lipid oxidation in oil-in-water emulsions

7/17/2019 Lipid Oxidation in Oil-In-Water Emulsions

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Lipid Oxidation in Oil-in-Water Emulsions:

Involvement of the Interfacial LayerClaire C. Berton-Carabin, Marie-Helene Ropers, and Claude Genot

Abstract: More polyunsaturated fats in processed foods and fewer additives are a huge demand of public health agencies

and consumers. Consequently, although foods have an enhanced tendency to oxidize, the usage of antioxidants, especially

synthetic antioxidants, is restrained. An alternate solution is to better control the localization of reactants inside the food

matrix to limit oxidation. This review establishes the state-of-the-art on lipid oxidation in oil-in-water (O/W) emulsions,

with an emphasis on the role of the interfacial region, a critical area in the system in that respect. We first provide a

summary on the essential basic knowledge regarding (i) the structure of O/W emulsions and interfaces and (ii) the general

mechanisms of lipid oxidation. Then, we discuss the factors involved in the development of lipid oxidation in O/W

emulsions with a special focus on the role played by the interfacial region. The multiple effects that can be attributed toemulsifiers according to their chemical structure and their location, and the interrelationships between the parameters that

define the physicochemistry and structure of emulsions are highlighted. This work sheds new light on the interpretation

of reported results that are sometimes ambiguous or contradictory.

Keywords: colloids, emulsion, food quality, lipid oxidation, microstructure

IntroductionTo address some of the current major health issues and fol-

low the most recent nutritional recommendations (Gebauer andothers 2006; EFSA Panel on Dietetic Products 2010), increasing

amounts of polyunsaturated fats are now often incorporated infoods (Ganesan and others 2014). At the same time, consumers

look for foods containing fewer additives, and among them thesynthetic antioxidants. This is why lipid oxidation in formulated

foods has become a renewed concern for manufacturers.Formulated foods often contain a lipid phase dispersed in an

aqueous medium. Thus, they can be schematically described asoil-in-water (O/W) emulsions. Such emulsions are stabilized by

surface-active molecules adsorbed at the oil–water interface. Fromtheir manufacture to their end-use, including their deconstruction

in the digestive tract, food emulsions are subjected to a broad rangeof physical–chemical treatments. Under these conditions, and in

the presence of oxygen, chemically reactive components may be-

come oxidized. Among them, polyunsaturated fatty acids (PUFAs)are particularly prone to oxidation. Lipid oxidation has a deleteri-

ous effect on the technological, sensory, and nutritional qualitiesof food (Pokorny 2003; Frankel 2005). The reaction generates

odorant compounds characterized by low detection thresholdsand generally unpleasant perceptions, which damages the sensory

quality of the products (Grosch 1982; Frankel 2005; Villiere and

MS 20140363 Submitted 5/3/2014, Accepted 12/5/2014. Author Berton-Carabin is with Food Process Engineering Group, Wageningen Univ., 6700 AA

Wageningen, The Netherlands. Authors Ropers and Genot are with INRA, UR1268 Biopolymeres Interactions Assemblages, F-44316 Nantes, France. Direct inquiries toauthor Genot (E-mail: [email protected] ).

others 2007). Lipid oxidation also causes a loss of components of

nutritional interest, and leads to the formation of free radicals andpotentially toxic compounds (Ursini and others 1998; Kanazawa

and others 2002; Riemersma 2002; Liu and others 2003; Petersen

and Doorn 2004; Turner and others 2006; Long and others 2008;Eder and Ringseis 2010; Serini and others 2011; Awada and others2012).

The strategies currently carried out by industry to counteractlipid oxidation include the use of vacuum-packaging or controlled

atmosphere exposure, low-temperature storage, encapsulation of sensitive added compounds, or addition of antioxidants. Despite

these efforts, the products containing PUFAs are still subject torisks of unpredicted development of oxidation at some stage in

their lifespan, which may impact the economic viability of theentire processing chain. This results from unknown or poorly

controlled factors, which stresses the need for better disseminationof the current knowledge and also a deeper understanding of the

oxidation reactions in complex food systems.As recently reviewed as a brief history of lipid oxidation by

Hammond and White (2011), the general mechanisms of lipidoxidation have been documented for decades and summed-up in

previous reviews (Frankel 1985, 2005; Porter 1986; Chan 1987;Gardner 1989; Hsieh and Kinsella 1989; Schaich 2005; Cheng

and Li 2007). Since the late 1980s, substantial work has fur-ther been performed with respect to lipid oxidation in emulsions

and, more generally, in multiphase systems. The main composi-tional and structural factors that influence lipid oxidation in emul-sions and food formulations have been deduced from studies per-

formed on more or less simplified model emulsions (Fritsch 1994;Coupland and McClements 1996; McClements and Decker 2000;

C 2014 Institute of Food Technologists®

doi: 10.1111/1541-4337.12097 Vol. 13, 2014 ComprehensiveReviewsinFoodScienceandFoodSafety 945



Lipid oxidation: an interface outlook . . .

Genot and others 2003, 2013; Frankel 2005; Jacobsen and others2008; Jacobsen 2010a, 2010b; Waraho and others 2011a). These

studies have notably emphasized that the interfacial region, whichis the contact region between the oil phase and the aqueous phase,

represents a particularly critical area in the system with regard tothe development of lipid oxidation. This assumption mostly relies

on the observed ability of factors such as oil droplet size (and there-fore interfacial area) or emulsifier type (and therefore interfacial

composition) to substantially affect lipid oxidation in emulsions.

Yet, many apparent contradictions regarding the role of the inter-face can be found in the related literature, which can be, at leastin part, attributed to the structural complexity of emulsion sys-

tems and to the complex behavior and partitioning of amphiphiliccompounds.

Based on an extensive review and analysis of the literature,including our own work, the present review proposes a critical

discussion of the available data about the influence of the inter-facial layer on lipid oxidation in O/W emulsions. The role of

antioxidants and associated chemistry is shortly reviewed here,as extensive information about the performance of antioxidants

in multiphase systems can already be found in the reviews of Jacobsen and others (2008), Laguerre and others (2007), Romsted

and Bravo-Diaz (2013), and Shahidi and Zhong (2011). Yet, wegive special attention to examples where the partitioning of an-tioxidants, and in particular, their natural or targeted localization

at the interface, affects lipid oxidation in emulsions.In the present work, we first summarize the essential basic

knowledge regarding (i) the structure of O/W emulsions and in-terfaces and (ii) the general mechanisms of lipid oxidation. Then,

we discuss the factors involved in the development of lipid oxida-tion in O/W emulsions with a special focus on the role played by

the interfacial region and the emulsifiers. Finally, we focus on theinterrelationships between relevant physical–chemical properties

of O/W emulsions, which shed new light on the interpretation of reported results that are sometimes ambiguous or contradictory.

Prerequisites on O/W Emulsions and InterfacesPhysical structure of emulsions

Many review articles and books describe the physical organi-zation of emulsions and, more generally, of multiphase systems.

Among them, we notably relied on previous works (Dickinson1992; McClements 2005).

Common food emulsions are constituted of 2 immiscible liq-uids, one being dispersed in the other in the form of droplets

(Figure 1). These droplets constitute the dispersed phase, whilethe liquid around them constitutes the continuous phase. The

border between the dispersed and continuous phase is the inter-face. A system constituted of oil droplets dispersed in an aqueous

phase is an O/W emulsion (for example, salad dressing, or mayon-

naise). Conversely, a system constituted of water droplets dispersedin an oil phase is a water-in-oil (W/O) emulsion (for example,butter or margarine). In the present work, we focus solely on

O/W emulsions.The droplet size distribution, often characterized by an average

diameter, determines the number of droplets and the total inter-facial area. The smaller the droplets, the higher their number and

the developed surface. A broad range of droplet sizes, from lessthan 0.1 to 20 µm or more can usually be encountered in food

emulsions.O/W emulsions are thermodynamically unstable systems. Thus,

oil droplets have to be stabilized physically to avoid the sponta-neous separation between the oil and aqueous phases. Stabilizing

agents include texture modifiers and emulsifiers. On the one hand,texture modifiers stabilize emulsions by increasing the viscosity of

the continuous phase. They limit the rising of the oil droplets(that leads to creaming) and their eventual flocculation and coa-

lescence. On the other hand, emulsifiers adsorb at the oil–water interface during the homogenization process, which decreases the

surface tension between oil and water and thus the total free energyof the system. The adsorbed emulsifiers can prevent the floccu-

lation and coalescence of oil droplets via steric or electrostatic

repulsions.

EmulsifiersEmulsifiers are generally classified into low-molecular-

weight emulsifiers (LMWEs), high-molecular-weight emulsi-fiers (HMWEs), and solid particles. They may be used either

alone (model emulsions), mixed together (the general case for food emulsions), or associated with polysaccharides (many food

products).

LMWEs. This category comprises surfactants, which are gener-

ally synthetic molecules, and also polar lipids from natural sources.They are small surface-active molecules constituted of a hy-

drophilic headgroup and a hydrophobic tail. The headgroup of

LMWEs may be anionic, cationic, zwitterionic, or nonionic. Thehydrophobic tail is generally constituted of 1 or several hydro-carbon chains with 10 to 20 carbon atoms. Several food-grade

surfactants exist, including mono- and diglycerides (monoacyland diacyl glycerols), Tweens (polyoxyethylene sorbitan esters),

polysorbates, Spans (sorbitan esters), sucrose esters, and citric, lac-tic, and acetic acid esters of mono- and diglycerides (Genot and

others 2013). In the category of polar lipids, lecithins are widelyused as food emulsifiers. Lecithins are generally complex mixtures

of polar lipids (phospholipids, glycolipids, and sphingolipids) andresidual triacyglycerols, and they can be obtained from various

sources. The most common phospholipids encountered in dietarylecithins and in foods are the zwitterionic phosphatidylcholine(PC) and phosphatidylethanolamine (PE).

HMWEs. High-molecular-weight amphiphilic biopolymers arealso widely used to stabilize O/W emulsions. In this category,

we distinguish proteins from hydrocolloids, the latter being repre-sented by gum arabic ( Acacia senegal ), modified starches, modified

celluloses, certain pectins, and galactomannans (Dickinson 2009).The interfacial activity of these biopolymers results from the pres-

ence of hydrophobic and hydrophilic domains distributed alongtheir polymeric chain. When amphiphilic biopolymers adsorb at

an oil–water interface, they adopt a conformation allowing thehydrophobic parts to locate in contact with the oil phase and the

hydrophilic parts to locate in the aqueous phase. Depending ontheir intrinsic characteristics, especially the presence of covalent

intramolecular bonds and their subsequent flexible or rigid struc-

ture, biopolymers adopt an extended or globular conformationat the interface. Biomolecule aggregates such as protein aggre-gates, often generated during food processes, can also adsorb at

the interface and participate in emulsion stabilization (Euston andHirst 1999; Chapleau and de Lamballerie-Anton 2003; Relkin

and others 2006; Mahmoudi and others 2011; Audebrand andothers 2013).

Solid particles. Oil droplets can be stabilized by particles thatadsorb at the interface (Pickering 1907; Aveyard and others 2003;

Leal-Calderon and Schmitt 2008; Dickinson 2010). In other stud-ies, silica particles were mixed with protein or surfactant emulsifiers

to stabilize emulsions (Pichot and others 2010; Kargar and oth-ers 2011). Recently, Gupta and Rousseau (2012) also developed

946 Comprehensive Reviews in Food Science and Food Safety Vol. 13, 2014 C 2014 Institute of Food Technologists®




Figure 1–Schematic representation (not to scale) of the main physical properties, compartments, and possible components of an oil-in-water (O/W)emulsion.

surfactant-based solid lipid nanoparticles that effectively kinetically

stabilized oil–water interfaces in O/W emulsions.

Physical properties of interfacesThe oil–water interfacial layer in emulsions is the thin layer

or boundary (a few nm) between oil and water. For relativelysmall droplets (for example, around 0.1 µm), the interfacial re-

gion comprises a significant volume of the total droplet (Mc-Clements and Decker 2000). Its characteristics depend to a great

extent on the type and concentration of the adsorbed molecules.LMWEs and HMWEs usually form a monolayer at the oil–

water interface. However, biopolymers such as caseins, whenpresent in sufficient amounts, can form multilayers at the interface

(Dickinson 1999). Multilayered interfaces can also be constructedby the consecutive electrostatic deposition of emulsifiers and

biopolymers (Guzey and McClements 2007; Grigoriev and Miller 2009).

When proteins are used as emulsifiers, only a part of the

polypeptidic chain is actually in direct contact with the oil–water interface, the other protein segments being located in the aqueousphase. Accordingly, only 30% to 40% of the interface is covered

with proteins, whereas almost 100% are covered with surfactants(Dickinson 1992). LMWE molecules are more tightly packed at

the interface than proteins, which is particularly marked for poly-oxyethylene sorbitan esters such as Tween 20 (Wilde and oth-

ers 2004; Grigoriev and Miller 2009). However, for monolayer-and/or single emulsifier-covered interfaces, the interface thickness

varies from 0.5 to 1 nm, for surfactant-stabilized interfaces, to 1to 15 nm for protein emulsifiers due to the formation of loops

inside the water phase (Dalgleish 1993; Fang and Dalgleish 1993;Atkinson and others 1995; Dickinson 2009; Singh 2011).

Surfactants form fluid interfaces with a substantial surface lateral

diffusion coefficient (Wilde and others 2004). This is also the casefor phospholipids. Their lateral diffusion coefficient at 26 °C in

mixed monolayers adsorbed at the oil–water interface, as com-puted by molecular dynamics simulation, was higher (9.3 × 10−8

cm2

/s) than that calculated for mixed bilayers (3.4 × 10−8

cm2

/s)(Hennere and others 2009b). Mixed phospholipid monolayers

would also have a looser packing, smaller order parameters andslightly larger geometrical areas par molecule (57.4 A2 compared

with 55 A2) than in bilayers, while a different trend was foundwhen a homogeneous phospholipid monolayer was compared to

the corresponding bilayer (Hennere and others 2009a). The acylchains of the triacylglycerols were found to interact deeply with

the phospholipid monolayer, one phospholipid methyl interactingwith 15.4 triacylglycerol methylene (Hennere and others 2009b).

In contrast, proteins form immobile, viscoelastic interfacial filmsexhibiting non-Newtonian behavior. Surface viscosity increases

with intra- and intermolecular cohesion within the film, andit is greater for proteins that can easily aggregate, such as β-

lactoglobulin (BLG), than for disordered proteins, such as β-casein (BCN) (Dickinson 1992; Murray and Dickinson 1996;

Damodaran 2004; Wilde and others 2004). Proteins can undergostructural and conformational rearrangements after adsorption,

which implies that the physical properties of the interface layer may evolve (Dickinson 1992; Tcholakova and others 2008).

Effect of interface properties on the physical stabilityof emulsions

The properties of the oil droplet surface control, to a large

extent, the physical stability of emulsions. The nature and con-centration (surface load) of adsorbed emulsifiers, together with the

C 2014 Institute of Food Technologists® Vol. 13, 2014 Comprehensive Reviews in Food Science and Food Safety 947




physical chemistry of the aqueous phase (pH, ionic strength, andions) determine the electrostatic charge and the thickness of the in-

terfacial layer, both parameters being deeply involved in the phys-ical stability of emulsions. Interfacial proteins can generate both

steric and electrostatic repulsion, and therefore, form a physicalbarrier to droplet coalescence (Dickinson 1992; Wilde and others

2004; Tcholakova and others 2008; Grigoriev and Miller 2009).For instance, bovine serum albumin (BSA) induces long-ranged

steric repulsions, whereas BCN exerts mainly electrostatic repul-

sions, and BLG combines electrostatic repulsions at large distancesand steric repulsions at short distances between the surfaces of 2nearby droplets (Dimitrova and Leal-Calderon 1999; Dimitrova

and others 2001, 2004). In a certain concentration range, poly-mers can also be responsible for droplet–droplet bridging, leading

to flocculation (Dickinson 1992; Dimitrova and Leal-Calderon1999; Tcholakova and others 2008; Grigoriev and Miller 2009).

With regard to LMWEs, ionic surfactants adsorbed at interfaces in-duce electrostatic repulsions between droplets, whereas nonionic

surfactants such as Tween 20 can exert steric repulsion becauseof their polyoxyethylene chains. However, adsorbed LMWEs are

generally considered as less efficient than proteins in avoiding theinterfacial film-breaking and droplet coalescence. This is mainly

because LMWEs form thinner interfacial films that provide weaker steric repulsion as compared to proteins (Dickinson 1992; Wildeand others 2004).

Partitioning of emulsifiers and associated measurementsThe amounts of emulsifiers used to stabilize emulsions are gen-

erally widely higher than the actual amounts required to only cover

the interfacial surface. As a consequence, a substantial fraction of the emulsifiers remains unadsorbed, and therefore, partitions be-

tween the oil and the water phases.

Distribution of single emulsifiers. Depending on their hy-

drophilicity, unadsorbed emulsifiers will locate preferentially inthe oil or the aqueous phase of emulsions. Biopolymer emulsifiers(such as proteins) are generally water-soluble and their unadsorbed

fraction will therefore be mostly present in the aqueous phaseas monomers, oligomers, or holo- or mixed aggregates. Surfac-

tants can be either water-dispersible or oil-dispersible dependingon their hydrophilic–lipophilic balance (HLB), which will deter-

mine the preferred location of their unadsorbed fraction. Beyond acertain concentration (the critical micelle concentration [CMC])

unadsorbed surfactants will form micelles (in the aqueous phase)or reverse micelles (in the oil phase), which constitutes a distinct

pseudophase within the emulsion (Capek 2004; Oehlke and oth-ers 2008; Pastoriza-Gallego and others 2009). The dynamics of

the molecules in these structures is very fast, the exchange of sur-factants between micelles and the aqueous solution taking place in

around 10−6 s, the mean lifetime of a micelle being around 10−3 s

(Charvolin 1983). Diacyl phospholipids do not form micelles inaqueous media because of their molecular geometry, but they caninstead organize as larger colloidal structures such as lamellae or

vesicles. These structures exhibit fairly slow dynamics of exchanges(hours to days) but rapid lateral movements (10−3 to 10−6 s; diffu-

sion coefficient around 1×10−8 to 15×10−8 cm2/s), provided thatthe temperature is above the phase transition temperature (Lange

1986; Mouritsen 2005). Accordingly, lateral diffusion coefficientof phospholipid bilayers computed by molecular dynamics sim-

ulation varied from 3.4 to 6.1 10−8 cm2/s (Hennere and others2009a, 2009b).

