be more surface active than the individual protein or polysaccharide; therefore, they are regarded as better emulsifier and stabilizers in emulsion systems (Benichou, Aserin, Lutz, & Garti, 2007). This characteristic of complex coacervates consequently helps to obtain increased encapsulation efficiency and results into higher oxidative stability of the core. In the above context, this review mainly focuses and attempts to provide a detailed overview of the complex coacervation method of microencapsulation that has been employed to stabilize PUFAs-rich oils. Various aspects of microencapsulation of PUFAs-rich plant oils using complex coacervation technology have been reviewed elsewhere (Ducel, Richard, Saulnier, Popineau, & Boury, 2004). Complex coacervation of plant proteins and polysaccharides and their

use in microencapsulation of various core materials, particularly PUFAs-rich oils, are the two foci of this review. Recent advances in complex coacervation science involve the preparation of complex coacervates from plant protein and gum extracted from a single plant and utilization of such plant protein-gum complex coacervates to microencapsulate PUFAs-rich oil from the same plant. This review paper will also evaluate such complex coacervation techniques of microencapsulation. Digestion of oils and fats and bioavailability of PUFAs are critical factors to be considered in food fortification and functional food development. Therefore, an attempt has also been made to review relevant information on the digestion of PUFAs in the physiological conditions. Section 2 of this review provides an overview of the structural organization of fatty acids and triacylglycerols and compares fatty acid profiles of some important plant oils. Sections 3 and 4 provide a comprehensive overview of the important aspects of prevailing microencapsulation technologies, principles and process of complex coacervation between proteins and polysaccharides and

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application of resultant complex coacervates in encapsulating PUFAs-rich oils. Recent advancement in complex coacervation technology using plant protein-gum system is presented in Section 5, whereas available literature on the physiological digestion process involved in lipid metabolism is reviewed in Section 6. The final section (Section 7) provides concluding remarks and points to the future direction/scope of complex coacervation research. 2. Fatty acids and PUFAs profile and content in plant oils Major lipids present in the plant oils are triacylglycerols comprising more than 95% of the total oil mass (Singh, Ye, & Horne, 2009). These triacylglycerols are comprised of three fatty acids joined together onto a glycerol backbone through an ester linkage (Fig. 2). Length of fatty acid chain, number and position of double bonds in them and positional distribution of fatty acids in the triacylglycerol molecule determine the physicochemical properties of oils. Fatty acids are long molecules comprising a non-polar hydrocarbon chain with a carboxyl acid group (COOH) at one end and a methyl group (CH3) at the other end. The fatty acids can be saturated, monounsaturated or polyunsaturated depending on the absence and presence of one or more double bonds (Kralovec, Zhang, Zhang, & Barrow, 2012). For example, oleic acid (OA) contains one double bond at the ninth position from the methyl terminus and docosahexaenoic acid (DHA) contains six double bonds with the first double bond occurring at the third carbon atom from the methyl terminus. Therefore, OA and DHA are known as u-9 and u-3, respectively. The composition of triglycerides is different in different fats and oils. The triglyceride containing single type of fatty acid is known as monoacid triglyceride while the one containing different types of fatty acids is known as mixed triglycerides. A mixed triglyceride can exhibit different stereoisomeric forms depending on the type and position of fatty acid [a (outer) (sn-1 and sn-3) and b (middle) (sn2) position] in the glycerol molecule. EPA and DHA in fish oil are generally found in the sn-2 position, whereas they are present in the sn-1 or sn-3 positions in case of oils derived from marine mammals such as whales and seals (Genot, Meynier, BernoudHubac, & Michalski, 2016; Wan, 2000). Thus positional distribution of fatty acids in the triglyceride backbone not only provides information about the origin of the oil but also important information about the stability and bioavailability of the oil. For example, fatty acids located at sn-2 position are reported to be more resistant to autoxidation and exhibit superior bioavailability, compared to those distributed in sn-1 or sn-3 position (Bandarra et al., 2016; Wang et al., 2015). It is also reported that saturated fatty acids and monounsaturated fatty acids are usually present in

Fig. 2. (A): Structure of a triacylglycerol. R, R0 and R00 are the positions at which fatty acids are linked (Adapted from Christie, 1986).

