Effects of Free Fatty Acids, Mono- and Diacylglycerols on ...

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Effects of Free Fatty Acids, Mono- and Diacylglycerols on Effects of Free Fatty Acids, Mono- and Diacylglycerols on

Oxidative Stability of Soybean Oil-In-Water Emulsions Oxidative Stability of Soybean Oil-In-Water Emulsions

Thaddao Waraho University of Massachusetts Amherst, [email protected]

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EFFECTS OF FREE FATTY ACIDS, MONO- AND DIACYLGLYCEROLS ON

OXIDATIVE STABILITY OF SOYBEAN OIL-IN-WATER EMULSIONS

A Dissertation Presented

by

THADDAO WARAHO

Submitted to the Graduate School of the

University of Massachusetts Amherst in partial fulfillment

of the requirements for the degree of

DOCTOR OF PHILOSOPHY

May 2011

Food Science

© Copyright by Thaddao Waraho 2011

All Rights Reserved



A Dissertation Presented

by

THADDAO WARAHO

Approved as to style and content by:

_______________________________________

Eric A. Decker, Chair

_______________________________________

D. Julian McClements, Member

_______________________________________

Yeonhwa Park, Member

_______________________________________

Young-Cheul Kim, Member

____________________________________

Eric A. Decker, Department Head

Food Science

DEDICATION

I would like to dedicate this doctorial dissertation to my parents, Preecha and

Chaweewan Waraho, who taught me that the greatest assets in life are knowledge and

education. There is no doubt in my mind that without their love, continued support and

encouragement, I could not have completed this process. I would also like to dedicate

this work to my brother, Dome, and my sister, Dujduan Waraho, who gave me strength

through their confidence and were always there for me during difficult times. Lastly, I

would like to dedicate my work to my loving husband, Keith Ogren, and my mother-in-

law, Valerie Ogren, who have always stood by me and have been great sources of

motivation and inspiration.

v

ACKNOWLEDGMENTS

First of all, I would like to express my deepest gratitude to my advisor, Professor

Eric Decker for giving me the opportunity to pursue my PhD through his teaching and

research assistantships. His support helped me to be successful in numerous ways. His

sage advice and patient encouragement helped me to accomplish my academic goals. I

also would like to thank my committee members, Dr. Julian McClements, Dr. Yeonhwa

Park, and Dr. Young-Cheul Kim for all their guidance, encouragement and support that

were instrumental in helping me to complete my research.

I would also like to give a special thanks to Jean Alamed who gave her time,

effort and experience in support of my lab work and most of all for being a great friend

who always gives me wonderful support and advice as well as sharing all the good times

and tough times with me.

I also would like to give a special thank to my previous and current lab members

especially, Tuk, Pom, Tang, Kla, Get, Lauren, Ann-Dorit, Ricard, Mette, Mickeal,

Vladimiro, Ryan, Bingcan and everyone who have been very generous in their support of

my academic pursuits and have contributed ideas, feedback, advice and above all, for

their friendship and providing me with an excellent atmosphere that gave me the

enthusiasm to carry on my work. I am grateful to have worked with you all.

Special thanks to my big sister, Chanti Chanthawong and the Thai Mafia for their

friendship and for sharing many wonderful times with me. They made me feel like I had

a big family away from home and made my time at UMass very joyful.

vi

I would like to thank Dan, Ruth, Fran, Beverly and Darlene who were ever ready

to lend a hand when I needed it and for their wonderful friendship throughout my

studying.

Lastly, I heartily thank Barbara Decker who always supported me with wonderful

advice and the warmest hugs through all the good and difficult times.

vii

ABSTRACT



MAY 2011

THADDAO WARAHO

B.Sc., KASETSART UNIVERSITY

M.S., UNIVERSITY OF MASSACHUSETTS AMHERST

Ph.D., UNIVERSITY OF MASSACHUSETTS AMHERST

Directed by: Professor Eric A. Decker

Even though edible oils undergo refining processes to remove undesirable

components, commercial oils still contain small amounts of minor components that can

contribute to either prooxidant and antioxidant pathways which ultimately affect the

quality of the oils. The objective of this research was to determine the role of free fatty

acids and mono- and diacylglycerols on the oxidative stability of oil-in-water emulsions.

Free fatty acids acted as a strong prooxidants in stripped soybean oil-in-water

emulsions. Concentrations as low as 0.1% of the lipid accelerated lipid oxidation rate by

both shortening the lag phase of lipid hydroperoxide and hexanal formation. The results

showed that the most likely mechanisms for the prooxidant activity of free fatty acids is

through their ability to increase the negatively charge on emulsion droplets that in turn

could attract the cationic transition metals to the emulsion droplet surface where they can

interact with lipid and thus promote oxidation. The prooxidant activity of free fatty acids

was dependent on fatty acid type with lipid oxidation rates being in the order of linolenic

< linoleic < oleic. Surprisingly, an increase in the degree of unsaturation of the free fatty

viii

acids lowered the ability of the free fatty acids to promote oxidation which may be due to

their differences in geometric shape thus influencing their ability to access the emulsion

droplet interface and increase the negative charge. Overall, free fatty acids are strong

prooxidants in oil-in-water emulsions. This prooxidant activity is dependent not only on

their concentration but also on the molecular structure of the fatty acid.

Addition of mono- and diacylglycerols in oil-in-water emulsions showed an

antioxidative effect in both non-stripped and stripped soybean oil. Addition of 1-

monooleoylglycerol only had a small impact on the oxidative stability of non-stripped

soybean oil-in-water emulsions but did inhibit lipid oxidation in emulsions prepared with

stripped soybean. Much stronger antioxidant activity was observed upon the addition of

1,2-dioleoyl-sn-glycerol to both non-stripped and stripped soybean oil-in-water

emulsions. Both lipid hydroperoxide and hexanal formation decreased with increasing

1,2-dioleoyl-sn-glycerol concentrations with 2.5% 1,2-dioleoyl-sn-glycerol almost

completely preventing hydroperoxide and hexanal production over the course of the

study. Overall, these results suggest that diacylglycerols could be an effective antioxidant

in oil-in-water emulsions which possibility due to their ability to form a liquid crystal

phase which could form a physical barrier that decreases interactions between

unsaturated fatty acids in the emulsion droplet core and prooxidants or oxygen in the

aqueous phase of the emulsion. However, the antioxidant mechanism of diacylglycerols

is not currently understood and needs further investigation.

ix

TABLE OF CONTENTS

Page

DEDICATION ................................................................................................................... iv

ACKNOWLEDGEMENTS .................................................................................................v

ABSTRACT ...................................................................................................................... vii

LIST OF TABLES ........................................................................................................... xiv

LIST OF FIGURES .........................................................................................................xvv

CHAPTER

1. INTRODUCTION ...................................................................................................1

1.1. Introduction .......................................................................................................1 1.2 Objectives .........................................................................................................5

2. LITERATURE REVIEW ........................................................................................6

2.1 Emulsions: Preparation, Properties and Characterization ..................................6

2.1.1 Emulsion Types ..................................................................................6

2.1.1.1 Conventional Emulsions ......................................................6

2.1.1.2 Nanoemulsions .....................................................................9

2.1.1.3 Multiple Emulsions ............................................................10

2.1.2 Droplet Characteristics......................................................................12

2.1.2.1 Droplet Concentration ........................................................13

2.1.2.2 Particle Size Distribution ...................................................13

2.1.2.3 Droplet Charge ...................................................................13

2.1.2.4 Interfacial Characteristics ..................................................13

2.1.2.5 Physical State .....................................................................14

2.1.3 Physicochemical Properties of Emulsions ........................................14

2.1.3.1 Optical Properties...............................................................14

2.1.3.2 Rheology ............................................................................15

x

2.1.3.3 Physical Stability ...............................................................16

2.2 Lipid Oxidation in Emulsified Food Products .................................................16

2.2.1 Lipid Oxidation Mechanisms in Emulsions ......................................18

2.2.2 Lipid Oxidation Mechanism .............................................................19

2.2.2.1 Initiation .............................................................................20

2.2.2.2 Propagation ........................................................................20

2.2.2.3 Termination ........................................................................22

2.2.3 Monitoring Lipid Oxidation ..............................................................22

2.2.4 Lipid Oxidation of Bulk oil vs. Emulsified Oil ................................23

2.2.5 Factors that Impact Lipid Oxidation in Emulsions ...........................25

2.2.5.1 Droplet Interface Characteristics .......................................25

2.2.5.1.1 Interfacial Area ...................................................26

2.2.5.1.2 Droplet Charge ....................................................28

2.2.5.1.3 Interfacial Thickness ...........................................32

2.2.5.1.4 Interfacial Permeability .......................................33

2.2.5.1.5 Interfacial Chemical Composition ......................34

2.2.5.2 Antioxidants .......................................................................35

2.2.5.2.1 Chain Breaking Antioxidants ..............................36

2.2.5.2.2 Transition Metal Chelators .................................36

2.2.5.2.2.1 Ethylenediaminetetraacetic Acid .........38

2.2.5.2.2.2 Phosphates............................................39

2.2.5.2.3 Impact of Physical Location on Antioxidant

Effectiveness .......................................................39

2.2.5.3 Influence of Minor Oil Components on Lipid Oxidation in

Emulsions ...........................................................................44

2.2.5.3.1 Minor Oil Components .......................................44

2.2.5.3.2 Influence of Other Emulsion Ingredients

on Lipid Oxidation in Emulsions ........................48

2.2.5.3.2.1 Continuous Phase Proteins ...................48

2.2.5.3.2.2 Polysaccharides ....................................51

2.2.5.3.2.3 Surfactant Micelles ..............................54

xi

2.2.6 Controlling Lipid Oxidation Using Structured Emulsions ...............55

2.2.6.1 Filled Hydrogel Particles ...................................................57

2.2.6.2 Solid Lipid Particles ...........................................................58

2.2.6.3 Multilayer Emulsions .........................................................59

2.2.6.3.1 One-Step Mixing .................................................60

2.2.6.3.2 Two-Step Mixing ................................................61

2.3 Conclusions ......................................................................................................62

3. PROOXIDANT MECHANISM OF FREE FATTY ACIDS IN

STRIPPED SOYBEAN OIL-IN-WATER EMULSIONS ....................................65

3.1. Abstract ...........................................................................................................65 3.2 Introduction .....................................................................................................66

3.3 Materials and Methods .....................................................................................68

3.3.1 Materials ...........................................................................................68

3.3.2 Methods.............................................................................................68

3.3.2.1 Preparation of Stripped Soybean Oil .................................68

3.3.2.2 Preparation of Free Fatty Acids .........................................69

3.3.2.3 Emulsion Preparation and Storage Conditions .................70

3.3.2.4 Measurement of Particle Size Distributions and

Zeta Potential (ζ) ................................................................71

3.3.2.5 Measurement of Lipid Oxidation .......................................71

3.3.2.6 Statistical Analysis .............................................................72

3.4 Results and Discussion ....................................................................................73

3.4.1 Physical Stability of Emulsions ........................................................73

3.4.2 Effect of Oleic Acids Concentrations on the Physical and

Chemical Properties of Oil-in-Water Emulsions ..............................73

3.4.3 Effect of Methyl Oleate and Oleic Acid on the Physical and


3.4.4 The Effect of pH on Physical and Chemical Properties of Oil-

in-Water Emulsions Containing Oleic Acid .....................................79

3.4.5 The Effect of EDTA and Fatty Acid Hydroperoxides ......................83

3.5 Conclusions ......................................................................................................88

4. IMPACT OF FATTY ACID CONCENTRATION AND STRUCTURE

ON LIPID OXIDATION IN OIL-IN-WATER EMULSIONS .............................89

xii

4.1 Abstract ............................................................................................................89

4.2 Introduction ......................................................................................................90

4.3 Materials and Methods .....................................................................................93

4.3.1 Materials ...........................................................................................93

4.3.2 Methods.............................................................................................93

4.3.2.1 Removing of Polar Minor Components from the Oils

and Free Fatty Acids ..........................................................93

4.3.2.2 Emulsion Preparations and Storage Conditions .................94


Zeta Potential (ζ) ................................................................95

4.3.2.4 Measrement of Interfacial Tension ....................................95

4.3.2.5 Measurement of Lipid Oxidation .......................................96

4.3.2.6 Statistical Analysis .............................................................97

4.4 Results and Discussion ....................................................................................98

4.4.1 Physical Stability of Emulsions ........................................................98

4.4.2 Impact of Low Oleic Concentrations on the Physical and


4.4.3 Effect of Degree and of Free Fatty Acids Unsaturation on the

Physical and Chemical Properties of Oil-in-Water Emulsions .......102

4.4.4 The Effect of cis vs. trans Double Bonds of Free Fatty Acids

on Physical and Chemical Properties of Oil-in-Water

Emulsions ........................................................................................106

4.5 Conclusions ....................................................................................................109

5. ANTIOXIDANT EFFECTS OF MONO- AND DIACYLGLYCEROLS IN

NON-STRIPPED AND STRIPPED SOYBEAN OIL-IN-WATER

EMULSIONS .......................................................................................................111

5.1. Abstract .........................................................................................................111 5.2 Introduction ...................................................................................................112

5.3 Materials and Methods ...................................................................................114

5.3.1 Materials .........................................................................................114

5.3.2 Methods...........................................................................................115

5.3.2.1 Removing Polar Minor Components from Soy Bean Oil

(Stripping Oils) ................................................................115

5.3.2.2 Emulsion Preparation and Storage Conditions ................116


Zeta Potential (ζ) ..............................................................117

xiii

5.3.2.4 Measurement of Interfacial Tension ................................117

5.3.2.5 Measurement of Lipid Oxidation .....................................118

5.3.2.6 Statistical Analysis ...........................................................119

5.4 Results and Discussion ..................................................................................119

5.4.1 Physical Stability of Emulsions ......................................................120

5.4.2 Effect of Mono- and Diacylglycerols on Interfacial Tension .........121

5.4.3 Effect of Mon- and Diacylglycerols on the Physical and

Chemical Properties of Non-stripped and Stripped Oil-in-

Water Emulsions .............................................................................123

5.5 Conclusions ....................................................................................................133

6. OVERALL CONCLUSIONS ..............................................................................135

BIBLIOGRAPHY ............................................................................................................139

xiv

LIST OF TABLES

Table Page

2.1. Average compositions of crude and refined soybean oils ....................................45

3.1. The droplet surface charge or zeta potential (ζ) of 1.0% stripped soybean oil-in-

water emulsions without (control) and with addition of 1.0, 2.5, and 5.0% oleic

acids and from 1.0% methyl oleate (oil wt.) at pH 7.0 ..........................................74

xv

LIST OF FIGURES

Figure Page

2.1. The three regions of an oil-in-water emulsion .........................................................7

2.2. Structure of multiple emulsions (W1/O/W2) ..........................................................11

2.3. Inhibition of iron-promoted lipid oxidation by a cationic emulsion droplet

interface created by using proteins as an emulsifier at pH values below

the pI of the protein ................................................................................................31

2.4. The proposed mechanism for free fatty acid promoted oxidation on

emulsified oil droplets. Mn+

= transition metal ......................................................47

2.5. Examples of different kinds of structured emulsion systems that can

be utilized in foods .................................................................................................56

2.6. Locations where the chemical and physical properties of oil-in-water

emulsions can be altered to impact lipid oxidation reactions ...............................64

3.1. Formation of lipid hydroperoxide concentration (a) and hexanal (b) in

1.0% stripped soybean oil-in-water emulsions at pH 7.0 without (control)

and with addition of 1.0, 2.5, and 5.0% oleic acids (oil wt.) during storage

at 15ºC in the dark for 6 days .................................................................................76

3.2. Formation of lipid hydroperoxide concentration (a) and hexanal (b) in

1.0% stripped soybean oil-in-water emulsions without (control) and

with addition of 1.0% oleic acids and 1.0% methyl oleate (oil wt.)

at pH 7.0 during storage at 15ºC in the dark for 8 days .........................................78

3.3. The droplet surface charge or zeta potential (ζ) of 1.0% stripped soybean

oil-in-water emulsions with addition of 1.0 % oleic acids (oil wt.) at

pH 2.0, 4.0, 6.0, and 8.0 .........................................................................................81

3.4. Formation of lipid hydroperoxide concentration (a) and hexanal (b)

in 1.0% stripped soybean oil-in-water emulsions with addition of

1.0% oleic acids (oil wt.) at pH 2.0, 4.0, 6.0, and 8.0 during storage

at 15ºC in the dark for 10 days ...............................................................................82



1.0% oleic acids (oil wt.) without and with 200 m EDTA at pH 7.0

during storage at 15ºC in the dark for 21 days .......................................................84

xvi



1.0% oleic acids with low and high in hydroperoxides (oil wt.) at

pH 7.0 during storage at 15ºC in the dark for 8 days .............................................87

4.1. The structure of a) oleic acid (18:1, cis), b) elaidic acid (18:1, trans), c) linoleic

acid (18:2, cis-cis), d) linoelaidic acid (18:2, trans-trans) and e) linolenic acid

(18:3, cis-cis-cis) ....................................................................................................92


water emulsions with addition of 0-1.0 % (wt.% of oil) oleic acids at pH 7. Data

points represent means (n=3) + standard deviations (some error bars may lie

within the data points) .........................................................................................100

4.3. Formation of lipid hydroperoxide concentration (a) and headspace hexanal (b)

in 1.0% stripped soybean oil-in-water emulsions without (control) and

with 0-1.0 % (wt.% of oil) oleic acids at pH 7 during storage at 10ºC in

the dark for 7 days...............................................................................................101


water emulsions without (control) and with 0.50% (wt.% of oil) oleic, linoleic

and linolenic acids at pH 7 ...................................................................................103

4.5. Formation of lipid hydroperoxide concentration (a) and headspace hexanal (b) in

1.0% stripped soybean oil-in-water emulsions without (control) and with 0.50%

(wt.% of oil) oleic, linoleic and linolenic acids at pH 7.0 during storage at

15ºC in the dark for 7 days ..................................................................................105

4.6. Formation of lipid hydroperoxide concentration (a) and headspace hexanal (b) in

1.0% stripped soybean oil-in-water emulsions without (control) and with 0.50%

(wt.% of oil) oleic, linoleic, elaidic and linoelaidic acids at pH 7.0 during

storage at 15ºC in the dark for 8 days ..................................................................108

5.1. Influence of addition of 0.01-2.50% 1-monooleoylglycerol or 1,2-dioleoyl-sn-

glycerol in medium chain triacylglycerols on interfacial tension at ambient

temperature. Data represents means (n=3) standard deviations. Some error bars

lay within data points ...........................................................................................122

5.2. The droplet surface charge or zeta potential (ζ) of: (a) 1.0% non-stripped soybean

oil-in-water emulsions with addition of 0.01-2.50% 1-monooleoylglycerol (oil

wt.) at pH 7.0 after 24 hour of storage in the dark at 250C and (b) 1.0% stripped

soybean oil-in-water emulsions with addition of 0.01-2.50% 1-

monooleoylglycerol (oil wt.) at pH 7.0 after 24 hour of storage in the

dark a150C ............................................................................................................124

xvii

5.3. The droplet surface charge or zeta potential (ζ) of: (a) 1.0% non-stripped soybean

oil-in-water emulsions with addition of 0.01-2.50% 1,2-dioleoyl-sn-glycerol (oil

wt.) at pH 7.0 after 24 hour of storage in the dark at 250C and (b) 1.0% stripped

soybean oil-in-water emulsions with addition of 0.01-2.50% 1,2-dioleoyl-sn-

glycerol (oil wt.) at pH 7.0 after 24 hour of storage in the dark at 150C .............125

5.4. Formation of lipid hydroperoxides (a) and hexanal (b) in 1.0% non-stripped

soybean oil-in-water emulsions without (control) and with addition of 0.01-2.50%

1-monooleoylglycerol (oil wt.) at pH 7.0 during storage at 25ºC in the dark for 17

days. Data represents means (n=3) standard deviations. Some error bars lay

within data points .................................................................................................128

5.5. Formation of lipid hydroperoxides (a) and hexanal (b) in 1.0% stripped soybean

oil-in-water emulsions without (control) and with addition of 0.01-2.50% 1-

monooleoylglycerol (oil wt.) at pH 7.0 during storage at 15ºC in the dark for 8



5.6. Formation of lipid hydroperoxides (a) and hexanal (b) in 1.0% non-stripped

soybean oil-in-water emulsions without (control) and with addition of 0.01-2.50%

1,2-dioleoyl-sn-glycerol (oil wt.) at pH 7.0 during storage at 25ºC in the dark for

14 days. Data represents means (n=3) standard deviations. Some error bars lay


5.7. Formation of lipid hydroperoxides (a) and hexanal (b) in 1.0% stripped soybean

oil-in-water emulsions without (control) and with addition of 0.01-2.50% 1,2-

dioleoyl-sn-glycerol (oil wt.) at pH 7.0 during storage at 15ºC in the dark for 9



1

CHAPTER 1

INTRODUCTION

1.1 Introduction

Many lipid containing processed foods are either oil-in-water or water-in-oil

emulsions such as milk, infant formula, salad dressing, mayonnaise, sauces, soups,

beverages, cream, and some desserts (McClements and Decker, 2000; Okuda et al., 2005;

McClements, 2005; Friberg et al., 2004). As well as the food industry, the cosmetics,

pharmaceutical and medical industries also utilize oil-in-water emulsions as a means to

encapsulate, protect, and release bioactive lipids in their products. However, these

industries face a major problem regarding utilizing an oil-in-water emulsion because they

can undergo lipid oxidation which then causes a deterioration of the product.

Lipid oxidation is a great concern for the food industry because it causes

deterioration to lipid containing food products, even in foods that contain only small

amount of lipids such as vegetable products. Not only does lipid oxidation cause

undesirable changes in appearance, texture and development of rancidity that shortens

product shelf life, but it also causes losses in important nutrients and formation of

potentially toxic reaction products (such as aldehydes and ketones) which cause

important health concern for consumers (Frankel, 1998; Coupland and McClements,

1996; McClements and Decker, 2000; Chaiyasit et al., 2007a; Decker and McClements,

2008). Therefore, retarding lipid oxidation is necessary in order to extend the shelf life of

the products as well as to maintain nutrition functionality of the lipid with a benefit of

reduction of raw material wastes (Chaiyasit et al, 2007a). It is very important for the lipid

2

chemists to understand the mechanisms of the lipid oxidation thoroughly to be able to

utilize it as a basic fundamental to develop the proper methods to retard lipid oxidation.

The oil-in-water emulsions can be differentiated into three different regions; the

emulsion droplet’s lipid core, the interfacial membrane of the emulsion droplet, and the

continuous phase. It has been suggested that the polar molecules are located in the

aqueous phase while non-polar molecules are mostly located in the oil droplets and

surface active or amphiphillic molecules are accumulated at the interface (McClements

and Decker, 2000; Decker et al., 2005; Chaiyasit et al., 2007a). The reactants that

influence in lipid oxidation can partition in these different regions resulting in different

lipid oxidation rates and mechanisms in oil-in-water emulsions than in bulk oils (Nuchi et

al, 2001).