Competitiveadsorptionandprocess. In complex systems, whenemulsions contain mixtures of emulsifiers, competitive adsorption

phenomena may modify the adsorption at the oil–water interfaceand the partitioning of individual molecules. For instance, LMWEs

can compete with proteins for adsorption at the oil–water inter-face and even displace already adsorbed proteins (orogenic dis-

placement). This is explained by the usually f aster adsorption rateof surfactants as compared to proteins during the emulsification

process (Tcholakova and others 2008) and also because surfactantsare more surface-active (Courthaudon and others 1991a, 1991b;

Dickinson 1991, 1992, 1998; Mackie and others 2000; Bos and

van Vliet 2001; van Aken 2003; Damodaran 2004; Kotsmar andothers 2008; Morris and Gunning 2008; Day and others 2010).The partitioning of surface-active molecules within emulsions is

also dependent on kinetic or process-related factors in complexemulsions. For instance, Waninge and others (2005) assessed the

distribution of proteins and phospholipids in emulsions of identi-cal compositions, but prepared through varied procedures: (i) oil

emulsification was performed in the presence of proteins, thenphospholipids were added to the emulsion; (ii) oil emulsification

was performed in the presence of phospholipids, then proteinswere added to the emulsion; or (iii) oil emulsification was per-

formed in the presence of both proteins and phospholipids. Theinterfacial composition measured in these emulsions was different,

even after up to 48 h of storage, and was dominated by the emulsi-fier that was present during the homogenization step. This suggeststhat nonequilibrium interfacial structures may exist within a sub-

stantial timescale, and that not only the type and concentrationof surface-active molecules, but also the procedure for emulsion

preparation, are important.

Dynamic evolution of emulsifier partitioning. The amount of

unadsorbed emulsifiers may evolve during the storage of emul-sions (Euston and Mayhill 2001; Dalgleish and others 2002;

Bongard and others 2009). This phenomenon, linked to dynamicexchanges between adsorbed and unadsorbed emulsifiers, can take

place in minutes to days, depending on the composition of thesystem and on the temperature. Lipid oxidation may also in-

duce changes in the interfacial composition because surface-activemolecules are formed in the oil phase (Moberger and Larsson

1985). For example, linoleic acid (LA), methyl linoleate, and trili-nolein hydroperoxides are more surface-active than their non-

peroxidized counterparts (Abousalham and others 2000; Nuchiand others 2002). This may lead to reorganization and even des-

orption of some emulsifiers from the interface and contribute tophysical destabilization of the system (Genot and others 2003). It

is therefore interesting to measure unadsorbed emulsifiers not onlyin the freshly prepared emulsions, but also along the incubation

period.It is thus clear that the nature and concentration of molecules

present at the interface do not reflect the global compositionof emulsifiers used to prepare the emulsions. The actual parti-

tioning of emulsifiers has to be carefully considered since unad-sorbed emulsifiers present in the aqueous phase can dramatically

affect the physical and chemical stability of emulsions. Unadsorbedproteins or surfactants can induce depletion flocculation, leading

to the physical destabilization of the emulsion (Dickinson andGolding 1997; Casanova and Dickinson 1998; Dickinson and

others 2003; Dauphas and others 2008; Ye 2008). The role of unadsorbed emulsifiers on the chemical stability of emulsified oil

is detailed below.The actual partitioning of emulsifiers should be experimentally

measured and taken into account for understanding the physical

and chemical stability of the emulsions. Two main strategies havebeen developed to measure proportions and concentrations of





adsorbed and unadsorbed emulsifiers in emulsions: “in situ” and“indirect” measurements.

In situ measurements. This approach consists of evaluating di-rectly the relative proportions of adsorbed and unadsorbed emul-

sifiers through in situ measurement of a physical parameter thatis modified upon the emulsifier adsorption and quantitative cali-

bration. Accordingly, front-surface fluorimetry has been proposedto evaluate directly the unadsorbed fraction of BSA in dodecane

emulsions (Castelain and Genot 1994; Granger and others 2005).

Unfortunately, the protein fluorescence characteristics in emul-sions depends both on the oil and the protein used (Rampon andothers 2004) and the method failed to be generalized. Oehlke and

others (2008) used small-angle neutron scattering (SANS) to dis-tinguish between surfactants adsorbed at the interface, as aqueous

phase micelles or monomers in O/W emulsions. The results wereconsistent with other methods involving a physical separation and

analysis of the emulsion phases. SANS requires, however, con-trasting the aqueous phase and surfactant using D2O, which might

slightly alter the behavior of surfactants.

Indirect measurements. The 2nd and generally used strategy

to assess unadsorbed emulsifiers consists of physically separatingthe aqueous phase from the oil droplets covered by the adsorbed

fraction (that is, emulsion fractionation) and then quantifyingthe emulsifier in one or both phases (Patton and Huston 1986;Courthaudon and others 1991b, 1991c; James-Smith and others

2007; Ye and Singh 2007). The oil droplets are generally separatedthrough centrifugation (Rampon and others 2003). This step can

be performed directly if the size of the oil droplets is sufficient toallow their creaming. If the emulsion contains small droplets with

a density very close to that of the aqueous phase density, thesedroplets do not cream. They can be separated by either dilution

of the emulsion in sucrose solution to increase the density of theaqueous phase prior to centrifugation, which is then performed on

sucrose gradient (Patton and Huston 1986; Le Denmat and oth-ers 2000; Bongard and others 2009), or by sequential filtrations

through cellulose acetate membrane filters from 0.45 down to0.1 µm (Berton and others 2011a). The usage of nanoporous

filters combined with centrifugation has also been proposed andapplied to separate unadsorbed LMWEs (namely, sodium docecyl

sulfate [SDS]) from alkane oil droplets (James-Smith and others2007). The emulsifier amount is then quantified either in one

phase only, or in both phases (aqueous phase and creamed layer),which makes possible to construct a mass balance. However, it

must be pointed out that spectrophotometric methods used toquantify proteins in the aqueous phase may not be relevant in ox-

idizing conditions due to interferences with new light-absorbingcompounds arising from oxidative reactions or to alterations of

some amino acids involved in the quantification reaction (Bertonand others 2011a). The mass charge of the interfacial layer is finally

calculated by taking into account the total surface area developedby the oil droplets as measured by the light diffraction technique.

The underlying assumption is that the separation of the phasesdoes not modify the partitioning of the emulsifier, which is prob-

ably false especially when water-soluble LMWEs, allowing rapiddynamics of exchange between aqueous phase, micelles, and oil

droplets, are involved.

Overview of emulsifier partitioning in literature dataTo provide an overview of the common concentrations used

in the literature, we have reported the estimated concentrations

(C excess, g/L) of unadsorbed emulsifiers in various emulsionsand the corresponding proportion of nonadsorbed emulsifiers

(P excess, %) (Table 1A and 1B). The values were either extractedfrom the original research articles or estimated from the amounts

of emulsifiers used to prepare the emulsions, the reported aver-age droplet size, and the surface load of emulsifiers as found in

the literature. For instance, typical surface loads encountered for protein-stabilized interfaces range from 1.5 to 4.5 mg/m2, de-

pending on the protein type and the pH (Atkinson and others1995; Bos and van Vliet 2001), while surface loads are lower for

surfactant-stabilized interfaces, 1 to 2 mg m−2 (Bos and van Vliet

2001).Table 1C gives the estimated surface loads in articles where the

unadsorbed concentration was experimentally determined. C excess

and P excess in a given mass of emulsion were estimated accordingto the following calculations (Eq. 1 to 3):

C excess=m em total − [ A× ϕ × em )]

V aqueous phase

(1)

where mem total is the total mass of emulsifier (g), V aqueous phase is thevolume of aqueous phase (g), φ is the oil mass fraction (%), and

em is the theoretical surface load of the emulsifier at the oil–buffer interface (mg/m2). A is the specific surface area (m2/g oil), and is

defined as:

A=3

r × ρ(2)

where r is the average oil droplet radius deduced from d 3,2 (µm)and ρ is the density of the oil (0.92 g/mL at room temperature).

P excess =C excess × 100

C initial

(3)

where C initial is the initial concentration of emulsifier in aqueoussolution (g/L).

These calculations were applied to 80 examples of emulsion for-

mulations found in original research articles dealing with lipid ox-idation in emulsions (Table 1A and 1B). A broad variety of systems

was therefore covered with regard to emulsion design: the type of emulsifier, oil mass fraction, and droplet size. Twenty eight formu-

lations were associated with an estimated proportion of unadsorbedemulsifier higher than 90%; 20 resulted in an unadsorbed propor-

tion between 70% and 90%; and 13 resulted in an unadsorbedproportion between 50% and 70%. In other words, the great ma-

jority (76%) of the formulations gave emulsions in which the mainpart (more than one-half) of the emulsifier molecules were pre-

sumably unadsorbed. This proportion may even be slightly under-estimated, since the present calculations, based on the assumption

of a single layer of adsorbed emulsifiers, were done considering

rather high estimates for em among the values available in theliterature.

Negative values for the unadsorbed concentrations and propor-

tions of emulsifiers were obtained for 7 formulations, meaning thatthe actual surface load was lower than the theoretical value used

for the calculation. For these examples, we subsequently calculatedan estimated surface load (mg/m2), considering that all the emul-

sifier molecules used were adsorbed at the interface (Table 1B).The surface loads estimated accordingly were still consistent with

values available in the literature (Atkinson and others 1995;Dickinson 1998; Bos and van Vliet 2001; Ye 2008), indicating

that these systems may be reasonably considered as examples werethe adsorbed fraction of emulsifiers was predominant.





Table 1A–Estimated concentrations and proportions of unadsorbed emulsifiers in the aqueous phase of O/W emulsions for practical examplesencountered in the literature. Theoretical surface loads have been extracted from previous studies—Please note that estimates in the highest rangewere used for surface loads, meaning that the unadsorbed concentrations and fractions may be slightly underestimated rather than overestimated.When oil fractions were indicated as volume fractions in the original research articles, an oil density of 0.92 was applied to calculate the oil massfractions.

Initial Estimatedconcentration Average Theoretical unadsorbed Estimated

in aqueous Oil fraction surface surface load concentration (g/L unadsorbedArticle Emulsifier pH solution (g/L) (% wt) diameter (µm) (mg/m2)a aqueous phase) proportion (%)

Imai and others (2008) Sucrose ester 7.4 30.0 0.8 1.70 1.5 30.0 99.9Gohtani and others

(1999)Sucrose ester n/a 10 1 6.4 1.5 10.0 99.8

Gohtani and others(1999)

Sucrose ester n/a 10 1 3.4 1.5 10.0 99.7

Atares and others (2012) WPI 7 15.0 18.7 51.00 2.0 14.9 99.6Kargar and others (2011) NaCas n/a 20.0 5.0 8.90 3.0 19.9 99.4Kargar and others (2011) Tween 20 n/a 20.0 5.0 4.10 1.5 19.9 99.4Nakaya and others (2005) Sucrose ester 6.6 11.1 10.0 10.70 1.5 11.0 99.1Mora-Gutierrez and others

(2010)Casein 7 5.0 0.1 0.22 3.0 4.9 98.2

Ries and others (2010) WPI n/a 100.0 10.6 0.65 2.0 97.6 97.6Nakaya and others (2005) Sucrose ester 6.6 11.1 10.0 3.28 1.5 10.8 97.0Mora-Gutierrez and others

(2010)Casein 7 5.0 0.2 0.25 3.0 4.8 96.9

Lethuaut and others(2002)

BSA 4.3 20.0 28.3 8.10 2.0 19.4 96.8

Kargar and others (2011) Tween 20 n/a 20.0 30.0 6.50 1.5 19.4 96.8

Kargar and others (2011) NaCas n/a 20.0 30.0 12.50 3.0 19.3 96.6Ries and others (2010) NaCas n/a 100.0 10.6 0.65 3.0 96.4 96.4Kiokias and others (2006) Tween 20 7 10.0 10.0 2.35 1.5 9.5 95.4Ries and others (2010) WPI n/a 100.0 10.6 0.31 2.0 95.0 95.0Mora-Gutierrez and others

(2010)Casein 7 5.0 0.5 0.36 3.0 4.7 94.5

Mora-Gutierrez and others(2010)

Casein 7 5.0 1.0 0.67 3.0 4.7 94.1

Sun and Gunasekaran(2009)

WPI 7 21.0 4.6 0.50 2.0 19.7 94.0

Kiokias and others (2006) WPI 7 10.0 10.0 2.30 2.0 9.4 93.7Ries and others (2010) NaCas n/a 100.0 10.6 0.31 3.0 92.5 92.5Kiokias and others (2006) NaCas 7 20.0 10.0 1.40 3.0 18.4 92.2Mancuso and others

(1999)Tween 20 3 20.9 5.0 0.30 1.5 19.2 91.8

Kargar and others (2011) Tween 20 n/a 1.0 5.0 5.90 1.5 0.9 91.3Kiokias and others (2006) NaCas 7 10.0 10.0 2.32 3.0 9.1 90.6Kargar and others (2011) NaCas n/a 1.0 5.0 10.40 3.0 0.9 90.1


WPI 7 10.5 4.6 0.60 2.0 9.4 90.0

Lee and others 2011 WPI 7 9.0 0.5 0.07 2.0 8.0 89.0Nakaya and others 2005 Sucrose ester 6.6 11.1 10.0 0.81 1.5 9.8 87.9Sun and Gunasekaran

(2009)WPI 7 24.6 18.7 1.00 2.0 21.6 87.8

Imai and others (2008) Sucrose ester 7.4 30.0 0.8 0.02 1.5 26.3 87.7Kiokias and others (2006) NaCas 7 5.0 10.0 3.04 3.0 4.3 85.7Hu and others (2003b) WPI 3 15.0 5.0 0.26 2.0 12.4 82.4Atares and others (2012) WPI 7 15.0 18.7 1.10 2.0 12.3 81.8Sun and Gunasekaran

(2009)WPI 7 12.3 18.7 1.20 2.0 9.8 79.7


Casein 7 15.0 5.0 0.33 3.0 11.9 79.2


BSA 4.3 20.0 28.3 1.20 2.0 15.7 78.5

Mancuso and others(1999)

DTAB 3 5.2 5.0 0.30 1.0 4.1 78.0


WPI 7 32.3 38.0 1.10 2.0 25.0 77.5

Hu and others (2003b) Casein 3 15.0 5.0 0.30 3.0 11.6 77.1Mancuso and others

(1999)SDS 3 4.9 5.0 0.30 1.0 3.8 76.6

Osborn and Akoh (2004) WPI 7 5.6 10.0 1.07 2.0 4.2 75.8Hu and others (2003b) Casein 3 10.0 5.0 0.42 3.0 7.5 75.5Mora-Gutierrez and others

(2010)Casein 7 10.0 5.0 0.38 3.0 7.3 72.9

Hu and others (2003b) WPI 3 10.0 5.0 0.25 2.0 7.3 72.5Osborn and Akoh (2004) WPI 7 7.1 30.0 2.69 2.0 5.0 70.7Villiere and others (2005) BSA 6.5 20.0 28.3 0.87 2.0 14.1 70.4Fomuso and others

(2002a)Tween 20 7 10.0 10.0 0.35 1.5 6.9 68.9

(Continued)





Table 1A–Continued

Initial Estimatedconcentration Average Theoretical unadsorbed Estimated

in aqueous Oil fraction surface surface load concentration (g/L unadsorbedArticle Emulsifier pH solution (g/L) (% wt) diameter (µm) (mg/m2)a aqueous phase) proportion (%)

Haahr and Jacobsen(2008)

Citrem 3 11.2 10.0 0.31 1.5 7.6 68.2

Taherian and others(2011)

WPI 3.4 10.3 3.0 0.41 7.0 b 6.8 66.4

Sun and Gunasekaran

(2009)

WPI 7 2.1 4.6 0.80 2.0 1.3 62.3

Kargar and others (2011) Tween 20 n/a 1.0 30.0 10.50 1.5 0.6 60.1Taherian and others

(2011)WPI 6.8 10.3 3.0 0.34 7.0 b 6.1 59.6


WPI 7 16.1 38.0 1.20 2.0 9.5 58.7

Fomusoand others(2002a)

WPI 7 10.0 10.0 0.35 2.0 5.9 58.6

Villiere and others (2005) NaCas 6.5 20.0 28.3 0.92 3.0 11.6 58.0Hu and others (2003b) Casein 3 5.0 5.0 0.47 3.0 2.8 56.2Lee and others (2011) WPI 7 0.5 0.5 0.33 2.0 0.2 55.2Ries and others (2010) WPI n/a 5.0 10.6 0.65 2.0 2.6 52.4Haahr and Jacobsen

(2008)Tween 20 3 11.2 10.0 0.20 1.5 5.7 50.5

Hu and others (2003b) WPI 3 5.0 5.0 0.27 2.0 2.5 49.1Haahr and Jacobsen

(2008)Tween 20 7 11.2 10.0 0.19 1.5 5.4 48.4


Casein 7 5.0 5.0 0.38 3.0 2.3 45.8

Kargar and others (2011) NaCas n/a 1.0 30.0 15.10 3.0 0.4 44.5Haahr and Jacobsen

(2008)Citrem 7 11.2 10.0 0.17 1.5 5.0 44.2

Haahr and Jacobsen(2008)

NaCas 3 11.2 10.0 0.31 3.0 4.2 37.8


BSA 4.3 20.0 28.3 0.40 2.0 7.1 35.6

Ries and others (2010) NaCas n/a 5.0 10.6 0.65 3.0 1.4 28.6Haahr and Jacobsen

(2008)NaCas 7 11.2 10.0 0.24 3.0 2.2 19.5

Ries and others (2010) WPI n/a 5.0 10.6 0.31 2.0 0.0 0.2

aFrom Atkinsonand others(1995),BonfillonandLangevin (1993), Bosand vanVliet(2001),Dickinson (1998), Oehlkeand others(2008), Tanakaand Ikeda(1991),Ye (2008).bFromTaherianand others(2011).Abbreviations: BSA, bovine serum albumin; NaCas, sodium caseinate; WPI, whey protein isolate; SDS, sodium dodecyl sulfate; DTAB, dodecyl trimethylammoniumbromide; Citrem, ester of citric acid and fattyacids;n/a, not adjusted.