sn-1 position (Lei et al., 2012). Plant seeds such as flax, chia, hemp, and perilla are rich in PUFAs (Table 1). The proportion of PUFAs in chia seed oil is higher than others listed in this table. The relative content of a-linolenic acid (>60%) and linoleic acid (~20%) is also superior to these oils. A comparison of the fatty acid profile of different plant seed oils (Table 1) indicates that chia seed oil provides a healthy balance of two important essential fatty acids, i.e., alpha-linolenic acid and linoleic acid. The u-3 to u-6 fatty acid ratio of chia oils (3.18e4.18)
was also found to be higher than many vegetable oils (Alvarez
vez, Valdivia-Lo
pez, Aburto-Juarez, & Tecante, 2008). Chia Cha seed oil was found to contain twelve species of triacylglycerols which comprise of more than 90% of the oil. Trilinolenin was found to be the prevalent triacylglycerol and a-linolenic acid was found to present in most triacylglycerols of chia seed oil (Ixtaina et al., 2011; Timilsena, Vongsvivut, Adhikari, & Adhikari, 2017). 3. Complex coacervation process Coacervation is a term used in colloidal chemistry to denote the associative phase separation induced by altering the pH, ionic strength, temperature or solubility of the dissolving medium. If the coacervation is induced in a single polymer system, it is known as simple coacervation, while the one taking place between two or more polymers is regarded as complex coacervation process. In complex coacervation process, electrostatic interaction takes place between two oppositely charged polymer molecules and finally leads to phase separation. In many cases, the interaction between protein and polysaccharide is used to produce complex coacervates for food applications (Schmitt & Turgeon, 2011). Although electrostatic force between oppositely charged macromolecules is the main driving force, van der Walls intermolecular forces and hydrophobic interactions in proteins also affect the complex coacervation process (Turgeon, Schmitt, & Sanchez, 2007). The coacervation process was systematically studied first by de Jong using gelatin and gum Arabic as reacting polymers (de Jong, 1949). Since then, a number of studies have been carried out to better elucidate the complex coacervation process. However, the mechanism of interaction between two biopolymers which leads to the formation of the complex coacervates is so intricate and complex that it is not yet adequately understood. It has been established that both the material properties such as molecular weight of polymers, their charge density, concentration and mixing ratio and process parameters such as method used for producing emulsion/ degree of homogenisation, pH, temperature, and ionic strength affect the complex coacervation process to a considerable extent (Weinbreck, De Vries, Schrooyen, & De Kruif, 2003). Therefore, the optimum condition for complex coacervation for one set of polymers cannot be simply used for another set of polymers. Because of this reason, the optimization of the coacervation parameters is the essential priori step for its successful application in microencapsulation. Yield of complex coacervates as a function of dispersant pH and polymer ratio is the most commonly used parameter for determining optimum condition for complex coacervation between two polymers. This is because the coacervate yield can be easily determined by separating the precipitated complex coacervates from the aqueous phase by centrifugation. Surface charge density (zeta potential) and turbidity as a function of pH and polymer ratio are recently and effectively used to optimise the complex coacervation conditions between two biopolymers (Eratte, Wang, Dowling, Barrow, & Adhikari, 2014; Timilsena, Wang, Adhikari, & Adhikari, 2016; Timilsena, Adhikari, Barrow, & Adhikari, 2016; Kaushik, Dowling, Barrow, & Adhikari, 2015; Wang et al., 2014). Surface electrostatic charge is the fundamental property of polymers. It is

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373

Table 1 Monounsaturated (MUFAs) and polyunsaturated fatty acids (PUFAs) content of plant oils.a Fatty acids

Chia

Flax

Hemp

Olive

Sunflower Sesame

Rapeseed Canola

C16:1 (u-7) C18:1 (u-9) C18:2 (u-6) C18:3 (u-3) C20:1 (u-9) C20:2 Trans-fats MUFAs PUFAs u-3 u-6 u-6: u-3 Reference

0.2

0.09

0.11

1.8

0.12

0.11

0.21

10.5

18.1

11.5

66.4

28.0

41.5

20.4

15.3

59.4

16.4

62.2

60.0

58.2

0.36

1.6

0.2

0.2

16.5

0.07 e 11.0 80.4 e e 0.35 Ciftci, Przybylski,
ska, 2012 & Rudzin

0.05 e 18.5 73.6 e e 0.27 Ciftci et al., 2012

e e 28.1 62.8 0.4 62.4 156.0 Orsavova et al., 2015

a

Walnut

Coconut

Perilla

Peanut

e

e

e

0.07

0.07

63.3

59.5

27.3

6.2

16.2

71.1

40.9

19.6

18.8

50.0

1.6

17.9

18.2

0.16

0.21

1.2

11.9

15.0

e

60.4

e

0.30

0.18

0.32

9.1

e

e

e

e

e

e e 68.2 18.0 1.6 16.4 10.3 Orsavova et al., 2015

e e 28.3 62.4 0.4 62.2 155.5 Orsavova et al., 2015

e e 42.0 41.2 0.2 40.9 204.5 Orsavova et al., 2015

e 0.14 72.8 20.9 1.2 19.6 16.3 Orsavova et al., 2015

e e 59.5 30.7 e e e Kostik, Memeti, & Bauer, 2013

e e 27.3 65.0 e e e Dogan & Akgul (2005)

e e 6.2 1.6 0.0 1.6 e Orsavova et al., 2015

0.05 e 16.6 75.9 e e 0.22 Osakabe et al., 2002; Ciftci et al., 2012

e e 71.1 18.2 0.0 18.2 e Orsavova et al., 2015

Data are expressed as percentages of total fatty acid methyl esters (FAMEs).