Lipid oxidation is favored in oil-in-water emulsions because of the large contact

surface between the oxidizable lipid droplets and water-soluble compounds including

oxygen and prooxidants, which contribute to the initiation and propagation of oxidation

reactions (Frankel, 1998; Lethuaut et al., 2002; Villiere et al., 2005). There are several

factors that impact the rate of lipid oxidation in oil-in-water emulsions. For example,

fatty acid composition, aqueous phase pH and ionic composition, type and concentration

of antioxidants and prooxidants, oxygen concentration, lipid droplet characteristics such

as particle size, concentration and physical state as well as emulsion droplet interfacial

properties such as thickness, charge, rheology, and permeability (McClements and

Decker, 2000). The susceptibility of emulsified lipids to oxidation also depends on the

surrounding molecular environment and interactions with other molecules within the

immediate vicinity of the droplets (Kellerby et al., 2006a).

3

Even though the commercial oils undergo a refining process, normally they still

contain some minor components which are polar lipids such as free fatty acids,

phospholipids, mono- and diacylglycerols, sterols, cholesterols, tocopherols as well as

some polar oxidation products such as lipid hydroperoxides, alcohols, aldehydes and

ketones (Chaiyasit et al., 2007a). For examples, according to Pryde (1980), after refining,

bleaching and deodorizing processes, soybean oils still contain about 0.3%

unsaponifiable matter including of 0.13% phytosterols, 0.11-0.18% tocopherols, 0.01%

hydrocarbons as well as less than 0.05% free fatty acids and about 0.1-0.3 ppm of iron

and 0.02-0.06 ppm of copper. These polar minor components could impact lipid

oxidation by affecting the physical properties of the oils (Chaiyasit et al., 2007a). Several

studies have found that lipid oxidation of oil-in-water emulsions are influenced by the

properties of oil-water interface (Donnelly et al., 1998; Mei et al., 1998a; 1998b; 1999;

Silvestre et al., 2000; Chaiyasit et al., 2000). Therefore, it might be possible that these

polar minor components from the oils could influence lipid oxidation in the oil-in-water

emulsions as well because they tend to accumulate at the oil-water interface.

Free fatty acids are formed during lipid extraction and refining by hydrolysis of

triacylglycerides by lipases and high temperature in the presence of water. They are

removed from crude oils by neutralization and deodorization. However, these refining

steps are not 100% efficient with commercial oils typically containing 0.05-0.70% free

fatty acids depending on the type of oils and the refining process (Pryde, 1980; Jung et

al., 1989; Chiyasit et al., 2007a). Several researchers showed that free fatty acids act as

prooxidants in bulk oils. These studies suggest that the prooxidant activity of free fatty

acids is due to the ability of the carboxylic acid group of free fatty acids to form

4

complexes with transition metals and or the ability of the acid group to directly promote

hydroperoxide decomposition (Miyashita and Takagi, 1986; Mistry and Min, 1987;

Yoshida et al., 1992; Frega et al., 1999). Free fatty acids could be important prooxidants

in oil-in-water emulsions because they are surface active compounds since they are more

polar than triacylglycerols due to the presence of an unesterified carboxylic acid groups.

The surface activity of free fatty acids allows them to diffuse and concentrate at the

water-lipid interface of the oil-in-water emulsions (Nuchi et al., 2002). The pKa’s of fatty

acids are in the range of 4.8-5.0 for medium- and long-chain (C ≥10) acids in aqueous

solution (White, 1950; Spector, 1975; Lieckfeldt et al., 1995). Thus free fatty acids could

potentially make the emulsion droplet more negatively charged when pH values are

above their pKa’s. Previous researchers have shown that negatively charged oil-in-water

emulsion droplets can attract prooxidant transition metals that can increase metal-lipid

interactions thus accelerating oxidation (Yoshida and Nikki, 1992; Fukuzawa et al. 1995;

Mei et al., 1998a; 1998b). Therefore, having free fatty acids in food where the pH is

above their pKa could cause a critical problem in oil-in-water emulsion system.

1.2 Objectives

Free fatty acids, monoglycerols and diglycerols are found in vegetable oils as

minor components. They are surface active compounds that accumulate at the oil-water

interface which is a physical location where lipid oxidation reactions are prevalent in oil-

in-water emulsions. While there are several studies on the impact of free fatty acids,

monoglycerols and diglycerols in bulk oils, there is almost no studies on the impact of

free fatty acids , monoglycerols and diglycerols on lipid oxidation of oil-in-water

5

emulsions. Therefore, the objective of this research was to clarify the effects of free fatty

acids, monoglycerols and diglycerols and their prooxidant or antioxidant mechanisms on

the lipid oxidation in oil-in-water emulsions. The understanding of how free fatty acids,

monoglycerols and diglycerols influence lipid oxidation in oil-in-water emulsions could

provide fundamental knowledge that could be used to improve the oxidative stability of

oils in emulsion and other food dispersions.

6

CHAPTER 2

LITERATURE REVIEW

2.1 Emulsions: Preparation, Properties and Characterization

Many varieties of emulsion-based delivery system have been developed for use in

the food, pharmaceutical, cosmetic and medical industries to protect bioactive lipid

components (such as omega-3 fatty acids, oil-soluble vitamins and carotenoids) from

chemical degradation during production, storage, and transportation (McClements et al.,

2007). There are several emulsion-based technologies that are widely used in the food

and other industries, including conventional emulsions, nanoemulsions and multiple

emulsions.

2.1.1 Emulsion Types

2.1.1.1 Conventional Emulsions

Conventional emulsions consist of two immiscible liquids (such as oil and water)

with one of the liquids being dispersed as small spherical droplets in the other liquid

(Dickinson and Stainsby, 1982; Dickinson, 1992). The mean droplet diameter in food

emulsions ranges from less than 100 nm to greater than 100 m (McClements et al.,

2007). Conventional emulsion can be classified as either water-in-oil (W/O) or oil-in-

water (O/W) depending on the spatial arrangement of the two immiscible liquids. Water-

in-oil emulsion consists of water droplets dispersed in an oil phase, while oil-in-water

emulsion consists of oil droplets dispersed in a water phase. Emulsions can be

7

conveniently divided into three different regions: the continuous phase, the interfacial

region, and the interior of the droplets (McClements and Decker, 2000). The three

different regions of an oil-in-water emulsion are shown in Figure 2.1.

Figure 2.1. The three regions of an oil-in-water emulsion.

8

Emulsions are thermodynamically unfavorable systems that tend to break down

over time. To create emulsions that are kinetically stable for a reasonable period of time,

stabilizers such as emulsifiers or texture modifiers need to be added to prevent

gravitational separation, flocculation, coalescence and Oswald ripening (Friberg et al.,

2004; McClements, 2005; Dickinson, 1992). The most commonly used emulsifiers in the

food and beverage industries are small molecule surfactants that are very mobile at the

interface which they reduce the interfacial tension efficiently as well as rapidly coat the

freshly created oil-water interface during emulsification (e.g., Tweens, Spans, and esters

of fatty acids), phospholipids (e.g., egg, soy or dairy lecithin), while surface-active

proteins (e.g. casein, whey, egg, and soy) and surface-active polysaccharides (e.g., gum

Arabic and modified starch) are considered as high-mass surfactants (Decker et al., 2005,

Kralova and Sjoblom, 2009).

Conventional O/W emulsions can be prepared by homogenizing an oil phase and

an aqueous phase together in the presence of a water-soluble emulsifier. Wide varieties of

homogenizers can be used, including high shear mixers, high pressure homogenizers,

colloid mills, ultrasonic homogenizers and membrane homogenizers depending on the

characteristics of the materials being homogenized (e.g., product viscosity, interfacial

tension, shear sensitivity) and the desired emulsion properties (e.g., droplet concentration,

particle size distribution). The desired droplet characteristics can be manipulated by

careful selection of homogenizer type, homogenizer operating conditions, and emulsifier.

For example, the droplet size of O/W emulsions produced by high-pressure homogenizers

can be reduced by increasing the homogenization pressure or number of passes through

the homogenizer. The electrical charge on the droplets can be controlled by selecting an

9

appropriately charged emulsifier, which may be positive, neutral or negative

(McClements et al., 2007; McClements, 2005; Walstra, 1993; Walstra, 2003).

2.1.1.2 Nanoemulsions

Nanoemulsions are conventional emulsions that contain very small droplets, with

the appropriate size range being defined differently by different authors, e.g., 1-100 nm

(Mason et al., 2006), 100-600 nm (Bouchemal et al., 2004) or 400-800 nm (Sarker,

2005). In this chapter, we prefer to use a definition where nanoemulsions exhibit

distinctly different bulk physicochemical characteristics to conventional emulsions, i.e.,

when the particle size becomes so small that they do not scatter light strongly and they

appear clear, which generally occurs when the diameter is less 50 nm. Nanoemulsions are

metastable systems that can be designed to persist for many months or years if they are

stabilized appropriately, e.g., by reducing droplet aggregation and Ostwald ripening.

Nanoemulsions can be formed using high or low intensity methods. High

intensity methods involve the application of intense mechanical energy to a system to

break up the disperse phase liquid into smaller portions. The mechanical energy needed

to break large droplets into small droplets increases as their initial radius decreases

because of the Laplace pressure (P = 2/r) that tends to oppose particle deformation and

rupture. Hence, only mechanical devices that are capable of generating extremely high

intensity disruptive forces can be used to form nanoemulsions, such as sonicators and

high-pressure homogenizers (especially microfluidizers). Sonicators use high-intensity

ultrasonic waves to generate intense disruptive stresses (particularly cavitational, shear

and turbulent forces) that break up the droplets (Landfester et al., 2000). In

10

microfluidizers, a premixed emulsion is divided into two streams that are forced through

separate microchannels and then made to impinge on each other at high velocity, which

generates intense disruptive forces (particularly strong extensional flow) that break up the

droplets (Meleson et al., 2004). Low intensity methods rely on the spontaneous phase

separation of two immiscible liquids under certain conditions, leading to the formation of

a dispersion of one liquid in the other liquid. These methods include the phase inversion

temperature (PIT), emulsion inversion, and solvent displacement methods. The

advantage of low intensity methods is that they require much less mechanical energy to

form emulsions, however, they often require the use of organic solvents, synthetic

surfactants or co-surfactants.

2.1.1.3 Multiple Emulsions

The most commonly used water-dispersible multiple emulsions are water-in-oil-

in-water (W/O/W) emulsions, which consist of small water droplets contained within

larger oil droplets that are dispersed within an aqueous continuous phase (Garti, 1997a;

Garti, 1997b; Garti and Benichou, 2004; Garti and Bisperink, 1998, McClements et al.,

2007) (Figure 2.2). More accurately this type of emulsion should be referred to as a

W1/O/W2 emulsion, where W1 is the inner water phase and W2 is outer water phase, since

the composition of the two water phases is usually different. There are also two different

interfacial layers in this type of emulsion: the W1-O layer surrounding the inner water

droplets, and the O-W2 layer surrounding the oil droplets. Therefore, two different types

of emulsifier are normally used to stabilize W/O/W emulsions: an oil-soluble emulsifier

11

is used for the inner water droplets and a water-soluble emulsifier is used for the oil

droplets.

Figure 2.2. Structure of multiple emulsions (W1/O/W2).

12

Usually, W/O/W emulsions are created using a two-step procedure. The first step

is to create a W1/O emulsion by homogenizing water, oil and an oil-soluble emulsifier,

and then a W1/O/W2 emulsion is created by homogenizing the W1/O emulsion with an

aqueous solution containing a water-soluble emulsifier. Similar techniques to

homogenize O/W emulsions can be used to produce W1/O/W2 emulsions such as high

shear mixers, high pressure homogenizers, colloid mills, ultrasonic homogenizers and

membrane homogenizers (McClements, 2005). However, the homogenization conditions

used in the first stage are usually more intense than those used in the second stage, in

order to avoid disruption or expulsion of the W1 droplets formed within the oil phase.

The size of the water droplets in the W1/O emulsion and of the final W1/O/W2 emulsion

can be controlled by varying emulsifier type, emulsifier concentration and

homogenization conditions. Multiple emulsions have not been widely used in food

products because they are highly susceptible to breakdown during processing, e.g.,

mechanical forces, thermal processing, chilling, freezing and dehydration or during

storage (McClements et al., 2007). Nevertheless, recent advances in understanding the

physicochemical basis of the stability of these systems are leading to more applications in

foods being implemented.

2.1.2 Droplet Characteristics

Knowledge of the most important properties of the droplets within emulsions is

useful for determining the best strategy to control their oxidative stability.

13

2.1.2.1 Droplet Concentration

The droplet concentration is usually expressed as the number, mass or volume of

droplets per unit volume or mass of emulsion and can be controlled by varying the

proportions of the two immiscible liquids used to prepare it (McClements et al., 2007;

McClements, 2005).

2.1.2.2 Particle Size Distribution

The particle size distribution (PSD) of an emulsion represents the fraction of

droplets falling into different size categories. It is usually represented as either a table or a

plot of particle concentration (e.g., volume or number percent) versus droplet size (e.g.,

radius or diameter) (McClements et al., 2007; McClements, 2005).

2.1.2.3 Droplet Charge

The electrical properties of a droplet are usually characterized in terms of its

potential (), which can be conveniently measured (Hunter, 1986). The potential of

a droplet depends on the surface charge density (i.e., the number of charges per unit

area), as well as the prevailing environmental conditions (i.e., ionic strength and

dielectric constant).

2.1.2.4 Interfacial Characteristics

Each droplet in an emulsion is usually coated by a thin layer of adsorbed material

to protect it against aggregation with other droplets. The composition and properties of

this interfacial region are defined by the type, concentration and interactions of any

14

surface-active species before, during and after emulsion formation, e.g., emulsifiers,

biopolymers, and minerals (Dickinson, 2003).

2.1.2.5 Physical State

Normally, the droplets that make up the dispersed phase of an emulsion are liquid.

Nevertheless, in some emulsion-like systems the dispersed phase is either partially or

fully solidified, e.g., solid lipid particles (McClements et al., 2007; McClements, 2005;

Walstra, 2003; Muller and Keck, 2004; Wissing et al., 2004). The nature, location and

concentration of the fat crystals within the lipid droplets in an O/W emulsion can be

controlled by proper selection of oil type (e.g., solid fat content vs. temperature profile),

thermal history (e.g., temperature vs. time profile), the presence of additives (e.g., crystal

structure modifiers), emulsifier type, and droplet size (Walstra, 2003; Muller and Keck,

2004; Muller et al., 2000).

2.1.3 Physicochemical Properties of Emulsions

It should be stressed that any strategy used to retard or inhibit lipid oxidation in

emulsions should not adversely affect the bulk physicochemical and sensory properties of

the final product.

2.1.3.1 Optical Properties

Opacity and color are the most important optical properties of emulsions, and can

be quantitatively described using tristimulus color coordinates, such as the L*a*b*

system (McClements, 2005). The optical properties of emulsions are determined by the

15

droplet size, droplet concentration, and relative refractive index (McClements, 2005;

McClements, 2002a; 2002b). The lightness of an emulsion tends to increase and the color

intensity decrease with increasing droplet concentration and refractive index contrast, and

they have a maximum value at a particular droplet size. The optical properties of an

emulsion may affect its susceptibility to oxidation. Ultraviolet and visible light can

penetrate further into optically clear emulsions than into optically opaque ones, thereby

accelerating light-catalyzed lipid oxidation. This has important consequences for the

development of transparent beverages containing chemically labile lipids, such as -3

fatty acids or carotenoids.

2.1.3.2 Rheology

Food emulsions vary widely in their rheological behaviors depending on the

nature of the food. They may be viscous liquids, visco-elastic liquids, visco-elastic solids,

plastics, or elastic solids depending on their composition, structure and interactions

(Walstra, 2003; McClements, 2005; Genovese et al., 2007). Generally, when the droplet

concentration of an emulsion increases, the viscosity will increase gradually at first and

then steeply as the droplets become more closely packed. When the droplet concentration

is around 50-60% (for a non-flocculated O/W emulsion), the droplets pack so closely

together that the emulsion exhibits solid-like characteristics, such as visco-elasticity and

plasticity (McClements, 2005). Flocculated emulsions may exhibit these solid-like

characteristics at much lower droplet concentrations due to particle-particle interactions.

The rheology of emulsions may be altered by oxidation reactions that lead to covalent

cross-linking of adsorbed or non-adsorbed proteins.

16

2.1.3.3 Physical Stability

Emulsions are thermodynamically unfavorable systems that tend to break down

over time because of physicochemical mechanisms such as gravitational separation,

flocculation, coalescence and Ostwald ripening (Friberg et al., 2004; McClements, 2005;

Dickinson, 1992). One of the most common causes of instability in food emulsions is

gravitational separation, which can be either creaming or sedimentation depending on the

relative densities of the dispersed and continuous phases. The shelf life of many

emulsion-based food products is determined by their physical stability. The physical

stability of an emulsion may be altered by lipid oxidation, since there will be a change in

the chemistry and organization of the molecules present in the system. For example,

oxidation may change the nature of the molecules at the oil-water interface surrounding

the droplets, which will alter their ability to stabilize the droplets against aggregation.

2.2 Lipid Oxidation in Emulsified Food Products

Many lipid containing processed foods are either oil-in-water or water-in-oil

emulsions (McClements, 2005; Friberg et al., 2004; Richards et al., 2002). While

oxidation is a problem in both, the majority of research has been done in oil-in-water

system. Therefore, in this chapter we mainly focus on lipid oxidation in oil-in-water type

emulsions, which represents products such as milk, infant formula, salad dressing,

mayonnaise, sauces, soups, beverages, cream, and some desserts, etc (McClements and

Decker, 2000; Okuda et al., 2005; McClements, 2005; Friberg et al., 2004). As well as

the food industry, the cosmetics, pharmaceutical and medical industries also utilize oil-in-

water emulsions as means to encapsulate, protect, and release bioactive lipids in their

17

products. For this reason, the number of studies attempting to understand the

physicochemical mechanisms underlying lipid oxidation in oil-in-water emulsions has

increased dramatically during the past decade.

Lipid oxidation is a great concern in the food industry because it causes physical

and chemical deteriorations, such as losses in important nutrients, formation of

potentially toxic reaction products (such as aldehydes and ketones), undesirable changes

in appearance and texture, and development of rancidity that shortens product shelf life

(Frankel, 1998; Coupland and McClements, 1996a; McClements and Decker, 2000;

Chaiyasit et al., 2007a; Decker and McClements, 2008). Lipid oxidation is favored in oil-

in-water emulsions because of the large contact surface between the oxidizable lipid

droplets and water-soluble compounds including oxygen and pro-oxidants, which

contribute to the initiation and propagation of oxidation reactions (Frankel, 1998;

Lethuaut et al., 2002; Villiere et al., 2005). There are many factors that can potentially

influence the rate of lipid oxidation in oil-in-water emulsions: fatty acid composition;

aqueous phase pH and ionic composition; type and concentration of antioxidants and pro-

oxidants; oxygen concentration; lipid droplet characteristics such as particle size,

concentration and physical state; interfacial characteristics such as thickness, charge,

rheology, and permeability (McClements and Decker, 2000; Okuda et al., 2005, Villiere,

et al., 2005; Kiokias et al., 2006). The susceptibility of emulsified lipids to oxidation also

depends on the surrounding molecular environment and interactions with other molecules

within the immediate vicinity of the droplets (Kellerby et al., 2006b).

18

2.2.1 Lipid Oxidation Mechanisms in Emulsions

Lipid oxidation is known to cause both physical and chemical deteriorations in

foods that contain lipids, such as milk, salad dressing, meat products, oils, nuts and even

in foods that contain relatively small amounts of lipids such as vegetable products.

Oxidation reactions occur due to the interaction of oxygen with unsaturated fatty acyl

groups in lipids. The rate of lipid oxidation in a particular product depends on a number

of factors: The presence of oxygen which is required for the development of oxidative

rancidity; the chemical composition of the lipids, - polyunsaturated fatty acids are more

susceptible than monounsaturated fatty acids; temperature – oxidation usually occurs

more rapidly at higher temperatures except in some conditions where high temperature

limit oxygen solubility; the presence of prooxidants; the nature of the reaction

environment (Frankel, 1985; Kim and Min, 2008). The quality attributes of lipid

containing food products can be dramatically decreased by lipid oxidation (Frankel,

1998; McClements and Decker; 2000; Min and Boff, 2002; Laguerre et al., 2007;

Chaiyasit et al., 2007a, Decker and McClements, 2008): flavor modifications due to

formation of hydroxyl acids; aroma changes due to formation of new volatile odorous

compounds; color changes (darkening) due to condensation reactions between proteins

and oxidation products; and texture changes due to cross-linking reactions between lipids,

lipids and proteins or protein-protein interactions induced by free radicals originating

from lipid oxidation. From a health and safety perspective, lipid oxidation is of particular

concern because it can lead to loss of valuable nutrients and formation of potentially toxic

reaction products (Kanner and Rosenthal, 1992). There are some in vivo studies that show

adverse affects of lipid oxidation products that could lead to Alzheimer’s disease

19

(Markesbery and Lovell, 1998), cancers (Boyd and Mcguire, 1998), atherosclerosis

(Glavind et al., 1990; Esterbauer et al., 1992; Esterbauer et al., 1993), inflammation, and

aging (Laguerre et al., 2007).

2.2.2 Lipid Oxidation Mechanism

Traditionally, lipid oxidation is assumed to be autocatalytic “free radical chain

reaction”. However, food products often contain prooxidants that can initiate lipid

oxidation reactions, such as transition metals (e.g., iron and copper), photosensitizers, and

enzymes (e.g., lipoxygenases). In addition, food products are often exposed to harsh

environmental conditions that can initiate lipid oxidation reactions, such as thermal

processing or exposure to UV light. Lipid oxidation involves a complex series of

chemical reactions that can be conveniently divided into three stages; initiation –

formation of free radicals; propagation – free-radical chain reactions; and termination –

formation of non-radical products (Pryor, 1976; Kanner and Rosenthal, 1992; Frankel,

1998; McClements and Decker; 2000; Min and Boff, 2002; Chaiyasit et al., 2007a;

Laguerre et al. 2007; Decker and McClements, 2008).

The classical lipid oxidation pathway can be described by the following reaction

scheme (Chaiyasit et al., 2007a; Erickson, 2002; Frankel, 1998, 2005, Kanner and

Rosenthal, 1992; Laguerre et al., 2007; Kim and Min, 2008; Decker and McClements,

2008):

20

Initiation LH + In•

L• + InH (1)

Propagation L• + O2 LOO•

(2)

LOO• + LH LOOH + L•

(3)

Termination LOO• + LOO• LOOL + O2

(4)

LOO• + L• LOOL

(5)

L• + L• LL

(6)

2.2.2.1 Initiation

According to Equation (1), the initiation step begins with the abstraction of

hydrogen from a free fatty acid molecule to form a free alkyl radical (L•), which is

normally considered to be the “rate-limiting step” in lipid oxidation. This reaction is

endothermic and usually occurs very slowly in the absence of initiators (e.g., heat, metal

ions, free radicals, reactive oxygen species, ultraviolet light, metallo-proteins, etc.) due to

its very high activation energy. The initiation of radical formation on a lipid normally

occurs at the carbon that requires the least energy to remove the hydrogen atom. The

alkyl radical formed is a free radical with an unpaired electron, which is therefore highly

unstable.