Table 1B–Estimated surface loads of emulsifiers in examples where the calculations performed in Table 1A led to negative unadsorbed concentrationsandfractions.Hence,weassumeherethatlowconcentrationsandproportionsofunadsorbedemulsifiersarepresentintheaqueousphase(approximately0). When oil fractions were indicated as volume fractions in the original research articles, an oil density of 0.92 was applied to calculate the oil massfractions.

Initial Estimatedconcentration Average Calculated unadsorbed Estimated

in aqueous Oil fraction surface surface concentration unadsorbedArticle Emulsifier pH solution (g/L) (% wt) diameter (µm) load (mg/m2) (g/L aqueous phase) proportion (%)

Mora-Gutierrez and others (2010) Casein 7 1.0 5.0 0.42 1.2 0 0Mora-Gutierrez and others (2010) Casein 7 2.0 5.0 0.40 2.3 0 0Hu and others (2003b) Casein 3 2.0 5.0 0.48 2.8 0 0Hu and others (2003b) WPI 3 2.0 5.0 0.29 1.7 0 0Sun and Gunasekaran (2009) WPI 7 2.5 18.7 1.10 1.8 0 0Sun and Gunasekaran (2009) WPI 7 3.2 38.0 1.60 1.3 0 0Ries and others (2010) NaCas n/a 5.0 10.6 0.31 2.0 0 0

Abbreviations: NaCas,sodium caseinate; WPI,whey protein isolate; n/a,not adjusted.

Finally, 2 studies (Faraji and others 2004; Berton and others

2011a) directly reported experimental values for the concentra-tions and proportions of unadsorbed emulsifiers. For these studies,

we calculated the actual surface load of emulsifiers (Table 1C). Inthe work of Berton and others (2011b), surface loads ranging from1.8 to 2.8 mg/m2 were found depending on the type of emulsi-

fier, which fits with theoretical values (Bos and van Vliet 2001).

In the work of Faraji and others (2004), surface loads of 4.1 to16.9 mg/m2 were found for whey protein isolate (WPI)-stabilized

emulsions. Some of these values are therefore high as compared

to the usually described surface loads of whey proteins; hence, theunadsorbed concentrations may have been underestimated.

To conclude, our estimate of unadsorbed emulsifier concentra-tions and proportions in several studies dealing with oxidation of

O/W emulsions points out that the contribution of the unad-sorbed fraction of emulsifiers to the physical and chemical stability

can be substantial and may often overcome the effect of the emul-sifiers adsorbed at the oil–water interface.





Table 1C–Estimated surface loads of emulsifiers in examples where the concentrations and fractions of unadsorbed emulsifiers were experimentallymeasured. When oil fractions were indicated as volume fractions in the original research articles, an oil density of 0.92 was applied to calculate the oilmass fractions.

Initial Average Measuredconcentration Oil surface Calculated unadsorbed unadsorbed

in aqueous fraction diameter load concentration proportionArticle Emulsifier pH solution (g/L) (% wt) (µm) (mg/m2) (g/L aqueous phase)a (%)

Berton and others (2011b) Tween 80 6.7 5.0 30.0 1.70 2.0 1.6 33.0Berton and others (2011b) BLG 6.7 5.0 30.0 1.50 1.9 1.5 30.0Berton and others (2011b) BLG 3 5.0 30.0 2.20 2.3 2.1 42.0

Berton and others (2011b) BSA 6.7 4.0 30.0 1.80 1.8 1.2 29.0Berton and others (2011b) Tween 20 6.7 5.0 30.0 1.40 2.3 0.4 9.0Berton and others (2011b) Tween 20 3 5.0 30.0 1.60 2.5 0.5 11.0Berton and others (2011b) BCN 6.7 5.0 30.0 1.70 2.8 0.4 9.0Faraji and others (2004) WPI 7 2.5 5.0 0.59 4.1 0.1 5.0Faraji and others (2004) WPI 7 5.0 5.0 0.50 6.6 0.5 10.0Faraji and others (2004) WPI 7 10.0 5.0 0.50 12.5 1.4 14.0Faraji and others (2004) WPI 7 15.0 5.0 0.46 16.9 2.4 16.0

aValueswere determined experimentally.Abbreviations: BSA,bovine serumalbumin; WPI,whey protein isolate; BLG,β-lactoglobulin; BCN,β-casein.

The General Rules of Lipid OxidationGeneral mechanisms of lipid oxidation

Lipid oxidation consists of the reaction of molecular oxygenwith unsaturated fatty acids (Chevreul 1823; Lea 1939). The reac-

tion schemes and rate constants for oxidation of unsaturated fattyacids and bulk oils and fats can be found in several books and

reviews (Labuza 1971; Frankel 1985, 2005; Porter 1986; Chan1987; Gardner 1989; Hsieh and Kinsella 1989; Denisov 1995;

Kamal-Eldin 2003; Schaich 2005; Cheng and Li 2007; Laguerreand others 2007; Kamal-Eldin and Min 2010; Schaich and others

2013). Lipid oxidation is described as a free radical chain reac-tion divided into 3 stages: initiation, propagation, and termination

(Farmer and others 1942).

Initiation. The direct reaction between acyl chain (LH) and

triplet oxygen (3O2) cannot occur spontaneously because the lipidground state has an opposite spin direction from that of 3 O2. The

activation energy of this direct reaction would therefore be too

high (Chan 1987; Belitz and others 2004). The spin barrier can beovercome in the presence of initiators that produce lipid radicals or convert 3 O2 in reactive oxygen species (ROS), including hydroxyl

radical (OH·) and singlet oxygen (1O2).The initiation mechanism occurs through different pathways:

autoxidation, enzymatic oxidation, and singlet oxygen (1O2) oxi-dation. The latter is encountered during photoinitiation, the sys-tem being exposed to light in the presence of a sensitizer. It is a

nonradical reaction because 1O2 reacts directly with the doublebonds. Enzymatic oxidation is not usual in food emulsions, and

therefore, not under the scope of this review. Both enzymatic andsinglet oxygen reactions have specificities with regard to primary

reaction products, but the latest stages of the reactions are similar to autoxidation.

The 1st step of autoxidation is described as follows. In thepresence of initiators, unsaturated fatty acids (LH) lose a hydrogen

atom (H) in an allylic position relative to a fatty acid double bond.It forms a lipoyl or alkyl free radical (L•), according to reaction

(1):

LH → H+ L

(1)

This reaction is supposed to occur via different mechanisms:- Direct reaction between LH and transition metal ions (M):

LH +M3+ → L

+ H++ M2+ (2)

- Reaction between LH and oxygen radicals arising from metal(such as iron) autoxidation (Schaich 2005):

Fe2+

+ O2 → Fe3+

+ O−

2 ←→ HOO

+ H+

→ L

+ H2O2 (3)

2O−

2 or O−

2 /HOO

→ H2O2 + O2 (4)

H2O2 + Fe2+ → HO

+ OH−+ Fe3+ (5)

HO

+ LH → H2O+ L

(6)

Lipid matrices often contain trace hydroperoxides (LOOH) thatcan undergo oxidation, reduction, or decomposition, leading to

lipid radicals involved in the early stages of lipid oxidation:- Oxidation or reduction of hydroperoxides (LOOH) catalyzed

by metals:

LOOH+M2+ → LO

+ OH−+ M3+ (7)

LOOH+M3+ → LOO

+ H++ M2+ (8)

- Thermal decomposition of hydroperoxides:

LOOH → LO

+ O

H (9)

The activation energy of this reaction is high (Frankel 2005);

another mechanism, in agreement with kinetic observations, in-volves 2 molecules of hydroperoxides:

2LOOH → LO

+ H2O+ LOO

(10)

Propagation. The alkyl radicals (L•) produced during initiationreact very quickly with triplet oxygen to generate peroxyl radicals

(LOO•), according to reaction (11):

L

+ O2 → LOO

(11)





Since peroxyl radicals are unstable, they abstract hydrogen atomsfrom other unsaturated fatty acids to form hydroperoxides and

other alkyl radicals:

LOO

+LH → LOOH+ L

(12)

The newly formed alkyl radicals can then react with triplet oxy-

gen, and so on. This is why the measurement of oxygen uptake,even if not a very sensitive method, can be a good alternative to

evaluate oxidation development for samples put in closed vessels(Villiere and Genot 2006; Berton and others 2012b). The radical

chain reaction thereby engaged propagates at a high rate and theincrease of hydroperoxides formed becomes exponential. These

primary products of oxidation are generally quantified with col-orimetric methods and often expressed as peroxide value (PV), the

detected amounts reflecting the balance between both formationand further reactions of these unstable compounds. The formation

of hydroperoxides from nonconjugated PUFAs goes with the sta-bilization of free radicals via double-bond rearrangement, which

gives rise to conjugated dienes. The UV absorption of these chem-ical structures around 233 nm allows an easy quantification of most

hydroperoxides from linoleic and linolenic oils, but may reveal to

be inappropriate in other oils containing long-chain PUFAs.Many alternate reaction pathways to these simplified schemes

also take place during lipid oxidation. These pathways were re-

cently summed up by Schaich (2005) and Schaich and others(2013). They comprise β -scission of oxygen, internal rearrange-

ment to epidioxides, addition and disproportionation of lipid per-oxyl radicals, internal rearrangement to epoxides, and addition and

α- or β-scissions of alkoxyl radicals. According to Schaich (2005),these reactions depend highly on environment conditions (oxygen

pressure, protic or aprotic solvent, lipid concentration, mediumviscosity, presence of external abstractable hydrogen, level of

PUFAs, and presence of water or of proton donors). They giverise to various reaction products, among them compounds whichare rarely measured. Thus, in the current literature related to oxi-

dation in emulsified and dispersed systems, these alternate reactionpathways to the classical hydroperoxide route are not taken into

account. This could explain some discrepancies between availabledata or difficulties to explain the kinetics of formation of oxidation

products.In the classical reaction scheme, the next step, which involves

radical and nonradical pathways, is the decomposition of hydroper-oxides. The reaction is favored by the presence of metal catalysts

and by an increased temperature. The prominent mechanism con-sists of the homolytic scission of the double-bond adjacent to

the hydroperoxyl group to give secondary products of oxidation(Gardner and others 1974). It is a complex phenomenon, which

leads to the formation of various compounds. The main com-

pounds are carbonyl compounds, alcohols, aldehydes, and hy-drocarbons, a wide range of them being low-molecular-weightcompounds. Among them, volatile lipid oxidation products are

responsible for the flavor deterioration of food lipids. Some of the produced molecules of higher molecular weight, such as core

aldehydes, are less hydrophobic than the initial compounds thatmay change their location toward a more polar environment when

produced in a multiphase system. It is generally advised to mea-sure at least 1 marker of secondary lipid oxidation products, along

with hydroperoxides and/or conjugated dienes. In that regard, themost used methods include the determination of the ρ-anisidine

value (a measure of total aldehydes, through a spectrophotometricassay), volatile compounds (such as propanal, hexanal; generally

achieved through static or dynamic headspace combined withgas chromatography), or thiobarbituric acid-reactive substances

(TBARS) (a measure of various compounds including malondi-aldehyde, through a spectrophotometric assay).

Termination. During this stage, which is generally less exten-sively described than the others, the radicals react together to form

stable nonradical compounds:

LOO

+ LOO

→ nonradicals products (13)

These nonradical compounds cannot participate in the reactioncycle anymore, which stops the radical chain reaction. Because of

the varied decomposition pathways of hydroperoxides, differentradicals are produced, and there are many possibilities of combina-

tions between them. Some antioxidant compounds can promotetermination reactions. To our knowledge, these compounds have

never been measured during the oxidation of O/W emulsions.

Effects of themolecular structureand physical state of lipidson their oxidizability in bulk phases

Apart from environmental conditions, including temperature,

oxygen concentration, and the presence of light, radiations, and

so on, lipid oxidation is highly dependent on the characteristicsof the substrate, that is, the fatty acid composition of the oil andthe molecular structures of lipids. The presence of oil-soluble or

oil-dispersible minor compounds can also dramatically affect theoxidation of bulk oils.

Fatty acid unsaturation. The susceptibility of fatty acids to hy-drogen abstraction (Eq. 1, 2, and 6) depends on the dissociation

energies of C-H bonds. In the presence of a double bond inthe fatty acid chain, the dissociation energy of the C-H bonds

located on the allylic carbons is weakened (from around 100 to74 kcal/mol), which makes the hydrogen removal easier, and for a

bisallylic C-H bond, the dissociation energy is substantially lower (65 kcal/mol) (Reich and Stivala 1969; Porter 1986; Gardner 1989;Schaich and others 2013). Consequently, the susceptibility of mo-

nounsaturated fatty acids such as oleic acid to oxidation is muchlower than that of PUFAs, and the rates of lipid oxidation in-

crease with lipid unsaturation. The most oxidizable fatty acids arethe long-chain PUFAs, since they contain several bis-allylic C-H

bonds in their hydrocarbon chain. For example, the rate of autox-idation of ethyl linolenate (C18:3 n-3) was found 2.4 times that of

ethyl linoleate (C18:2 n-6), and the rate of autoxidation of methylarachidonate (C20:4 n-6) was about twice that of ethyl linolenate

(Holman and Elmer 1947). The introduction of 1 additional dou-ble bond into a fatty acid at least doubles the rate of oxidation

of the fatty acid or its ester: oxidizabilities (M−1/2 sec−1/2) of LA,ALA, ARA, and DHA at 37 °C were 2.03×10−2, 4.07×10−2,

5.75×10−2, and 10.15×10−2, respectively (Cosgrove and others

1987). The absolute rate constants for the reactions of radicalswith unsaturated fatty acids also increase with increasing unsatu-ration (Forni 1990) and the induction period for the autoxidation

of fatty acids decreases with increasing unsaturation. For instance,at 50 °C, induction periods of 700, 22, and 10 h were observed

during the autoxidation of oleic acid (C18:1, n-9), LA, and ALA,respectively (Martin and others 1990).

Total unsaturation of the lipid phase. Oxidizability of bulkphase lipids also increases with the concentration in unsaturated

fatty acids. Increasing slightly the oil content in long-chain PUFAs(ARA, all -cis-7,10,13,16,19-docosapentaenoic acid [DPA, C22:5

n-3] and DHA) through the addition of low proportions (between0.5% and 8% w/w) of fungal, tuna, and egg oil to a vegetable oil





mix resulted in a significant decrease of the oxidation lag phase(Bartee and others 2007). Other studies showed a decrease of lipid

oxidation in polyunsaturated oils (LA, ALA) blended with oilscontaining oleic acid or, more generally, monounsaturated fatty

acids (Torres and others 2006; Anwar and others 2007; Mezouariand Eichner 2007).

In an attempt to quantitatively predict the susceptibility of lipidsto undergo oxidation depending on their degree of unsaturation,

empirical calculations based on the fatty acid composition of oils

have been developed. For example, the double-bond index can becalculated as the sum of the fraction of each fatty acid (%, w/w)times the number of double bonds in each acid (Pietrangelo and

others 1990). The peroxidizability index assesses the contributionof each fatty acid according to the following equation (Eq. 4)

(Arakawa and Sagai 1986; Cosgrove and others 1987; Hu andothers 1989):

Peroxidizability index = (% dienoic × 1)

+ (% trienoic× 2)

+ (% tetraenoic× 3)

+ (% pentaenoic× 4)

+ (% hexaenoic× 5) (14)

Triacylglycerol structure. The position of fatty acids on tria-cylglycerols can also impact their oxidative stability. This has been

studied both with pure synthetic tr iacyglycerols and with naturaloils.

Working with synthetic triacylglycerols, Neff and ElAgaimy(1996) observed that sn-1(3)-palmitoyl-sn-1,2(2,3)-dilinoleoyl-

glycerol (PLL) had lower oxidative stability than sn-2-palmitoyl-

sn-1,3-dilinoleoyl-glycerol (LPL). They attributed the effect to an

easier propagation of the chain reaction between the adjacent LAchains of PLL compared to nonadjacent LA of LPL. Endo and oth-ers (1997) found that in synthetic triacylglycerols, EPA and DHA

were more oxidizable when located at the sn-1,2(2,3)-positionof glycerol than at the sn-1,3-position. For instance, sn-1,2(2,3)-

dipalmitoyl-3(1)-eicosapentaenoyl-glycerol oxidized f aster than

sn-1,3-dipalmitoyl-2-eicosapentaenoyl-glycerol. Other experi-

ments showed that PUFAs were more stable to oxidation whenlocated at the sn-2 position of triacylglycerol compared to sn-1(3)

position (Wijesundera and others 2008; Wang and others 2010;Wang and Shahidi 2011).

Physical state of lipids (liquid compared with solid state). Thecrystallization state of vegetable bulk oils can also affect the devel-

opment of lipid oxidation (Calligaris and others 2006, 2007). Inthe former study, the oxidation rate of olive oil below its melting

point was higher than the theoretical value determined from the

Arrhenius equation. The authors proposed that an increased con-centration of unsaturated triacylglycerols in the liquid areas sur-rounding fat crystals could explain, at least in part, this result. The

latter study did not highlight any impact of the oil physical stateon the peroxide formation in a water-in-sunflower oil emulsion,

but the formation of hexanal was slightly greater, when comparedto the expected value, at the lowest temperatures tested (down

to −30 °C). The cryoconcentration of prooxidant compounds inthe aqueous phase was mentioned to explain this deviation.

Oil-phase minor components. Commercial oils naturally con-tain minor components that intervene in the lipid oxidation re-

action in bulk oils. Indeed, tocopherols, carotenoids, or phenolicsexert antioxidant activities in bulk oils through various chemi-

cal mechanisms, including quenching of free radicals by electrontransfer or hydrogen donation, and metal chelation (Porter 1980;

Laguerre and others 2007). According to the polar paradox, theseactivities depend not only on the chemical structure of the antiox-

idants, but also on their possible location in the bulk oil, at the air– oil interface, or even in colloidal structures dispersed in the bulk

oil (Huang and others 1996a, 1996b; Cuvelier and others 2000).Phospholipids present in nonrefined oils also exhibit antioxidant

properties, partly attributed to synergistic effects with tocopherols

(Kashima and others 1991; Bandarra and others 1999; Doert andothers 2012). However, they are also amphiphilic molecules thatorganize as colloidal structures within the oil phase in the presence

of traces of water and promote the prooxidant activity of metalions. For instance, colloidal structures formed by the association

of trace water and phospholipids were prooxidant in soybean oil(Chen and others 2010a, 2010b).