determined by measuring electrophoretic mobility of biopolymer molecules in a dispersant at a given temperature using combined Doppler electrophoresis and phase analysis light scattering (Amin, Rega, & Jankevics, 2012). In general, gums are negatively charged in a wide range of pH while proteins can be positively or negatively charged, depending on the pH of the medium. The optimum complexation is usually observed at a pH value where the polymers have opposite charges and the magnitude of charge difference between the polymers is the highest (Liu, Elmer, Low, Nickerson, 2010; Liu, Low Nickerson, 2010). Likewise, turbidity can be determined by measuring the optical density or transmittance of a polymer-solvent mixture using UVeVis spectrophotometer. When the complex coacervate particles (micro coacervates) start forming, the turbidity of initially clear solution starts to increase. Once the coacervation occurs to its highest degree the complex coacervates start to coalescence giving rise to a visibly insoluble coacervate phase; hence two phase system is formed. This phase separation stage of complex coacervation is known as macro-coacervation. The denser phase or polymer-rich portion is known as coacervate phase and the lighter polymer deficient portion is called equilibrium phase (Nairm, 1995) (Fig. 3 A). Complex coacervation is a reversible process and the complex coacervates formed by the interaction of a protein and a polysaccharide can dissociate when the conditions, especially the pH and temperature are unfavourable. Complex coacervates are shelfstable within a narrow range of pH, ionic strength and temperature. Therefore, careful monitoring of the process is essential to arrive at optimum complex coacervation conditions and to produce stable complex coacervates. Many complex coacervates, especially the gelatin-based complex coacervates require further consolidation when they are intended to be used as microencapsulating shell materials (Timilsena, Wang, et al., 2016; Wang et al., 2014). Hence, complex coacervates are stabilised by either cross-linking or by the application of mild heat (Kamyshny & Magdassi, 2006). A number of synthetic and natural materials, including glutaraldehyde, tannic acid, gallic acid and transglutaminase (Table 2), are used for crosslinking the complex coacervates. Glutaraldehyde forms covalent binding between amino groups. This type of cross-linking is irreversible and the mechanical integrity of the cross-linked shell is enhanced in such a way that it can resist the effect of pH, temperature and mechanical shear to a greater extent (Mwangi &

Ofner, 2004). However, due to toxic nature of glutaraldehyde, it is not considered a suitable cross-linking agent for food application. Therefore, natural substances such as polyphenols and transglutaminase enzyme find increased application in food industries € ny-Meyer, 2013). The polyphenols (Heck, Faccio, Richter, & Tho could form both covalent and non-covalent complexes with proteins (e.g. zein), and their structural and functional properties depended on the nature of the complexes formed (Liu, Ma, McClements, & Gao, 2016). On the other hand, transglutaminase catalyzes the formation of protein networks by introducing glutamyl-lysyl isopeptide bonds between target proteins (Fig. 4). 4. Microencapsulation of PUFAs-rich oil using complex coacervation Microencapsulation of PUFAs-rich oils by simple emulsification and spray drying (Shaw, McClements, & Decker, 2007), complexation with proteins and polysaccharides (Augustin & Hemar, 2009) and complex coacervation followed by spray drying (Eratte et al., 2014; Timilsena, Adhikari, et al., 2016) has successfully been carried out. This process encapsulates the oils and fats in solid (powder) matrix. These solid forms of microcapsules can readily be incorporated into other food and pharmaceutical formulations and make the fortification process more effective. In general, complex coacervation-based microencapsulation process consists of four steps. Firstly, aqueous solution of two polymers is prepared. Protein solution is prepared at a temperature beyond its gelling temperature and pH above its isoelectric point. Secondly, hydrophobic core (oil) is blended with the protein solution first and the mixture is homogenised in order to make fine oilin-water (o/w) emulsion. Thirdly, aqueous solution of the other biopolymer (polysaccharide, in general) is mixed with the o/w emulsion. If two biopolymers have optimum density of opposite charges, complex coacervation occurs instantly and the complex coacervates form a layer surrounding the oil droplets. If the biopolymers do not have satisfactory level of opposite charge density, adjustment of pH or temperature is required to induce opposite charge density in order to initiate the formation of complex coacervate. These complex coacervates are usually insoluble in water. Adsorption or deposition of the insoluble coacervates on the surface of the oil droplets leads to precipitation (phase separation) of

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Fig. 3. Complex coacervation and phase separation process. (A): protein-protein complex coacervation (BSA: Bovine serum albumin, b-Lg: b-lactoglobulin and GB: gelatin B) [Adapted from Pathak, Rawat, Aswal, & Bohidar, (2015), (B): Precipitation of the complex coacervate microcapsules of chia seed oil demonstrating distinct two phases [Adapted from Timilsena, Adhikari, et al. (2016)].