2.2.2.2 Propagation

The first step of propagation occurs when the alkyl radical (L•) formed during the

initiation stage interacts with an oxygen biradical to form a peroxyl radical (LOO•) (Eq.

(2)). The peroxyl radical has higher energy than alkyl radical. Therefore, it is more likely

to abstract hydrogen from another unsaturated fatty acid to form lipid hydroperoxide

21

(LOOH) and another alkyl radical (Eq. (3)). This stage is referred to as the “self-

sustained” radical chain reaction, and it occurs at a high rate that is characterized by a

rapid increase of hydroperoxide formation. The lipid hydroperoxides formed are

considered to be a primary oxidation product. However, once the lipid hydroperoxides

are formed, they can decompose which is induced by high temperature during thermal

processing or by various types of prooxidants such as transition metals and UV light.

Hydroperoxides themselves do not contribute to an off-flavor, but they are substrates for

rancidity due to the fact they are decomposed into low molecular weight volatile

compounds (Decker and McClements, 2008). Decomposition of hydroperoxides (LOOH)

can involve a “homolytic cleavage” between the two oxygen atom of the hydroperoxide

to form an alkoxyl (LO•) and a hydroxyl (•OH) radical (Min and Boff, 2002).

Alternately, reduced transition metals can decompose lipid hydroperoxide in a

reaction where an electron is transferred to the lipid hydroperoxide to form an alkoxyl

(LO•) and a hydroxyl anion (-OH). The alkoxyl radical (LO•) is more energetic than the

alkyl (L•) or peroxyl (LOO•) radicals, therefore, they can abstract a hydrogen from

another unsaturated fatty acids to further propagate the reaction, attack a pentadiene

group within the same fatty acid which can produce cyclic compounds, or abstract an

electron from the covalent bonds adjacent to the alkoxyl radicals to cleave the fatty acid

chain in what is known as “-scission reactions”. The -scission reaction is the main

pathway responsible for decomposition for unsaturated fatty acids into the low molecular

weight, volatile compounds contributing to rancid odors, including aldehydes, ketones,

alcohols, and short-chain hydrocarbons. Some nonvolatile secondary compounds are also

22

formed such as oxidized compounds still esterified to triacylglycerols, fatty acids and

triacylglycerol polymers and high molecular weight, nonvolatile oxidation products.

2.2.2.3 Termination

Termination steps occur when two free radicals interact turning into nonradical

oxidation products. The end products can vary depending on the type of radicals

interacting together [Eqs. (4), (5), (6)]. Most of the time, the food is already rancid before

termination reactions are highly prevalent. One exception can be frying oils where low

oxygen concentrations favor termination reactions versus lipid hydroperoxide formation

and decomposition.

2.2.3 Monitoring Lipid Oxidation

The extent of lipid oxidation can be investigated by measuring the 1) losses of

unsaturated fatty acids by GC or HPLC, 2) the formation of primary oxidation products

such as lipid hydroperoxides, and 3) the formation of secondary products such as

carbonyls and hydrocarbon gases (Halliwell and Chirico, 1993). To obtain an adequate

picture of the overall oxidative quality of foods, oxidation should be monitored with both

primary and secondary reaction products (Frankel and Meyer, 2000). Primary oxidation

products are allylic hydroperoixdes [-CH=CHCH(COOH)]. These hydroperoxides are

unstable which can decompose into different kinds of secondary oxidation products

including rearrangement to the products that have similar molecular weight, dimerization

to have a higher molecular weight materials, and fusion to give shorter-chain compounds

such as aldehydes and acids. Lipid oxidation can be inhibited by antioxidants but cannot

23

be prevented for the long term. Therefore, to monitor the extent of lipid oxidation should

be investigated by both primary and secondary lipid oxidation products because primary

products can provide important information about the initial stages of oxidation.

However, the primary oxidation products are decomposed over time, therefore, the

monitoring of secondary products can provide the information of the oxidation stage.

Besides, primary oxidation products are flavorless and odorless and therefore measuring

secondary oxidation products is important for correlation with rancid odors (Osborn and

Akoh, 2004).

2.2.4 Lipid Oxidation of Bulk oil vs. Emulsified Oil

The oxidation of emulsified lipids is different from that of bulk lipids for a

number of reasons: the presence of an aqueous phase containing prooxidants and

antioxidants; the presence of an oil-water interface and the partitioning of antioxidants,

prooxidants and oxidizable substrates between oil, interfacial, and water phases. The

emulsion droplet interface can attract or repel prooxidants and antioxidants through their

surface charges and by forming a physical barrier that influences the interactions between

lipid and water soluble prooxidants (Frankel et al., 1994; McClements and Decker, 2000;

Richards et al., 2002). Fritsch (1994) suggested that the impact of oxygen on the rate of

lipid oxidation is similar in water-in-oil emulsions and bulk oils due to the direct

exposure of the bulk oil to air. However, most oil-in-water emulsions are much more

prone to oxidation than bulk oils (Chaiyasit et al., 2007a). This is likely due to their large

surface areas which expose the lipids to aqueous phase prooxidants.

24

Appreciable differences have been reported between the mechanisms of lipid

oxidation in colloidal dispersions of lipids and in bulk lipids. In bulk oils, fats with a high

degree of unsaturated fatty acids are more vulnerable to lipid oxidation (Nawar, 1996;

Shahidi and Wanasundara, 1998). However, the opposite was found in micelle systems

with free fatty acids where oxidation rates increased when the degree of unsaturation of

fatty acids decreased (e.g. linolenic acid was more susceptible than arachadonic,

Miyashita et al., 1993; 1994). This effect may be due to differences in the molecular

packing of the fatty acids within the micelles: the greater the degree of unsaturation of the

fatty acids, the deeper they are buried within the hydrophobic core of the micelles. Thus,

they were less exposed to the prooxidants in the surrounding aqueous phase.

Many studies indicate that transition metals originating in the aqueous phase are

the most common cause of oxidative degradation of emulsified lipids. These water-phase

prooxidants are capable of interacting with lipid hydroperoxides located at the droplet

surface (Yoshida and Niki, 1992; Mei et al., 1998a; 1998b; Mancuso et al., 2000; Nuchi

et al. 2001, Dimakou et al., 2007). The interaction of lipid hydroperoxides (ROOH) with

both reduced and oxidized forms of transition metals can produce highly reactive peroxyl

(LOO•) and alkoxyl (LO•) radicals (Eqs. 1 and 2) that either attack other unsaturated

lipids (LH) within the oil droplets or at the oil-water interface to promote oxidation or in

the case of the alkoxyl radical promote -scission reactions that decompose fatty acids

into the low molecular weight volatile compounds that cause rancidity (Eqs. 3-5). These

lipid radicals could then react with other lipids in their immediate vicinity leading to a

chain reaction propagation of lipid oxidation (Eq. 6). The termination reaction occurs

when the lipid radicals react with each other (Eq. 7).

25

Fe 3+

+ LOOH Fe 2+

+ LOO• + H+

(1)

Fe 2+

+ LOOH Fe 3+

+ LO• + OH-

(2)

In general, iron is thought to be the most important transition metal prooxidant as

can be seen by the ability of iron binding proteins (e.g. transferrin and lactoferrin) to

strongly inhibit lipid oxidation in lipid dispersions (Huang et al., 1999; Mancuso et al.,

1999). Ferrous (Fe2+

) is much more prooxidative than ferric (Fe3+

) due to its higher

solubility and reactivity (Halliwell and Gutteridge, 1990). Ferric ions were only found to

be effective at decomposing lipid hydroperoxides when they are concentrated at the oil-

water interface (Mancuso et al., 2000). Since metals are important prooxidants in the

oxidative stability of emulsions, minimizing their concentrations is an effective method to

decrease oxidation rates (Mei et al., 1998a; Nuchi et al., 2002; Katsuda et al., 2008;

Wang and Wang, 2008).

2.2.5 Factors that Impact Lipid Oxidation in Emulsions

In this section we highlight some of the most important factors that impact lipid

oxidation in oil-in-water emulsions since these factors are critical for developing

technologies to inhibit lipid oxidation in food dispersions.

2.2.5.1 Droplet Interface Characteristics

Three different physical environments can be conveniently defined in oil-in-water

emulsions: the lipid inside the emulsion droplets; the interfacial layer surrounding the

26

droplets; and the aqueous continuous phase surrounding the interfacial layer. The

chemical composition of the interfacial layer may be fairly complex, and include

emulsifiers, antioxidants, minor lipid components (e.g. sterols and triacylglycerol

hydrolysis products) biopolymers, mineral ions, and lipid oxidation products (Dickinson

and McClements, 1995; McClements and Decker, 2000; Chaiyasit et al., 2007a). The

thickness of the interfacial layer is usually in the range of a few nanometers (e.g., 1 to 40

nm), and can be controlled by careful selection of emulsifiers and other ingredients. The

physical location of the various chemical reactants in emulsions therefore depends on

their lipid and water solubility characteristics and surface activities (Hiemenz and

Rajagopalan, 1997). For example, polar molecules tend to be located in the aqueous

phase, non-polar molecules in the oil phase, and surface active molecules in the

interfacial region (Chaiyasit et al., 2007a). A key place to modulate the initiation of lipid

oxidation in oil-in-water emulsions is therefore at the oil/water interface since this is the

place where lipid- and water-soluble components interact with each other, and where

surface-active reactants such as lipid hydroperoxides concentrate (Nuchi et al., 2002;

Villiere et al., 2005; Haahr and Jacobsen, 2008). The properties of the interfacial region

can be controlled in a variety of different ways to control the lipid oxidation reaction in

emulsions (Decker et al., 2005).

2.2.5.1.1 Interfacial Area

The interfacial area of an emulsion depends on the droplet concentration and

particle size: A = /6d32, where A is the interfacial area per unit volume of emulsion, is

the disperse phase volume fraction, and d32 is the surface-weighted mean diameter. The

27

size of the droplets in a food emulsion, and therefore the interfacial area, vary in different

food products. Droplet diameters can vary from larger than 100 m in salad dressings and

mayonnaise to less than 0.2 m in cream liqueurs and soft drinks. Since lipid oxidation

reactions in emulsions are greatly influenced by surface interactions between metals and

hydroperoxides, would expect droplet surface area to also be important factors

(McClements and Decker, 2000; Lethuaut et al., 2002). Nevertheless, studies of the effect

of droplet size on lipid oxidation in O/W emulsions are conflicting:

Some studies have found that the rate of lipid oxidation increased when

the surface area increased (Gohtani et al. 1999).

Some studies have found that the rate of lipid oxidation increased when

the surface area decreased (Hegenauer et al. 1979, Lethuaut et al., 2002,

Nakaya et al. 2005, Imai et al., 2008).

Some studies found that the lipid oxidation rate was fairly independent

of surface area (Coupland et al., 1996b; Shimada et al., 1996; Osborn

and Akoh, 2004; Dimakou et al., 2007; Kiokias et al., 2007;

Paraskevopoulou et al., 2007; Sun and Gunasekaran, 2009).

There are a number of physicochemical mechanisms that might influence the

affect of droplet size and surface area on the lipid oxidation rate: (i) as the interfacial

surface area increases more of the lipid phase is exposed to the surrounding aqueous

phase, which should promote lipid oxidation; (ii) as the interfacial area increases the

28

partitioning of reactants, pro-oxidants and antioxidants between the oil, water and

interfacial regions is altered, which may either promote or retard oxidation; (iii) as the

interfacial area increases the amount of surfactant at the interface increases, which may

decrease the surfactant present in the aqueous phase; (iv) more mechanical energy must

be supplied to increase the interfacial area of an emulsion containing small droplets

during homogenization, which may promote oxidation (Nakaya et al. 2005). At present,

the relative importance of these and possibly other mechanisms is not clearly understood,

and further work is required. It is likely that the effects of particle size will depend on the

precise nature of the system, e.g., the type and concentration of emulsifiers, pro-oxidants

and antioxidants present. However, research to date suggests that emulsion droplet size

and thus interfacial surface area is not a major factor in the oxidative stability of O/W

emulsions. This could be due to the extremely large surface area of all of the emulsions

used in these studies. These large surface areas could mean that surface area never limits

reaction rates.

2.2.5.1.2 Droplet Charge

The oxidative stability of oil-in-water emulsions depends on the electrical charge

on the droplet surfaces (Mei et al., 1998a; 1998b; Mancuso et al., 1999; 2000; Silvestre et

al., 2000; Boon et al., 2008). Surface charge determines electrostatic interactions, either

attractive or repulsive, between emulsion droplets and charged metals (Mei et al., 1998a;

Mancuso et al., 1999; Villiere et al., 2005; Haahr and Jacobsen, 2008). Droplet charge

can be manipulated by selecting appropriately charged emulsifiers (e.g., cationic, anionic

or neutral), or by using the electrostatic layer-by-layer (LbL) deposition method to

29

deposit charged biopolymers onto oppositely charged droplets (Shaw et al., 2007;

Klinkesorn et al., 2005a; 2005b; Djordjevic, et al., 2007). Several studies have shown that

anionic surfactants (such as sodium dodecyl sulfate, SDS) at droplet surfaces promote

lipid oxidation by attracting cationic transition metals to the surfaces (e.g., Fe2+

or Fe3+

),

whereas cationic surfactants (such as dodecyl trimethyl ammonium bromide, DTAB)

retard lipid oxidation by repelling these transition metals away from the surface (Mei et

al., 1998a; 1998b; Mancuso et al., 1999; 2000; Silvestre et al., 2000; Boon et al., 2008).

The impact of droplet surface charge has also been observed in oil-in-water

emulsions stabilized by proteins, where the rate of lipid oxidation was faster when the

protein-coated droplets were anionic (pH > pI) than when they were cationic (pH < pI)

(Figure 2.3) (Donnelly et al., 1998; Mei et al., 1998a; 1998b; Mancuso et al., 1999; 2000;

Hu et al., 2003a; 2003b; 2004b; Trunova et al., 2007; Djordjevic et al., 2008). Hu and

coworkers (2003b) found that the oxidation of cationic emulsion droplets produced by

emulsifying oil with proteins at pH 3.0 varied as a function of protein type. In this

experiment, oxidative stability was in the order sodium caseinate > whey protein isolate >

soy protein isolate. The density of the cationic charge of the emulsion droplets did not

correlate with oxidative stability suggesting that other factors such as droplet interfacial

thickness and/or the antioxidant properties of the protein were also involved in the ability

of the interfacial proteins to inhibit oxidation at pH 3.0. The impact of negative surface

charge on the rate of lipid oxidation in protein-stabilized emulsions was reported by

Villiere and coworkers (2005). This study compared stripped sunflower oil-in-water

emulsions (30 vol %) stabilized by sodium caseinate (NaCas) or bovine serum albumin

(BSA) at pH 6.5. The droplets in the NaCas-stabilized emulsions had a substantially

30

higher negative charge than those in the BSA-stabilized emulsions. Presumably the

transition metals irons were more strongly attracted to the surfaces of the lipid droplets in

the NaCas-stabilized emulsions. In the presence of EDTA, the oxidation rate was actually

lower in the emulsions stabilized by NaCas which was attributed to its higher ability to

scavenge free radicals.

31

Figure 2.3. Inhibition of iron-promoted lipid oxidation by a cationic emulsion droplet

interface created by using proteins as an emulsifier at pH values below

the pI of the protein.

Droplet charge also affects the location and activity of antioxidants via

attractive/repulsive electrostatic interactions (Mei et al., 1999). The activity of charged

antioxidants is often improved when they are located at the surface of charged lipid

particles because of electrostatic attraction. An anionic antioxidant (ascorbic acid) was

more effective at retarding lipid oxidation in the presence of positively charged lipid

micelles (Pryor et al. 1993). A negatively charged antioxidant (Trolox C) had higher

antioxidant activity in the presence of positively charged phospholipids (Barclay and

32

Vinqvist 1993), while a positively charged antioxidant (spermine) had higher antioxidant

activity in the presence of negatively charged phospholipids (Kogure et al. 1993). Mei et

al. (1999) found that emulsion droplets coated with an anionic surfactant (SDS) were less

oxidatively stable than those coated by a non-ionic surfactant (Brij) in the presence of an

anionic gallic acid. This phenomenon may be attributed to electrostatic repulsion of the

anionic antioxidants away from the anionic droplets thereby making the antioxidants

ineffective.

Controlling the electrical charge on emulsion droplets is therefore one of the most

important potential means of impacting lipid oxidation in oil-in-water emulsions. If the

droplets in an emulsion can be made to be neutral or positive, then they are less likely to

attract the cationic transition metal ions that frequently catalyze lipid oxidation in

emulsions.

2.2.5.1.3 Interfacial Thickness

Interfacial thickness can be manipulated by selecting emulsifiers with different

molecular dimensions (e.g., molecular weights, conformations, head group sizes, or tail

group sizes), or by using the LbL deposition method to deposit one or more biopolymer

layers around droplets (Shaw et al., 2007; Klinkesorn et al., 2005a; 2005b; Djordjevic et

al., 2007). Multilayer emulsions are discussed in more detail later in this review.

Emulsifiers with large molecular dimensions can be used to form thick interfacial

coatings around droplets that may protect against lipid oxidation. For example, the

coating could form a barrier that decreases interactions between lipids and

hydroperoxides or between lipids and aqueous phase prooxidants e.g. transition metals

33

(Silvestre, et al., 2000; Chaiyasit, et al., 2000). The influence of surfactant head-group

size on lipid oxidation in salmon oil-in-water emulsions was studied using Brij 76 and

Brij 700 as surfactants (Silvestre et al. 2000). The results showed that Fe2+

-promoted

decomposition of cumene hydroperoxide was lower in emulsions made with Brij 700 (10

times more polyoxyethylene groups than Brij 76), which was attributed to a thicker

interfacial layer on the emulsion droplets. The effect of surfactant tail group size has also

been studied using Brij-lauryl (contains 12 carbon atoms) and Brij-stearyl (contains 18

carbon atoms) (Chaiyasit et al. 2000). This study suggested that surfactant tail group size

played a minor role in lipid oxidation in oil-in-water emulsions, with increasing tail group

size slightly increasing oxidative stability.

2.2.5.1.4 Interfacial Permeability

Different surfactants have different packing properties at the oil-water interface

depending on their molecular dimensions, which may impact the diffusion of oxygen,

free radicals, and prooxidants through them (Villiere et al., 2005). One might expect that

an interfacial layer where the emulsifier molecules were closely packed or cross-linked

would provide more resistance to molecular diffusion into or out of the droplets.

Kellerby and coworkers (2006a) cross-linked casein on the interface of menhaden oil-in-

water emulsions which resulted in a cohesive interfacial protein layer that could not be

removed from the emulsion droplet by Tween 20. Although transglutaminase cross-

linked the interfacial casein, these emulsions did not show increased oxidative stability

when compared to untreated emulsions. In another study, O/W emulsions stabilized with

-lactoglobulin were heated to induce disulfide cross-links that produce a cohesive

34

protein layer. In this case, cross-linking the protein had no impact on the ability of iron

to decompose lipid hydroperoxides (Kellerby et al., 2006b). These studies suggest that

cohesive protein layers at the emulsion droplet surface do not increase oxidative stability

which could be due to the protein interface still being highly porous thus allowing iron to

diffuse through the anionic emulsion droplet interface where it can react with lipid

hydroperoxides. The impact of the density and packing behavior of small molecular

surfactants in O/W emulsions is unknown.

2.2.5.1.5 Interfacial Chemical Composition

The chemical composition of the interfacial layer surrounding lipid droplets may

influence the oxidative stability of food emulsions because of its ability to participate or

alter lipid oxidation reactions, e.g., by scavenging free radicals, chelating transition

metals, or interfering with hydroperoxides-transition metal interactions (Haahr and

Jacobsen, 2008). It is therefore possible to control lipid oxidation by controlling the

chemical composition of the interfacial layer surrounding the droplets, e.g., by selecting

appropriate emulsifiers and/or by adsorbing other materials onto the droplet surfaces.

Very little research has been done in this area with the exception of surface active

antioxidant compounds which are discussed in more detail below. Rampon and

coworkers (2001) reported that adducts between proteins and lipid oxidation products can

occur at the emulsion droplet interface during oxidation. Headspace propanal

concentrations have also been reported to decrease in protein-stabilized O/W emulsions

again suggesting interactions between lipid oxidation products in proteins at the emulsion

droplet interface (Shen et al. 2007). Finally, Leaver and coworkers (1999) found that

35

casein isolated from the interface of an oxidized soybean oil-in-water emulsion exhibited

an increased molecular weight which was suggested to be due to casein-lipid oxidation

adducts. Since formation of lipid-protein adducts will decrease the volatility of oxidation

products (Zhou and Decker, 1999), this could decrease rancidity.

2.2.5.2 Antioxidants

Antioxidants have been defined as “any substance that, when present at low

concentrations compared to those of an oxidisable substrate, significantly delays or

prevents oxidation of that substrate” (Halliwell and Gutteridge 1990). In food, the

definition of antioxidants was defined by Chipault (1962) as “substances that in small

quantities are able to prevent or greatly retard the oxidation of easily oxidizable materials

such as fats”. Addition of antioxidants maintains the nutritional quality and prolongs the

shelf life of lipid-containing foods (Halliwell et al., 1995) with compounds that scavenge

free radicals and/or chelate prooxidative metals being the most common food

antioxidants (Frankel, 1998; Decker and McClements, 2008). The effectiveness of

antioxidants in heterogeneous food systems such as emulsions depends on both chemical

and physical factors such as overall antioxidant concentration; distribution of antioxidants

and reactants in oil, water and interfacial phases; interactions with other food

components; and, environmental conditions such as pH, ionic strength and temperature

(Frankel et al., 1994; Mei et al., 1998a; Frankel and Meyer, 2000; Xie et al., 2007;

Medina et al., 2009).

Antioxidants generally work by inhibiting the formation of new radicals and/or

reducing the rate at which free radicals are formed. The two most common antioxidants

36

in emulsions are free radical scavengers also know as chain breaking antioxidants and

metal chelators.

2.2.5.2.1 Chain Breaking Antioxidants

Chain breaking antioxidants can inhibit free radical chain reactions by

scavenging radicals such as the peroxyl (LOO), alkyl (L) and alkoxyl radicals(LO) by

hydrogen donation (Eq. 1-3). These hydrogen donation reactions result in the formation

of an antioxidant radical. Each antioxidant can inactivate two free radicals since the

antioxidant radical can react with another free radical through a termination reaction (Eq.

4) (Decker and McClements, 2008).

AH + LOO

LOOH + A (1)

AH + L

LH + A (2)

AH + LO

LOH + A (3)

A + LO

LOA (4)

2.2.5.2.2 Transition Metal Chelators

As discussed above, transition metals are strong prooxidants in food emulsions.

Their prooxidative activity is related to their ability to react directly with triplet oxygen to

form superoxide radical anion (a potential source of free radicals at low pH; Kanner and

Rosenthal, 1992), decompose lipid hydroperoxides into free radicals via several redox

cycling pathways (Halliwell and Gutteridge, 1990) and produce alkyl radicals by

37

abstracting hydrogen from unsaturated fatty acids (Frankel, 1998). However, the latter

pathway is believed to occur very slowly and thus may not be important in promoting

lipid oxidation in foods (Reische, 1998).