Lipid Oxidation in Emulsions: Preliminary StatementsLipid oxidation in emulsified oil compared with bulk oil

Lipid oxidation occurs generally earlier and faster in O/W emul-sions than in bulk oil, for a given type of oil (van Ruth and others

2000; Frankel and others 2002; Lethuaut and others 2002; Chee

and others 2006; Cercaci and others 2007). Such a favored oxida-tion induced by emulsification is attributed to several causes. First,the creation of interfacial area between the oil and the aqueous

phase is assumed to promote the contacts between unsaturatedlipids and prooxidant compounds, such as metal ions, dissolved

in the aqueous phase. Also, a high interfacial area could favor theaccessibility of the oil phase to oxygen dissolved in the aqueous

phase, which may be an important parameter when the oxygeninitially dissolved in the oil phase has been consumed. Second,

the emulsification process itself could promote oxidation throughincorporation of oxygen, overheating due to shear stress (Mao and

others 2009), or direct production of free radicals by acoustic cav-itation in the case of sonication (Jana and others 1990; Riesz and

Kondo 1992). Such an effect of the emulsification step is supportedby experimental evidence that lipid oxidation in emulsions can dif-

fer according to the initial homogenization procedure. Horn andothers (2012b, 2013) recently investigated the influence of the ho-

mogenization equipment (microfluidizer or 2-stage valve homog-enizer) and the homogenization pressure on lipid oxidation in fish

O/W emulsions stabilized with sodium caseinate (NaCas), BLG,or WPI. The oxidative stability of emulsions prepared with NaCaswas not influenced by the homogenization procedure, whereas the

microfluidizer homogenization resulted in lower oxidation levelswhen WPI was used as the emulsifier. Presumably, the homoge-

nization equipment, as well as the homogenization pressure, affectsthe relative adsorption of whey proteins (in particular, BLG and

α-lactalbumin), leading to different interfacial compositions.

Conversely, some studies did not show any increase of lipidoxidation induced by emulsification in comparison with lipidsin the bulk phase (Khan and Shahidi 2000), or even showed an

improved oxidative stability of long-chain PUFA oils after emulsi-fication (Belhaj and others 2010). In the former example, it could

have resulted from the presence of hydrophobic antioxidants thatwere more efficient in the multiphase system than in the bulk oil

according to the polar paradox. In the latter example, the resultscould be explained by an antioxidant effect of the phospholipid

emulsifier. One can also notice that, as calculated by molecular dynamics simulation, diffusion coefficients of triacylglycerols at

26 °C are lower in the core of emulsion droplets (6.9×10−8 to10.8×10−8 cm2/s) than in bulk oil (8.4×10−8 to 17.2 10−8 cm2/s)





(Hennere and others 2009a, 2009b). Similarly, 1 H NMR diffusion

coefficients of the acyl chains of tr iacylglycerols in marine oil andin a blend of vegetable and marine oils varied between 1×10−8

and 1.4×10−7 cm2/s. They decreased to 0.45×10−7 cm2/s in the

core of the oil droplets of freshly prepared submicron emulsionsstabilized by LMWEs (phospholipids and Tween 80) (Kabri and

others 2013). Even if the apparent reaction rates deduced fromglobal measures of oxidation are, by far, slower than such diffusion

rates, the effects of the decreased mobility of the triacylglycerol

molecules inside the oil droplets should be appreciated withregard to the most rapid and/or limiting steps of oxidation.

These contrasted results illustrate the fact that it is difficult to get

a clear picture of the effect of oil emulsification on lipid oxidation,as other paired factors affecting the reaction (such as homoge-

nization conditions, aqueous phase components, and emulsifiers)vary as well. We will see in the following how difficult it is to

deconvolute these effects to propose clear rules with regard to theinfluence of selected parameters on lipid oxidation.

The huge diversity of the dispersed model systems used inthe literature to study lipid oxidation

An impressive and increasing amount of work has been pub-

lished about the oxidative stability of emulsions for decades. Ac-cording to the ISI Web of Knowledge database, more than 100papers containing the terms “lipid, oxidation, emulsion” in their

title, abstract, or keywords were published in the year 2012 alone,which was more than an 8-fold increase as compared to 1992, for

instance.The most widely used dispersed model systems to study lipid

oxidation are O/W emulsions. In most of the studies, the oil phaseis composed of natural or synthetic triacylglycerols, which already

gives a broad range of possible molecular compositions.Edible oils are used as crude or refined commercial oils. They

accordingly contain varied concentrations of minor components(including phospholipids, antioxidants, free f atty acids (FFAs),mono- and diacylglycerols, and pigments). This is why many

studies are performed with oils separated from these minor compounds. The emulsifiers used in these studies also widely

vary, reflecting the variety of the available natural and syntheticsources, but also the level of simplification of the model.

Aqueous dispersions of FFAs or of methyl or ethyl esters of fattyacids with aqueous surfactant may also be used, as the sole lipid

phase or mixed with triacylglycerol or alkane oil (Miyashita andothers 1994, 1997; Hirano and others 1997; Okuda and others

2005; Imai and others 2008). The colloidal structures of such dis-persed systems are different from these of triacylglycerol emulsions.

Indeed, FFAs are surface-active (Waraho and others 2009), formmicelles or mixed micelles with surfactants (Lim and others 2005)

or, when mixed with a nonpolar oil, preferentially locate at the oil

droplet surface (Okuda and others 2005). When the pH is greater than their pKa, FFAs are negatively charged, which impacts thesurface charge of micelles or oil droplets, and consequently, the

electrostatic interactions with ionic species present in the aqueousphase. Other authors used, as model systems, liposomes where un-

saturated fatty acids are esterified under the form of phospholipids(Araseki and others 2002; Mozuraityte and others 2006; Lu and

others 2011). Only occasionally, more complex emulsions, suchas double or multiple emulsions, have also been investigated with

respect to lipid oxidation (O’ Dwyer and others 2013; Poyato andothers 2013).

Real or model food matrices incorporating unsaturated oils aspreformed emulsions or as oil droplets dispersed during the process

have also been investigated. Examples of food matrices range frommilk (Let and others 2004, 2007a; Sorensen and others 2007),

infant formulas (Zou and Akoh 2013), yogurt (Nielsen and others2007), salad dressing (Let and others 2007b), mayonnaise (Sorensen

and others 2010; Raudsepp and others 2014), cereal bars (Nielsenand Jacobsen 2009), cream cheese (Horn and others 2012a), or

meat-based products (Delgado-Pando and others 2010; Salminenand others 2013b).

In all these systems, the composition and purity of the oil phase

and the emulsifiers can largely vary from 1 study to another andaffect the kinetics of oxidation. However, the contents in im-purities of the materials, such as metal ions and lipid oxidation

products, are rarely known, although hydroperoxides may alreadybe present in LMWEs or in the freshly emulsified oils (Mancuso

and others 2000; Nuchi and others 2001). The oil phase fraction,the homogenization conditions, the formulation steps, and so on,

vary largely from 1 study to another. Huge differences can also befound in the physical–chemical characteristics of the systems, such

as the pH, ionic strength, type of salts, and so on, of the aqueousphase. How the composition of the aqueous phase may affect the

lipid oxidation kinetics, at least in part due to its interactions withthe interfacial layer, is detailed below. We also mention later on

the importance of the agitation conditions.The storage temperature, oxygen availability (close or open ves-

sels), the addition of oxidation initiators such as iron salts, heme

proteins, and hydrophilic or lipophilic radical initiators are, amongothers, possible variation factors. Various incubation temperatures

or oxidation catalysts have been reported in the literature. In-cubation temperature may range from typical food storage tem-

peratures (2 to 4 °C) to accelerated lipid oxidation conditions(50 to 60 °C) or higher temperatures when cooking or frying

are the targeted process. Recent studies also report lipid oxi-dation in emulsions incubated in simulated digestive conditions

(Kenmogne-Domguia and others 2012, 2014). Even though somestudies have been performed without any intentional addition of

oxidation initiator, iron has been widely used to accelerate lipidoxidation in emulsions (Donnelly and others 1998; Mei and oth-

ers 1998a, 1998b; Fomuso and others 2002a; Haahr and Jacob-sen 2008; Guzun-Cojocaru and others 2011), even if other met-

als such as copper have been sometimes encountered (Fomusoand others 2002a; Osborn-Barnes and Akoh 2003; Branco and

others 2011). Water-soluble azo initiators such as 2,2-azobis(2-amidinopropane)-dihydrochloride (AAPH) or 2,2-azobis[2-(2-

imidazolin-2-yl)propane] (ABIP) have also been shown to pro-mote efficiently lipid oxidation in O/W emulsions or fatty acid

dispersions (Hanlon and Seybert 1997; Kubouchi and others 2002;Gotoh and others 2010). Heme proteins such as myoglobin, one of

the main forms of dietary iron, efficiently promote lipid oxidation(Baron and Andersen 2002). Myoglobin has therefore been used as

a lipid oxidation catalyst in several multiphase systems (Roginskyand others 2007; Lorrain and others 2010b; Min and others 2010).

Recently, 2 studies highlighted the importance of the incuba-tion conditions and the possible interactions between initiators and

emulsifiers regarding their effects on emulsion oxidation (Bertonand others 2012b; Mosca and others 2013). In the first one, lipid

oxidation was followed in emulsions stabilized by either BLG,BCN, BSA, or Tween 20 and incubated either at 33 °C with-

out any added chemical initiator, or at 25 °C in the presenceof iron/ascorbate, metmyoglobin (MetMb), or AAPH at concen-trations chosen to obtain roughly the same rates of oxidation.

The kinetic parameters (oxygen uptake rates and the estimated lagphases) indicated that the emulsions incubated in the presence of





MetMb were characterized by an early beginning of oxygen up-take. We assumed that this rapid initiation of oxidation by MetMb

was due to its high efficiency to decompose trace hydroperox-ides and its surface activity (Berton and others 2012b). In the

other incubation conditions, lipid oxidation kinetics were clearlyaffected by certain combinations of emulsifiers and incubation

conditions. The incubation of emulsions at 33 °C without initia-tor led to 2 opposite behaviors: either the highest rates of oxygen

uptake with short lag phases (BLG- and BCN-stabilized emul-

sions) or low rates of oxygen uptake with the longest lag phases(BSA- and Tween 20–stabilized emulsions). In the presence of AAPH, oxidation behaviors of BLG- and BCN-stabilized emul-

sions also distinguished from the 2 others, but mainly by their lag phases (Berton and others 2012b), while with iron/ascorbate,

the emulsions stabilized by BLG, BSA, and BCN exhibited sim-ilar kinetics, in contrast with the emulsion stabilized by Tween

20, which was significantly more oxidatively stable. In the 2ndstudy, Mosca and others (2013) also observed different lipid ox-

idation kinetics in a Tween 80–stabilized emulsion when vari-ous free radical initiators were used to promote oxidation (the

hydrophilic AAPH; the hydrophilic 2,2-azobis[2-(2-imidazolin-2-yl) propane] dihydrochloride [AIPH]; or the lipophilic 2,2-

azobis(2,4-dimethylvaleronitrile) [AMVN]). AAPH was the mostefficient initiator, but, interestingly enough, when AAPH andAMVN were used together a synergistic prooxidant effect was

observed.This great variety of models and experimental conditions ex-

plains the difficulty of identifying generic trends regarding therules that govern lipid oxidation in dispersed systems: it is often

difficult to gather a large number of experimental results involving1 single variation factor and, in different systems, the same vari-

ation of a determined factor can lead to opposite results, due towithin-factors interactions.

Specific factors related to the oil phase that influence lipid

oxidation in emulsionsThe properties of the oil phase that affect lipid oxidation inemulsions are mostly similar to those reported in bulk oil, even

though a few specificities associated with the colloidal state of oiland molecular arrangement of the acyl chains can be pointed out.

Oil unsaturation. As observed in bulk oil, in emulsions, lipidoxidation is generally favored by an increase of the unsaturation of

the lipid material. For instance, Kiokias and others (2006) observedthat in emulsions preparedwith different dietary oils, the formation

of conjugated diene hydroperoxides increased with the LA contentof the oil. Conversely, a better oxidative stability of emulsified nat-

ural or synthetic triacylglycerols containing docosahexaenoic acylchains (DHAs) oxidized more slowly than emulsified triacylglyc-

erols containing LA (Azuma and others 2009; Gotoh and others

2010). The molecular conformation of the long-chain PUFAs inmultiphase systems, and the subsequent reduced accessibility of bis-allylic hydrogen atoms, was evoked to explain these results.

Hence, in multiphase systems, the oxidizability of marine oils,with a high content in long-chain PUFAs, does not necessarily

increase with their degree of unsaturation.Such a particular behavior of long-chain PUFAs with regard to

lipid oxidation was also reported for ethyl esters and nonesteri-fied forms of the fatty chains. Ethyl esters of DHA solubilized in

nonionic emulsifier micelles dispersed in an aqueous phase wereaccordingly found more stable to oxidation than ethyl ALA, which

was itself more stable than ethyl LA (Hirano and others 1997).Similar results were obtained with FFAs (Miyashita and others

1994, 1997). The authors suggested that the long-chain PUFAsare buried more deeply within the hydrophobic core of the mi-

celles when dispersed in an aqueous phase. They would thereforebe less susceptible to attack by aqueous phase prooxidants.

Lipophilic minorcomponents. Trace components naturally con-tained in commercial oils also modify the development of oxida-

tion in emulsified oils. First of all, antioxidants naturally present inthe oils and in the ingredients used to prepare the emulsions are

deeply involved in the oxidative stability of emulsions. For instance,

a substantial lag phase was observed for an emulsified commercialsunflower oil that contained about 700 mg α-tocopherol/kg, butnot when stripped oil was used instead (Villi ere and others 2005).

Other minor polar compounds removed from the oil phase dur-ing refining or during stripping also influence the development of

emulsified oil oxidation, due to their surface activity (Decker andothers 2010; Chen and others 2011c). The presence of FFAs in

emulsified oil was shown to increase lipid oxidation, which couldbe related to their location at the interface and the increased neg-

ative net charge of the oil droplets, which is postulated to attractmetal ions onto the oil droplet surface (Waraho and others 2009,

2011b).

Physical state of emulsified lipids. The effect of the solid fat

content on oxidation has been investigated in an emulsified oc-tadecane/methyl linolenate mixture (Okuda and others 2005).This study highlighted that the emulsion was more prone to metal-

catalyzed oxidation when octadecane was in solid state instead of in liquid state. The authors suggested that octadecane crystalliza-

tion induced migration of methyl linolenate toward the exterior of oil droplets, the unsaturated substrate, becoming more accessible

to the metal ions in the aqueous phase. More recently, the rate of transport of oxygen and free radicals from the aqueous phase to

the core of oil droplets as a function of the physical state of theoil phase was investigated (Tikekar and Nitin 2011). This study

highlighted that the solid state of the oil phase decreased onlyslightly the rate of oxygen transport as compared to the liquid

state, whereas the transport of peroxyl radicals was not affected.As previously stated by Okuda and others (2005), this poor pro-

tective effect of crystallizing the oil phase can be explained bythe expulsion of the oxidizable lipophilic compounds towards the

periphery of lipid droplets, thus exposing them more easily to theprooxidant compounds of the aqueous phase. Such an expulsion

has been clearly shown for a spin-labeled stearic acid in eicosanedroplets by electron paramagnetic resonance (Berton-Carabin and

others 2013b).

Oil-phase volume fractionThe volume fraction of the oil phase influences lipid oxidation

in O/W emulsions. When oil volume fraction increased from

10% to 30% (Osborn and Akoh 2004), from 5% to 40% (Sun and

Gunasekaran 2009), or from 5% to 30% (Kargar and others 2011),lipid oxidation decreased significantly. Kiokias and others (2006)also observed a decrease of lipid oxidation in O/W emulsions

when increasing the oil-phase volume fraction from 10% to 40%.In these studies, the size of the oil droplets varied with the oil

fraction. For instance, volume-surface mean diameter (d 3,2) of theoil droplets increased from 0.26 to 0.37 µm, or from 1.09 to 1.98

µm with increasing oil fraction from 10% to 30% v/v, dependingon the emulsifier nature and homogenization conditions (Osborn

and Akoh 2004), which corresponds to a 1.7- to 2.1-fold increaseof the total surface area developed by the oil droplets. The in-

crease in the mean droplet diameter from approximately 0.5 to 1.5

µm, when the oil-phase volume fraction increased from 10% to





40%, also corresponds to an 1.3-fold increase of the interfacial area(Kiokias and others 2006). In the work of Sun and Gunasekaran

(2009), the size of the oil droplets decreased from 1.60 to 0.80

µm or 1.10 to 0.60 µm with increasing oil fraction from 5% to

40% v/v, depending on the emulsifier concentration, which cor-responded to a dramatic increase of the total surface area from

14.7- to 16-fold. To take into account these variations of the in-terfacial area, Kargar and others (2011) normalized the amounts of

detected oxidation compounds by the total interfacial area devel-

oped by the oil droplets. This procedure amplified the differencesbetween emulsions with low and high oil volume fractions regard-ing the formation of hydroperoxides. The authors concluded that

this indicates that the droplet size had no significant effect. Theyassumed that when the oil fraction increased, the aqueous phase

fraction decreased proportionately, and so the amount of water-soluble prooxidants, such as metal ions, thereby decreasing the

number of free radicals generated per droplet. One may also no-tice that a higher oil volume fraction and/or a decreased droplet

size will lead to a decrease in the distance separating droplets.The propagation of the oxidation reaction could thus benefit

from direct exchanges of reacting lengths species (hydroperoxidesand free radicals) between neighboring oil droplets. In addition,

as previously suggested by Gohtani and others (1999), Sun andGunasekaran (2009) assumed that the better oxidative stability of the emulsions, when the oil-phase volume fraction increased from

5% to 40%, resulted from their improved creaming stability, oxida-tion being favored in the creamed phase, which is closer to ambient

air than in droplets present in the bottom of the containers, dueto the rate of oxygen diffusion. Differences were accordingly ob-

served in the rates of formation of conjugated dienes according tothe agitation mode of sunflower oil emulsions (30% w/w) (Genot

and others 2003). In agitated conditions, the type and speed of agitation determine the rate of transport of the reactants (oxygen,

free radicals, and oil droplets) and thus influence the oxidationrate.