Table 2 Cross-linkers used to consolidate the gelatin-based complex coacervates. Active ingredient

Cross linker

MEE (%)

References

Turmeric oleoresin Peppermint oil Peppermint oil Allyl isocyanate Ascorbic acid Garlic oil Flaxseed oil Vitamin A Aspartame Microalgal oil Jasmine essential oil Fish oil Orange oil

e FA or TG TA TA e FA e FA e TG TG e

49e73 90 82 84 97e99 63e99 e 64e83 e e e 53e77

Zuanon, Malacrida, and Telis (2013) Dong et al. (2011) Pakzad, Alemzadeh, and Kazemi (2013) Zhang, Pan, and Chung (2011) Comunian et al. (2013) Siow and Ong (2013) Liu, Elmer, et al. (2010) and Liu, Low, et al. (2010) Junyaprasert, Mitrevej, Sinchaipanid, Boonme, and Wurster (2001) Rocha-Selmi, Bozza, Thomazini, Bolini, and F
avaro-Trindade (2013) Zhang, Zhang, Hu, Bao, and Huang (2012) Lv, Yang, Li, Zhang, and Abbas (2014) Habibi, Keramat, Hojjatoleslamy, and Tamjidi (2016) Dong, Zhao, Sun, and Li (2013)

Abbreviations: MEE: Microencapsulation efficiency, FA: Formaldehyde, TG: Transglutaminase, GA: Glutaraldehyde, TA: Tannic acid.

Fig. 4. Protein crosslinking through transamidation reactions catalyzed by transglutaminase [Adapted from Heck et al. (2013)].

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375

the microcapsules. Finally, this complex coacervate shell is hardened or consolidated, if required, by heating, desolvation, or crosslinking in order to improve the mechanical and thermal stability of the microcapsules (Timilsena, Wang, et al., 2016; Turgeon et al., 2007). A microencapsulation technology can be considered successful when it yields relatively higher encapsulation efficiency (MEE) and microencapsulation yield (MEY) with enhanced oxidative stability of the core and comparatively very low surface oil. Among these important parameters, MEE and MEY are calculated using Equations (1) and (2).

MEE ¼

Wt
Ws  100 % Wt

(1)

MEY ¼

Wt Wi

(2)

 100 %

where, MEE is microencapsulation efficiency (%), MEY is yield of the microcapsules (%), Wt and Ws are the mass (g) of total oil and surface oil of the microcapsules, Wi is the mass (g) of oil introduced into microcapsule and Wm is the mass (g) of the microcapsules taken. Lipid oxidation produces hydroperoxides, ketones and alcohols are oxidation products. Therefore, the efficacy of microencapsulation is measured as delaying the onset of oxidation. Oxidative stability is usually measured at elevated temperature using Rancimat® or Oxypress®. In Rancimat® test, stability is measured as the time required to attain the inflection point of the conductivity curve indicated by the abrupt increase in the slope of the curve and is used as the oxidative stability index (OSI) (Timilsena, Adhikari, et al., 2016). In ambient temperature, oxidative stability is measured in terms of peroxide value, p-Anisidine value or Totox value (Totox ¼ p
AVþ2(PV)). OSI is dependent on various factors such as microcapsule shape, size, porosity and the surface oil of the microcapsule. It has been found that higher the surface oil, lower is the OSI. It is because the surface oil gets rapidly oxidised as it comes into the contact with air. In general, microcapsules prepared using complex coacervates as shell can be either mononucleated or multinucleated. The size and morphology of microcapsules can be highly affected by the processing conditions. Wang et al. (2014) demonstrated that microcapsules produced through the complex coacervation between gelatin and sodium hexametaphosphate possess a core-shell structure in which each droplet of oil is covered by the thin layer of shell and multiple core particles are surrounded by a second layer of shell producing a highly stable microcapsules of tuna oil (Fig. 5). Barrow & Shahidi (2007) reported that complex coacervation is the most effective methods of microencapsulation of u-3 fatty acids and delivering them into foods. Timilsena, Adhikari, et al. (2016) demonstrated that microcapsules obtained by complex coacervation followed by spray drying possess higher encapsulation efficiency and increased oxidative stability than the microcapsules obtained by simple emulsification followed by spray drying. It suggests that complex coacervation-based microencapsulation better protects PUFAs-rich oils than a simple spray drying. Higher encapsulation efficiency in complex coacervation-based microencapsulation is resulted from the better surface active nature of the complex coacervates. It has been reported that complex coacervates migrate rapidly to the surface of the lipid droplets during the emulsification stage and form a uniform layer around the oil droplets (Benichou, Aserin, & Garti, 2002). These coacervates can be converted into powder form either by freeze drying or spray drying. The powder form of microcapsules produced using complex

Fig. 5. Multicore complex coacervate microparticles of tuna oil encapsulated into gelatin-sodium hexametaphosphate complex coacervates (Adapted from Wang et al., 2014).