Metal chelators are the main strategy by which the food industry inhibits lipid

oxidation due to transition metals. The antioxidant mechanisms of chelating agents

include sterically preventing the metal from interacting with the oxidizable lipids and

hydroperoxides, preventing metal redox cycling, preferentially binding the oxidized, less

reactive state of the metal and decreasing metal solubility (Kanner et al., 1987; Dunford,

1987; Graf and Eaton, 1990; Decker and McClements, 2001). In addition, chelators can

inhibit lipid oxidation by changing the physical location of metals so they are not

associated with lipids. For example, a study by Cho et al. (2003) showed that chelating

agents can increase the transfer of iron from inside lipid droplets into the surrounding

aqueous phase, thereby reducing the prooxidant activity of iron.

Since transition metals are toxic to most living systems, their reactivity is tightly

controlled in biological tissues. Control of iron reactivity is primarily accomplished by

proteins with highly specialized iron binding sites such as lactoferrin, transferrin,

phosvitin and ferritin. Unfortunately, during the processing of foods many of the metal

control mechanisms in biological tissues are destroyed and iron is released of iron can

enter the food from avenues such as water or processing equipment. A good example of

this is phosvitin. Phosvitin is the major iron storage protein in egg yolk. Iron bound to

phosvitin is largely inactive thus protecting the lipid in the egg. However, when egg yolks

are used to produce mayonnaise, the low pH environment causes phosvitin to release its

iron and promote lipid oxidation (Jacobsen et al., 2001). Therefore additional iron control

38

is often needed which is accomplished by the addition of food additives such as

polyphosphates, flavonoids, organic acids (e.g. citric acid) and ethylenediaminetetracetic

acid (EDTA). These chelators can be effective on their own as well as in combinations

with free radical scavengers since reduction in metal reactivity will decrease free radical

generation and thus spare free radical scavengers from decomposition (Decker and

McClements, 2001).

The most common metal chelators in foods contain multiple carboxylic acids such

as EDTA, polysaccharides (e.g. pectin and alginate) and citric acid, or contain phosphate

groups such as polyphosphates, phosphorylated proteins (e.g. casein and phosvitin) and

phytate (Decker, 1998). The effectiveness of chelating agents can decrease with

decreasing pH as their chelating groups become protonated and loss their metal binding

activity. In addition, while many of these compounds have the potential to inhibit metal

promoted lipid oxidation, not all are suitable as food additives since some could reduce

the bioavailability of essential minerals (e.g. phytate).

2.2.5.2.2.1 Ethylenediaminetetraacetic Acid

EDTA is a synthetic antioxidant that contains four carboxylate groups and two

amine groups that can form strong complexes with metal ions. EDTA is an extremely

potent antioxidant when present at concentration greater than the prooxidant metal

concentrations (Mahoney and Graf, 1986; Mei et al., 1998b; Mancuso et al., 1999;

Frankel et al., 2002; Alamed et al., 2006). However, when EDTA concentrations are less

than prooxidant metals, it can accelerate lipid oxidation presumably by increase the

39

solubility of the metal without reducing its chemical reactivity (Mahoney and Graf, 1986;

Decker, 1998; Jacobsen et al., 2001; Frankel et al., 2002).

2.2.5.2.2.2 Phosphates

Polyphosphates are also widely used as chelating agents in the food industry,

however, they do have some limitations in emulsions such as poor chelating efficiency

(Hu et al., 2004b), poor stability (Li et al., 1993), and a possibility of adversely affecting

protein functionality (Sofos, 1896). In general, the chelating efficiency of phosphates

increases with an increasing number of phosphate groups (Sofos, 1896). However,

neither sodium tripolyphosphate nor hexametaphosphate were able to inhibit lipid

oxidation in fish oil-in-water emulsions at pH 3 or 7. However, phosphorylated proteins

and peptides originating casein are natural polyphosphates that can be effective

antioxidants when used in foods. They have polar domains that contain phosphorylated

serine residues such as -SerP-SerP-SerP-Glu-Glu that can form complexes with calcium,

iron and zinc (Baumy and Brule, 1988; Bennett et al., 2000). Peptides from hydrolyzed

casein are effective at retarding lipid oxidation in oil-in-water emulsions, however, this

activity is likely to be due to both metal Chelation and free radical scavenging by amino

acids in the peptides (Diaz et al., 2003).

2.2.5.2.3 Impact of Physical Location on Antioxidant Effectiveness

The effectiveness of an antioxidant depends on its physical location within an

emulsion, e.g., oil, water and interfacial regions (Castle and Perkins, 1986; Yi et al.,

1991; Koga and Terao, 1994; 1995). This is because an antioxidant should be present at

40

the site of the lipid oxidation reaction to be effective. The effectiveness of antioxidants as

a function of their physical location has been described by the antioxidant “polar

paradox” (Porter et al., 1989; Porter, 1993; Frankel, 1998). In this hypothesis, polar

antioxidants are more effective in bulk oils and non-polar antioxidants are more effective

in oil-in-water emulsions. In bulk oils, polar antioxidants where thought to be more

effective since they would accumulate at the oil-air interface. However, recent

investigations have shown that due to the low polarity of air, polar antioxidants do not

tend to accumulated at oil-air interfaces (Chaiyasit et al., 2007a). Therefore, an

alternative hypothesis has been proposed to suggest that the increased effectiveness of

polar antioxidants is due to their ability to accumulate at the interface of associations

colloid structures in bulk oils such as reverse micelles and lamellar structures. These

association colloids can be formed by minor lipid components such as phospholipids and

free fatty acids in the presence of the small amounts of water naturally found in bulk oils

(Koga and Terao, 1994; 1995, Chaiyasit et al., 2008).

The antioxidant “polar paradox” also states that nonpolar antioxidants are more

effective oil-in-water emulsions since they are more highly retained in the oil droplet

where oxidation is most prevalent. This observation has been supported by studies

showing that predominantly non-polar antioxidants (-tocophorol, ascorbyl palmitate,

carnosol) are more effective antioxidants than their polar counterparts (Trolox, ascorbic

acid, carnosic acid and rosmarinic acid) in oil-in-water emulsions (Frankel et al., 1994,

1996a; 1996b; Hopia et al., 1996; Huang et al., 1996a;1996b).

Many researchers proposed that the ability of nonpolar antioxidants to be more

effective in O/W emulsions was not only due to their ability to be retained within

41

emulsion droplets but also their ability to accumulate at the oil-water interface where

oxidation is most prevalent. Most effective antioxidant compounds have structure that

allows them to act as surface active agents. In fact, antioxidants such as -tocopherol, -

tocopherol, TBHQ and propyl gallate have been found to accumulate at oil-water

interfaces as measured by their ability to decrease interfacial tension (Chaiyasit et al.,

2007a). The association of antioxidants with surfactants such as SDS and Brij has also

been observed by NMR and EPR (Heins et al., 2007a and b). Gunaseelan et al. (2006)

reported that 73% of -tocopherol is located in the interfacial region of a Brij 30-

stabilized octane-in-water emulsion.

Several recent studies have shown that not all antioxidants behave according to

the polar paradox hypothesis, indicating that antioxidant activity in complex systems is

more complicated than previous assumed (Laguerre et al., 2009). This can be seen from a

series of studies on the effectiveness of surface active antioxidant in O/W emulsions. The

idea that the effectiveness of antioxidants could be improved by increasing their

concentration at the oil-water interface prompted several researchers to synthesize surface

active antioxidants by covalently attaching a lipophilic hydrocarbon chain onto

antioxidants. Hunneche and coworkers (2008) found that the antioxidant activity of

ferulic acid could be improved in O/W emulsions oxidized by metmyoglobin when the

ferulic acid was attached to C11 or C12 hydrocarbons through a serine linkage. Medina

and coworkers (2009) also reported an increase in the antioxidant activity of

hydroxyltyrosol in O/W emulsions when it was esterified to hydrocarbon chains.

However, when p-hydroxyphenylacetic acid (HPA) was conjugated with either a butyl or

dodecyl group, both its retention in the oil droplet and it surface activity increased but its

42

antioxidant effectiveness was less than free HPA in O/W emulsions (Yuji et al., 2007).

One potential reason for this could be due to the process of covalently attaching a

hydrocarbon chain onto the antioxidant could decrease its ability to scavenge free

radicals. Therefore a subsequent study added C4, C8, or C12 esters of chlorogenic acid

(CGA) to O/W emulsions at equal free radical scavenging activity Sasaki et al. (2010).

Even under conditions of equal free radical scavenging activity, the antioxidant esters

were not more effective than the free CGA even though they had higher surface activity

and retention in the oil droplets. In another study, antioxidant activity in O/W emulsion

did not increase when hydrocarbons from C1 to C8 were esterified to CGA. However,

C12 CGA ester show enhanced activity but further increasing the hydrocarbon chain

length resulted in a decrease in the ability of the antioxidant to inhibit lipid oxidation

even though more of these antioxidant oxidant esters where retained in the emulsion

droplets (Laguerre et al., 2009).

The overall results of these studies indicate that while the antioxidants polar

paradox hypothesis has been a useful way to help understand antioxidant behavior in

emulsions, polarity cannot be used to consistently predict the ability of an antioxidant to

inhibit lipid oxidation in oil-in-water emulsions. This is likely due to the fact that most

antioxidants do not act alone and many antioxidants often have chemical properties that

can promote as well as inhibit lipid oxidation (Alamed et al., 2009). For example, some

phenolic antioxidants are able to chelate iron while others like ferulic acid which do not

have a galloyl moiety do not bind iron (Alamed et al., 2009). This chelating activity could

help explain why water-soluble polyphenols like those found in grape seed extract can

strongly inhibit lipid oxidation in O/W emulsions (Hu et al., 2004a). Another possible

43

reason why polarity might not predict antioxidant activity in O/W emulsions is that many

antioxidant compounds can participate in redox reactions with iron resulting in the

formation of ferrous ions which are stronger prooxidants than their oxidized counterpart,

ferric ions (Decker and McClements, 2001; Decker and Hultin, 1992; Mei et al., 1999).

Such prooxidant activity has been reported for ascorbic acid (Mahoney and Graf, 1986;

Decker and Hultin, 1992) gallic acid (Mei et al., 1999), caffeic acid (Sorenson et al.,

2008) and lycopene (Boon et al., 2009). Difficulty in prediciting antioxidant activity as a

function of polarity could also be due to interactions between antioxidants. One example

of this type of relationship is ability of ascorbic acid to regenerate oxidized -tocopherol

to reactivate -tocopherol in biological membranes (Porter, 1993; Buettner, 1993). This

could explain why the polar antioxidant propyl gallate was not effective in emulsion

containing stripped corn oil (Schwarz et al., 2000) but was effective in oil-in-water

emulsions made with commercial corn oil (Alamed et al., 2009). This difference could be

due to the ability of propyl gallate to interact with antioxidants naturally found in the

commercial corn oil.

The polar paradox is also complicated by the fact that the physical location of

antioxidants can also depend on their electrical charge characteristics. Charged

antioxidants are more water-soluble than their uncharged counter-parts, and therefore

they may be located in the aqueous phase away from the site of action. In contrast,

uncharged antioxidants often have low water-solubility, and are therefore located in the

oil phase or at the oil-water interface which increases their antioxidant activities

(Schwarz et al., 1996; Huang et al., 1999; Mei et al., 1999; McClements and Decker,

2000). The electrical charge, physical location and therefore activity of an antioxidant

44

that is a weak acid or base depends on solution pH (Mei, et al., 1999). When the pH is

near the pKa, the charge status and partitioning behavior of antioxidants alters (Wedzicha,

1988). Because of the myriad of physical and chemical properties of antioxidants, it is not

surprising that a single hypothesis such as the polar paradox can singly predict the ability

of a compound to inhibit lipid oxidation in emulsions.

2.2.5.3 Influence of Minor Components on Lipid Oxidation in Emulsions

Emulsions typically contain a number of functional ingredients, some of which

may either retard or promote lipid oxidation, e.g., non-adsorbed proteins, thickening

agents, minerals, sugars, colorants, flavors, buffering agents and alcohol. In this section,

we review the current understanding of some important ingredients on lipid oxidation in

emulsions.

2.2.5.3.1 Minor Oil Components

Even though, the crude oils undergo refining process to remove non-

triacylglycerols compounds, there are some minor components left in the oils that could

impact the oxidative stability of the oils. Some of these non-triacylglycerols compounds

are known as unsaponifiable fraction that consists of various components such as sterols,

carbohydrates, phenols (e.g. tocopherols), proteins, pigments (e.g. chlorophyll,

carotenoids, gossypol), trace metals, and pesticides (Chiyasit et al., 2007a). Some of

minor components from crude and refined soybean oils are shown in Table 2.1.

45

Table 2.1 Average compositions of crude and refined soybean oils.

Components Crude oil Refined oil

Triacylglycerols (%)

Phospholipids (%)

Unsaponifiable matter (%)

Phytosterols

Tocopherols

Hydrocarbons

Free Fatty Acids (%)

Trace metals

Iron (ppm)

Copper (ppm)

95-97

1.5-2.5

1.6

0.33

0.15-0.21

0.014

0.3-0.7

1.0-3.0

0.03-0.05

> 90

0.003-0.045

0.3

0.13

0.11-0.18

0.01

< 0.05

0.1-0.3

0.02-0.06

Adapted from (Pryde, 1980)

46

Commercial, refined oils typically containing 0.05-0.70% free fatty acids

(Chaiyasit et al., 2007; Jung et al., 1989; Pryor, 1976). Free fatty acids are well know

prooxidants in bulk oils (Frankel, 1998) but until recently there impact on oxidation in

O/W emulsions was unknown (Waraho et al., 2009). Addition of oleic acid (0 to 5.0% of

oil) to O/W emulsions increases both the negative charge of the emulsion droplets and the

formation of lipid hydroperoxides and headspace hexanal. Methyl oleate did not increase

oxidation rates and EDTA strongly inhibited lipid oxidation. These data suggest that free

fatty acids are able to migrate to the emulsion droplet surface where they make the

droplet anionic allowing attraction of transition metals that promote oxidation (Figure

2.4).

47

Figure 2.4. The proposed mechanism for free fatty acid promoted oxidation on

emulsified oil droplets. Mn+

= transition metal.

48

Phytosterols are also an important minor component of commercial, refined oils.

No studies have been conducted to determine if varying phytosterols impact lipid

oxidation rates in O/W emulsions. However, Cercaci and coworkers (2007) found that

phytosterols oxidize faster in emulsions than bulk oils. This was postulated to be due to

the surface activity of phytosterols which allowed them to concentrate at the oil-water

interface where they were more susceptible to oxidation.

Phospholipids are widely reported to inhibit lipid oxidation in bulk oils. Their

antioxidant activity has been attributed to their ability to chelate metals, increase the

partitioning of other antioxidants at the oil-water interface and form complexes with lipid

oxidation products that reduce the volatility of the compounds that cause rancidity. When

phospholipids are at the interface of the emulsion droplet they generally accelerate lipid

oxidation by producing an anionic interface (Shaw et al., 2007; Klinkesorn et al., 2005a;

2005b).

2.2.5.3.2 Influence of Other Emulsion Ingredients on Lipid Oxidation in Emulsions

2.2.5.3.2.1 Continuous Phase Proteins

There is growing interest within the food industry in replacing synthetic food

additives with more natural alternatives, which is mainly driven by consumer concerns.

Proteins are considered to be natural food additives that are generally recognized as safe

(GRAS), which are commonly added to stabilize food because of their ability to absorb to

oil droplet surfaces and prevent droplet aggregation (McClements, 2005). As discussed

earlier, these adsorbed proteins form a thin coating around the lipid droplets that can help

49

inhibit lipid oxidation when they are positively charged (pH < protein pI) and can repel

cationic transition metals and when they form thick interfacial layers that inhibit

transition metal- lipid interactions.

When proteins are used to make emulsions, they absorb to the surface of the lipid

droplet until the droplet surface is saturated with excess protein partitioning into the

continuous phase (Faraji et al., 2004). A number of studies have been shown that these

continuous proteins (e.g. whey protein isolate, sweet whey, casein, -lactoglobulin,

lactoferrin, and soy protein isolate) are capable of inhibiting lipid oxidation in O/W

emulsions (Taylor and Richardson, 1980; Tong et al., 2000; Hu et al., 2003; Faraji et al.,

2004; Elias et al., 2005; 2006; 2007). Several physicochemical mechanisms have been

proposed for the antioxidant activity of these proteins. Some proteins contain appreciable

amounts of amino acids that act as free radical scavengers, e.g., tyrosine, cysteine, and

tryptophan (Taylor and Richardson, 1980; Ostdal et al., 1996). In addition, many proteins

are able to bind transition metals when the pH is greater than the pI of the protein and

thus the protein is negatively charged (Faraji et al., 2004). This binding activity can

inhibit lipid oxidation by limiting the access of metal to the lipid or decreasing metal

reactivity (Elias et al., 2008). The antioxidant activity of globular proteins is often

increased by changing their confirmation by thermal processing or enzymatic hydrolysis

since this exposes antioxidant amino acid residues that are usually buried in the protein

interior (Taylor and Richardson, 1980; Tong et al., 2000; Pena-Ramos et al., 2004; Elias

et al., 2005; 2006; 2007; Peng et al., 2009).

Taylor and Richardson (1980) found an increase in the antioxidant activity of

various milk fractions containing whey proteins after heating. The antioxidant activity of

50

the whey proteins decreased after a sulfhydryl blocker (iodoacetic acid, IAA) was added

to the emulsions, which suggested that the sulfhydryl groups from whey protein played

an important role in the antioxidant mechanism. A later study by Tong et al. (2000) on

the impact of non-adsorbed whey protein fractions (WPF) on lipid oxidation in salmon

oil-in-water emulsions attempted to identify the physicochemical origin of the protein’s

antioxidant activity. These workers found that whey proteins could act as free radical

scavengers and chelating agents, and that cysteine and tyrosine were particularly

important for this function.

Faraji et al. (2004) found that when whey protein isolate (WPI), soy protein

isolate (SPI), or sodium caseinate (CAS) were used to stabilize oil-in-water emulsions,

only a fraction of them adsorbed to the droplet surfaces, while the rest remained in the

aqueous continuous phase. The non-adsorbed proteins had good antioxidant activity at

pH 7 but not at pH 3, which may have been due to greater binding of cationic transition

metals to anionic non-adsorbed proteins at neutral pH. This study highlighted the

importance of solution pH in determining the antioxidant capacity of proteins.

The influence of free radical scavenging and chelation of transition metals by -

lactoglobulin in surfactant-stabilized oil-in-water emulsions was studied by Elias et al.

(2005). Surface exposed antioxidant amino acids were found to be preferentially

oxidized prior to the emulsified lipids, thereby retarding the rate of lipid oxidation. When

the whey proteins were subjected to limited enzymatic hydrolysis, which increased the

exposure of antioxidant amino acid residues, their free radical scavenging and metal

chelating abilities increased (Elias et al., 2006).

51

The oxidative stability of amino acid residues varies greatly (Stadtman, 1993).

The pseudo-first-order rate constants of amino acid side chains oxidized with hydroxyl

radicals were studied by Sharp et al. (2004). The relative reactivity rate was ranked Cys >

Trp, Tyr > Met > Phe > His > Iie > Leu > Pro. A study from Hernandez-Ledesma et al.

(2005) also showed different radical scavenging antioxidant activity of several amino

acids and peptide fragments using the oxygen radical absorbance capacity (ORAC)

method. They found that tryptophan exhibited a higher antioxidant activity than

methionine and cysteine, respectively. Therefore, proteins with different amino acid

compositions can interact with lipid hydroperoxides and lipid-derived free radicals

differently, which will affect their overall ability to act as antioxidants in emulsions

(Viljanen et al., 2005).

2.2.5.3.2.2 Polysaccharides

Polysaccharides are widely used as functional ingredients in food emulsions

because of their ability to thicken, gel, or stabilize them (Matsumura et al., 2003; Kishk

and Al-Sayed, 2007). Many polysaccharides have also been found to have antioxidant

activity in oil-in-water emulsions, which has been attributed to mechanisms such as free

radical scavenging, transition metal binding, and viscosity enhancement (Shimada et al.,

1992; 1994; 1996; Matsumura et al., 2003; Paraskevopoulou et al., 2007; Chen et al.,

2010). On the other hand, some polysaccharide ingredients may increase the rate of lipid

oxidation because they contain high levels of transition metal impurities, such as iron

(Katsuda et al., 2008).

52

Xanthan gum was found to act as an antioxidant in soybean oil-in-water

emulsions, which was attributed to its ability to bind Fe2+

ions at anionic pyruvate sites

along the polysaccharide chain (Shimada et al., 1992). Another study by Shimada et al.

(1996) suggested that the ability of certain polysaccharides to inhibit lipid oxidation in

oil-in-water emulsions may be due to an increase in continuous phase viscosity, thus

lowering the diffusion rate of oxygen and oil droplet collision probability. Nevertheless,

it is important to distinguish between the macroscopic and microscopic viscosities of

biopolymer solutions. A biopolymer solution may have an extremely high macroscopic

viscosity, but the molecular diffusion of small molecules (like oxygen, free radicals or

transition metals) is not restricted because of the large space between the polysaccharides

chains in solution. In other words, the microscopic viscosity of the aqueous phase is not

greatly increased by the presence of the polysaccharide. Indeed, studies have shown that

pullulan and maltodextrin can greatly increase the viscosity of O/W emulsions but do not

greatly retard lipid oxidation rate, while the glycoproteins, gum arabic and soluble

soybean polysaccharides (SSPS) do not greatly increase the viscosity but have good

antioxidant activity (Matsumura et al., 2000; Matsumura et al., 2003). SSPS was found to

suppress the initiation stage and the breakdown of lipid hydroperoxides more efficiently

than gum arabic, which was attributed to the presence of more antioxidant amino acids on

its protein component. The same group found that pectin had a stronger free radical

scavenging activity than SSPS, but that its overall antioxidant activity was less

(Matsumura et al., 2003). Tragacanth gum has been shown to have the ability to act as a

radical chain-breaker due to its ability to donate hydrogen atoms (Shimada et al., 1992).

53

While polysaccharides can increase the physical stability of O/W emulsions by

increasing viscosity they can also cause emulsion destabilization by causing depletion

flocculation. Therefore, it polysaccharides are going to be used to inhibit oxidation, this

must be accomplished at concentrations that do not negatively impact the physical

stability of emulsions. Using emulsion physical stability as a selection criteria for the

upper allowable concentration of polysaccharides (0.1 wt %), Chen and coworkers (2010)

investigated the ability of continuous phase low methoxyl pectin, high methoxyl pectin,

α-carrageenan and sodium alginate to inhibit lipid oxidation in polyoxyethylene (23)

lauryl ether (Brij 35) stabilized O/W emulsions at neutral pH. All polysaccharides were

able to inhibit lipid oxidation but low methoxyl pectin was the most effective. This was

thought to be due to the higher ferrous binding capacity low methoxyl pectin since none

of the polysaccharides were effective free radical scavengers.