If emulsions are stored under steady conditions, the self-diffusion of reactants and oil droplets, as affected by the viscosity of

the medium, can be a limiting factor. It decreases with increasingoil volume fraction, which therefore limits the propagation of the

reaction from one droplet to another. Unfortunately, the condi-tions of storage of the emulsions, namely, in steady or agitated

conditions, are rarely mentioned.

The Role of the Aqueous Phase in the Oxidationof Emulsified Systems

As mentioned above, not only the oil phase, but also the compo-

sition of the aqueous phase, namely, the presence of unadsorbedwater-soluble emulsifiers and other soluble biomolecules, salts,

buffers, oxidation initiators, and antioxidants, may influence the

oxidative reactions. The aqueous phase can be involved through 2main mechanisms. On the one hand, the aqueous phase may favor or inhibit the formation and diffusion of reactive species originally

distributed in this phase. On the other hand, it can modify somecharacteristics of the interfacial layer, which may either favor or prevent the migration of reactive species initially present or formed

in the aqueous phase up to the oxidizable substrate.

Unadsorbed emulsifiers can protect the lipid phase againstoxidation

Unadsorbed protein emulsifiers may strongly interact with

water-soluble reagents such as free radicals and metal ions(Donnelly and others 1998; Ponginebbi and others 1999;

McClements and Decker 2000; Hu and others 2003a, 2003b;Faraji and others 2004; Villiere and others 2005; Cheng and oth-

ers 2010). Micelles in the aqueous phase, even if there are dynamicstructures with very fast exchange rates, can also trap molecules

of various lipophilicities (antioxidants, hydroperoxides, and fattyacids), which affects their overall microlocalization and partition-

ing in the multiphase systems. The presence of these unadsorbedsurfactant or protein emulsifiers has been accordingly repeatedly

shown to greatly influence the chemical reactivity of emulsions

(Nuchi and others 2002; Richards and others 2002; Almajano andothers 2007a; Berton-Carabin and others 2012, 2013c; Panya andothers 2012).

As revealed by Table 1A, the concentration of unadsorbedemulsifiers in the aqueous phase of O/W emulsions can reach

great levels. This fraction of unadsorbed emulsifiers has been ex-perimentally shown to affect the oxidation of emulsified lipids.

For example, washed O/W emulsions oxidized faster than thecorresponding unwashed emulsions containing unadsorbed pro-

teins (Faraji and others 2004; Ries and others 2010). In the sameway, the addition after emulsification with excess emulsifiers in

the aqueous phase of emulsions delayed lipid oxidation, as com-pared to the original emulsion containing a low concentration

of unadsorbed emulsifier (Berton and others 2011b; Kargar andothers 2011) (Figure 2). This antioxidant activity of unadsorbedemulsifiers, or emulsifiers added in excess, has been attributed to

different mechanisms.

Proteins as metal chelators. The ability of proteins to act as

metal chelators or metal binders in dispersed systems has beendemonstrated for caseins and β-casein-phosphopeptide (Kansci

and others 1997, 2004; Rival and others 2001; Diaz and others2003; Hu and others 2003a, 2003b; Sorensen and others 2007),

WPI (Tong and others 2000; Faraji and others 2004), BLG (Eliasand others 2007) and NaCas (Faraji and others 2004; Villiere and

others 2005; Haahr and Jacobsen 2008; Sugiarto and others 2010;Guzun-Cojocaru and others 2011). Metal chelation would in-

hibit hydroperoxide decomposition by preventing cationic proox-idants from reaching hydroperoxides located close to the emulsion

droplet surface and by decreasing the reactivity of iron. This abil-ity of proteins to bind metal ions depends on their charge, which

is influenced by pH. Indeed, above their isoelectric point, pro-teins are anionic, hence able to bind positively charged ferrous

ions (Faraji and others 2004; Kellerby and others 2006b). Themetal-chelating activity of BCN and its hydrolysates is linked to

the presence of phosphoseryl groups. Even at low pH conditionsthese groups remain negatively charged, and unadsorbed casein

in the continuous phase of O/W emulsions keeps protecting thelipid phase against oxidation through its metal chelating activity

(Hu and others 2003a).

Proteins as free radical scavengers. Proteins can trap free rad-

icals produced in the aqueous phase and/or issued from lipid ox-idation. Due to radical transfer reactions, proteins are known to

exhibit free radical scavenging activity and to have antioxidant ef-fects (Roubal 1970; Dean and others 1997; Stadtman and Levine

2003; Schaich 2008). According to some authors, this activitywas involved in delaying oxidation in emulsions stabilized by ca-

seins, NaCas (Villiere and others 2005; Clausen and others 2009),and potato proteins (Cheng and others 2010; Habeebullah and

others 2010). At neutral pH, soy protein isolate (SPI) exhibiteda better antioxidative capacity than casein, which was itself moreeffective than WPI (Faraji and others 2004). The antioxidant ac-

tivities of SPI and WPI in the aqueous phase were attributed tothe sulfhydryl groups of the proteins and to the presence of an





0

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0 20 40 60 80

Time (h)

m m o l O 2

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m m o l O 2

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-casein, no excess

-casein, excess

Tween 20, n o excess

Tween 20, excess

Figure 2–Lipid oxidation, measured through monitoring headspace oxygen consumption, in rapeseed oil emulsions stabilized by various emulsifiers (A,β-casein; B, Tween 20) without or with excess emulsifiers added to the aqueous phase. The pH of the aqueous phase was 6.7, and lipid oxidation wasinitiated by an equimolar complex of FeSO4 and EDTA (final concentration in emulsion, 200 µM of each). Adapted with permission from Berton,Ropers, Viau and Genot (2011). Contribution of the interfacial layer to the protection of emulsified lipids against oxidation. J Agric Food Chem59:5052–61. Copyright (2011) American Chemical Society.

isoflavone associated with SPI (Faraji and others 2004). Elias andothers (2005, 2006) showed that tryptophanyl, cysteyl, methionyl,and tyrosyl residues are able to scavenge aqueous phase radical

species, thereby leading to improved oxidative stability in O/Wemulsions. Methionyl residues could reduce hydroperoxides to

low-reactive species (Garner and others 1998). However, the abil-ity of amino acid residues to scavenge free radicals also depends on

the tertiary structure of the protein. When not sufficiently surface-exposed, the residues may be physically unable to participate in

free radical scavenging (Elias and others 2005). For example, theaddition of a mixture of Tween 20 and WPI in the aqueous phase

of an emulsion stabilized with Tween 20 decreased lipid oxidation,whereas the addition of either Tween 20 or WPI did not moderate

oxidation development (Donnelly and others 1998). According tothe authors, Tween 20 altered protein conformation and increased

the accessibility of certain amino acid residues.

Binding of secondary lipid oxidation products by proteins. Inmultiphase systems containing oxidizing lipids, proteins can, in

turn, undergo a range of chemical modifications. Such modifi-cations are partly due to secondary oxidation products that par-

tition between the oil and aqueous phases and can covalentlybind to proteins. Typically involved reactants here are nucle-

ophilic amino acids (such as histidine and lysine) and unsatu-rated aldehydes (including malondialdehyde, 4-hydroxyhexenal,

and 4-hydroxynonenal), which may form Schiff base complexesand Michael adducts (Leaver and others 1999a; Genot and others

2003, 2013; Meynier and others 2004; Schaich 2008; Lund and

others 2011; Berton and others 2012c). Noncovalent interactionsmay also be established, for instance, hydrophobic interactionsbetween volatile oxidation products and the hydrophobic parts

of proteins. Such interactions between proteins and lipid oxida-tion products can result in substantial alteration of the functional

properties of proteins, especially those adsorbed at the oil–water interface. They may also lead to underestimation of the amount of

formed lipid oxidation products, particularly if the assay directlyrelies on the partitioning of volatile oxidation products between

the emulsion and its headspace without taking into account their binding on the protein. However, quantitative data regarding the

extent and significance of such an underestimation are not avail-able, to our knowledge.

Colloidal structures formed by surfactants in excess. At con-centrations above the CMC, surfactant molecules in the aqueousphase are present as monomers, micelles, mixed micelles, or very

small droplets in which small amounts of lipids are incorporated (aso-called Winsor I system [Winsor 1948]). These colloidal struc-

tures present in the aqueous phase of emulsions have been proposedto alter the oxidation process in various ways. Excess Tween 20 in

the aqueous phase of Tween 20-LA emulsions had a protective ef-fect against the oxidation of LA (Ponginebbi and others 1999). The

authors proposed that some LA was incorporated in the surfac-tant micelles, and thus, was more protected against oxidation than

when located at the droplet interface because of the presence of acompact noncharged layer separating the negatively charged sub-

strate from the positively charged iron dissolved in water. Micellesof Brij 76 also inhibited lipid oxidation of corn O/W emulsions,

which was attributed to the solubilization of lipid hydroperoxidesby surfactant micelles, making them incapable to propagate oxi-dation in the droplets, on the basis of experiments showing that

the micelles are able to displace hydroperoxides toward the aque-ous phase (Nuchi and others 2002). Similarly, excess Tween 20

added in the aqueous phase of a rapeseed O/W emulsion delayedlipid oxidation and decreased the reaction rate (Berton and oth-

ers 2011b). This explanation relative to a possible sequestration of oxidation-sensitive material, or of intermediate reaction products,

proposed by several authors, fits well with the presented data. It isuseful to tentatively predict the effect of unadsorbed surfactants on

oxidation in emulsions. One may, however, notice that micelles are

highly dynamic systems characterized by rapid rates of moleculeexchanges with the external medium (other micelles, aqueousmedium, or possibly oil droplets), and by rapid diffusion rates of

the individual molecules as compared to the rates of most reactionsinvolved in the oxidation pathways (Denisov 1995; Romsted and

Bravo-Diaz 2013). This led authors to conclude that the chemicalreactivity of colloids, and generally of emulsions, could be mod-

eled by using the pseudophase kinetic model, with the assumptionthat the distribution of reactants in emulsions is rapid as compared

to the timescale of oxidation, and that the system can be consideredin dynamic equilibrium. The concept has been developed for a

better understanding of the efficiency of antioxidants in modelemulsions (Romsted and Bravo-Diaz 2013). It can give some





insights on the effects of ionic surfactants on oxidation of emul-sions (Gu and others 2013). However, applications of this model

to real food emulsions, and to explain the effect of unadsorbedsurfactants on oxidation kinetics, have not been proposed yet.

Conversely to the above-mentioned protective effects, Donnellyand others (1998) noticed that the addition of Tween 20 in the

aqueous phase of a Tween 20–stabilized emulsion increased lipidoxidation as determined by both lipid peroxides and TBARS val-

ues. The authors suggested that the increased oxidation rates could

result either from the displacement to the surfactant micelles fromthe emulsion droplets of antioxidants, that became less efficient toinhibit oxidation, or from the solubilization in the micelles of lipid

oxidation products where they could have increased interactionswith prooxidants in the aqueous phase. They also thought that

the surfactant itself would be an additional source of peroxides.One may also evoke the possible influence of unadsorbed sur-

factants on direct exchanges of hydrophobic molecules betweenneighboring droplets. Such exchanges would favor the propaga-

tion of the reaction by transfer of reactive species from dropletsto droplets. This question of interdroplet molecular exchanges has

been addressed on model emulsions made of different pure lin-ear alkanes. In such emulsions stabilized by Tween 20, complete

mixing of the oils in the droplets took from 4 to over 7 d, de-pending on the presence of additional surfactant in the aqueousphase, which is a timescale in the same range as oxidation kinetics

in emulsions (McClements and others 1992; Samtlebe and others2012). The presence of surfactant micelles was assumed to facilitate

the transfer of molecules from 1 droplet to another. The rate of transfer decreased with increasing NaCl concentration and droplet

diameter (lower surface area) (McClements and Dungan 1993). Itwas drastically increased by rehomogenization of mixed emulsions

(Elwell and others 2004). More rapid exchanges (<10 min) of hydrophobic fluorescent dyes were observed for toluene or octane

emulsions stabilized by Triton X-100 (Malassagne-Bulgarelli andMcGrath 2013). It was attributed to a transient hole mechanism, or

to a continual reversible coalescence (fusion–fission mechanism).By comparison, it was recently demonstrated that the content of

droplets of mayonnaises prepared separately with saturated and un-saturated oils did not mix together (Raudsepp and others 2014).

The oxidation kinetics of the mixed mayonnaises differed fromthose observed for the mayonnaise prepared with a mixture of

both oils, and from those in the mayonnaise made of the unsatu-rated oil. It could be relevant to perform similar experiments with

emulsions containing various concentrations of surfactants in theaqueous phase to test the above hypothesis of facilitated transfers of

reacting species by micelles, and their effect on oxidation kinetics.Interpreting the effect of the concentration of emulsifiers

on lipid oxidation in O/W emulsions is also difficult, becauseincreasing the total emulsifier concentration (without changing

the emulsification procedure) tends to decrease the droplet size(Donnelly and others 1998; Ponginebbi and others 1999; Kiokias

and others 2006; Sun and Gunasekaran 2009). To get similar droplet size distributions with varied emulsifier concentrations,

the emulsification procedure and/or the input energy should beadapted, which may, in turn, influence oxidation.

Other unadsorbed biopolymers can delay lipid oxidation. Theeffect of polysaccharides incorporated in the aqueous phase of

O/W emulsions on lipid oxidation has been investigated by sev-eral authors (Shimada and others 1994, 1996; Sirendi and others1998; Matsumura and others 2000; Kishk and Al-Sayed 2007; Sun

and others 2007; Chen and others 2010b). Some of the testedpolysaccharides showed antioxidant effect, for instance, xanthan,

bean gum, glucomannan, curdlan, gum arabic, and pectin. This ef-fect was generally attributed to the metal-chelating and free-radical

scavenging properties of the polysaccharides.

pH, salts, chelators, and reducing agents modulate lipid ox-idation by interacting with the reactants in the aqueousphase and at the interface

The pH influences the charge, solubility, partitioning, redoxstate, and chemical stability of major actors in oxidative reactions

such as metal ions, antioxidants, and biopolymers (proteins andcharged polysaccharides). For instance, the net charge of proteins

is negative when they are dissolved at a pH above their isoelectricpoint (pI), and the reverse is observed at pH < pI. Electrostatic

interactions at the molecular or colloidal levels, therefore governedbypH, can modify oxidation kinetics. The influence of pH on lipid

oxidation in O/W emulsions has been investigated with contrastedresults.

In some studies, the oxidativestability of emulsions prepared atneutral pH was better than in emulsions at acidic pH. Haahr and

Jacobsen (2008) showed that, irrespective of the emulsifier type(Tween 80, ester of citric acid and fatty acids (Citrem), NaCas or

lecithin) and iron addition, the formation of peroxides and volatiles

was faster at pH 3.0 than at pH 7.0. An increased oxidation, whendecreasing pH, was also found in Tween 80 and Citrem-stabilizedemulsions (Sorensen and others 2008), and in sodium dodecyl sul-

fate (SDS)-stabilized emulsions (Mei and others 1998b). Similar results were found by Branco and others (2011): in SDS-stabilized

emulsions, changing the pH from 3.0 to 7.0 reduced the decompo-sition of primary oxidation products (peroxides) and the formation

of secondary oxidation products (TBARS). The higher solubilityof iron at low pH levels was suggested to be partly responsible for

the increased oxidation. In addition, in the presence of both chela-tors and reducing agents, a low pH could promote the activation of

metal ions due to their displacement from the chelators that madethem inactive at higher pH. Oxidation was also favored at pH4.0 as compared to pH 6.5 in BSA-stabilized emulsions (Villi

`ere

and others 2005; Villiere and Genot 2006). This could result fromthe presence of high protein concentrations in the aqueous phase

(14.1 g/L, Table 1A), as electrostatic interactions between metalions and unadsorbed proteins would have screened any effect of

electrostatic interactions at the interface; but differences in proteinconformation and accessibility of radical scavenging amino acids

were also considered by the authors as a possible explanation.

In contrast, other studies provided evidence that a neutral pHfavored lipid oxidation as compared to acidic pH. In emulsionsstabilized with LMWEs such as Tween 20 or SDS, a prooxidant

effect was observed at the highest tested pH values (Huang andothers 1996b; Mancuso and others 1999). It was hypothesized that

the low iron solubility at pH 7.0 could result in the precipitation of

metal onto the lipid droplet interface, thereby bringing iron intocloser contact with the lipid phase compared to pH 3.0 (Mancusoand others 1999). Osborn-Barnes and Akoh (2003) found that

copper-catalyzed lipid oxidation in WPI-stabilized emulsions waslower at pH 3.0 than at pH 7.0. In accordance with Donnelly

and others (1998), they suggested that at pH 7.0, the negativelycharged WPI-stabilized droplets were able to attract positively

charged transition metals, which promoted lipid oxidation. Theantioxidants used in this study (α-tocopherol and citric acid) were

also suggested to be more effective at pH 3.0 than at pH 7.0,a low pH enhancing the hydrogen-donating capacity of the an-

tioxidants (effect on their redox potential). Lipid oxidation alsoproceeded faster at pH 6.7 than at pH 3.0 in BLG- and Tween





20-stabilized emulsions containing low amounts of unadsorbedemulsifiers (Berton and others 2011b). As Tween 20–stabilized

emulsions exhibited a ζ -potential close to zero independently of pH, differences in electrostatic interactions at the interface can be

hardly relied on to explain the changes in the oxidation kinetics.As phosphoric acid was used to prepare the pH 3.0 buffer, instead

of pH 6.7 PIPES buffer, the acid could have interacted with metalions in the aqueous phase.

Interestingly, when WPI-stabilized emulsions were washed and

their aqueous phases replaced by the buffer solutions, the emulsionprepared at pH 7.0 oxidized dramatically faster than the pH 3.0emulsion, whereas the differences were moderate in the emulsions

containing unabsorbed proteins in the aqueous phase (Faraji andothers 2004). These data stress the importance of the interactions

between the intrinsic effects of pH on oxidation and the effects of unadsorbed proteins.

To summarize, in multiphase systems, pH may have a positive or negative effect on lipid oxidation through a wide range of under-

lying mechanisms. This effect, which depends on the modulationof the interactions between reactants at the interface, depends,

among others, on the composition of the system.