coacervates as shell materials have significantly higher encapsulation efficiency and longer storage life compared to the solid microcapsules produced using either individual protein or gum as shell material (Timilsena, Adhikari, et al., 2016). In most cases complex coacervation-based microencapsulation produces multinuclear microcapsules surrounded by double layers of shell materials, thus making it less porous and more robust. As a consequence, it offers higher oxidative stability to PUFAs-rich oils than simple emulsification and drying process. 5. Recent advances in complex coacervation Although the complex coacervation between various proteins and opposite charged biopolymers such as polyphosphates and polysaccharides has been well reported, gelatin is probably so far the only commercialised protein for complex coacervation purpose (Table 2) due to its excellent functional properties such as solubility, emulsifying and gelling capability. At Ocean Nutrition Canada (now DSM), commercial omega-3 oil powders with 60% oil loading, MEG3®, are currently produced with gelatin-sodium hexametaphosphate complex coacervates as the encapsulant (Yan & Zhang, 2014, chap. 12). In Wang et al. (2014)’s work, the preparation of this MEG3® encapsulation matrix was systemically investigated and optimised. The resulted fish oil microcapsule exhibited high oil loading (approx. 60%), low surface oil (<0.01%) and significantly improved oxidative stability. As a result, MEG-3® has been successfully incorporated into various food products including bakery, health supplements and infant formula. However, gelatin is relatively expensive and cannot be consumed by the increasing vegetarian and vegan population. Consequently, there is an increasing trend of plant protein use in the food industry, partly driven by their lower tendency to stimulate an allergic response compared to animalderived alternatives (Choi et al., 2010). Hence, novel encapsulation matrices based on complex coacervation technique require to be developed. The development of alternative proteinpolysaccharide complex coacervates requires rigorous optimization of the complex coacervation parameters because each pair of protein and polysaccharide form stable complex coacervates at specified pHs and biopolymer mixing ratios and the optimised conditions for one set of biopolymers cannot be replicated to another set of biopolymers. Considerable research has been carried out in recent past to produce complex coacervates using milk and plant proteins such as whey, soy and pea proteins by optimizing

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coacervation parameters (Table 3). These complex coacervates have been utilised to microencapsulate u-3 oils. However, their stability and bioavailability of encapsulated oils have yet to be adequately understood. This area of research is currently receiving increased attention. Soybean protein and pea protein isolates are two of the most commonly used plant proteins as encapsulating shell materials (Ducel et al., 2004; Elmer, Karaca, Low, & Nickerson, 2011; Gai, Li, & Jiang, 2012; Jun-xia, Hai-yan, & Jian, 2011; Klemmer, Waldner, Stone, Low, & Nickerson, 2012; de Conto, Grosso, & Gonçalves, 2013). Recently complex coacervates of lentil protein isolate (Aryee & Nickerson, 2012; Chang, Gupta, Timilsena, & Adhikari, 2016; Chang, Varankovich, & Nickerson, 2016) and canola protein isolate (Chang, Gupta, Timilsena, & Adhikari, 2016; Chang, Varankovich, & Nickerson, 2016) with chitosan or gum Arabic were developed and used to encapsulate oils and other unstable and bioactive food ingredients (Table 4). These studies used protein from one plant and polysaccharide from another to produce complex coacervates. There is a paucity of information on the formation of complex coacervates between proteins and gums from the same plant source. Until now, there are only two studies which report the science of formation of complex coacervates between protein and gum of same plant species and subsequent application of these unique complex coacervates in microencapsulation of PUFAs-rich oils from the same plant (Kaushik et al., 2015, 2016; Timilsena, Wang, et al., 2016; Timilsena, Adhikari, et al., 2016). Kaushik et al. (2015; 2016) studied the complex coacervation between flaxseed protein isolate (FPI) and flaxseed gum (FSG) and used such FPI-FSG complex coacervates as shell material to encapsulate PUFAs-rich flaxseed oil (FSO). Similarly, Timilsena, Wang, et al. (2016) and Timilsena, Adhikari, et al. (2016) investigated the complex coacervation formation between chia seed protein isolate (CPI) and chia seed gum (CSG). The authors also used these CPI-CSG complex coacervates to microencapsulate PUFAs-rich chia seed oil. It can be expected that this types of complex coacervation and microencapsulation system will provide better core-shell compatibility. In both studies, the authors compared the effect of different core-toshell ratio and different drying methods in the microencapsulation efficiency and oxidative stability of the PUFAs-rich oils (Table 4). It was shown that the microencapsulated CSO was at least 6 times more stable against oxidation than the unencapsulated one

under same test conditions. Similarly, the microencapsulated FSO was about 9 times more stable compared to the unencapsulated one under same conditions. These two studies showed that plant protein-gum complex coacervates can be effectively used to encapsulate and stabilize PUFAs-rich oils. Characteristics of shell and core material are important factors determining encapsulation efficiency, core stability and controlled release of the core. The bioavailability and bioefficiency of active substances also depend on the encapsulation and delivery system. The design of an effective encapsulation system requires an understanding of mechanisms of encapsulation, chemical composition of shell matrix and possible interaction of the active compounds with the shell matrix (Augustin & Hemar, 2009). Bioavailability refers to the fraction of an ingested nutrient that gets released at specified spots of alimentary canal and becomes available for absorption. As the digestion process is affected by the nature of the food structure, the design of a suitable food matrix is important to ascertain bioavailability of the nutrient. In order to maximize the benefits of PUFAs fundamental understanding of their release from microcapsule during their passage through gastrointestinal tract and subsequent absorption is vitally important.