Some polysaccharides are surface active, and may therefore be located at the oil-

water interface, rather than in the continuous aqueous phase. For example, gum arabic,

modified starch and propylene glycol alginate are widely used by the food and beverage

industries as functional ingredients to emulsify oils (Chanamai and McClements, 2002;

Minemoto et al., 2002; Djordjevic et al., 2007; Paraskevopoulou et al., 2007). These

polysaccharide emulsifiers generally produce anionic emulsion droplets and therefore can

increase oxidation rates. However, the high local concentration of these polysaccharides

at the site of the lipid oxidation reaction, may enable them impact lipid oxidation

reactions if they contain components that can act as antioxidants.

However, not all carbohydrates act as antioxidants. Some studies from Mabrouk

and Dugan (1961), Mabrouk (1964), and Yamaguchi and Yamada (1981) showed that

54

sugars such as pentose, hexose, and reducing disaccharides could act as strong

prooxidants in methyl linoleate and linoleic acid in O/W emulsions. Another study from

Yamauchi, et al. (1984) showed that reducing sugars have the ability to reduce transition

metal ions from Fe3+

to Fe2+

, which is a stronger prooxidant, thus accelerating lipid

oxidation. On the other hand, the sugar alcohols have been shown to exhibit antioxidant

activity in safflower oil emulsions (Sims et al., 1979; Yamauchi et al., 1982).

2.2.5.3.2.3 Surfactant Micelles

Like proteins, when small molecule surfactants are used to stabilize emulsions,

they absorb onto the emulsion droplet surface until the interface is saturated with the

remaining surfactant partitioning into the continuous phase. Surfactant micelles form in

the continuous phase of the emulsions when the concentration of surfactants exceeds the

critical micelle concentration or CMC. While continuous phase surfactant micelles are

too small to solubilize triacylglycerols they can can solubilize lipophilic or amphiphilic

components, such as antioxidants and prooxidants, which alters their distribution between

the oil, water and interfacial regions (Nuchi et al., 2002, Cho et al., 2002; Richards et al.,

2002). Overall, the presence of surfactant micelles in oil-in-water emulsions inhibits lipid

oxidation (Richards, et al., 2002). Nuchi and coworkers (2002) found that surfactant

micelles are able to solublize lipid hydroperoxides and promote their movement from

lipid droplets into the surrounding aqueous phase. The presence of surfactant micelles

therefore could inhibit lipid oxidation in the emulsions by preventing the free radicals

formed by decomposing hydroperoxides from attacking unsaturated lipids in the droplet

core. A study from Cho et al. (2002) suggested that surfactant micelles could promote

55

movement of lipid-phase iron out of oil droplets which could inhibit lipid oxidation. In

contrast, a study from Richards et al. (2002) showed that Brij micelles could solubilize

antioxidant out of the emulsion droplet into the continuous phase. Polar antioxidants such

as propyl gallate and TBHQ where susceptible to micelle solubilization while the

concentration of the non-polar antioxidant BHT in the emulsion droplet was not influence

by surfactant micelles. This study suggested that surfactant micelles could increase the

oxidative stability of emulsions by removing antioxidants from the site of oxidation.

2.2.6 Controlling Lipid Oxidation Using Structured Emulsions

A number of researchers have examined the use of structured emulsions to

overcome some of the limitations of using conventional emulsions to inhibit lipid

oxidation. Figure 2.5 showed several kinds of structured emulsions that can be used in

food industry. Methods of preparing some types of these emulsions were discussed in an

earlier section. Some of the potential benefits of these systems are discussed below.

56

Figure 2.5. Examples of different kinds of structured emulsion systems that

can be utilized in foods.

57

2.2.6.1 Filled Hydrogel Particles

Filled hydrogel particles consist of oil droplets contained within hydrogel

particles that are dispersed within an aqueous continuous phase, and can therefore be

referred to as oil-in-water-in-water (O/W1/W2) emulsions. The concentration, particle

size distribution and spatial location of the oil droplets within the hydrogel particles can

be varied. Also the properties of the hydrogel particles themselves can be varied such as

their composition, charge, digestibility, stability, permeability, and environmental

responsiveness (McClements et al., 2007).

In general, a variety of different methods can be used to form filled hydrogel

particles (Pich and Adler, 2007; Chen et al., 2006; Norton and Frith, 2001; Zhang et al.,

2007). These include methods based on phase separation of biopolymer solutions,

injection of biopolymer solutions into gelling solutions, fragmentation of macroscopic

biopolymer gels, and formation of biopolymer gels within double emulsions

(McClements et al., 2007). Typically, these methods include mixing a pre-formed oil-in-

water emulsion with an aqueous solution containing one or more biopolymers, and then

causing the biopolymers to phase separate and/or gel by altering the environmental

conditions.

Filled hydrogel particles have been studied as a means of improving the physical

and chemical stability of emulsified lipids. For example, Lamprecht et al. (2001) created

filled hydrogel particles using an aggregative phase separation technique to encapsulate

and protect -3 fatty acids. They found that the filled hydrogel particles hardened by

ethanol were the most stable to lipid oxidation because the biopolymer shell could be

formed to help prevent lipid oxidation as well as improve the physical stability of the

58

emulsion. Wu et al. (2005) also used filled hydrogel particles to encapsulate fish oils

while Weinbreck et al. (2004) used them to stabilize flavor oils. Filled hydrogel particles

may be designed to retard lipid oxidation by concentrating antioxidant hydrogel

molecules (such as proteins or polysaccharides) in close proximity to the encapsulated

lipid droplets.

2.2.6.2 Solid Lipid Particles

Solid lipid particle (SLP) emulsions are similar to conventional emulsions because

they consist of emulsifier-coated lipid droplets dispersed in an aqueous continuous phase.

However, the lipid phase in SLP is either fully or partially solidified. The morphology

and packing of the crystals within the lipid phase can often be controlled to obtain

particular functional attributes (Saupe et al., 2005; Souto et al., 2004; Uner et al., 2004;

Wissing et al., 2004; Wissing and Muller, 2002). As with conventional emulsions, the

size and concentration of the lipid droplets can be controlled, as can the nature of the

interfacial coating surrounding the lipid phase.

SLP emulsions can be formed using the same methods as for conventional

emulsions or nanoemulsions depending on the particle size required, e.g., high or low

intensity methods. The main difference is that the lipid phase (or at least a part of it) will

be solid rather than liquid at the application temperature. To prepare an emulsion it is

therefore necessary to heat the lipid and aqueous phases above the melting point of any

crystalline material in the lipid phase prior to homogenization (Saupe et al., 2005; Souto

et al., 2004; Uner et al., 2004; Wissing et al., 2004; Wissing and Muller, 2002; Schubert

and Muller-Goymann, 2005). It is then important to always maintain the temperature of

59

the emulsion above the crystallization temperature of the highest melting lipid to prevent

any fat solidification within the homogenizer. After formation, the emulsion can then be

cooled in a controlled manner to promote crystallization of some or all of the lipids

within the droplets.

Solid lipid particles may be able to improve the stability of chemically labile

lipophilic components by trapping them inside structured solid matrices so the

oxidatively sensitive lipids are protected from aqueous phase prooxidants and oxygen

(McClements et al. 2007). However, a study on the influence of carrier lipid physical

state on the oxidative stability of octadecane oil-in-water emulsions found that methyl

linolenate oxidized more quickly in emulsions containing solid droplets than those

containing liquid droplets (Okuda et al. 2005). It was postulated that crystallization of the

octadecane caused methyl linolenate molecules to migrate from the interior of the lipid

droplets where they were partly protected against oxidation, to the exterior of the lipid

droplets where they were more exposed to water-soluble transition metals. If this

approach is going to work, then it is important that the polyunsaturated oils are trapped

within the interior of crystalline lipids, which may be achieved by controlling carrier oil

composition, surfactant type and cooling conditions.

2.2.6.3 Multilayer Emulsions

Multilayer emulsions are similar to conventional emulsions, but the interfacial

layer surrounding the lipid droplets is engineered using a layer-by-layer deposition

method. Multilayer oil-in-water (OM/W) emulsions are composed of oil droplets

dispersed in an aqueous medium, with each oil droplet being surrounded by a multilayer

60

interfacial coating. This coating usually consists of emulsifier and biopolymer molecules.

A major advantage of using multilayer emulsions is the ability to manipulate the

properties of the interfacial layer surrounding the oil droplets, e.g., its chemical

composition, charge, thickness, internal structure, permeability, rheology and

environmental responsiveness (McClements et al., 2007).

A multiple step process is usually used to prepare multilayer emulsions. A

“primary” oil-in-water emulsion is first prepared by homogenizing an oil and aqueous

phase together in the presence of a charged water-soluble emulsifier. The primary

emulsion contains small charged oil droplets dispersed in an aqueous continuous phase.

A “secondary” emulsion is created by adding an oppositely charged polyelectrolyte that

adsorbs to the droplet surfaces, thereby producing a two-layer emulsifier-polyelectrolyte

coating. The oil droplets can be coated by nano-laminated interfaces containing three or

more layers by successive deposition of oppositely charged polyelectrolytes. Each

polyelectrolyte layer can be deposited onto the droplet surfaces using either a one or two-

step mixing procedure as described below (McClements et al., 2007):

2.2.6.3.1 One-Step Mixing

An oil-in-water emulsion containing electrically charged droplets is mixed with

an aqueous solution of oppositely charged polyelectrolyte molecules, leading to direct

absorption of the polyelectrolyte onto the droplet surfaces through electrostatic attraction.

61

2.2.6.3.2 Two-Step Mixing

An oil-in-water emulsion is mixed with a polyelectrolyte solution at a pH where

the polyelectrolyte molecules do not absorb onto the droplet surfaces (e.g., where the

droplets and polyelectrolyte are both negative). Then the pH of the solution is adjusted to

change the electrical charge on the droplets and/or polyelectrolyte to allow the

polyelectrolyte to adsorb onto the droplet surfaces via electrostatic attraction.

Any excess non-adsorbed polyelectrolyte molecules remaining in the continuous

phase can be removed by a washing step done between each electrostatic deposition step,

e.g. by centrifugation or filtration.

Emulsions stabilized by multiple layers of emulsifiers have the potential to

increase the oxidative stability of lipids. As mentioned previously, emulsion droplets with

thick interfacial membranes exhibit increased oxidative stability (Silvestre et al. 2000) as

do emulsion droplets with a cationic surface (Donnelly et al. 1998; McClements and

Decker 2000). Stabilizing emulsion droplets with multiple layers of emulsifiers have the

potential to inhibit lipid oxidation by forming both thick and cationic emulsion droplet

interfacial membranes by using 2 or more emulsifier layers to increase thickness and by

using a cationic biopolymer for the most outer layer to produce a positive charge. A

number of studies have shown that lipid oxidation can be retarded in multilayer

emulsions by making the net charge on the droplets positive (Shaw et al., 2007;

Klinkesorn et al., 2005a; 2005b; Djordjevic et al., 2007).

In studies on the oxidative stability of tuna oil-in-water emulsions stabilized by a

multilayer system consisting of lecithin and chitosan it was found that emulsion droplets

coated with only lecithin oxidized quicker than the combination of lecithin-chitosan as

62

determined by measuring both lipid hydroperoxide and thiobarbituric acid reactive

substances (Klinkesorn et al. 2005a). Cationic SDS-chitosan layers are also able to

inhibit the degradation of flavor oil components such as limonene (Djordjevic et al.

2007). The improved stability of the lecithin-chitosan emulsion compared with the

lecithin emulsion is possibly due to cationic repulsion of iron and other prooxidative

metals by the positively charged lecithin-chitosan emulsion droplet interfacial membrane.

This trend was also seen in an emulsion stabilized by SDS and chitosan where the

formation of thiobarbituric acid reactive substance (TBARS) was faster in the anionic

SDS-stabilized emulsion compared to a cationic SDS-chitosan multilayer emulsion.

Addition of pectin to add a third layer on to the emulsion droplets resulted in the

formation of an anionic interfacial membrane. The oxidative stability of this tertiary

emulsion was similar to the secondary, cationic, SDS-chitosan emulsion (Shaw et al.,

2008). Katsuda et al. (2008) found that anionic emulsions with β-lactoblobulin-citrus

pectin multilayer had similar oxidative stability as the cationic emulsions stabilized with

β-lactoblobulin alone. These data suggest that by increasing the thickness of the

interfacial membrane of the emulsion droplet, it can be possible to overcome the

prooxidative effects of an anionic emulsion droplet interface.

2.3 Conclusions

Lipid oxidation is a major problem leading to deterioration of polyunsaturated

lipids, which needs to be prevented because it would cause undesirable changes in flavor,

texture, appearance, and nutritional quality of food products. Since many lipid containing

foods are in emulsified form, a thorough understanding of lipid oxidation mechanisms in

63

emulsions should be develop in order to develop innovative technologies to solve this

problem. Numerous techniques (Figure 2.6) can be applied to inhibit lipid oxidation in

oil-in-water emulsions including interfacial engineering to control the composition,

thickness or charge of the interfacial layer that separates the encapsulated lipids from the

surrounding aqueous phase. The selection of antioxidants is also important to ensure that

they are located at the major site of the lipid oxidation reaction. Structured emulsions,

such as filled hydrogel particles, multilayer emulsions or solid lipid particles, may be

used to protect lipids against chemical degradation, but further work is needed in this

area. Due to the high susceptibility of emulsified lipids to oxidation, it might be necessary

to use combinations of the techniques mentioned in this chapter to effectively retard lipid

oxidation and improve the shelf-life, utilization and quality of food emulsion systems.

For example, use of a combination of cationic interfacial membrane, emulsion droplet

antioxidant (tocopherols) and metal chelators (EDTA) can be an extremely effective way

to protect omega-3 fatty acids in emulsions (Djordjevic et al. 2004).

64

Figure 2.6. Locations where the chemical and physical properties of oil-in-water

emulsions can be altered to impact lipid oxidation reactions.

65

CHAPTER 3

PROOXIDANT MECHANISMS OF FREE FATTY ACIDS IN STRIPPED

SOYBEAN OIL-IN-WATER EMULSIONS

3.1 Abstract

Prooxidant role of free fatty acids was studied in soybean oil-in-water emulsions.

Addition of oleic acid (0 to 5.0% of oil) to the emulsions increased lipid hydroperoxides

and headspace hexanal formation and increased the negative charge of the emulsion

droplet with increasing oleic acid concentration. Methyl oleate (1.0% of oil) did not

increase oxidation rates. The ability of oleic acid to promote lipid oxidation in oil-in-

water emulsions decreased with decreasing pH with dramatic reduction in oxidation

observed when the pH was low enough so that the oleic acid was not able to increase the

negative charge of the emulsion droplet. Ethylenediaminetetraacetic acid (EDTA, 200

m) strongly inhibited lipid oxidation in emulsions with oleic acid indicating that

transition metals were responsible for accelerating oxidation. Oleic acid hydroperoxides

did not increase oxidation rates suggesting that hydroperoxides on free fatty acids are not

strong prooxidants in oil-in-water emulsion. These results indicate that the oxidative

stability of oil-in-water emulsions could be greatly improved by maintaining low levels

of free fatty acids.

66

3.2 Introduction

Lipid oxidation is a common cause of quality deterioration in lipid-containing

food products resulting in changes in quality attributes such as taste, appearance, texture,

and shelf life as well as the changes in nutritional quality with very much concern in

developing of potentially toxic reaction products (Nawar, 1996; Frankel, 1998; Coupland

and McClements, 1996a; McClements and Decker, 2000). Many lipid containing food

products are in the form of oil-in-water emulsion such as milk, fruit and nutritional

beverages, salad dressing, soups and sauces. There are a lot of factors that affect lipid

oxidation rates in oil-in-water emulsions including fatty acid composition, oxygen

concentration, type and concentration of antioxidants, interfacial characteristics of

emulsion droplet such as electrical charge, and the ability of aqueous phase prooxidants

such as transition metals to interact with oxidizable lipids (McClements and Decker,

2000).

Even though edible oils are refined to remove undesirable components,

commercial oils still contain small amounts of minor components including free fatty

acids, monoacylglycerols, diacylglycerols, phospholipids and sterols. These minor

components are surface active compounds that could affect lipid oxidation by altering the

chemical and physical properties of oils. Free fatty acids are formed during lipid

extraction and refining by hydrolysis of triacylglycerides by lipases and high temperature

in the presence of water. Free fatty acids are removed from crude oils by neutralization

and deodorization. However, these refining steps are not 100% efficient with commercial

oils typically containing 0.05-0.70% free fatty acids (Pryde, 1980; Jung et al., 1989;

Chaiyasit et. al., 2007a). Besides negatively affecting oil quality by causing foaming and

67

reducing the smoke point of the oils, free fatty acids can also act as prooxidant in bulk

oils. Several researchers have reported that the prooxidant effect of free fatty acids in

bulk oils is the result of the carboxylic acid group since methyl esters of free fatty acids

are not prooxidative (Miyashita and Takagi, 1986; Mistry and Min, 1987; Yoshida et al.,

1992; Frega et al., 1999). The current hypothesis for the prooxidant activity of free fatty

acids is their ability to form complexes with trace metals and promote the acid-catalyzed

decomposition of lipid hydroperoxides.

Free fatty acids are surface active compounds since they are more polar than

triacylglycerols due to the presence of an unesterified carboxylic acid groups. The surface

activity of free fatty acids allows them to diffuse and concentrate at the water-lipid

interface of the oil-in-water emulsions (Nuchi et al., 2002). Thus free fatty acids could

potentially make the emulsion droplet more negatively charged when pH values are

above their pKas (4.8-5.0 for medium- and long-chain (C ≥10) fatty acids in aqueous

solution; White, 1950; Spector, 1975; Lieckfeldt, et al., 1995). Previous researches have

shown that negatively charged oil-in-water emulsion droplets can attract prooxidant

transition metals that can increase metal-lipid interactions thus accelerating oxidation

(Yoshida and Nikki, 1992; Fukuzawa et al. 1995; Mei et al., 1998a; 1998b).

While there are several studies on the prooxidant effects of free fatty acids in bulk

oils, there is almost no studies on the impact of free fatty acids on lipid oxidation of oil-

in-water emulsions. Since the mechanisms of lipid oxidation in oil-in-water emulsions

can be very different than bulk oils, (McClements and Decker, 2000), this study as

conducted to investigate the role free fatty acids on oxidation in emulsions as a function

of free fatty acid concentration and pH as well as in the presence of free fatty acid

68

hydroperoxides and metal chelators. Understanding how free fatty acids impact lipid

oxidation in oil-in-water emulsions could provide fundamental knowledge that could be

used to improve the oxidative stability of oils in emulsion and other food dispersions.

3.3 Materials and Methods

3.3.1 Materials

Soybean oil was purchased from a local retail store. Oleic acid and methyl oleate

were purchased from Nu-Chek Prep, Inc. (Elysian, MN). Ethylenediaminetetraacetic acid

(EDTA), potassium phosphate monobasic, and potassium phosphate dibasic

heptahydrate, silicic acid (100-200 mesh, 75-150 m, acid washed), activated charcoal

(100-400 mesh), polyoxyethylene (20) sorbitan monolaurate (Tween 20), ammonium

thiocyanate and iron (II) sulfate heptahydrate were obtained from Sigma Chemical Co.

(St. Louis, MO., U.S.A.). Iso-octanol, n-hexane, 2-propanol, methanol, 1-butanol were

purchased from Fisher Scientific (Fair Lawn, NJ). All the chemicals used in this

experiments were analytical grade or purer. Glassware was incubated in 3 mM HCL

overnight to remove metals followed by rinsing with double-distilled water before use.

Double-distilled water was used throughout the study.

3.3.2 Methods

3.3.2.1 Preparation of Stripped Soybean Oil

Stripped soybean oils as prepared according to Boon et al. (2008) was used in all

experiments. In short, silicic acid (100 g) was washed three times with a total of 3 L of

69

distilled water followed by filtering with Whatman filter paper in a Buchner funnel and

drying at 110C for 20 h. The washed silicic acid (22.5 g) and activated charcoal (5.625

g) were suspended in 100 and 70 ml n-hexane, respectively. A chromatographic column

(3.0 cm internal diameter x 35 cm height) was then packed sequentially with 22.5 g of

silicic acid followed by 5.625 g activated charcoal and then another 22.5 g silicic acid.

Thirty grams of soybean oil was dissolved in 30 ml of n-hexane and passed through the

column by eluting with 270 ml of n-hexane. To retard lipid oxidation during stripping,

the collected triacylglyerols were held in an ice bath which was covered with an

aluminum foil. The solvent in the stripped oils was removed with a vacuum rotary

evaporator (RE 111 Buchi, Flawil, Switzerland) at 37C and traces of the remaining

solvent were removed by flushing with nitrogen. Then three grams of the stripped oil

were transferred into 3-ml vials, flushed with nitrogen and kept at -80 C for subsequent

studies.

3.3.2.2 Preparation of Free Fatty Acids

Hydroperoxides are primary products from lipid oxidation and are substrates for

decomposition of secondary products such as aldehydes and ketones. Therefore, in these

studies, the initial hydroperoxides in commercial oleic acid were removed to insure that

the effects of added free fatty acids on oxidation in emulsions were not due to the

addition of lipid hydroperoxides. Hydroperoxide reduction process was adapted from

Miyashita and Takagi (1986) using silicic acid. Column chromatography was set up using

a glass syringe (2.0 cm internal diameter x 10.5 cm height) whose outlet was covered

with three layers of nylon membrane filters (Nylaflo Nylon membrane filters 47 mm

70

0.45 m, GelmanSciences, Ann Arbor, MI). Silicic acid was pretreated as described in

the preparation of stripped soybean oil section. Silicic acid (5.0 g) was suspended in 22.5

mL n-hexane and then poured into the column. Three grams of oleic acids diluted in 3

mL of n-hexane were loaded onto the column and followed by elution with 40.0 ml of n-

hexane. The eluent was collected in an ice bath covered with an aluminum foil to retard

lipid oxidation. The solvent with oleic acid was kept in a glass tube with a seal cap at

-80oC until use. Solvent was removed by flushing with nitrogen prior to use.

Hydroperoxide residues in oleic acids after the treatment were reduced from 6.8 mmol to

< 0.05 mmol of hydroperoxides /kg of fatty acids. For experiments on the effect of oleic

acid hydroperoxides on oxidation rates, purified oleic acid was incubated at 55oC in the

dark to allow for formation of hydroperoxides (5.0 mmol/kg of oleic acid final

concentration) and was then added to the 1.0% stripped soybean oil-in-water emulsions,

keeping total oleic acid concentration at 1.0% of the oil.

3.3.2.3 Emulsion Preparation and Storage Conditions

Oil-in-water emulsions were prepared using 1.0% (wt) stripped soybean oil in a

10 mM phosphate buffer solution (pH 7.0). Tween 20 was used as an emulsifier at a 1:10

emulsifier: oil ratio. The emulsion was prepared by adding purified oleic acid in n-hexane

into a beaker and flushing with nitrogen gas to remove the solvent. Phosphate buffer,

Tween 20, and stripped soybean oil were then added to the beaker and a coarse emulsion

was made by blending with a hand-held homogenizer (M133/1281-0, Biospec Products,

Inc., Bartlesville, OK) for 2 min. The coarse emulsion was then homogenized with a

microfluidizer (Microfluidics, Newton, MA) at a pressure of 9 kbar for three passes.