Salt (NaCl) and other inorganic ions dissolved in the aqueous

phaseaffect the oxidativestabilityof O/Wemulsions. In the pres-ence of low concentrations of iron (50 µM), NaCl (0.17 M) wasshown to decrease lipid oxidation in emulsions by less than 20%,

which was attributed to a decrease of the binding of metal ions atthe droplet surface due to shielding of the surface charge by the salt

ions, or by formation of iron–chloride complexes (Mei and others1998b). Conversely, in the presence of higher iron amounts (500

µM), the same concentration of NaCl increased lipid oxidation,some increase of the catalytic activity of iron being held responsible

(Mei and others 1998a). The prooxidant effect of NaCl (0.5 M)in the aqueous phase of emulsions was also observed in the pres-

ence of copper (Osborn-Barnes and Akoh 2003). Mozuraityteand others (2006) investigated the effect of different salts (NaCl,

KCl, NaNO3

, CaCl2

, Na2

SO4

, K2

SO4

, Ca(NO3

)2

, Mg(NO3

)2

,and NaH2PO4) on the oxidation of liposomes catalyzed by iron.

Neither the tested cations (Na+, K+, Ca2+, and Mg2+) nor theSO4

2− and NO3− anions influenced lipid oxidation, whereas Cl−

and H2PO4− anions decreased the oxidation rate of the liposomes.

The effect of phosphate salts could result from their iron chelation

properties. In the case of chloride salts, the authors brought thatthe rate of O2 consumption decreased linearly with increasing the

ζ -potential of liposomes.

Buffers may interact in the radical chain reactions taking placein the aqueous phase. Due to the iron-chelating properties of phosphate ions, phosphate buffers have an antioxidant activity

(Yoshimura and others 1992). Similar activity is found with citratebuffers due to the chelating properties of citric acid (Fomuso and

others 2002a; Serfert and others 2009), even though its chelatingpower is less than for EDTA (Cho and others 2003; Djordjevic and

others 2004). Other buffers, for example, Tris, Hepes, or Mops,may also intervene in lipid oxidation through their free-radical

scavenging activity (Tadolini 1987; Fiorentini and others 1989; Yoshimura and others 1992).

The aqueous phase of O/W emulsions can contain moleculeswith metal-chelating activities, which largely prevent lipidoxidation. The antioxidant effect of ethylenediaminetetraaceticacid (EDTA) against metal-catalyzed oxidation has extensivelybeen described (Jacobsen and others 2001; Cho and others 2003;

Djordjevic and others 2004; Nielsen and others 2004; Villiere andothers 2005; Lee and Decker 2011; Polavarapu and others 2012).

This is attributed to its ability to chelate metal ions present in theaqueous phase and to keep them away from the interface. EDTA

or sodium tripolyphosphate added to O/W emulsions were evenshown to induce the transfer of iron initially contained in the oil

phase to the aqueous phase, thus decreasing lipid oxidation (Choand others 2003).

Yet, the antioxidant efficiency of EDTA is dependent on sev-eral factors. When used in concentrations sufficient to inhibit the

iron–oil droplets association, EDTA has a maximum antioxidant

efficiency (Mei and others 1998a; Lee and Choe 2011). Con-versely, an equimolar mixture of EDTA and ferrous iron favorsthe development of lipid oxidation in O/W emulsions (Mahoney

and Graf 1986; Samokyszyn and others 1990; Berton and oth-ers 2011b, b). The equimolar complex allows the concomitant

presence of different valence states of iron, which is necessary toinitiate efficiently lipid oxidation (reactions 3, 4) (Minotti and Aust

1992; Cheng and Li 2007), especially at neutral pH where it hasa free coordination site available for redox reaction (Mahoney and

others Graf; Haahr and Jacobsen 2008). The efficiency of EDTAcan also be affected by emulsifiers (Haahr and Jacobsen 2008). In

Tween 80- or Citrem-stabilized emulsions, EDTA decreased theformation of volatile oxidation compounds, whereas it increased

their amounts in emulsions stabilized by NaCas or lecithin. Ithad, however, a limited effect on the peroxide formation. The au-thors suggested that the chelator may not necessarily prevent the

formation of peroxides in the initial oxidation stages, but rather the metal-catalyzed breakdown of already present peroxides. The

efficiency of EDTA is also conditioned by pH.

Reducing compounds such as ascorbate can either favor orprevent lipid oxidation. This depends to a large extent on thepresence of iron and on the ascorbate/iron ratio. When present

in low enough concentrations (about equimolar ratio), ascorbatebehaves as a prooxidant in the presence of iron because of its ability

to reduce Fe3+ to Fe2+ (Buettner and Jurkiewicz 1996; Halliwell1996). Ferrous iron can then be involved in the formation of ROS

(reactions 3 and 5) and in the decomposition of hydroperoxides(reaction 7). Accordingly, ascorbate favored oxidation in fish-oil-

enriched mayonnaise and in SDS-stabilized emulsions (Jacobsenand others 2001; Branco and others 2011). Conversely, when

ascorbate concentration exceeds to a large extent that of metals,ascorbate has an antioxidant role through the direct scavenging of

hydrophilic free radicals (Buettner and Jurkiewicz 1996; Halliwell1996).

Interfacial Properties Affecting Lipid Oxidation inEmulsions

The interface is the place where unsaturated acyl chains and thechemical species dissolved in the aqueous phase, such as diffusing

atmospheric oxygen and hydrophilic pro- and antioxidants, get

into contact. Hence, the interfacial region is postulated as beingthe critical area where the oxidation of the lipid phase is pro-moted (Labuza 1971). Indeed, the formulation strategy and the

partitioning of the emulsifiers are cr itical to highlight unambigu-ously the effect of interface. If the emulsifier concentration in the

aqueous phase is high, as was presumably the case in a large num-ber of studies analyzed on Table 1A, unadsorbed emulsifiers may

hide the contribution of the emulsifiers located at the interface tolipid oxidation. In these conditions, we tried to reinterpret data in

taking into account this parameter. To unambiguously assess therole of adsorbed emulsifiers on lipid oxidation, emulsions shall be

formulated to minimize the concentration and fraction of unad-sorbed emulsifiers, and hence their contribution to the oxidative





fate of the system (Donnelly and others 1998; Faraji and others2004; Berton and others 2011b, 2012b).

The contrasted effects of interfacial area and droplet sizeon lipid oxidation

Depending on the emulsification conditions, average dropletsizes and droplet size distributions can deeply vary, even for a

similar overall composition of the emulsions. For a same oil volumefraction and total amount of emulsifier, a smaller droplet size means

more droplets, more interfacial area, and more adsorbed and lessunadsorbed emulsifier. The partitioning of other surface-active

species, including amphiphilic antioxidants, is accordingly affected.Emulsification itself, which requires a large energy input, may

produce free radicals. Hence, it is difficult to unambiguously findevidence and interpret the data addressing the effect of droplet

size on the oxidative stability of O/W emulsions. In that respect,the findings reported in studies dealing with this question are

summarized in Table 2.Some results showed that increasing the oil droplet size led

to a better oxidative stability of emulsions (Gohtani and others1999; Lethuaut and others 2002; Azuma and others 2009; Lee and

others 2011). This was usually related to the change in interfacial

area. A smaller droplet size corresponds to a larger interfacial area,which is expected to favor the contacts between the oil phaseand the water-soluble prooxidant compounds (free radicals, met-

als) and oxygen diffusing through the aqueous phase. It also im-plies a higher ratio of oxidizable fatty acid chains located near

the interface to fatty acid chains embedded in the hydrophobiccore of the droplets (Schuster and others 1995; Coupland and

McClements 1996). As the increase of oxidation rate was lesser than the increase of the surface area in the finest emulsions,

Lethuaut and others (2002) proposed that the protein (BSA)present at the interface had a protective effect, which would

have partly counterbalanced the greater accessibility of unsatu-rated fatty acids to the aqueous prooxidants. Accordingly, in thesame emulsions, protein fluorescence decreased faster with the

smaller droplets, which could be related to the protective effectagainst lipid oxidation of the adsorbed proteins (Rampon and oth-

ers 2001). Additionally, in these emulsions, the estimated concen-trations of unadsorbed emulsifiers ranged from 96.8% (19.4 g/L)

for the largest droplet size to 35.6% (7.1 g/L) for the small-est droplet size (Table 1A). The protective effect of the un-

adsorbed proteins, if any, more limited in the finest emulsionthan in the coarsest, should also be taken into account to de-

convolute the effects of droplet size on oxidation. In the workof Gohtani and others (1999), the estimated fraction of unad-

sorbed emulsifier (sucrose ester, about 10 g/L) was very high(>99%, Table 1A), whatever the droplet size (3.4 or 6.4 µm).

The difference in oxidative stability according to droplet sizes

can thus hardly be explained by differences in concentrations of unadsorbed emulsifiers. The larger surface developed with smaller droplets may have favored lipid oxidation. However, in this study,

the oil phase was made of nonesterified DHA. FFAs are am-phiphilic and surface-active molecules that locate preferentially at

the interface, which could have emphasized any interface-relatedeffect, even though aqueous dispersions of long-chain PUFAs are

not very prone to oxidation due to the molecular conformationof the acyl chain (Miyashita and others 1994, 1997). Lee and

others (2011) also detected higher amounts of TBARS dur-ing the storage of a menhaden oil nanoemulsion (droplet

size = 66 nm) than in an emulsion of 0.325 µm aver-age droplet diameter. The amount of WPI used to prepare

both emulsions was varied and, according to our calculation(Table 1A), the fraction of unadsorbed emulsifier was about 89%

(8 g/L) of in the nanoemulsion compared with 55% (0.2 g/L) inthe largest droplet emulsion. Yet, lipid oxidation was favored in

the small-droplet emulsion, which could indicate a greater im-portance of the developed surface area (multiplied by about 5 in

the nanoemulsion) over the unadsorbed protein emulsifier in thissystem.

Perhaps, surprisingly, a few studies did not report any signifi-

cant effect of the droplet size on lipid oxidation in O/W emul-sions (Osborn and Akoh 2004; Dimakou and others 2007). Finally,some papers report a better oxidative stability of O/W emulsions

with smaller droplets than with larger ones. These examples refer,for instance, to purify fish-oil-based emulsions stabilized by su-

crose esters (Nakaya and others 2005), methyl linoleate emulsifiedwith decaglycerol monolaurate (Imai and others 2008), LA with

WPI of Na-Cas (Ries and others 2010), sunflower oil with WPI(Atares and others 2012), or fish oil with sucrose or polyglyc-erol esters (Azuma and others 2009). According to our calculation

(Table 1A), in the emulsions prepared by Nakaya and others (2005),Imai and others (2008), and Atares and others (2012), unadsorbed

emulsifiers represented more than 80% of total emulsifier (more

than 10 g/L) independently of the droplet size. Yet, the authorsproposed that their results could be explained by a protective effectprovided by the interface against oxidation of the lipid phase. For

example, Nakaya and others (2005) and Imai and others (2008)assumed that the hydrophobic tail of surfactants would induce

a wedge effect, resulting in a lower mobility of the oxidizablefatty acids in the droplets. Ries and others (2010) observed that

at high protein concentrations, the antioxidant effect of proteinsin the emulsions overcame the effects of droplet size and protein

type. They attributed the better oxidative stability of emulsionswith small droplets, as compared to emulsions with large droplets,

to both a physical barrier effect of the interfacial protein layer and an antioxidant activity of interfacial proteins. These proteins

were presumed to undergo dynamic exchanges with the proteinmolecules in the continuous phase (Ries and others 2010). It can

be also noticed that Azuma and others (2009) obtained oppositeeffects of droplet size when they prepared emulsions of fish or

soybean oil, which was assumed to result from a protection of DHA located near the interface due to the penetration of water

close to the acyl chains of the long-chain PUFAs belonging tothe triacylglycerols of the fish oil, inhibiting the abstraction of bis-

allylic hydrogen by free radicals; the wider the interface layer, thelower the oxidation. Conversely, linoleic acyl chains of soybean

triacylglycerols would not benefit from this protection linked tothe molecular conformation and dynamics of the fatty acids.

These contrasted results obtained with different dispersed sys-tems, and their various interpretations by the authors, underline

the fact that droplet size and interfacial area probably influencelipid oxidation in O/W emulsions through several mechanisms

depending on various structural and compositional parameters of the systems. The underlying mechanisms are far from completely

understood, and their effects far from quantified. It can be alsoconcluded that the frequent apparent discrepancy between stud-

ies could originate from the lack of full characterization of thesystems.

Surface charge is generally considered as a main parameterin the oxidative stability of O/W emulsions

The electriccharge of theinterface is determined bythe compo-sition of the droplet interfacial layer and the pH and ionic strength





Table 2–Effects of droplet size on lipid oxidation in O/W emulsions as observed in several studies. Data of droplet size indicated in bold correspond tothe best oxidative stability for a given study.

Oil Estimated Mean droplet Mean droplet size Droplet sizephase Aqueous unadsorbed of the finest of the coarser effect size

Reference (fractiona) phase Emulsifier fraction (%)b emulsionc(µm) emulsiond (µm) effect

Lethuaut andothers (2002)

Sunflower oil(30%v/v)

pH 4.3 BSA 36 to 97 0.40 8.1 Better oxidativestability withlargerdroplets(S < L)

Gohtani andothers (1999) DHA, freefattyacid (1%) Xanthan solution(0.5%) Decaglycerolmonostearate 99 to 99 3.4 6.4 S < L

Lee andothers(2011)

Fishoil(0.5%) Phosphate buffer,pH 7.0

WPI 89 to 55 0.07 0.33 S < L

Azuma and others(2009)

Soybean oil(0.5%)

Nonbufferedwater

Sucrose ester orpolyglycerolester

nd 6.6 – 7.4 37.2 to 37.5 S < L

Osborn andAkoh(2004)

Triglyceride oile

(10 or 30%)Phosphate buffer,

pH 7.0WPI or sucrose

esters79 to 71 1.1 2.7 No significant

effectDimakou and

others (2007)Sunflower oil

(30%)Nonbuffered

waterNaCas nd 0.67 3.2 No significant

effectRies and others

(2010)Linoleic acid

(10.6%)Nonbuffered

waterNaCas o r WPI 0 t o 95 t o 29 t o 98 0.31 0.65 Better oxidative

stability withsmallerdroplets(S > L)

Nakaya and others(2005)

Purified fish oil(10%)

Citrate-phosphatebuffer, pH 6.6

Sucrose esters 88 to 99 0.80 12.8 S > L

Azuma and others(2009) Fishoil(0.5%) Nonbufferedwater Sucrose ester orpolyglycerolester

nd 6.6 to 7.4 37.2 to 37.5 S > L

Imai andothers(2008)

Methyl linoleate(0.75% v/v)

Phosphate buffer,pH 7.4

Decaglycerolmonolaurate

88to99 0.02 8.0 S > L

Atares and others(2012)

Sunflower oil(20%v/v)

Phosphate buffer,pH 7.0

WPI 82 to 99 0.3 to 1.2 45 to 51 S > L

aOilfractionis expressedin g/100g unlessotherwise stated.bEstimated unadsorbedfractionsfor the smallest and largest droplet sizes,respectively (seeTable 1A).cDroplet sizesare reported as volume-surface mean diameter (d 3,2).eCanolaoil transesterified withcaprylicacid.nd: not determined because full required informationnot available.

of the aqueous phase. It has long been considered as one of themain factors controlling lipid oxidation in emulsions (McClements

and Decker 2000). In some studies, lipid oxidation being less in-tense when the droplets were positively charged than in the reverse

situation, it was assumed that the positive surface charge repelledpositively charged prooxidant metal ions (Hu and others 2003a;

Kellerby and others 2006b). Negatively charged droplets wouldattract the charged metal ions and favor oxidation, as observed

in emulsions stabilized by anionic, cationic, and nonionic surfac-tants (Mei and others 1998b; Mancuso and others 1999; Choi

and others 2010) (Figure 3). However, other findings contradictthis statement. For instance, a WPI-stabilized emulsion with a

net charge twice greater than that of a casein-stabilized emulsionhad a worse oxidative stability than the latter emulsion (Hu and

others 2003a). Similarly, BSA-stabilized emulsions were found tooxidize faster at pH 4.0, where BSA is positively charged, than at

pH 6.5 where it is negatively charged (Villiere and others 2005;

Villiere and Genot 2006). The anionic surfactant Citrem, whichgave a strongly negatively charged interface, decreased oxidationsignificantly in emulsions in comparison with nonionic Tween

surfactants (Sorensen and others 2008; Berton and others 2011b),which was attributed to the metal-chelating properties of citric

acid esters (Sorensen and others 2008). Similarly, when emulsionswere prepared with various emulsifiers, mostly located at the inter-

face, the oxidative stability of emulsions did not correlate with the

ζ -potentials of the droplets (Figure 4) (Berton and others 2011b).

These findings show that the surface charge of the interfaceis not systematically such a main parameter controlling the ox-

idative stability of O/W emulsions. First, interactions other thanelectrostatic interactions (for instance, chelation or free radical

Figure 3–Lipid oxidation, measured through monitoring the formation of conjugated diene hydroperoxides, in corn oil emulsions stabilized byvarious surfactants: the anionic surfactant sodium dodecyl sulfate (SDS);

the nonionic surfactant Brij 35; and the cationic surfactantdodecyltrimethylammonium bromide (DTAB). The pH of the aqueousphase was 4.0, and lipid oxidation was initiated by ascorbate (150 µM)and FeCl3 (50 µM). Reprinted from Food Chemistry, 61, Mei, McClements,Wu & Decker, Iron-catalyzed lipid oxidation in emulsion as affected bysurfactant, pH and NaCl, 307–12, Copyright (1998), with permission fromElsevier.

scavenging) may exist between the droplet surface and prooxidantcompounds, and limit the oxidative reaction even when metal ions

are bound to the interface. Second, changing the pH in order toreverse the charge of proteins may result in side effects (such as the

modification of the solubility of metals), which also affect the ox-idative reactions. The change of pH also affects the electric charge





Figure 4–An example where the oxidative stability of emulsions does not correlate with the droplet surface charge ( ζ -potential, mV). The parameterused to compare the oxidative stability of the different emulsions is the time necessary to reach half the maximum level of oxygen consumption ( t 1/2).Lipid oxidation was initiated by an equimolar complex of FeSO 4 and EDTA (final concentration in emulsion, 200 µM of each). Tw20 and Tw80 standfor Tween 20 and Tween 80, respectively; other abbreviations are explained in the body text. Adapted with permission from Berton, Ropers, Viau and

Genot (2011). Contribution of the interfacial layer to the protection of emulsified lipids against oxidation. J Agric Food Chem 59:5052–61. Copyright(2011) American Chemical Society.

of the unadsorbed protein fraction. A strongly negatively chargedprotein will not only attract metal cations at the interface; the

unadsorbed fraction will also electrostatically interact with metalcations, which may prevent them from rapidly reaching the oil

droplets. Thus, the lack of an effect of the droplet surface chargeon lipid oxidation in some conditions may be due to the contri-

bution of unadsorbed proteins (Table 1A).