6. Physiological digestion of encapsulated oils and fats Majority of hitherto lipid digestion research (Mun, Decker, & McClements, 2007; Hur, Decker, & McClements, 2009; Sarkar, Goh, Singh, & Singh, 2009; Sarkar, Horne, & Singh, 2010) has focussed studying the digestion of emulsified oil using a single or multi-compartment in vitro digestion model. These studies were aimed at understanding the extent and nature of digestion of lipids in each stage of the gastro-intestinal tract. Most of the reported studies deal with the digestion of oil-in-water emulsion systems. In these emulsion systems, both the rate of lipolysis and their in vivo absorption are affected largely by the size of emulsion droplets and the effective area available for lipase adsorption (Golding et al., 2011). However, there are limited number of studies which investigated the impact of individual microencapsulation technologies on the digestibility and bioavailability of encapsulated oils and fats (Augustin et al., 2014; Chung, Sanguansri, & Augustin, 2011). These previous studies have been discussed in this section.

Table 3 Optimum pH and biopolymer ratio for the formation of complex coacervates between some biopolymers. Biopolymers

pH range

Cationic biopolymers

Anionic biopolymers

WPI SPI Chitosan Gelatin Gelatin Chitosan SPI LPI PPI PPI PPI Alpha gliadins Pea globulins BSA Gelatin b-lactoglobulin Gelatin Gum Arabic

Gum Arabic Gum Arabic Gum Arabic CMC Chitosan CMC Pectin Gum Arabic Gum Arabic Chitosan Alginate Gum Arabic Gum Arabic Pectin Alginate Gum Arabic Pectin Chitosan

3.0e4.5 2.5e4.5 2.0e4.0 9.0e11.0 4.5e6.5 3.0e5.0 e 1.5e8.0 2.4e4.3 e 1.55e2.98 2.0e4.0 2.0e4.0 1.6e4.7 2.0e5.0 3.5e4.4 4.0e5.5

Optimum conditions pH

Biopolymer ratio

4.0 4.0 3.6 9.0 5.25e5.50 4.0 4.4 3.5 3.6 6.2 2.10 2.75 3.0 4.7 3.5e3.8 3.76 3.8 e

2:1 1:1 4:1 1:1 10:1e20:1 1:1 1:1 2:1 7.5:1 4:1 3:7 1:1 5:1 3.5:1 2:1 1:1 5.5:1

References

Weinbreck, Minor, and De Kruif (2004) Jun-xia et al. (2011) Butstraen and Salaün (2014) Lii, Tomasik, Zaleska, Liaw, and Lai (2002) Remunan-Lopez and Bodmeier (1996) Tiyaboonchai & Ritthidej (2003) Mendanha et al. (2009) Aryee and Nickerson (2012) Liu, Elmer, et al. (2010) and Liu, Low, et al. (2010) Elmer et al. (2011) Klemmer et al. (2012) Ducel et al. (2004) Ducel et al. (2004) Ru, Wang, Lee, Ding, and Huang (2012) Shinde and Nagarsenker (2009) Sanchez, Mekhloufi, and Renard (2006) Liu, Low, and Nickerson (2009) Espinosa-Andrews et al. (2013)

Note: CMC ¼ Carboxymethyl cellulose, SPI ¼ Soy protein isolate, PPI ¼ Pea protein isolate, BSA ¼ Bovine serum albumin, WPI ¼ Whey protein isolate, LPI ¼ Lentil protein isolate.

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Table 4 Microencapsulation of bioactive ingredients through complex coacervation of non-gelatin proteins and polysaccharides. Polymer combinations

Core

Opt pH

Biopolymer ratio

MEE (%)

OSI (h)

Drying method

References

WPI/GA SPI/GA SPI/Pectin SPI/Pectin CPI/CSG

Tuna oil Sweet orange oil Casein hydrolysate Propolis extract Chia seed oil

3.75 4.0 4.4 4.0 2.7

3:1 1:1 1:1 e 6:1

73 65e78 79e92 66e72 93.6

22.1 e e e 12.3

Spray & freeze drying Spray drying Freeze drying Freeze drying Spray & freeze drying

FPI/FSG

Flaxseed oi

3.1

3:1

87.6

8.8

Spray & freeze drying

Eratte et al. (2014) Jun-xia et al. (2011) Mendanha et al. (2009) Nori et al. (2011) Timilsena, Wang, et al. (2016) and Timilsena, Adhikari, et al. (2016) Kaushik et al. (2015, 2016)

Abbreviations: MEE: Microencapsulation efficiency, OSI ¼ oxidative stability index, SPI: Soy protein isolate, GA: Gum Arabic, PPI: Pea protein isolate, GG: Guar gum, CPI¼ Chia seed protein isolate, CSG ¼ Chia seed gum, FPI ¼ Flaxseed protein isolate, FSG ¼ Flaxseed gum.