71

During homogenization, ice was used to cover the homogenizer chamber and coil in

order to maintain the emulsion temperature at ≤ 25oC. One milliliter of each emulsion

was transferred into 10 ml GC vials (Supelco, Bellefonte, PA), capped with aluminum

caps with PTFE/Silicone septa and stored in the dark at 15oC.

3.3.2.4 Measurement of Particle Size Distributions and Zeta Potential (ζ)

Samples for droplet size distribution and zeta potential measurements were

diluted into 10 mM phosphate buffer at the same pH as the emulsions at an

emulsion:buffer ratio of 1:50. Both particle size distributions and zeta potential of the

emulsions were analyzed in a ZetaSizer Nano-ZS (Malvern Instruments, Worcestshire,

UK). Each measurement was repeated twice at room temperature.

3.3.2.5 Measurement of Lipid Oxidation

Lipid hydroperoxides which are primary products of lipid oxidation were

measured using the method adapted from Shantha and Decker (1994). Each sample (0.3

mL) was vortexed three times (10 second each) with 1.5 mL of an isooctanol +

isopropanol (3:1 v:v) solution. The samples were then centrifuged for 2 min at 3400 g

(Centrific TM Centrifuge, Fisher Scientific, Fairlawn, NJ) and 0.2mL of the upper

organic layer or diluted with methanol/butanol (depending on the extent of lipid

oxidation) was mixed with 2.80 mL of methanol + butanol solution (2:1 v:v), 15 L of

3.94 M ammonium thiocyanate and 15 L of ferrous iron solution (prepared by mixing

0.132 M BaCl2 and 0.144 M FeSO4). The absorbance of the samples was measured at 510

nm using a Genesys 20 spectrophotometer (ThermoSpectronic, Waltham, MA) 20 min

72

after the addition of the iron. Hydroperoxide concentrations were quantitated based on a

cumene hydroperoxide standard curve.

Hexanal was measured as a secondary lipid oxidation product as described by

Boon et al. (20) using a GC-17A Shimadzu gas chromatography equipped with an AOC-

5000 autosample (Shimadzu, Kyoto, Japan). Emulsions (1 mL) in 10 mL glass vials

capped with aluminum caps with PTFE/Silicone septa were shaken and heated at 55°C

for 13 min in an autosampler heating block before measurement. A 50/30 m

DVB/Carboxen/PDMS solid-phase microextraction (SPME) fiber needle from Supelco

(Bellefonte, PA) was injected into the vial for 1 min to absorb volatiles and then was

transferred to the injector port (250oC) for 3 min. The injection port was operated in split

mode, and the split ratio was set at 1:5. Volatiles were separated on a Supleco 30 m x

0.32 mm Equity DB-1 column with a 1 m film thickness at 65oC for 10 min. The carrier

gas was helium at 15.0 mL/min. A flame ionization detector was used at a temperature of

250oC. Hexanal concentrations were determined from peak areas using a standard curve

prepared from authentic hexanal.

3.3.2.6 Statistical Analysis

All experiments were conducted in triplicate samples and were repeated at least

two times. Data were presented as mean + standard deviation. Data results were analyzed

by analysis of variance (ANOVA) using SPSS (SPSS Inc., Chicago, IL). The differences

between mean values were compared using Duncan’s multiple range test with

significance defined as p 0.05.

73

3.4 Results and Discussion

3.4.1 Physical Stability of Emulsions

The droplet size of the emulsions was measured immediately after emulsion

preparation and every 24 hr throughout storage. Emulsion droplets size ranged from 165

– 185 nm and did not change significantly during the course of the experiments (data not

shown). The stability of the emulsions was also confirmed by no visual observation of

creaming during storage (data not shown). These indicated that the emulsions were stable

to droplet aggregation, flocculation, or coalescence (McClements, 2007).

3.4.2 Effect of Oleic Acids Concentrations on the Physical and Chemical Properties

of Oil-in-Water Emulsions

Previous studies by Miyashita and Takagi (1986) and Mistry and Min (1987)

showed that free fatty acids are prooxidative in bulk soybean oil. To determined how free

fatty acids influence the oxidation of oil-in-water emulsions, different concentrations of

oleic acid (1.0, 2.5, and 5.0% of oil concentration) were added during emulsification of

the Tween 20-stabilized soybean oil-in-water emulsions at pH 7.0. Tween 20 was chosen

because it is a nonionic surfactant so it would have a lower impact on emulsions droplet

surface charge than ionic emulsifiers. Since commercial free fatty acids can contain lipid

hydroperoxides, they were purified prior to addition to the emulsions.

The droplet surface charge or zeta potential (ζ) of the emulsions with varying

concentrations of oleic acid are shown in Table 3.1. Control emulsions had a surface

charge of -9.54 mV compared to – 27.50, -44.95, and – 53.70 mV when the emulsions

74

contained 1.0, 2.5 and 5.0% oleic acid, respectively. Tween 20 stabilized oil in water

emulsions have been previously reported to be negatively charged (Hur et al., 2009;

Malhotra and Coupland, 2004; Li et al., 2001). It is unclear if this negative charge is due

to impurities in the Tween 20. The decrease in surface charge with increasing oleic acid

concentrations suggested that acid groups of the oleic acid were migrating to and

concentrating at the lipid-water interface of the emulsion droplet.

Table 3.1. The droplet surface charge or zeta potential (ζ) of 1.0% stripped soybean

oil-in-water emulsions without (control) and with addition of 1.0, 2.5, and

5.0% oleic acids and from 1.0% methyl oleate (oil wt.) at pH 7.0.

.

Sample Zeta potential (mV)

control

addition of 1.0% oleic acid



addition of 1.0% methyl oleate

- 9.54

- 27.50

- 44.95

- 53.70

- 12.2

The rates of lipid oxidation of 1.0% stripped soybean oil-in-water emulsions with

added oleic acid were followed by monitoring lipid hydroperoxide formation as an

75

indictor of primary oxidation products and hexanal formation as an indicator of

secondary oxidation products (Figures 3.1). Increasing oleic acid concentrations

significantly increased both lipid hydroperoxides and headspace hexanal formation. After

6 h of storage, 5.0% oleic acid exhibited a significantly higher hydroperoxide

concentration than all other emulsions. After 1 day of storage, there were dramatic

increases in hydroperoxide formations in all the emulsions containing added oleic acid

(Figure 3.1a). A similar trend was observed for hexanal formation (Figure 3.1b). From

this study, free fatty acids were clearly shown to act as powerful prooxidant in oil-in-

water emulsions with as little as 1.0% oleic acid in the oil phase accelerating both lipid

hydroperoxide and hexanal formation.

76

Figure 3.1. Formation of lipid hydroperoxide concentration (a) and hexanal (b) in

1.0% stripped soybean oil-in-water emulsions at pH 7.0 without (control)

and with addition of 1.0, 2.5, and 5.0% oleic acids (oil wt.) during storage

at 15ºC in the dark for 6 days.

0

200

400

600

800

1000

1200

0 1 2 3 4 5 6Hyd

rop

ero

xid

e c

on

ce

ntr

ati

on

(m

mo

l/k

g o

il)

Oxidation time (Days)

a

0% oleic acid

1.0% oleic acid

2.5% oleic acid

5.0% oleic acid

0

10

20

30

40

50

60

70

80

0 1 2 3 4 5 6He

xa

na

l c

on

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ntr

ati

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(m

mo

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g o

il)


b0% oleic acid

1.0% oleic acid

2.5% oleic acid

5.0% oleic acid

77

3.4.3 Effect of Methyl Oleate and Oleic Acid on the Physical and Chemical

Properties of Oil-in-Water Emulsions

The ability of the free fatty acids to accumulate at the oil-water interface can

decrease emulsion droplet surface charge (Table 3.1). Since cationic metals are strong

prooxidant in oil-in-water emulsion, negatively charged emulsion droplets could attract

metals thus accelerating lipid oxidation rates. However, Miyashita and Takagi, (1986)

also postulated that free fatty acids could promote lipid oxidation via acid catalyzed

hydroperoxide decomposition into free radicals. To determine if a free carboxylic acid

group was necessary for the prooxidant activity of fatty acids, both oleic acid and methyl

oleate were added to the stripped soybean oil-in-water emulsions at 1.0% of the oil

content. Table shows that the droplet surface charge was -9.54, -27.50 and -12.20 mV for

emulsions with no added fatty acids, oleic acid and methyl oleate, respectively. Figure

3.2 shows that while free oleic acid accelerated both lipid hydroperoxides and headspace

hexanal formation, the presence of methyl oleate did not change oxidation rates

compared to the control with no added fatty acids. These data show that a free carboxylic

acid group is necessary for the prooxidant activity of fatty acids in oil-in-water emulsions

which is in agreement with Miyashita and Takagi, (1986), Mistry and Min, (1987), and

Frega et al., (1999) who found similar results in bulk oils.

78


1.0% stripped soybean oil-in-water emulsions without (control) and with

addition of 1.0% oleic acids and 1.0% methyl oleate (oil wt.) at pH 7.0

during storage at 15ºC in the dark for 8 days.

0

100

200

300

400

500

600

700

800

900

0 1 2 3 4 5 6 7 8

Hyd

rop

ero

xid

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on

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(m

mo

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g o

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a

control

oleic acid

methyl oleate

0

10

20

30

40

50

60

70

80

0 1 2 3 4 5 6 7 8

He

xa

na

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(m

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bcontrol

oleic acid

methyl oleate

79

3.4.4 The Effect of pH on Physical and Chemical Properties of Oil-in-Water

Emulsions Containing Oleic Acid

Transition metals are found abundantly in nature and thus can end up in food from

a variety of sources including water, packaging, processing equipment and ingredients

including fats and oils (Taylor, 1987). Transition metals such as iron primarily accelerate

lipid oxidation by promoting the decomposition of lipid hydroperoxides into highly

reactive alkoxy and peroxy radicals which can abstract hydrogen from fatty acids thus

further propagating oxidation (Chaiyasit, et al., 2007a). One of the most important factors

that influence the prooxidant activity of iron is its physical location which dictates its

ability to interact with lipid hydroperoxides. For example, iron-lipid hydroperoxide

interactions and thus lipid oxidation increases dramatically in negatively charged

emulsion droplets where iron is attracted to the emulsion droplet surface (Mancuso et al.,

1999; 2000).

The pKa of medium- and long-chain (C ≥10) fatty acids in aqueous solution is

approximately 4.8-5.0 (White, 1950; Spector, 1975; Lieckfeldt, et al., 1995). If the ability

of free fatty acids to promote lipid oxidation in oil-in-water emulsions is due to their

ability to make emulsion droplet more negatively charged, then when the pH of the

emulsions is below the pKa of the free fatty acids, the emulsion droplet charge would

decrease as the acid group became protonated and lipid oxidation rates would decrease.

To test this hypothesis, the emulsion droplet charge and oxidative stability of 1.0%

stripped soybean oil-in-water emulsions with and without 1.0% oleic acid (wt. % in oil)

was measured over the pH range of 2.0 to and 8.0.

80

The droplet surface charge of the emulsions as a function of pH is shown in

Figure 3.3. The surface charge ranged from -2.22 to -42.10 mV and -2.05 to -19.50 mV

from pH 2.0 to 8.0 for emulsions with and without oleic acid, respectively. A significant

increase in emulsion droplet charge in the presence of oleic acid occurred at pH values

above 4.0. The large increase in droplet charge above the pKa of the free fatty acids is

likely due to the deprotonation of the fatty acids producing a highly polar anionic

carboxylic acid which can then migrate to the oil-water interface.

The influence of pH on the oxidative stability of stripped soybean oil-in-water

emulsions with and without 1.0% oleic acid (wt. % in oil) is shown in Figure 3.4. As the

pH of the emulsions without added fatty acids was decreased from 8.0 to 6.0, both

hexanal and hydroperoxide formation decreased. In emulsions without oleic acid at 2.0

and 4.0, the concentration of lipid hydroperoxides (< 55 and 95 mmol/kg oil,

respectively) and headspace hexanal (< 850 and 300 mmol/kg oil, respectively) was low

throughout the storage study. Addition of oleic acid only showed a prooxidant effect at

pH 6.0 and 8.0. Overall, oxidation rates in all the emulsions decreased as the negative

charge of the emulsion droplet decreased (Table 3.1). Added oleic acid was only

prooxidative at pH values above the pKa where the free fatty acids were able to make the

emulsion droplet more negatively charged. These data further support the notion that an

unprotonated carboxylic acid group is necessary for the prooxidant activity of fatty acids.

It is unclear why the negative charge of the emulsions droplet without added fatty acids

81

Figure 3.3. The droplet surface charge or zeta potential (ζ) of 1.0% stripped soybean

oil-in-water emulsions with addition of 1.0 % oleic acids (oil wt.) at pH

2.0, 4.0, 6.0, and 8.0.

pH

8 +

1.0

% o

leic

acid

pH

8

pH

6 +

1.0

% o

leic

acid

pH

4 +

1.0

% o

leic

acid

pH

4

pH

2 +

1.0

% o

leic

acid

pH

2

pH

6

-45

-40

-35

-30

-25

-20

-15

-10

-5

0

ze

ta p

ote

nti

al (m

V)

82


1.0% stripped soybean oil-in-water emulsions with addition of 1.0% oleic

acids (oil wt.) at pH 2.0, 4.0, 6.0, and 8.0 during storage at 15ºC in the

dark for 10 days.

0

200

400

600

800

1000

1200

0 1 2 3 4 5 6 7 8 9 10

Hyd

rop

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xid

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(m

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apH 2

pH 4

pH 6

pH 8

pH 2 + 1.0% oleic acid




0102030405060708090

100110120130

0 1 2 3 4 5 6 7 8 9 10He

xa

na

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on

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(m

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bpH 2

pH 4

pH 6

pH 8





83

decreased. It is possible that this could be due to the presence of free fatty acids in the

Tween 20.

3.4.5 The Effect of EDTA and Fatty Acid Hydroperoxides

From the above data it is unclear if the prooxidant activity of free fatty acids in

oil-in-water emulsions is due to acid catalyzed decomposition of preexisting lipid

hydroperoxides or is due to the ability of surface active free fatty acids to concentrate at

the emulsion droplet oil-water interface where they can make the droplet more anionic

thus attracting transition metals which promote the oxidation. To better understand which

mechanism is most prevalent, EDTA was added to the emulsions to evaluate the role of

transition metals in the prooxidant activity of free fatty acids. There was no significant

difference in emulsion droplet charge in stripped soybean oil-in-water emulsions in the

presence of 200 M EDTA (data not shown). However, lipid oxidation was significantly

inhibited by EDTA with only small amounts of hydroperoxides being formed and almost

complete suppression of headspace hexanal formation (Figure 3.5). Since EDTA did not

change the surface charge of the emulsion droplets, these data strongly suggest that the

prooxidant mechanism of free fatty acids is due to their ability to attract prooxidant

metals to the surface of the emulsion droplet where they can interact with oxidizable

lipids versus the acid catalyzed decomposition of preexisting lipid hydroperoxides.

84



acids (oil wt.) without and with 200 m EDTA at pH 7.0 during storage at

15ºC in the dark for 21 days.

0

100

200

300

400

500

600

700

800

900

0 1 2 3 4 5 6 7 8 9 101112131415161718192021

Hyd

rop

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(m

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a

control

oleic acid

oleic acid + EDTA

0

10

20

30

40

50

60

70

80

90

0 1 2 3 4 5 6 7 8 9 101112131415161718192021

He

xa

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(m

mo

l/k

g o

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bcontrol

oleic acid

oleic acid + EDTA

85

Hydroperoxides are primary products of lipid oxidation that are normally found in

fats and oils due to formation during processing, transportation, and during storage

(Paulose and Chang, 1973; Billek et al., 1978). In most lipid-containing foods contain

hydroperoxides, even high quality lipids still contain about 10-100 nmol/g lipid. In oil-in-

water emulsions, another source of hydroperoxides comes from the surfactant, for

example, phospholipids and Tween 20 which were found to have 4-35 mol

hydroperoxides/g surfactant (Nuchi et al., 2001). Lipid hydroperoxides are strong

prooxidants due to the ability to decompose into free radicals in the presence of light,

metals and high temperatures (Decker and McClements, 2008; Nuchi et al., 2001).

Hydroperoxides on free fatty acid, methyl esters and triacylglycerols increases the surface

activity of the parent molecules presumably due to the presence of oxygen which

increases polarity. Fatty acids hydroperoxides are more surface active than methyl esters

or acylglycerols hydroperoxides (Nuchi et al., 2002). Therefore, it is possible that the

presence of free fatty acid hydroperoxides could increase oxidation rates in oil-in-water

emulsions since the free fatty acid hydroperoxides would concentrate at the oil-water

interface where they can readily interact with iron bound to the emulsion droplet surface.

In the previous experiments in this study, oleic acid was purified so that

hydroperoxides concentrations were low (< 0.05 mmol/kg fatty acid). To determine if

oleic acid hydroperoxides accelerated lipid oxidation in oil-in-water emulsion, oleic acid

oxidized at 55oC in the dark was blended with purified oleic acid and added to the 1.0%

stripped soybean oil-in-water emulsions at 5.0 mmol hydroperoxides/kg oleic acid

(keeping total oleic acid concentration at 1.0% of the oil). The zeta potential of emulsion

droplets with oleic acid low in hydroperoxides (-31.3 ± 0.5) was not significantly

86

different (p<0.05) than emulsion droplets with oleic acid high in hydroperoxides (-29.2

0.4). According to the lipid oxidation results shown in Figure 3.6, neither lipid

hydroperoxides nor hexanal formation rates were different in oil-in-water emulsions with

low or high oleic acid hydroperoxides concentration. Many studies have shown that lipid

oxidation rates are very dependent of the concentration of lipid hydroperoxides. Studies

from Nuchi and coworkers (Nuchi et al., 2002; Nuchi et al., 2001) found that either

Tween 20 or linoleic acid hydroperoxides decreased the lag phase of hexanal formation

in oil-in-water emulsions. Therefore, it was somewhat surprising that the addition of oleic

acid hydroperoxides did not increase oxidation rates. However, the amount of

hydroperoxides in the original emulsions was 35 M/kg emulsion. Therefore the added

oleic acid hydroperoxides only increased total hydroperoxides concentrations 1.5%.

Even though the oleic acid hydroperoxide level is low in comparisons to hydroperoxides

on the triacylglycerols, it does show that a highly surface active oleic acid

hydroperoxides are not more prooxidative than hydroperoxides on the triacylglycerols.

87



acids with low and high in hydroperoxides (oil wt.) at pH 7.0 during

storage at 15ºC in the dark for 8 days.

0

100

200

300

400

500

600

700

800

900

0 1 2 3 4 5 6 7 8Hyd

rop

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acontrol

oleic acid with low peroxide

oleic acid with high peroxide

0

10

20

30

40

50

60

70

80

90

0 1 2 3 4 5 6 7 8Hexan

al co

ncen

trati

on

(m

mo

l/kg

oil)


bcontrol

oleic acid with low peroxide

oleic acid with high peroxide

88

3.5 Conclusions

Oil-in-water emulsions containing free fatty acids are extremely susceptible to

lipid oxidation when the pH of the emulsions is higher than the pKa of the free fatty

acids. Under these conditions the carboxylic acid groups on the free fatty acids are

negatively charged. Since the charged free fatty acids are surface active they migrate to

the oil-water interface of emulsion droplets where they decrease the negative charge of

the emulsion droplet. Inhibition of lipid oxidation in oil-in-water emulsions containing

free fatty acids by EDTA indicates that the most likely mechanisms for the prooxidant

activity of free fatty acids is the attraction of cationic transition metals to the emulsion

droplet surface where they can interact with lipid and promote oxidation. These results

indicate that the oxidative stability of oil-in-water emulsions could be greatly improved

by maintaining low levels of free fatty acids.

89

CHAPTER 4

IMPACT OF FREE FATTY ACID CONCENTRATION AND STRUCTURE ON

LIPID OXIDATION IN OIL-IN-WATER EMULSIONS

4.1 Abstract

Free fatty acids are strong prooxidants in both bulk and emulsified oils. Addition

of oleic acid to an oil-in-water emulsions increased lipid hydroperoxide and hexanal

formation at free fatty acid concentrations as low as 0.1% of the lipid. The prooxidant

effect of free fatty acids was dependent on fatty acid type with lipid oxidation rates being

in the order of linolenic < linoleic < oleic. There were no significant differences in lipid

oxidation rates when free fatty acid isomers with cis or trans double bonds were

compared. The prooxidant activity of the free fatty acids was postulated to be due to their

ability to attract prooxidant metals as well as co-oxidize the triacylglycerol in the oil.

Overall, these results show that the oxidative stability of oil-in-water emulsions is

strongly linked to both the concentration and type of free fatty acids present.

90

4.2 Introduction

Oil-in-water emulsions are widely utilized not only in the food industry but also

the cosmetics, pharmaceutical and medical industries as a means to encapsulate, protect

and release bioactive lipids (McClements et al., 2007). However, these industries face a

major problem that causes the deterioration of these products because of the susceptibility

to lipid oxidation. Therefore, unsurprisingly, numerous studies have been conducted in

order to gain a more thorough understanding of lipid oxidation processes in oil-in-water

emulsions with the goal of developing methods to control these detrimental reactions (for

review see Waraho et al., 2011). There are many types of minor components present in

edible oils after the commercial refining and deodorization process such as free fatty

acids, mono- and diacylglycerols, phospholipids, tocopherols, chlorophylls, carotenoids,

hydroperoxides, thermally oxidized compounds and metals. In bulk oils, these minor

components can act as antioxidants while others can be prooxidative (Choe, 2008).

A study from Kinsella et al. (1978) showed that free fatty acids are more

susceptible to autoxidation than esterified fatty acids in bulk oils. Free fatty acids contain

both a hydrophobic hydrocarbon tail and hydrophilic carboxylic acid head in the same

molecule. This combination of hydrophobic and hydrophilic groups allows the free fatty

acid molecule to concentrate at the surface of water-oil interfaces (Choe, 2008). Mistry

and Min (1987) found that free fatty acids had ability to reduce surface tension of bulk

soybean oil as well as increased the diffusion rate of oxygen from the headspace into the

oil thus increasing lipid oxidation rates. In oil-in-water emulsions, free fatty acids are

prooxidative due to their ability to concentrate at the emulsion droplet surface where they

attract prooxidative transition metals that promote oxidation (Waraho et al., 2009).

91

Commercial oils typically contain 0.05-0.70% free fatty acids (Pryde, 1980; Jung

et al., 1989; Chaiyasit et al., 2007a). While free fatty acids are known to promote lipid

oxidation in oil-in-water emulsions, very little is known about how fatty acid type

impacts their prooxidative activity. For instance, are unsaturated free fatty acids more

prooxidative than saturated fatty acids due to their increased susceptibility to oxidation?