How can the characteristics of single-layer interfacial filmsmodulate lipid oxidation in O/W emulsions?

The thickness and packing density of LMWE-stabilizedinterfaces may modulate the accessibility of the lipid phase towater-soluble prooxidants. The effect of the size of surfactants’

polar headgroup on lipid oxidation in emulsions has been inves-tigated in O/W emulsions stabilized by the nonionic surfactants

Brij 76 (polyoxyethylene 10 stearyl ether) and Brij 700 (poly-oxyethylene 100 stearyl ether), which differ in the number of

polyoxyethylene units by a factor of 10. Both emulsions were madewith equal molar amounts of surfactants. Lipid oxidation was faster

in the emulsions stabilized by Brij 76 (smaller headgroup), indi-cating that a larger surfactant headgroup decreases lipid oxidation

(Silvestre and others 2000). This protective effect could also arisefrom the ability of polyoxyethylene units to scavenge free radicals

(Kerwin 2008).

The influence of the size of the hydrophobic tail group of emulsifiers was investigated on emulsions stabilized by Brij-laurylor Brij-stearyl (Chaiyasit and others 2000). Lipid oxidation was

slightly lowered in the Brij-stearyl-stabilized emulsions as com-pared to the Brij-lauryl-stabilized emulsions. The authors sug-

gested that the thicker barrier provided by the longer hydrophobictail groups could prevent free radicals from reaching the fatty acids

in the lipid core. A better packing of the surfactant tail groupswith increased carbon numbers preventing the transfer of free rad-

icals to the oil droplets and moving away the unsaturated fattyacids from the interfacial region was also suggested (Chaiyasit and

others 2000). More recently, Salminen and others (2013a) com-pared lipid oxidation in fish-oil-based emulsions stabilized by high-

melting (HM) lecithin or low-melting (LM lecithin). Both kinds of lecithins differed by the length and unsaturation of their fatty acid

chains (mostly stearic acid for the HM lecithin, and mostly LA for the LM lecithin). No large difference in lipid oxidation between

emulsions with both kinds of lecithins was recorded when the oilphase was pure fish oil. However, when fish oil was mixed with

tristearin before emulsification, the emulsions were more resistantto lipid oxidation (as monitored by hydroperoxide and propanal

formation) when the emulsifier was HM lecithin, as compared toLM lecithin. The authors suggested that HM lecithin could act asan interfacial crystallization template for tristearin, which would

result in the formation of an interfacial shell that would physicallyprotect the core fish oil against oxidation.

Phospholipids exert antioxidant activities through severalmechanisms. Phospholipids primarily act as synergists in rein-

forcing the antioxidant activity of phenolic compounds becauseof their metal-chelating properties (Frankel 2005). Other pos-

sible mechanisms include reactivity of amine functions (Lu andothers 2011, 2013) and quenching of singlet oxygen (Lee and

Choe 2008). Emulsions stabilized by lecithins were accordinglyless oxidizable than emulsions stabilized by sucrose esters or mono-

diacylglycerols (Fomuso and others 2002b) or by Tween 80 or Citrem (Haahr and Jacobsen 2008). Similarly, Helgason and others

(2009) showed that lecithin can also protect lipophilic compounds

such as β -carotene against oxidative degradation in O/W emul-sions, as compared to Tween emulsifiers. Recently, Pan and others(2013) observed that the permeation of peroxyl radicals through-

out the oil–water interface in emulsions was reduced when oildroplets were stabilized by lecithin, as compared to Tween 20.

Interestingly, such a barrier effect did not apply anymore whenlecithin had preliminarily been oxidized, suggesting that it is pri-

marily due to a chemical barrier provided by fresh lecithin. Theantioxidant activity of lecithins is, however, pH-dependent. Car-

denia and others (2011) observed an antioxidant effect of phos-pholipids (synthetic PC and PE with various acyl chains) in O/W

emulsions mainly stabilized by Tween 20 at pH 7.0. But at pH 3.0,phospholipids exerted a prooxidant effect. Though the antioxidant





mechanism of phospholipids at neutral pH was not clearly eluci-dated, the authors suggested the formation of protective physical

structures within the oil phase, or a metal-chelating activity. Inemulsions stabilized by low amounts of caseinate, Garcıa-Moreno

and others (2014) observed an improved oxidative stability in thepresence of a lecithin composed of a mixture of phospholipids.

They proposed that this was due, among other mechanisms, tomodifications of the thickness and structure of the interfacial layer,

including changes in the conformation of caseins and their partial

desorption.Native proteins could create a protecting layer when adsorbed

at the droplet surface. Even though in most studies, the effect of

unadsorbed proteins could not be deconvoluted from that of theinterfacial fraction (Table 1A), adsorbed proteins were identified

as having several roles in protecting the oil phase against oxidationin emulsions.

The thickness of protein-stabilized interfaces has often beenconsidered as an important determinant with regard to the oxida-

tion of the dispersed lipids. The thicker interfacial layers formedby casein as compared to other proteins, such as whey proteins

(Atkinson and others 1995), have also been postulated to con-tribute to the better oxidative stability of casein-stabilized emul-

sions as compared with WPI- or SPI-stabilized emulsions (Huand others 2003a). However, this study lacked interfacial charac-terization. Kiokias and others (2006) also suggested that a thicker

interface layer and a stronger protein network acted as an effectiveinterfacial barrier against the prooxidant initiators to explain that

oxidative stability increased in emulsions prepared with increasingamounts of proteins (NaCas or whey protein concentrate (WPC)).

However, the actual surface coverage and interface thickness werenot measured, while, according to our calculation, the increase of

caseinate concentration from 5 to 20 g/L would have led to un-adsorbed concentrations of about 4 to 18 g/L (Table 1A). Hence,

the protective effect of the increased protein emulsifier could havebeen mostly due to the unadsorbed protein fraction. Hu and

others (2003a) also mentioned a possible antioxidant effect of spe-cific amino acid residues present at higher levels in casein than

in WPI and SPI (tyrosine and methionine). The exposure of sulfhydryl groups in WPC, due to partial denaturation of the pro-

teins during the thermal autoxidation of emulsions, favoring thehydrogen-donating antioxidant activity of the proteins was pro-

posed by Kiokias and others (2006) to explain the better oxidativestability of the emulsions they prepared with WPC than with

NaCas. Such controversial findings regarding the performanceof caseins compared with whey proteins at protecting emulsified

lipids against oxidation can also be seen in the article by Nielsenand others (2013). In this work, other formulation parameters were

also varied, such as the emulsifier concentration, pH, and additionof iron. In most cases, caseinate-stabilized emulsions were more

oxidatively stable as compared to whey protein-stabilized emul-sions, except at high emulsifier concentrations (0.75% w/w in

emulsions) and with added iron; in these particular cases, no differ-ence were observed or the trend was even slightly reversed. Finally,

Villiere and others (2005) observed a better oxidative stability of BSA-stabilized emulsions than NaCas-stabilized emulsions at pH

6.5, while in the presence of EDTA, oxidation was tremendouslydelayed in both systems, but developed faster in the BSA-stabilized

emulsions than in the caseinate-stabilized ones. The authors pro-posed that in the conditions where the metal ions were kept awayfrom the interface, due to the water-soluble chelator, the adsorbed

caseinate exerted free-radical quenching activity, as demonstratedby electron-spin resonance in the presence of a nitroxide probe.

In emulsions formulated to limit the contribution of unadsorbedemulsifier, BCN limited the development of oxidation as com-

pared to BLG, independent of the incubation conditions (Bertonand others 2011b, 2012b). This could indeed result from the ability

of caseins to form thicker and denser layers, as already mentioned,but other mechanisms such as the ability of casein to chelate and

inactivate metal ions or to quench free radicals cannot be excluded.We observed that the oxidative stability of BSA-stabilized emul-

sions depended to a large extent on the incubation conditions, in-

cluding the temperature and chemicals used to induce oxidation.This result gives evidence that any modification of the studiedsystem may change the chemical interactions between the entities

involved in the oxidative reaction, which will affect the efficiencyof interfacial proteins at protecting lipids against oxidation.

A recent study dealing with oxygen transfer across protein filmsat an air–water interface concluded that BLG provides a better

barrier to oxygen transfer than BCN. Possibly, a globular structureof proteins forming a viscoelastic layer is more effective than afluid-like structure to decrease oxygen permeability (Toikkanen

and others 2014). Nevertheless, the generally reported faster de-

velopment of lipid oxidation in emulsions stabilized by BLG incomparison to BCN led the authors to conclude that the differ-

ence in oxygen permeability may not substantially affect the rate of oxidation in emulsions and, due to the extremely thin nanometer-size interface, oxygen permeability is unlikely to be the limiting

step in controlling lipid oxidation in emulsions.Finally, as previously mentioned, adsorbed proteins undergo ex-

tensive oxidative modifications upon aging of an emulsion (Leaver and others 1999b; Rampon and others 2001; Hidalgo and Zamora

2002; Dalsgaard and others 2010; Mestdagh and others 2011). Pro-tein modifications may even precede lipid oxidation (Østdal and

others 2002; Salminen and others 2010; Berton and others 2012c).Altered proteins could transfer free radicals to the emulsified lipid

phase and participate in the initiation of lipid oxidation. Hence,adsorbed proteins could favor the development of lipid oxida-

tion while unadsorbed proteins can conversely act as antioxidants.This hypothesis for the double role of proteins has recently been

reviewed (Genot and others 2013).

Is it possible to alter the structure of interfacial proteins toimprove their barrier effect? Thermally, chemically, high-pressure-treated, or cross-linked proteins have been used in an

attempt to improve the protection of emulsified lipids against ox-idation. The underlying idea was to reinforce the cohesiveness of

the interfacial protein layer.Heat denaturation of BLG (80 °C, 30 min) prior to emul-

sion preparation did not modify lipid oxidation development inBLG-stabilized emulsions (Berton-Carabin and others 2013a). In

another study, protein-stabilized emulsions were thermally treated(75 °C or 80 °C, 30 min) to induce the postadsorption rearrange-

ment of protein molecules and the formation of a thicker and morecohesive layer surrounding the oil droplets. The treatment did not

affect lipid oxidation development for WPI-, casein- (Djordje-vic and others 2004), or BLG-stabilized emulsions (Kellerby and

others 2006b).A moderate increase of the oxidative stability of BLG-stabilized

emulsions homogenized at 28 to 77 °C, as compared to lower homogenization temperature (8 °C), was also recently reported

(Phoon and others 2013). In another recent study, Shao and Tang(2014) investigated the oxidative stability of emulsions stabilizedby SPI (native or heat-treated, 95 °C, 15 min). At the lowest

tested SPI concentration (0.5% w/w in aqueous solution), theoxidative stability of the emulsion was clearly improved when





SPI was heat-treated, as compared to native SPI. Interestingly,a comparable improvement was seen when native SPI was used

together with 300 mM of NaCl in the emulsion. In both cases,the improved oxidative stability of the emulsion was associated

with an increased protein surface load (from 2.1 mg/m2 to 3.1 or 4.9 mg/m2, respectively).

When homogenization was performed at 72 °C and combinedwith a high-pressure treatment (22.5 MPa), the oxidative stabil-

ity of fish-oil-enriched milk emulsions was better than after ho-

mogenization at lower pressure (5 MPa) and lower temperature(50 °C) (Let and others 2007c; Sorensen and others 2007). Thiswas related to changes in the protein conformation induced by

higher homogenization temperature and pressure. BLG, for in-stance, unfolds above 65 °C, which potentially exposes antioxi-

dant amino acid residues (sulfhydryl groups) (Let and others 2007c)and improves the coverage of the interface (Sorensen and others

2007).The chemical pretreatment of pea proteins (pH 12, 1 h, 20 °C)

prior to emulsification led to a reduction of the formation of pri-mary and secondary lipid oxidation products in emulsions. This

was associated with higher fractions of adsorbed proteins in emul-sions prepared with alkali-treated pea proteins (33%), compared

with 56% with the native proteins (Jiang and others 2014).A few studies have investigated the effect of protein cross-linking

on lipid oxidation in protein-stabilized emulsions, with contradic-

tory conclusions. Enzymatic cross-linking of interfacial casein after emulsification increased the cohesiveness of the interfacial layer of

a casein-stabilized O/W emulsion, but did not improve the oxida-tive stability of the emulsion (Kellerby and others 2006a). More

recently, protein enzymatic cross-linking was performed prior toemulsification (Ma and others 2012). This appreciably improved

both the physical stability of the emulsion and the oxidative sta-bility of the lipid phase. The authors hypothesized that the cross-

linked interfacial proteins could provide a more compact barrier to the diffusion of oxygen, although this was not supported by

experimental evidence and is questionable considering the di-mensions of oxygen and casein molecules. Besides, Tikekar and

others (2011) found that oxygen diffusion through the interfaciallayer in O/W emulsions proceeded at a similar rate with native or

cross-linked WPI as emulsifiers.

Adsorbedproteinemulsifiersdonotprotecttheoilphaseagainstoxidation better than surfactants. Protein emulsifiers have oftenbeen more effective at protecting lipids against oxidation than

many nonprotein emulsifiers (Donnelly and others 1998; Fomusoand others 2002b; Osborn and Akoh 2004; Haahr and Jacobsen

2008).However, most of these studies were conducted with high emul-

sifier concentrations, a large proportion of emulsifiers remainingin the aqueous phases of the emulsions (Table 1A). As a conse-

quence, the above-referred effects cannot be unambiguously at-tributed to the adsorbed emulsifiers. In emulsions formulated to

lower the contribution of unadsorbed emulsifiers, proteins wereless effective at protecting emulsified lipids against oxidation than

Tween 20, Tween 80, and Citrem (Berton and others 2011b).Such a lower efficiency of protein-stabilized interfaces at protect-

ing emulsified lipids against oxidation as compared to surfactant-stabilized interfaces was further confirmed independently of the

pH (6.7 or 3.0) and of the incubation conditions (Berton andothers 2012b). A possible explanation is the more structurally ho-mogeneous and compact interfacial films formed by surfactants,

whereas protein-based films are presumably more heterogeneousand mesoscopically porous.

Surface-active polysaccharides may act as a physical barrier.Polysaccharides such as gum arabic are surface-active and are there-

fore used as emulsifiers. Gum arabic has been shown to have an an-tioxidant effect as compared to BCN, which was attributed to the

formation of a steric barrier around the oil droplets (Matsumuraand others 2000). However, opposite conclusions were found by

Charoen and others (2012) who compared the oxidative stabil-ity of polysaccharide (modified starch or gum arabic)-stabilized

emulsions to that of WPI-stabilized emulsions. Here, gum-arabic-

stabilized emulsions were much more oxidizable than the 2 other formulations, whatever the pH. According to the authors, thiseffect would be explained by differences in the physicochemical

properties of the formed interfaces (thickness, porosity, and ironchelation properties), even if the specific involved mechanisms

remain unclear.Solid particles: potential effective barriers? Although the

use of solid particles to stabilize physically O/W emulsions hasbeen extensively described in the literature since the pioneer-

ing work of Ramsden and Pickering (Aveyard and others 2003;Leal-Calderon and Schmitt 2008; Dickinson 2010), little is known

about the effect of such solid particles on lipid oxidation in emul-sions. Recently, Kargar and others (2012) found that the incor-

poration of silica particles at the oil–water interface in protein-or surfactant-stabilized emulsions decreased lipid oxidation. Thiseffect was later on shown to depend on the characteristics of the

solid particles: in another study, the same authors highlighted abetter oxidative stability of O/W emulsions stabilized with micro-

crystalline cellulose particles than with modified starch particles(Kargar and others 2012). This antioxidative effect was attributed

to the ability of microcrystalline cellulose to scavenge free radicalsand to form thick interfacial layers.

Multicomponent interfacial layers: synergism or antago-nism?

As the interface is an area containing high local concentration of solutes, including emulsifiers, any incompatibility, nonmiscibility,

repulsive interactions, and phase separation (Figure 5A and 5B)may be enhanced as compared to in-bulk, diluted phases (Dickin-

son 2011). With respect to this incompatibility, the lipid oxidationof emulsions stabilized with such layers was analyzed.

In a recent study conducted in our laboratory, oil droplets werestabilized by proteins (BLG or BCN) used alone or mixed with

a pure phospholipid (dilauroylphosphatidylcholine, DLPC), in alow enough ratio to avoid any orogenic displacement of pro-

teins (Berton-Carabin and others 2013a). The microstructure andtopography of the interfacial layers, reconstructed as Langmuir-

Blodgett (LB) films and investigated by atomic force microscopy(AFM), showed that the incorporation of DLPC in the BCN

layer increased the structural heterogeneity of the films. In fact,

the mixed BCN-DLPC film had a more heterogeneous, coarser,and grainier structure as compared to pure BCN film. Interest-ingly, the lipid oxidation rate in the emulsion stabilized by the

BCN-DLPC mixture was higher than in the emulsion stabilizedby only BCN. We assumed that the homogeneity of pure protein

layers as compared to the mixed layers could ensure some protec-tion of the dispersed lipids against oxidation. However, this effect

is presumably strongly dependent on the kind, purity, and amountof the phospholipid. Recently, Garcıa-Moreno and others (2014)

found that the incorporation of soybean lecithin (a mixture of phospholipids) as a coemulsifier to caseinate-stabilized emulsions

decreased lipid oxidation as compared to the emulsion with onlycaseinate; however, this effect was not seen when pure PC or





Figure 5–Possible connection between the mesoscale lateral heterogeneity of interfaces and lipid oxidation (A) Illustration of the orogenicdisplacement phenomenon: Atomic force microscopy image of a Langmuir–Blodgett film (made by loading a Langmuir film formed at the air–waterinterface onto mica) of β-lactoglobulin (gray domains) partially displaced by Tween 20 (black domains). Reprinted from Journal of Colloid andInterface Science, 210, Mackie, Gunning, Wilde & Morris, Orogenic displacement of protein from the air/water interface by competitive adsorption,157–166, Copyright (1999), with permission from Elsevier. (B) Confocal microscopy image showing the formation of long-lived immiscible domains of

segregated polar lipids (from a mixture of cholesterol, dioleoylphosphatidylcholine, and biotinylated lipids; the latter were labeled by fluorescentstreptavidin) at the surfaceof oil droplets. Courtesy of of J. Brujic and L. Pontani, New York Univ., Dept. of Physics and Centerfor Soft Matter Research.(C) Evidence for repulsive interactions between Tween 20 and monolauroyl glycerol in mixed films formed at the air–water interface: the plot of themean molecular area as a function of the molar fraction of monolauroyl glycerol (orange solid line) shows a positive deviation from the black dottedline that linearly joins the molecular areas of both pure surfactants. (D) Lipid oxidation, measured through monitoring headspace oxygenconsumption, in rapeseed oil emulsions stabilized by pure Tween 20 or mixtures of Tween 20/Span 20, or of Tween 20/monolauroyl glycerol. Panels(C) and (D) have been adapted from Journal of Colloid and Interface Science, 377, Berton, Genot, Guibert & Ropers, Effect of lateral heterogeneity inmixed surfactant-stabilized interfaces on the oxidation of unsaturated lipids in oil-in-water emulsions, 244–50, Copyright (2012), with permissionfrom Elsevier.

pure PE was used instead of lecithin. In contrast to our study,

the incorporation of phospholipids led to a substantial increasein the concentration of unadsorbed caseinate (more than a 2-fold

increase), so a concomitant effect of unadsorbed caseinate on lipidoxidation cannot be excluded.