Ilyasoglu and El (2014) studied the bioavailability of EPA and DHA from sodium caseinateegum Arabic complex nanocapsules and found that only 56.2% of EPA and 47.4% of DHA became available after digestion of fish oil. These authors suggested that the oil encapsulated in double-layered complex coacervate based nanocapsules was only partially released and its digestion was greatly reduced. A significant portion of oil from these nanocapsules was passed to the large intestine without digestion. Shen, Apriani, Weerakkody, Sanguansri, and Augustin (2011) also reported that only 78.6% of oil encapsulated into a composite matrix comprising sodium caseinate, glucose and HylonVII and incorporated into orange juice was digested, whereas extent of digestion was reduced to 64% when it was incorporated into a cereal bar. This study provided a conclusive evidence that nature of food matrix influences the lipolysis process and that the presence of dietary fibres slows down the lipolysis of the encapsulated oil. Patten, Augustin, Sanguansri, Head, and Abeywardena (2009) studied in vivo digestibility of spray dried fish oil powders in mice and reported that only 4e6% of the oil was released from the microcapsules and small quantity of low chain PUFAs were present in the plasma indicating that large quantity of oil reached the bowel without being digested in the small intestine. Augustin et al. (2014) compared the digestion behaviour of 12 types of spray dried powders of canola oil-in-water emulsion and reported that lipolysis of canola oil in these powders varied between 12 and 68%. This study suggested that in vitro digestibility of oil encapsulated in powders depends on both formulation and the processing steps used in their manufacture. Then, these authors selected 5 different formulations of oil-containing microcapsule powders and incorporated them into a dairy beverage intended for in vivo human trial. Free or unencapsulated oil incorporated into the same beverage was used as the control. They found that triglyceride levels in blood of the subjects fed with the microencapsulated oil was higher than the subjects fed with unencapsulated oil. Most in vitro digestion studies of lipids have focussed on quantifying the release of the oil and the production of free fatty acids as an indicator of the lipolysis. The effect of non-lipid ingredients in slowing down the digestion of lipids has not been studied and explained to date. Comparison or validation of data obtained from in vitro studies with in vivo experiments is another challenge that has not been covered so far. Timilsena, Adhikari, Barrow, & Adhikari (2017) and Timilsena, Vongsvivut, et al. (2017) reported that pattern and kinetics of digestion of unencapsulated oils is quite different than the digestion of the encapsulated oil. These authors also showed that digestion of encapsulated oil is influenced by the nature of the shell materials in which they are encapsulated. Protein-based matrices get readily digested in the stomach releasing the oil from the matrix, whereas gum-based matrices are not digestible but get solubilised in the stomach and intestinal conditions. On the other hand, matrices prepared from the protein-gum complexation are more

resistant to digestion and thus less oil is released in the gastric stage allowing more oil in the intact form to reach the intestinal stage. These authors also argued that main reason for slower digestion of encapsulated oil is because the oils/fats must be released from the encapsulating matrix before it is acted upon by the enzyme (lipase). In addition, these shell materials affect the interfacial behaviour and interfere with the diffusion and adsorption of lipase to the oil droplets. Presence of protein, fibres and polysaccharides in digestion mixture interfered lipase activity which reduced the rate of digestion of lipid by pancreatic lipase (Mun et al., 2007). All the aforementioned studies suggest that physicochemical properties of microcapsules, such as their size, interfacial characteristics and degree of crystallization of lipid significantly affect the rate and extent of lipolysis in the gastro-intestinal tract (Augustin et al., 2014). Although a portion of the oil gets released from the microcapsules in the oral stage, it passes to the stomach without digestion. In human beings the actual digestion of lipids starts at stomach due to the action of gastric lipase. Gastric lipase being active only at a pH range between 4.0 and 6.0, it remains inactive at pH values of the normal conditions of stomach, where the pH normally lies between 1.5 and 3.0. However, the pH of human stomach increases upto the optimum value for gastric lipase due to buffering effect of foods. It has been reported that only 10e30% of lipids is digested in the gastric phase and produces a mixture of free fatty acids, diacylglycerols (DAGs) and monoacylglycerols (MAGs) (Bauer, Jakob, & Mosenthin, 2005; Gallier & Singh, 2012). After the completion of the gastric digestion, the oil along with other food (the partially digested food is known as chyme) enters into the small intestine where it is mixed with enzymes (pancreatic lipase, a-amylase and proteases-trypsin and chymotrypsin), salts (bicarbonates and bile salts) and electrolytes (sodium and calcium). The acids present in the chyme is neutralised by bicarbonates so that the pH of the intestinal content generally reach up to 7.5. Majority of the lipids (70e90% of the lipids) present in foods get digested in the small intestine by the action of pancreatic lipases (Wickham, Wilde, & Fillery-Travis, 2002). The ultimate products of lipid digestion are monoglycerides and fatty acids (Fig. 6). Lipid digestion and absorption is a remarkably efficient process with more than 95% efficiency and there is no feed-back regulation to reduce its assimilation (Patton et al., 1985). The products of lipid digestion are absorbed through the epithelial cells and carried into the blood stream (Seidel & Long, 2006). During lipid digestion, it is very important that oil droplets remain emulsified in the stomach and intestinal fluids so that they become accessible to the enzymes (e.g. gastric and pancreatic lipases). Emulsification is very essential in order to bring the hydrophobic (lipid) and hydrophilic (lipase) components together. The water soluble lipase has to be able to attach at the lipid (oil)water interface for lipolysis to occur. Therefore, any factor that affects the migration of lipases to oil-water interface also affects the