Also, the shape of fatty acids changes quite dramatically with the introduction of cis and

trans double bonds (Figure 4.1). Do these differences in geometric configuration alter

prooxidative activity due to the ability of linear saturated or trans fatty acids to more

efficiently pack at the emulsion droplet surface? Therefore, the objective of this study

was to gain a better understanding of how free fatty acids type impacts lipid oxidation in

oil-in-water emulsions.

92

a) Oleic acid b) Elaidic acid

c) Linoleic acid d) Linoelaidic acid

e) Linolenic acid

Figure 4.1. The structure of a) oleic acid (18:1, cis), b) elaidic acid (18:1, trans), c)

linoleic acid (18:2, cis-cis), d) linoelaidic acid (18:2, trans-trans) and e)

linolenic acid (18:3, cis-cis-cis).

93


4.3.1 Materials

Soybean oil was purchased from a local retail store. Oleic acid (18:1, cis), elaidic

acid (18:1, trans), linoleic acid (18:2, cis-cis), linoelaidic acid (18:2, trans-trans) and

linolenic acid (18:3) were purchased from Nu-Chek Prep, Inc. (Elysian, MN). Miglyol

812 (Medium chain triglyceride; MCT) was purchased from Sasol (Witten, Germany).

Potassium phosphate monobasic, potassium phosphate dibasic heptahydrate, silicic acid

(100-200 mesh, 75-150 m, acid washed), activated charcoal (100-400 mesh),

polyoxyethylene (20) sorbitan monolaurate (Tween 20), ammonium thiocyanate and iron

(II) sulfate heptahydrate were obtained from Sigma Chemical Co. (St. Louis, MO.,

U.S.A.). Iso-octanol, n-hexane, 2-propanol, methanol, 1-butanol were purchased from

Fisher Scientific (Fair Lawn, NJ). All the chemicals used in these experiments were

analytical grade or purer. Glassware was placed in 3 mM HCL overnight to remove

transition metals followed by rinsing with double-distilled water and then drying before

use. Double-distilled water was used throughout the study.

4.3.2 Methods

4.3.2.1. Removing of Polar Components form the Oils and Free Fatty Acids

Soybean oils were stripped prior to making emulsions to remove polar minor

components such as tocopherols, free fatty acids, mono- and diacylglycerols,

phospholipids and hydroperoxides by passing it through a silicic acid and activated

94

charcoal chromatographic column using n-hexane as eluent according to the method as

described in previous work (Waraho et al., 2009). Commercial oleic, linoleic and

linolenic acids were found to contain significant amounts of lipid hydroperoxides. Since

these hydroperoxides could impact lipid oxidation rates, they were removed by diluting

in n-hexane and then passing through a glass syringe packed with silicic acid using the

methods described in a previous study (Waraho et al., 2009). Elaidic and linoelaidic acids

contained less than 0.05 mmol hydroperoxides/kg of fatty acid (similar to the other fatty

acids after stripping) so they were used directly.


The preparation of a 1% (wt.) stripped soybean oil-in-water emulsion was similar

to prior a study (Waraho et al., 2009). All emulsions were made with 10 mM phosphate

buffer solution (pH 7.0) using Tween 20 as an emulsifier at a 1:10 emulsifier:oil ratio.

The emulsions were prepared by adding purified free fatty acids in n-hexane into a beaker

and flushing with nitrogen gas to remove the solvent. Then stripped soybean oil and

Tween 20 in phosphate buffer were added to the beaker and a coarse emulsion was made

by blending with a hand-held homogenizer (M133/1281-0, Biospec Products, Inc.,

Bartlesville, OK) for 2 min. The coarse emulsion was then homogenized with a

microfluidizer (Microfluidics, Newton, MA) at a pressure of 9 kbar for three passes.

Since the melting points of elaidic and linoelaidic acids are higher than ambient

temperature, these fatty acids were diluted into medium chain triacylglycerols (MCT)

followed by heating at 550C for 10 minutes in order to melt the fatty acids. In the study

with elaidic and linoelaidic acids, all the fatty acids were subjected to this same heat

95

treatment so all samples were handled the same. MCT was used as a nonoxidizable lipid

to minimize lipid oxidation at these elevated temperatures. The emulsions were prepared

by adding the fatty acid/MCT stock solution to stripped soybean oils and Tween 20 in 10

mM phosphate buffer and then emulsions were prepared by the same method described

above. After emulsion preparation, samples were transferred into 10 ml GC vials

(Supelco, Bellefonte, PA) at 1 mL per vial and then the vials were capped with aluminum

caps with PTFE/Silicone septa. All the samples were stored in the dark at 15oC except for

the experiment designed to study the prooxidant threshold level of free fatty acids. In this

experiment, samples were kept at 10 oC to slow down oxidation so that differences in

oxidation rates among the samples could be more clearly observed.


Emulsions were diluted into 10 mM phosphate buffer (pH 7) at an

emulsion:buffer ratio of 1:50. The droplet size distribution was reported as the mean

particle diameter (z average) and zeta potential was determined using a ZetaSizer Nano-

ZS (Malvern Instruments, Worcestershire, UK). Each measurement was repeated twice

with three replicates for each sample. All measurements were conducted at room

temperature.

4.3.2.4 Measurement of Interfacial Tension

The interfacial tension of selected free fatty acids was determined using

interfacial tensiometry (DSA 100, Kruss USA, Charlotte, NC). The method used was

adapted from Chaiyasit et al. (2008). Each free fatty acid was diluted in MCT at 0.5% (wt

96

%) concentration. MCT was used as a nonoxidizable lipid to minimize oxidation during

analysis. The tensiometer hypodermic needle was submerged in a 0.1% Tween 20 and 10

mM phosphate solution at room temperature. The sample was formed into a pendant drop

at the inverted tip of the needle which was positioned on an optical bench between a light

source and a high speed charge couple device (CCD) camera. The CCD camera was

connected to a video frame-grabber board to record the image at a speed of one frame per

second. The pendant drop shape was determined with numerical analysis of the entire

drop shape. The interfacial tension was calculated from the drop shape using the Young-

Laplace equation (Duhkin et al., 1995).


Production of lipid hydroperoxides, a primary product of lipid oxidation were

measured according to the method Shantha and Decker (1994). In summary, each sample

(0.3 mL) was vortexed three times (10 second each) with 1.5 mL of an isooctanol +

isopropanol (3:1 v:v) solution. The samples were then centrifuged for 2 min at 3400 g

(Centrific TM Centrifuge, Fisher Scientific, Fairlawn, NJ) and 0.2 mL of the upper

organic layer (or less if lipid oxidation was extensive) was mixed with methanol +

butanol solution (2:1 v:v) to obtain a total volume of 3.0 mL. Then, 15 L of 3.94 M

ammonium thiocyanate and 15 L of ferrous iron solution were added. The absorbance

of the samples was measured at 510 nm using a Genesys 20 spectrophotometer

(ThermoSpectronic, Waltham, MA) 20 min after addition of the iron. Hydroperoxide

concentrations were determined based on a cumene hydroperoxide standard curve.

97

Hexanal, a secondary lipid oxidation product was measured using the method

described by Boon et al. (2008) using a GC-17A Shimadzu gas chromatography equipped

with an AOC-5000 autosampler (Shimadzu, Kyoto, Japan). Emulsions (1 mL) in 10 mL

glass vials capped with aluminum caps with PTFE/Silicone septa were shaken and heated

at 55°C for 13 min in the autosampler heating block before measurement. A 50/30 m

DVB/Carboxen/PDMS solid-phase microextraction (SPME) fiber needle from Supelco

(Bellefonte, PA) was injected into the vial headspace for 1 min to absorb volatiles and

then was transferred to the GC injector port (250oC) for 3 min. The injection port was

operated in split mode and the split ratio was set at 1:5. Volatiles were separated on a

Supleco 30 m x 0.32 mm Equity DB-1 column with a 1 m film thickness at 65oC for 10

min. The carrier gas was helium at 15.0 mL/min. A flame ionization detector was used at

a temperature of 250oC. Hexanal concentrations were determined from peak areas using

a standard curve prepared from authentic hexanal.


All experiments were conducted on triplicate samples and were repeated at least



between mean values were compared using the Duncan’s multiple range test with


98



The physical stability of each emulsion was determined by a change in droplet

size and a visual observation of creaming (Waraho et al., 2009). There was no visual

observation of creaming during storage in every emulsion. The emulsion droplets size

ranged from 165 – 185 nm and there was no significant change in droplet size of each

emulsion over the course of study (data not shown). The combination of these

observations indicated that no major changes in the physical stability of the emulsions

were occurring over the course of the studies.

4.4.2 Impact of Low Oleic Concentrations on the Physical and Chemical Properties

of Oil-in-Water Emulsions

Our previous study showed a powerful prooxidant activity of free fatty acids at

oleic acid concentrations of 1.0 to 5.0% in a 1.0% stripped soybean oil-in-water

emulsions (Waraho et al., 2009). Typically, the commercial oils contain 0.05-0.70% free

fatty acids (Pryde, 1980; Jung et al., 1989; Chaiyasit et al., 2007a). Therefore, this study

was performed to determine the threshold level at which oleic acid (0.05 to 1.0% of oil)

was able to promote oxidation in a 1.0% stripped soybean oil-in-water emulsions. In this

study, the samples were kept at 100C to slow down the progress of lipid oxidation in

order to better differentiate the oxidation rates among all the samples since a similar

study at 150C showed small differences in oxidation rates (data not shown).

99

The droplet surface charge or zeta potential (ζ) of the emulsions were observed

after 24 hours of storage. There was significant decrease in droplet charge with increasing

oleic acid concentrations (Fig. 4.2). The droplet surface charge of the control emulsions

was –5.28 + 0.43 mV while the emulsions with added oleic acid ranged from – 6.36 +

0.18 mV to –24.7 + 0.47 mV. The observed decrease in droplet surface charge suggested

that acid groups of the oleic acid were migrating to and concentrating at the lipid-water

interface of the emulsion droplet (Waraho et al., 2009).

The lipid oxidation rate in the 1.0% stripped soybean oil-in-water emulsions

without (control) and with oleic acid (0.05 to 1.0% of oil) was observed by following

lipid hydroperoxide and headspace hexanal formation as indicators of primary and

secondary lipid oxidation products, respectively. The results showed that there was no

significant difference in lipid hydroperoxide formation in the emulsions containing 0.05

and 0.10% oleic acid compared to the control emulsions throughout the course of study

(Fig. 4.3). The emulsions that contained 0.50, 0.75 and 1.0% oleic acid exhibited a

significantly higher hydroperoxide and headspace hexanal concentration than the control

emulsions after 2 days of storage. Oleic acid at 0.25% showed significantly higher

hydroperoxide formation after 5 days of storage. While 0.10% of oleic acid did not

increase lipid hydroperoxide concentrations compared to the control, it did increase

headspace hexanal concentrations after 7 days of storage. From this study, we can

conclude that oleic acid concentrations as low as 0.1% of the oil were able increase lipid

oxidation rates.

100


oil-in-water emulsions with addition of 0-1.0 % (wt.% of oil) oleic acids at

pH 7. Data points represent means (n=3) + standard deviations (some error

bars may lie within the data points).

-30

-25

-20

-15

-10

-5

0

0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1

Oleic acid content (%)

Ze

tap

ote

nti

al

(mV

)

101

Figure 4.3. Formation of lipid hydroperoxide concentration (a) and headspace hexanal

(b) in 1.0% stripped soybean oil-in-water emulsions without (control) and

with 0-1.0 % (wt.% of oil) oleic acids at pH 7 during storage at 10ºC in the

dark for 7 days.

102

4.4.3 The Effect of Degree of Free Fatty Acids Unsaturation on the Physical and

Chemical Properties of Oil-in-Water Emulsions

In bulk oils, fats with a high degree of unsaturated fatty acids are more vulnerable

to lipid oxidation (Decker and McClements, 2008; Shahidi and Wanasundara, 1998). A

similar trend was found in a comparative study of oxidation rates of bulk free fatty acids

(Takagi and Miyashita; 1987). However, an opposite trend was found in a free fatty acid

surfactant micelle system where the oxidative stability of polyunsaturated fatty acids

increased with increasing degree of unsaturation (Miyashita et al., 1993). Since there are

major differences in the impact of unsaturated free fatty acids on lipid oxidation in

colloidal dispersions and bulk lipid this study was conducted in order to investigate the

effect of degree of unsaturation of free fatty acids on the physical and chemical properties

of oil-in-water emulsions.

The impact of 0.5% (of lipid content) of oleic, linoleic and linolenic acids in 1.0%

stripped soybean oil-in-water emulsions on zeta potential and lipid oxidation were

compared to emulsions without added free fatty acid (control emulsions). Figure 4.4

shows that the droplet surface charge measured after 24 hours of storage was -6.43+0.13,

-16.03+0.37, -14.80+0.18 and -15.35+ 0.11 mV for control, oleic, linoleic and linolenic

acids, respectively. The droplet surface charge of control emulsions was significantly

lower for emulsions containing oleic acid than linoleic and linolenic acids. There were no

significant differences between zeta potentials in emulsions with linoleic and linolenic

acids.

103


oil-in-water emulsions without (control) and with 0.50% (wt.% of oil)

oleic, linoleic and linolenic acids at pH 7.

104

To confirm that the free fatty acids had the ability to migrate to oil-water

interfaces, their impact of interfacial tension was also determined. In oil-in-water

emulsions, the ability of free fatty acids to concentrate at the droplet interface will be

dependent on their ability to compete with the surface active agents used to stabilize the

emulsion. Therefore these experiments were conducted in the presence of both Tween 20

and the free fatty acids. Using this model, both oleic (5.49+ 0.07 mN/m) and linoleic acid

(5.52+ 0.05 mN/m) were able to significantly decrease interfacial tension compared to the

control (5.88+0.02 mN/m) while linolenic acid (5.75+0.06 mN/m) had no effect. This

data confirmed that the free fatty acids could migrate to and concentrate at lipid-water

interfaces. It is not clear why linolenic acid was able to decrease zeta potential but was

not able to decrease interfacial tension but this could be due to the small changes in

interfacial tension seen for all the fatty acids which would make observing their impact

on interfacial activity more difficult. The results of these two studies indicate that zeta

potential would be a more powerful analytical technique than interfacial tension for

observing the surface activity of free fatty acids.

Emulsions containing oleic, linoleic and linolenic acids all had higher lipid

hydroperoxide and hexanal formation rates than the control (Figure 4.5). Surprisingly,

both lipid hydroperoxides and hexanal concentrations decreased with increasing degree

of unsaturation. The higher oxidation rate for emulsions with oleic acid could be due to

the higher negative charge on the droplets which could attract prooxidant metals (Waraho

et al., 2009). However, the zeta potentials for the linoleic and linolenic acid containing

emulsion were statistically the same suggesting that there was another mechanism

involved as these two emulsions

105



with 0.50% (wt.% of oil) oleic, linoleic and linolenic acids at pH 7.0

during storage at 15ºC in the dark for 7 days.

106

had different lipid hydroperoxide and hexanal concentrations. Miyashita et al. (1993)

observed that the stability of free fatty acids in a surfactant micelle system increased with

increasing degree of unsaturation. If the same were true in oil-in-water emulsions, it is

possible that linolenic acid would oxidize slower than linoleic acid and could co-oxidize

the fatty acids on the triacylglycerols in the stripped corn oil less than oleic and linoleic

acids thus explaining why it had the lowest prooxidant activity.

4.4.4 The Effect of cis vs. trans Double Bonds of Free Fatty Acids on the Physical

and Chemical Properties of Oil-in-Water Emulsions

To determine if fatty acid shape did impact lipid oxidation in oil-in-water

emulsions, studies were conducted with cis and trans fatty acids. For example oleic acid

(18:1, cis) was compared to elaidic acid (18:1, trans) and linoleic acid (18:2, cis-cis) was

compared to linoelaidic acid (18:2, trans-trans) at a level of 0.5% of the oil in 1.0%

stripped soybean oil-in-water emulsion. The droplet surface charge of each emulsion was

evaluated after 24 hours of storage. The droplet surface charge of the control emulsions

was -5.24+0.42 mV while the droplet surface charge decreased to -11.30+0.31, -

11.35+0.15, -10.65+0.51 and -10.90+0.06 mV with addition of oleic acid, elaidic acid,

linoleic acid and linoelaidic acid, respectively. As seen previously, the zeta potential of

control emulsions was significantly higher than emulsions with any of the free fatty acids.

The zeta potential of the emulsions with oleic and elaidic acid were significantly lower

than emulsions with linoleic and linoelaidic acid. However, there was no significant

difference in droplet surface charge between the cis and trans isomers.

107

Again, all of the free fatty acids increased lipid hydroperoxide and headspace

hexanal formation (Figure 4.6). Lipid oxidation was greatest for emulsions with 18:1 with

no difference in oxidation rates whether the double bond was in the cis or trans

configuration. Emulsions with linoelaidic acid had similar oxidation rates to oleic and

elaidic acids, however, emulsions containing linoleic acid had significantly lower lipid

hydroperoxide and hexanal concentrations than the other 3 fatty acids.

The increased oxidation rates with oleic, elaidic and linoelaidic acids are likely

due to a combination of factors. Oleic and elaidic were able to cause the greatest

reduction in the negative charge of the emulsion droplet which could increase the ability

of transition metals to interact with the emulsion droplet and promote oxidation. The

mechanism by which linoelaidic acid promotes oxidation is more difficult to understand

as it had less impact on the emulsion droplet charge than oleic and elaidic acids yet

accelerated lipid oxidation in a similar manner. As discussed above, it is also possible

that the fatty acids could promote lipid oxidation by being oxidized themselves and then

co-oxidizing the fatty acids on the triacylglycerols in the stripped corn oil. Silwiok and

Kowalski (1974) showed that elaidic was more stable to oxidation than oleic acid in a

bulk solution of free fatty acids. If susceptibility of free fatty acids to oxidation in

colloidal dispersions is the opposite to what is observed in bulk solution as has been

proposed by Miyashita et al. (1993), it is possible that linoelaidic would oxidized faster

than linoleic acid and thus could more rapidly accelerate oxidation of the stripped corn

oil.

108



with 0.50% (wt.% of oil) oleic, linoleic, elaidic and linoelaidic acids at pH

7.0 during storage at 15ºC in the dark for 8 days.

109

4.5 Conclusions

Free fatty acids are minor components found in commercial oils that are strong

prooxidants since as little as 0.1% can accelerate hexanal formation in 1.0% stripped

soybean oil-in-water emulsions. Surprisingly, an increase in the degree of unsaturation of

the free fatty acids lowers the ability of the free fatty acids to promote oxidation. The

ability of monounsaturated fatty acids to promote lipid oxidation more than di- and

triunsaturated fatty acids could be due to their geometric shape which is more linear. The

linear fatty acids could allow them easier access to the emulsion droplet interface as

evidenced by their ability to decrease the negative charge of the emulsion droplets more

than the polyunsaturated fatty acids. A more negatively charged emulsion droplet would

be expected to be more susceptible to lipid oxidation was it would attract transition

metals which could then more readily interact with the lipid in the emulsion droplet core.

However, the prooxidant activity of the free fatty acids could not be solely explained by

the ability of the fatty acids to impact the surface charge of the emulsion droplet.

Therefore, it is possible that oxidation of the free fatty acids themselves could be

involved in the observed prooxidant activity. Free fatty acids in surfactant micelles show

an opposite tendency to oxidation with increasing unsaturation resulting in increasing

oxidative stability. Therefore, if the free fatty acids themselves were oxidizing and then

promoting co-oxidation of the triacylglycersols in the stripped soybean oil, it would be

expected that linolenic acid would be the least prooxidative of the fatty acids studied, as it

would oxidize the slowest.

Overall, free fatty acids are strong prooxidants in oil-in-water emulsions. This

prooxidant activity is dependent not only on their concentration but also on the molecular

110

structure of the fatty acid. Therefore, the prooxidant impact of free fatty acids would be

expected to be dependent on both the quality of the refined oils as well as its fatty acid

composition.

111

CHAPTER 5

ANTIOXIDANT EFFECTS OF MONO- AND DIACYLGLYCEROLS IN NON-

STRIPPED AND STRIPPED SOYBEAN OIL-IN-WATER EMULSIONS

5.1 Abstract

Antioxidant activity of mono- and diacylglycerols (0.01-2.50% of oil) was

observed in 1.0% non-stripped and stripped soybean oil-in-water emulsions by

monitoring lipid hydroperoxide and headspace hexanal formation. Addition of 1-

monooleoylglycerol only had a small impact on the oxidative stability of non-stripped

soybean oil-in-water emulsions but did inhibit lipid oxidation in emulsions prepared with

stripped soybean. Much stronger antioxidant activity was observed upon the addition of

1,2-dioleoyl-sn-glycerol to both non-stripped and stripped soybean oil-in-water

emulsions. To determine if ability of mono- and diacylglycerols to impact lipid oxidation

was due to their ability to alter the physical properties of the oil-in-water emulsions, zeta

potential and interfacial tension was measured. Both 1-monooleoylglycerol and 1,2-

dioleoyl-sn-glycerol reduced interfacial tension with 1-monooleoylglycerol being the

more surface active of the two. Both 1-monooleoylglycerol and 1,2-dioleoyl-sn-glycerol

also were able to increase the zeta potential of the emulsions although these increases

were small being less than 4 mV. Overall, these results suggest that diacylglycerols

could be an effective antioxidant in oil-in-water emulsions. However, the antioxidant

mechanism is not clear as 1,2-dioleoyl-sn-glycerol had little impact on the interfacial

properties of the emulsion droplet and they were effective antioxidant in emulsions with

112

stripped soy bean oil meaning that their activity is not related to interactions with other

antioxidants.

5.2 Introduction

Many lipid containing foods are either oil-in-water or water-in-oil emulsions

(McClements, 2005; Friberg et al., 2004; Richards et al., 2002). Lipid oxidation is of

great concern to the industry because it causes the physical and chemical deterioration of

food quality, such as losses in important nutrients, formation of potentially toxic reaction

products (such as aldehydes and ketones), undesirable changes in appearance and texture,

and development of rancidity (Frankel, 1998; Coupland and McClements, 1996a;

McClements and Decker, 2000; Chaiyasit et al., 2007a; Decker and McClements, 2008).

Lipid oxidation is favored in oil-in-water emulsions because of the large contact surface

between the oxidizable lipid in emulsion droplets and water-soluble compounds including

oxygen and pro-oxidants, which contribute to the initiation and propagation of oxidation

reactions (Frankel, 1998; Lethuaut et al., 2002; Villiere et al., 2005).

The major components in vegetable oils are predominately acylglycerols

including mono-, di- and triacylglycerols (approximately 95% of commercial oils). The

remaining of 5% consists of unsaponifiable compounds such as free fatty acids, sterols,

carbohydrates, phenols (e.g. tocopherols), proteins, pigments (e.g. chlorophyll,

carotenoids, flavonoids, gossypol), trace metals, and pesticides (Hamilton, 1994; Khan

and Shahidi, 2000; Abuzaytoun and Shahidi, 2006, Chaiyasit et al., 2007a). Mono- and

diacylglycerols are present in animal fats and vegetable oils at much smaller

concentrations than triacylglycerols. They exist in fats and oils because the oil extraction

113

process leads to a partial hydrolysis of triacylglycerols. For examples, grinding or

pressing oilseeds decompartmentalize lipases out of subcellular organels allowing them

to hydrolyze triacylglycerols. Heat and pressure can also accelerate fatty acid hydrolysis

from triacylglycerols (O’Brien, 2004). Some of these mono- and diacylglycerols are

removed by deodorization process (Johnson, 2002; Shahidi et al., 1997). The

concentration of diacylglycerols in refined commercial oils ranges from 0.8-5.8% while

monoacylglycerols are generally less than 0.2% of the oil concentration (Chaiyasit et al.,

2007a; D’Alonzo et al., 1982). Mono- and diacylglycerols are surface active compounds

because they contain both lipophilic (fatty acids) and hydrophilic (hydroxyl) groups.