A recent article (Berendsen and others 2014) also reported anattempt to connect lipid oxidation in emulsions and the physi-

cal properties of reconstituted model interfaces. In this work, theformation of TBARS, developed earlier and to a greater extent

in emulsions stabilized by a WPI-carboxymethyl cellulose com-plex, as compared to emulsions stabilized with solely WPI. The

biopolymer complex was shown to form a thicker but less denseinterfacial film, as compared to single WPI (1.4 compared with

2.6 g/cm3, as determined by surface plasmon resonance). Thisfinding, hence, suggests that interfacial density and packing, rather

than thickness, matters with respect to barrier properties againstlipid oxidation.

We also investigated the effect of the incorporation of a co-

surfactant in Tween 20–stabilized interfaces on lipid oxidationin emulsions (Berton and others 2012a) (Figure 5D). The tested

cosurfactants (Span 20 and monolauroyl glycerol) have a similar hydrophobic chain as Tween 20 (lauric acid), but a much smaller

headgroup. Lipid oxidation developed earlier in the emulsionsstabilized with Tween 20–cosurfactant mixtures than in emulsions

stabilized with solely Tween 20. We assume that this effect is linkedto the nonideal interfacial behavior of Tween 20-cosurfactant mix-

tures, as evidenced by surface-pressure isotherms (Berton andothers 2012a) (Figure 5C). Examples of lateral phase separation

in mixed interfacial films are readily available in the literature, notonly in planar films (Mackie and others 2000) (Figure 5A), but also

in real emulsion droplets (Pontani and others 2013) (Figure 5B).This could lead to structural heterogeneity of the interfacial layer,

thus favoring the accessibility of the lipid substrate to the prooxi-dants present in the aqueous phase. This effect may be even more





Table 3–Synthesis of the formulation parameters and of the oxidative stability of multilayer-stabilized O/W emulsions. Oil volume fractions areexpressed in g/100 g unless otherwise stated. Each reference is reported as the name of the 1st author and publication year.

Lipid Aqueous First Second Third/furtherReference phase phase layer layer layer(s) Conclusion

Ogawa andothers(2003)

Corn oil (1%) Acetate buffer,pH 3.0

Lecithin (−) Chitosan (+) – Better oxidative stabilitywhen the 2nd layer isapplied.

Klinkesorn andothers(2005a)

Tunaoil (5%) Acetate buffer,pH 3.0

Lecithin (−) Chitosan (+) – Better oxidative stabilitywhen the 2nd layer isapplied.

Djordjevic andothers(2007)

Hexadecane+citral orlimonene (3%total)

Acetate buffer,pH 3.0 SDS (−) Chitosan (+) – Better oxidative stabilitywhen the 2nd layer isapplied; better oxidativestability as compared witha gum arabic-stabilizedemulsion.

Katsuda andothers(2008)

Menhaden oil(1%)

Phosphatebuffer, pH3.5

BLG (+) Citrus pectin orsugar beetpectin(−)

– Better oxidative stabilitywhen the 2nd layer isapplied (except with sugarbeet pectin that containsmetal ions).

Gudipati andothers(2010)

Fish oil (5%) Acetate buffer,pH 3.5

Citrem (−) Chitosan (+) Alginate (−) Betteroxidative stabil ity withCitrem/chitosan than withCitrem alone orCitrem/chitosan/alginate.

Lesmes andothers(2010b)

Menhaden oil(10%)

Phosphatebuffer, pH7.0

Caseinate (−) Lactoferrin (+) – Better oxidative stabilitywhen the 2nd layer isapplied.

Lomova andothers(2010)

Linseed oil (5%v/v) Water BSA (−) Polyarginine (+) Dextran sulfate(−) or tannicacid

Efficient protection of oildroplets against oxidationby multilayer shellcontaining tannic acid.

Chen andothers(2011a)

Corn oil (5%) Phosphatebuffer, pH3.0 to 5.0

Silk fibroin(slightly+)

Beet pectin (−) – Better oxidative stabilitywhen the 2nd layer isapplied.

Taherian andothers(2011)

Fish o il ( 3%) Phosphatebuffer, pH3.4 or 6.8

WPI (+) pH3.4(−) pH6.8

Fish gelatin (FG)(+) pH3.4(approximately0)pH 6.8

– Better oxidative stabilitywhen the 2nd layer isapplied as compared toWPI- or FG-stabilizedemulsions, at both pHs.

(−), (+), (slightly+) or(approximately0) indicate theelectrostatic chargeof thelayerin theconsidered conditions.

marked when the alkyl chain of the cosurfactant is different fromthat of the main surfactant (Mosca and others 2013): the oxida-

tive stability for emulsions prepared with a Tween 80/Span 80mixture, as compared to Tween 60/Span 80 and Tween 20/Span

80 mixtures, is better. Alkyl chains with varied lengths and un-saturation can pack less easily, which may increase the structural

heterogeneity of the interface.This may be of general importance for the development of oxi-

dation as most of the technical emulsifiers used in food applicationsare actually composed of complex mixtures of molecules (WPI,

NaCas, lecithins, and so on).

Multilayered interfacesOver the past decade, interest has been growing in the for-

mulation of O/W emulsions stabilized with multilayered inter-

faces (Grigoriev and Miller 2009), and in the chemical properties

of such systems, notably regarding their oxidative stability. Anoverview of the main studies carried out on the oxidative stabilityof multilayered emulsions is presented in Table 3. These systems

are commonly prepared by electrostatic layer-by-layer (LbL) depo-sition. For the 2 layer-stabilized emulsions, the application of the

2nd layer generally improved the oxidative stability of emulsions(Ogawa and others 2003; Djordjevic and others 2004; Klinkesorn

and others 2005a, 2005b; Gudipati and others 2010; Lesmes andothers 2010a; Chen and others 2011a; Taherian and others 2011).

This effect was attributed to the electrostatic repulsions of metalions when the 2nd layer was positively charged. This hypothe-

sis was strengthened by the work of Gudipati and others (2010),who demonstrated that the coating of oil droplets with chitosan

(positively charged) had a protective effect against lipid oxidation,whereas the additional coating of the droplets with alginate (neg-

atively charged) decreased the oxidative stability of the emulsion.The coating of oil droplets with a 2nd layer constituted of anionic

pectin was as well protective against lipid oxidation. Here, thiswas attributed to a steric barrier effect (Chen and others 2011a).

Accordingly, Tikekar and others (2011) found that the additionof a chitosan layer onto SDS-stabilized emulsion droplets effec-

tively decreased the oxygen transport rate through the oil–water interface, as compared to a single SDS layer. The coating of oil

droplets with multilayers also allows trapping specific compoundsat the droplet surface, for example, antioxidants (Lomova and

others 2010).

Lipid oxidation products located at the interfaceFinally, it is worth mentioning again that the composition of the

lipid phase evolves during the lipid oxidation reaction, which leadsto the formation of numerous products with various reactivity andhydrophobicity. Nuchi and others (2002) demonstrated that the

incorporation of lipid hydroperoxides in hexadecane decreasedsubstantially the interfacial tension at the hexadecane–water inter-

face. Lipid hydroperoxides being surface-active, they are thus ableto migrate to the oil–water interface, where they could be accessi-

ble to water-soluble prooxidants (Decker and McClements 2001).At first sight, this phenomenon may be of importance in the devel-

opment of lipid oxidation in food emulsions, whose componentsgenerally contain trace hydroperoxides and metal ions. However,

a recent study could not confirm this hypothesis. Kittipongpittayaand others (2012) isolated polar products generated in frying oil





Figure 6–Overview of the main interrelationships between the formulation parameters of O/W emulsions and the associated emulsionphysical–chemical characteristics. Arrows connect the emulsion characteristics to their influencing formulation parameters. The orange “?’’ tags pointout emulsion parameters for which contradictory results have been obtained with respect to their effect on lipid oxidation, and that should be furtherinvestigated. The green ‘‘’’ tags point out emulsion properties that have been shown to affect lipid oxidation in a consistent and reproducible manner.

(FFAs, hydroperoxides) and further incorporated them in bulk or emulsified corn oil. Although their addition increased lipid oxida-

tion in bulk oil, no clear effect was seen in the emulsified system.

This could be because these polar compounds are yet less surface-active than the emulsifiers used to physically stabilize the emulsion(that is, Tween 20), thus they may not be able to locate at the

interface.

Antioxidant compounds located at the interfaceThe prevention of lipid oxidation through the use of antiox-

idants has been extensively described and discussed in the lit-erature for decades (Uri 1961; Porter 1980; Frankel 1996, 2005;

Laguerre and others 2007; Jacobsen and others 2008). Early works,which led to the paradigm of the polar paradox, demonstrated

that nonpolar antioxidants are generally more efficient than polar antioxidants in O/W emulsions, presumably because they locate

more easily at the oil–water interface (Porter 1980; Huang andothers 1996a, 1996b; Cuvelier and others 2000; Schwarz and

others 2000). However, this explanation was clearly not univer-

sal (Shahidi and Zhong 2011; Sorensen and others 2011; Zhongand Shahidi 2012) and, currently, the location of antioxidants isrecognized as a major factor governing their efficiency at prevent-

ing lipid oxidation in O/W emulsions (Heins and others 2007;Laguerre and others 2007; Laguerre and others 2013b). Cur-

rent research relates to the quantitative distribution of antioxidantswithin complex multiphase systems, and to the selection or design

of antioxidants that preferentially locate at the interfaces.

Quantifying antioxidants at the air–water interface. Conven-

tional methods have relied on physically separating the emulsion’sphases (for example, by centrifugation or filtration techniques),

then measuring the antioxidant concentration in the serum phase(that is to say, the aqueous antioxidant fraction) (Huang and others

1997; Oehlke and others 2010). However, this method does notgive any information about the partitioning of the antioxidants

between the oil–water interface and the core of the oil droplets.

Besides, as noticed about the measurement of the partitioning of emulsifiers, there is a risk that such a physical separation disturbsthe mesoscale distribution of antioxidants and other molecules.

To overcome these issues, the group of C. Bravo-Diaz, L. Rom-sted, and coworkers has developed a pseudophase kinetic method,

based on the reaction of antioxidants with arenediazonium ionprobes, which takes place exclusively at the oil–water interface.

Authors used 2 kinetic data sets of the observed rate constant of reduction of arenediazonium ions by antioxidants as a function of

surfactant volume fraction and at 2 oil-to-water volume ratios tofit and estimate the distribution of the antioxidants in emulsions

(Romsted and Zhang 2002; Gunaseelan and others 2004, 2006).This pseudophase model has been applied to numerous systems

and conditions, which allowed understanding the relationshipsbetween oxidative stability and antioxidant location (Sanchez-Paz

and others 2008; Pastoriza-Gallego and others 2009, 2011, 2012;

Lisete-Torres and others 2012; Losada-Barreiro and others 2012,2013a, 2013b).

Selection of antioxidant molecules with high affinityfor interfaces. Antioxidants can be chosen so that their physic-ochemical properties lead to a preferred location at the interface.

Some of them may intrinsically exhibit surface properties (DiMattia and others 2010, 2011), depending on their polarity, which

is controlled by the environmental conditions (for example, pH[Pastoriza-Gallego and others 2012; Costa and others 2013], and

temperature [Pastoriza-Gallego and others 2009; Losada-Barreiroand others 2012]). The partitioning of antioxidants is also altered

by the HLB of emulsifiers (Losada-Barreiro and others 2013b)and by the surfactant concentration through micelle formation





(Richards and others 2002). Antioxidants may also be selectedfor their ability to interact with the emulsifiers adsorbed at the

oil droplet surface (Almajano and others 2007b; Lorrain andothers 2010a, 2010b; Oehlke and others 2010; Conde and others

2011; von Staszewski and others 2014). However, the chemicalbinding of antioxidants to emulsifiers can also lead to a decrease of

the antioxidant activity, which could result from the formation of hydrogen bonds counteracting the hydrogen-donating activity of

the antioxidant (Pekkarinen and others 1999; Stockmann and

others 2000). These chemical activities are dependent on emulsi-fiers (Oehlke and others 2011). The use of multilayers of emul-sifiers to stabilize O/W emulsions can also allow incorporating

an antioxidant such as tannic acid within the emulsifier shell sur-rounding the oil droplets (Lomova and others 2010). The latter ex-

ample led to an improved emulsion oxidative stability as comparedwith multilayer-stabilized emulsions containing tocopherols in the

oil phase. Selected flavonoids could also locate at the oil–water interface in O/W emulsions as solid particles, thus contributing

to the physical stability of the system (Luo and others 2011).

Design of lipophilized antioxidants. Beyond the selection of

molecules with an appropriate polarity and oil–water partitioncoefficient (logP ), increasing interest is currently encountered for

the synthesis of antioxidants derivatives (Nenadis and others 2003;Lisete-Torres and others 2012), and particularly, lipophilized an-tioxidants (Hunneche and others 2008; Laguerre and others 2013a;

Losada-Barreiro and others 2013a; Liu and others 2014). Thelipophilization by alkyl chain esterification has been tested on var-

ious antioxidants: chlorogenic acid (Laguerre and others 2009,2010; Sasaki and others 2010), rosmarinic acid (Laguerre and

others 2010; Panya and others 2012), hydroxytyrosol fatty acids(Medina and others 2009; Lucas and others 2010), dihydrocaf-

feic acid (Sorensen and others 2012), and epigallocatechin gallate(Zhong and Shahidi 2011). These studies generally highlighted

that the esterified antioxidants exhibited a better antioxidativeeffect than the corresponding nonesterified molecules in O/W

emulsions, with an optimum activity when the alkyl chain con-tained typically between 8 and 12 carbons. A recent study by

Losada-Barreiro and others (2013a), in which the pseudophasekinetic model was applied to investigate the location of gallic acid

and its alkyl derivatives (3 to 12 carbons) in emulsions, confirmedthat the higher the antioxidant fraction located at the interface,

the greater the antioxidant activity. Further increase in the alkylchain length, though, led to a dramatic decrease in the antioxi-

dant activity. This phenomenon has been referred to as the cutoff effect. When a certain level of hydrophobicity is obtained for

lipophilized phenolic acids, the ester forms micelles in the aque-ous phase rather than being located at the interface or oil phase

(Sorensen and others 2012; Laguerre and others 2013a).

ConclusionsLipid oxidation is a multifactorial phenomenon that includes

a very large set of chemical reactions occurring in the different

phases of emulsions (oil–water interface, aqueous phase, and oilphase). Each factor intervening in the oxidation of O/W emul-

sions may impact other factors and vice versa, as shown in Figure 6.Hence, it is difficult to deconvolute the effects of the interacting

parameters, and most of them cannot be studied without anyothers. These interrelated effects could explain the discrepancies

often observed from one study to another. For these reasons, con-tributors must pay attention to the full characterization of O/W

emulsions in their future research. Despite these pitfalls, the dataabout lipid oxidation in O/W emulsions available in the literature

strongly support the hypothesis that the oil–water interface arecritically involved in the reaction.

Further research must be conducted to fully elucidate the phys-ical and chemical barrier effect of interfaces. For instance, only a

few recent studies directly related the development of lipid oxi-dation in emulsions to the physical and structural characterization

of the involved interfaces (interface heterogeneity, for example).More information about the physical and chemical barrier effect of

interfaces according to their molecular composition and physical

organization is now needed. This may be of general importanceas most of the technical emulsifiers used in food applications areactually composed of complex mixtures of molecules. It includes

being able to take into account the dynamics of these systems in-cluding the motions and interactions of involved molecules and

their colloidal assemblies, as well as the permeability of the inter-faces to oxidation substrates and products.

To unambiguously assess the role of adsorbed emulsifiers (asphysical or chemical barrier) on lipid oxidation, emulsions should

be formulated to minimize the concentration and fraction of unad-sorbed emulsifiers, and hence, their contribution to the oxidative

fate of the system. Alternatively, adequate models of oil–water in-terfaces and their tools for characterization must be developed. As

a clear contribution of the unadsorbed emulsifier fraction to lipidoxidation has been highlighted in this review, emulsified modelsystems and real products should be designed so that the partition-

ing of the emulsifier molecules can be managed, or at least char-acterized. The location of the oxidizable molecules, for example,

that of polyunsaturated chains in oil droplets made of mixtures of glyceride species, should also be more deeply considered, and also

the possible exchanges of oxidizable substrates between dropletsand the other available phases.

Finally, innovative strategies that integrate a deep understand-ing of the involved phenomena are required. Integrated numeric

models must also be built for a better understanding of the reactionpathways and for controlling the oxidative reactions in complex

multiphase systems.

AcknowledgmentThe financing of this work through a Ph.D. research grant for

C.B.-C. by INRA and Region Pays de la Loire (2008 to 2011)

is gratefully acknowledged. We also thank Karin Schroen (FoodProcess Engineer ing group, Wageningen Univ., The Netherlands)for interesting discussion and advice.

Author ContributionsAll 3 authors contributed equally in the conception, planning,

and writing of this article, and they reviewed the final version of the manuscript.

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