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Fig. 6. Breakdown of triacylglycerols (TAGs) into a monoacylglycerol (MAG) and two free fatty acids (FFAs) molecules.

lipid digestion process (Singh & Gallier, 2014). Emulsification in the stomach leads to the organization of dietary lipids in the form of droplets in the aqueous digestive system (Bauer et al., 2005). Size of the emulsified lipid droplets is also very important factor affecting lipid digestion. The smaller the droplet, the larger is the surface area available for lipase adsorption and faster is the lipid digestion. The nature of emulsifier and the molecular structure of triacylglycerol molecules such as the chain length and degree of saturation of the fatty acids and their position on the triacylglycerol backbone also influence the digestion of lipids (Gallier & Singh, 2012; Singh & Gallier, 2014). Lipid digestion in isolated conditions is quite different than that in the presence of non-lipid components such as proteins, polysaccharides and fibres. It has been reported that there is large influence of these non-lipid components in the degree and the kinetics of lipid digestion and bioavailability (Augustin et al., 2014; Singh et al., 2009). Better understanding of the mechanisms of lipid digestion and factors affecting its degree and kinetics will enable better design of encapsulating shell materials so that release and physiological digestion of lipids can be controlled according to bodily requirement, eventually reducing the risk of diseases that result from higher intake of fats (Bornhorst & Paul Singh, 2014; Sarkar et al., 2010; Singh et al., 2009; Zhang, Zhang, Zhang, Decker, & McClements, 2015; Zhu, Ye, Verrier, & Singh, 2013). In practice, food digestion studies are carried out using a relatively simpler in vitro models and more complex in vivo animal and human trials. During in vitro experiments, food is dispersed in simulated digestive fluids and mechanical movement and the digestion of targeted substance over time is measured. Although it is very difficult to mimic the complexity of real human digestion system, nevertheless these in vitro models provide insights into the fundamental mechanisms of detachment of the bioactive food ingredients from the complex food matrix and their subsequent hydrolysis in the gastro-intestinal environment (Chung et al., 2011; Mackie, 2012, pp. 49e70). Besides, these in vitro models allow independent study of many factors which cannot be done in in vivo. Furthermore, in vitro studies do not require stringent ethical approval; thus they can be carried out at much reduced cost (Kong & Singh, 2009). Due to these reasons, in vitro experiments are still popularly used in digestion studies before proceeding to the fullfledged in vitro trials. Ultimately, the results obtained from in vitro models have to be validated using data obtained from in vivo animal and/or human trials (Bornhorst & Paul Singh, 2014).

oxidative deterioration unless any protective measures are applied. Oxidation of PUFAs-rich oils not only lowers their nutritional value and sensory attributes but also generates toxic substances. Therefore, it is essential to innovate and adopt a suitable microencapsulation technology to maximize the health benefits of PUFAs. In general, they are encapsulated into edible and oxygen resistant food matrices. For this purpose, complex coacervation based microencapsulation is more preferable because of substantially high microencapsulation efficiency and increased oxidative stability achievable in this method even at high payload. Although fish oil is rich in PUFAs, the demand for plant based omega-3 rich oils is increasing due to the healthy image of the latter. Similarly, despite high effectiveness of gelatin to encapsulate PUFAs-rich oils, there is continued search for its alternative, preferably a plant protein due to the demand from vegetarian and vegan population. Recent trends are to utilize plant protein-gum complex coacervates in stabilization of PUFAs-rich oils and other sensitive bioactive food ingredients. Microencapsulation of chia seed oil and flaxseed oil using protein-gum complex coacervates produced from the same plant species has shown better compatibility between the core and the shell. Controlled release of PUFAs and their subsequent digestion in the gastro-intestinal tract are important consideration for successful design of PUFAs-rich microcapsules. However, there is acutely limited information on the digestion of microencapsulated oils. In vitro studies followed by in vivo trials would more accurately evaluate the effectiveness of microcapsules and microencapsulation technologies applied to PUFAs in protecting them from degradation and controlling their bioavailability in the body. Co-encapsulation of two or more bioactive food ingredients in a single matrix is a recently introduced concept in food industries. Wang et al. (2015) reported a successful co-encapsulation technology of omega-3 oil with vitamin A, D3, E, K2, curcumin and coenzyme Q10 in a gelatin-sodium hexametaphosphate complex coacervate. Eratte et al. (2015) also reported co-encapsulation of omega-3 rich oil with probiotic bacteria in whey protein isolategum Arabic complex coacervate shells. In both these studies, the authors used animal derived proteins. Replacing omega-3 rich fish oils with plant oils and replacing animal-based proteins (gelatin, whey protein) with plant proteins and application of the coencapsulation concept to more than one active functional ingredients can confer multiple benefits and could be important future research direction.

Acknowledgement 7. Concluding remarks and future perspectives Although PUFAs-rich oils contribute significantly in the health and wellbeing of consumers, they are highly susceptible to

The first author gratefully acknowledges the scholarship support from RMIT University, Melbourne, Australia and CSIRO, Australia.

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