Therefore, they are partially soluble in fat and water can reduce interfacial tension.

Mono- and diacylglycerols are the most commonly used emulsifiers being approximately

70% of the total emulsifiers used in the food industry (Garti, 2001; O’Brien, 2004).

The minor components that are naturally present in the oil such as fatty acids,

phospholipids, tocopherols and transition metals greatly impact oxidative stability (Yoon

et al., 1988; Shahidi and Shukla, 1996). Some of them can act as prooxidants such as free

fatty acids, metals and chlorophyll and some of them can act as antioxidants such as

tocopherols (Cort, 1974; Shahidi and Shukla, 1996; Chiyasit et al., 2007a). Even though

these components exist in small amounts, they can also affect the physicochemical

properties of the emulsions due to their surface activity (McClements, 2008). A study

from Holman and Elmer (1974) indicated that oxidation rates of bulk trilinolein and

trilinolenin were higher than the oxidation rates of ethyl linoleate and ethyl linolenate,

respectively. Similar results were reported by Miyashita and Takagi (1988) who showed

that the oxidation rates of bulk triacylglycerol were higher than diacylglycerol and

114

monoacylglycerol, respectively. They suggested that the order of the oxidation rates may

be due to intramolecular free radical transfer reactions occurring faster than

intermolecular radical transfer reactions. When mono- and diacylglycerols were added to

soybean oil they were found to act as prooxidants as determined by headspace oxygen

consumption (Mistry and Min, 1988). This prooxidant activity was proposed to be due to

the ability of mono- and diacylglycerols to decrease surface tension of the oil and

increase oxygen diffusion into the oil.

While there is research on the impact of mono- and diacylglycerols on lipid

oxidation in bulk oils, essentially no data has been reported on their impact on the

oxidative stability of oil-in-water emulsions. It is possible that the impact of mono- and

diacylglycerols on oxidation could be very different in bulk and emulsified oils due to

their surface activity that would impact their physical location differently in the two

systems.


5.3.1 Materials

Soybean oil was purchased from a local retail store. 1-Monooleoylglycerol was

purchased from Nu-Chek Prep, Inc. (Elysian, MN) and 1,2-dioleoyl-sn-glycerol was

purchased from Avanti Polar Lipids, Inc. (Alabaster, Alabama). Miglyol 812 (Medium

chain triglyceride; MCT) was purchased from Sasol (Witten, Germany). Potassium

phosphate monobasic, and potassium phosphate dibasic heptahydrate, silicic acid (100-

200 mesh, 75-150 m, acid washed), activated charcoal (100-400 mesh),

115

polyoxyethylene (20) sorbitan monolaurate (Tween 20), ammonium thiocyanate and iron

(II) sulfate heptahydrate were obtained from Sigma Chemical Co. (St. Louis, MO.,

U.S.A.). Iso-octanol, n-hexane, 2-propanol, methanol, 1-butanol were purchased from

Fisher Scientific (Fair Lawn, NJ). All the chemicals used in this experiments were

analytical grade or purer. Glassware was incubated in 3 mM HCL overnight to remove

transition metals followed by rinsing with double-distilled water and then drying before

use. Double-distilled water was used throughout the study.

5.3.2 Methods

5.3.2.1 Removing Polar Minor Components from Soybean Oil (Stripping oilsOils)

For some experiments, soybean oils were stripped prior to making emulsions to

remove polar minor components such as tocopherols, free fatty acids, mono- and

diacylglycerols, phospholipids and lipid oxidation products by passing through silicic

acid and activated charcoal chromatographic column using n-hexane as eluent (Waraho et

al., 2009). In summary, silicic acid (100 g) was washed three times with a total of 3 L of

distilled water then filtered with Whatman filter paper in a Buchner funnel and dried at

110C for 20 h. The washed silicic acid (22.5 g) and activated charcoal (5.625 g) were

suspended in 100 and 70 ml n-hexane, respectively. A chromatographic column (3.0 cm

internal diameter x 35 cm height) was then packed sequentially in three layers starting

with 22.5 g of silicic acid followed by 5.625 g activated charcoal and then another 22.5 g

silicic acid. Thirty grams of soybean oil was dissolved in 30 ml of n-hexane and passed

through the column by eluting with 270 ml of n-hexane. To retard lipid oxidation during

stripping, the collected triacylglyerols were held in an ice bath which was covered with

116

an aluminum foil. The n-hexane used as a solvent in the stripping oil method was

removed with a vacuum rotary evaporator (RE 111 Buchi, Flawil, Switzerland) at 37C.

The traces of the remaining solvent were removed by flushing with nitrogen. Then three

grams of the stripped oil were transferred into 3-ml vials, flushed with nitrogen and kept

at -80 C for subsequent studies. The effectiveness of stripping was determined by

reduction of tocopherol concentrations to below the level of detection using HPLC (Boon

et al., 2008). Since mono- and diacylglycerols used in this study contained low levels of

lipid hydroperoxides (< 0.05 mmol hydroperoxides/kg of mono/diacylglycerols) they

were used directly without prior stripping.


The preparation of a 1.0% (wt.) soybean oil-in-water emulsion was similar to

prior study (Waraho et al., 2009). All emulsions were made with 10 mM phosphate

buffer solution (pH 7.0) using Tween 20 as an emulsifier at a 1:10 emulsifier: oil ratio.

Soybean oil (non-stripped or stripped), 1- monooleoylglycerol or 1,2-dioleoyl-sn-

glycerol, Tween 20, and phosphate buffer were added to a beaker and a coarse emulsion

was made by blending with a hand-held homogenizer (M133/1281-0, Biospec Products,

Inc., Bartlesville, OK) for 2 min. The coarse emulsion was then homogenized with a

microfluidizer (Microfluidics, Newton, MA) at a pressure of 9 kbar for three passes. Then

each emulsion was divided into 10 ml GC vials (Supelco, Bellefonte, PA) at 1 mL per

vial then the vial was capped with aluminum caps with PTFE/Silicone septa. All the

samples made with non-stripped soybean oil were stored in the dark at 25oC while the

117

samples made with stripped soybean oil were stored at 15oC to slow down oxidation so

that differences in oxidation rates among the samples could be more clearly observed.


The emulsions were diluted into 10 mM phosphate buffer (pH 7) at an

emulsion:buffer ratio of 1:50. The droplet size distribution reported as the mean particle

diameter (z average) and zeta potential were invested using a ZetaSizer Nano-ZS

(Malvern Instruments, Worcestershire, UK). Each measurement was repeated twice with

three replicates for each sample. These measurements were conducted at room

temperature.

5.3.2.4 Measurement of Interfacial Tension

The interfacial tension of mono- and diacylglycerols was determined using

interfacial tensiometry (DSA 100, Kruss USA, Charlotte, NC). The method used was

adapted from Chaiyasit et al. (2008). Mono- and diacylglycerols were diluted in MCT at

the concentration ranged from 0.01-2.50% (wt %). MCT was used as a nonoxidizable

lipid to minimize oxidation during analysis. The tensiometer hypodermic needle was

submerged in a 0.1% Tween 20 and 10 mM phosphate solution at room temperature. The

sample was formed into a pendant drop at the inverted tip of the needle which was

positioned on an optical bench between a light source and a high speed charge couple

device (CCD) camera. The CCD camera was connected to a video frame-grabber board

to record the image at a speed of one frame per second. The pendant drop shape was

118

determined with numerical analysis of the entire drop shape. The interfacial tension was

calculated from the drop shape using the Young-Laplace equation (Duhkin et al., 1995).


Lipid oxidation was monitored with lipid hydroperoxides, a primary product of

lipid oxidation, and hexanal, is a secondary lipid oxidation product. Lipid

hydroperoxides were measured using a modified method of Shantha and Decker (1994).

In summary, each sample (0.3 mL) was vortexed three times (10 second each) with 1.5

mL of an isooctanol + isopropanol (3:1 v:v) solution. The samples were then centrifuged

for 2 min at 3400 g (Centrific TM Centrifuge, Fisher Scientific, Fairlawn, NJ) and 0.2

mL of the upper organic layer or diluted with methanol/butanol (depending on the extent

of lipid oxidation) was mixed with 2.80 mL of methanol + butanol solution (2:1 v:v), 15

L of 3.94 M ammonium thiocyanate and 15 L of ferrous iron solution (prepared by

mixing 0.132 M BaCl2 and 0.144 M FeSO4). The absorbance of the samples was

measured at 510 nm using a Genesys 20 spectrophotometer (ThermoSpectronic,

Waltham, MA) 20 min after the addition of the iron. Hydroperoxide concentrations were

quantitated based on a cumene hydroperoxide standard curve.

Hexanal was measured by the method described by Boon et al. (2008) using a

GC-17A Shimadzu gas chromatography equipped with an AOC-5000 autosample

(Shimadzu, Kyoto, Japan). Emulsions (1 mL) in 10 mL glass vials capped with aluminum

caps with PTFE/Silicone septa were shaken and heated at 55°C for 13 min in an

autosampler heating block before measurement. A 50/30 m DVB/Carboxen/PDMS

solid-phase microextraction (SPME) fiber needle from Supelco (Bellefonte, PA) was

119

injected into the vial for 1 min to absorb volatiles and then was transferred to the injector

port (250oC) for 3 min. The injection port was operated in split mode, and the split ratio

was set at 1:5. Volatiles were separated on a Supleco 30 m x 0.32 mm Equity DB-1

column with a 1 m film thickness at 65oC for 10 min. The carrier gas was helium at 15.0

mL/min. A flame ionization detector was used at a temperature of 250oC. Hexanal

concentrations were determined from peak areas using a standard curve prepared from

authentic hexanal.


All experiments were conducted in triplicate samples and were repeated at least



between mean values were compared using Duncan’s multiple range test with



Commercial food grade vegetable oils contain minor components such as

tocopherols, phospholipids, chlorophyll, free fatty acids, phospholipids, mono- and

diacylglycerols that can be either antioxidative or prooxidative (Khan and Shahidi, 2000;

Khan and Shahidi, 2002). This study was designed to examine the effects of mono- and

diacylglycerols on the oxidative stability of oil-in-water emulsions. In order to understand

the impact of just the mono- and diacylglycerols on oxidative stability, some experiments

were conducted with stripped soybean oil to remove minor lipid components that could

120

inhibit or promote lipid oxidation. Other studies were conducted with non-stripped

soybean oil to mimic the effect of mono- and diacylglycerols in a commercial food

emulsion.

The concentration of monoacylglycerols in refined vegetable oils is generally less

than 0.2% while diacylglycerols range from 0.8-5.8% (Chaiyasit et al., 2007a; D’Alonzo

et al., 1982). Therefore, to determine how mono- and diacylglycerols influence the

oxidative stability of oil-in-water emulsions, different concentrations of 1-

monooleoylglycerol or 1,2-dioleoyl-sn-glycerol (0.01, 0.10, 0.50, 1.00, and 2.50% of oil

concentration) were added during emulsification of the Tween 20-stabilized soybean oil-

in-water emulsions at pH 7.0. In this study, the oleic acid form of mono- and

diacylglycerols were used because they were easy to handle (liquid at room temperature)

and would be more stable to lipid oxidation than mono- and diacylglycerols with

polyunsaturated fatty acids.


The physical stability of each emulsion was determined by observing a change in

droplet size and a visual observation of creaming. There was no visual observation of

creaming during storage in every emulsion. The emulsion droplet size ranged from 165 to

185 nm and there was no significant change in droplet size of each emulsion over the

course of study (data not shown). The combination of these observations indicated that no

major changes in the physical stability of the emulsions were occurring over the course of

these studies.

121

5.4.2 Effect of Mono- and Diacylglycerols on Interfacial Tension

Mono- and diacylglycerols could impact lipid oxidation by changing the

properties of the emulsion droplet interface. To determine if mono- and diacylglycerols

were partitioning at the oil-water interface, their impact on interfacial tension was

determined. In oil-in-water emulsions, the ability of mono- and diacylglycerols to

concentrate at the droplet interface will be dependent on their ability to compete with the

surface active agents used to stabilize the emulsion. Therefore, these experiments were

conducted in the presence of both Tween 20 and mono- or diacylglycerols. Addition of 1-

monooleoylglycerol at concentrations higher than 0.10% of the oil significantly

decreased interfacial tension with increasing monoacylglyerol concentrations further

decreasing interfacial tension (Figure 5.1). Addition of 1,2-dioleoyl-sn-glycerol to MCT

did not signifacntly (p<0.05) decrease interfacial tension until concentrations greater than

1.0% and the reduction of interfacial tension by 1,2-dioleoyl-sn-glycerol was much lower

than 1-monooleoylglycerol. Overall, these data show that both 1-monooleoylglycerol and

1,2-dioleoyl-sn-glycerol can concentrate at lipid-water interfaces in the presence of

Tween 20 although the surface activity of 1,2-dioleoyl-sn-glycerol was much less than 1-

monooleoylglycerol. This difference in surface activity is likely due to the lower HLB

value of diacylglycerols (1.8) compared to monoacylglycerols (5.2; McClements, 2005).

122

Figure 5.1. Influence of addition of 0.01-2.50% 1-monooleoylglycerol or 1,2-

dioleoyl-sn-glycerol in medium chain triacylglycerols on interfacial

tension at ambient temperature. Data represents means (n=3) standard

deviations. Some error bars lay within data points.

0

1

2

3

4

5

6

7

0 0.5 1 1.5 2 2.5 3

MG

DG

Concentration (% in MCT)

Inte

rfac

ial T

en

sio

n (

mN

/m)

123

5.4.3 Effect of Mono- and Diacylglycerols on the Physical and Chemical Properties

of Non-stripped and Stripped Oil-in-Water Emulsions

The droplet surface charge or zeta potential (ζ) of the 1.0% oil-in-water emulsions

made with non-stripped soybean oil with varying concentrations of 1-monooleoylglycerol

from 0.01-2.50% (oil wt.) are shown in Figure 5.2a. The control emulsions made with

non-stripped soybean oil had a surface charge of -8.63+0.48 mV. The droplet surface

charge increased upon the addition of 0.1% 1-monooleoylglycerol to -7.08+0.28 with no

further increase in surface charge with increasing 1-monooleoylglycerol concentrations.

The droplet surface charge obtained from emulsions made with stripped soybean oil

showed a similar trend (Figure 5.2b). The control emulsions made with stripped soybean

oils was -5.97+0.05 mV. Addition of 1-monooleoylglycerol slightly increased zeta

potential to a maximum of -5.45+0.05 mV in the presence of 2.50% 1-

monooleoylglycerol. The lower negative charge in the emulsions made with stripped

soybean oil is likely due to the removal of minor components such as free fatty acids

(Waraho et al., 2009).

The droplet surface charge or zeta potential (ζ) of the 1.0% oil-in-water emulsions

made with non-stripped and stripped soybean oil and varying concentrations of 1,2-

dioleoyl-sn-glycerol are shown in Figure 5.3a and b, respectively. The control emulsions

made with non-stripped soybean oil had a surface charge of -12.47+0.90 mV. Droplet

surface charge increased with increasing 1,2-dioleoyl-sn-glycerol concentrations reaching

a maximum of -8.71+0.44 mV in the presence of 2.50% 1,2-dioleoyl-sn-glycerol in the

oil.

124

Figure 5.2. The droplet surface charge or zeta potential (ζ) of: (a) 1.0% non-stripped

soybean oil-in-water emulsions with addition of 0.01-2.50% 1-

monooleoylglycerol (oil wt.) at pH 7.0 after 24 hour of storage in the dark

at 250C and (b) 1.0% stripped soybean oil-in-water emulsions with

addition of 0.01-2.50% 1-monooleoylglycerol (oil wt.) at pH 7.0 after 24

hour of storage in the dark at 150C.

a

b

125

Figure 5.3. The droplet surface charge or zeta potential (ζ) of: (a) 1.0% non-stripped

soybean oil-in-water emulsions with addition of 0.01-2.50% 1,2-dioleoyl-

sn-glycerol (oil wt.) at pH 7.0 after 24 hour of storage in the dark at 250C

and (b) 1.0% stripped soybean oil-in-water emulsions with addition of

0.01-2.50% 1,2-dioleoyl-sn-glycerol (oil wt.) at pH 7.0 after 24 hour of

storage in the dark at 150C.

a

b

126

The droplet surface charge of emulsions made with stripped soybean oil showed

similar trend as the non-stripped soybean oil (Figure 5.3b) but overall was less negatively

charged again presumably due to the removal of minor oil components such as free fatty

acids. Droplet surface charge of the control emulsions made with stripped soybean oils

was -9.81+0.55 mV increasing to -6.95+0.27 mV in the presence of 2.50% 1,2-dioleoyl-

sn-glycerol in the oil. It is not all that unexpected that 1-monooleoylglycerol and 1,2-

dioleoyl-sn-glycerol did not dramatically change zeta potential as they are nonionic

emulsifiers. Their ability to increase zeta potential could be due to the displacement of

Tween 20 from the droplet surface as Tween 20 has a slight negative charge (Waraho et

al., 2009).

The rates of lipid oxidation of 1.0% soybean oil-in-water emulsions made with

both non-stripped and stripped soybean oils with added 1-monooleoylglycerol and 1,2-

dioleoyl-sn-glycerol were followed by monitoring lipid hydroperoxide and headspace

hexanal formation. The emulsions made with non-stripped soybean oil were stored at

250C (compared to 15C for stripped oils) to accelerate lipid oxidation rate due to their

higher oxidative stability. A study from Khan and Shahidi (2000) showed that oil-in-

water emulsions made from non-stripped oils were more stable than their corresponding

stripped oils due to the presence of minor components such as tocopherols.

127

Addition of 1-Monooleoylglycerol at concentrations ranging from 0.01 to 2.5% of

the oil had no effect on the lag phase of either lipid hydroperoxide or hexanal formation

in non-stripped soybean oil-in-water emulsions (Figure 5.4). When 1-monooleoylglycerol

was added to stripped soybean oil-in-water emulsions, lipid hydroperoxide (longer lag

times and lower hydroperoxide accumulation levels) and hexanal (lower hexanal

accumulation levels) formation was decreased (Figure 5.5). The stronger antioxidant

activity of 1-monooleoylglycerol in oxidative stability oil-in-water emulsions prepared

with stripped soybean oil may be due to the absence of antioxidants such as tocopherols

meaning that the 1-monooleoylglycerol would be the major component in the emulsion

that could impact lipid oxidation rates.

128

Figure 5.4. Formation of lipid hydroperoxides (a) and hexanal (b) in 1.0% non-

stripped soybean oil-in-water emulsions without (control) and with

addition of 0.01-2.50% 1-monooleoylglycerol (oil wt.) at pH 7.0 during

storage at 25ºC in the dark for 17 days. Data represents means (n=3)

standard deviations. Some error bars lay within data points.

0

100

200

300

400

500

600

700

0 2 4 6 8 10 12 14 16 18

Hyd

rop

ero

xid

ec

on

ce

ntr

ati

on

(m

mo

l/k

g o

il)


control (w/o MG)

0.01% MG

0.10% MG

0.50% MG

1.00% MG

2.50% MG

0

10

20

30

40

50

60

70

80

0 2 4 6 8 10 12 14 16 18

He

xa

na

l c

on

ce

ntr

ati

on

(m

mo

l/k

g

oil)


control (w/o MG)

0.01% MG

0.10% MG

0.50% MG

1.00% MG

2.50% MG

129

Figure 5.5. Formation of lipid hydroperoxides (a) and hexanal (b) in 1.0% stripped

soybean oil-in-water emulsions without (control) and with addition of

0.01-2.50% 1-monooleoylglycerol (oil wt.) at pH 7.0 during storage at

15ºC in the dark for 8 days. Data represents means (n=3) standard


0

100

200

300

400

500

600

700

800

900

0 1 2 3 4 5 6 7 8

Hyd

rop

ero

xid

e c

on

ce

ntr

ati

on

(m

mo

l/k

g o

il)


control (w/o MG)

0.01% MG

0.10% MG

0.50% MG

1.00% MG

2.50% MG

0

5

10

15

20

25

30

35

0 1 2 3 4 5 6 7 8

Hexan

al

co

ncen

trati

on

(m

mo

l/kg

oil)


control (w/o MG)

0.01% MG

0.10% MG

0.50% MG

1.00% MG

2.50% MG

130

The rates of lipid oxidation in the presence of 1,2-dioleoyl-sn-glycerol is shown in

Figures 5.6 and 5.7. In emulsions made with non-stripped oil, 0.01 and 0.1% 1,2-

dioleoyl-sn-glycerol only had a small effect on lipid hydroperoxide formation while

higher concentrations significantly inhibited hydroperoxide formation (Figure 5.6a). The

ability of 1,2-dioleoyl-sn-glycerol to inhibit headspace hexanal formation in emulsions

with non-stripped oil was even greater with all concentrations tested significantly

decreasing hexanal concentrations at all time points (Figure 5.6b). In emulsions made

with stripped oil, lipid hydroperoxide and hexanal formation both decreased with

increasing 1,2-dioleoyl-sn-glycerol concentrations with 2.5% 1,2-dioleoyl-sn-glycerol

almost completely preventing hydroperoxide and hexanal production (Figure 5.7a and

5.b). Again, the increased antioxidant activity of 1,2-dioleoyl-sn-glycerol in emulsions

made with stripped soybean oil could be due to 1,2-dioleoyl-sn-glycerol being the only

antioxidant in the oil.

131

Figure 5.6. Formation of lipid hydroperoxides (a) and hexanal (b) in 1.0% non-

stripped soybean oil-in-water emulsions without (control) and with

addition of 0.01-2.50% 1,2-dioleoyl-sn-glycerol (oil wt.) at pH 7.0 during

storage at 25ºC in the dark for 14 days. Data represents means (n=3)

standard deviations. Some error bars lay within data points.

0

100

200

300

400

500

600

700

800

0 2 4 6 8 10 12 14Hyd

rop

ero

xid

e c

on

ce

ntr

ati

on

(mm

ol/

kg

oil

)


control (w/o DG)

0.01% DG

0.10% DG

0.50% DG

1.0% DG

2.5% DG

0

10

20

30

40

50

60

70

80

90

0 2 4 6 8 10 12 14

He

xa

na

l c

on

ce

ntr

ati

on

(m

mo

l/k

g

oil)


control (w/o DG)

0.01% DG

0.10% DG

0.50% DG

1.0% DG

2.5% DG

132

Figure 5.7. Formation of lipid hydroperoxides (a) and hexanal (b) in 1.0% stripped

soybean oil-in-water emulsions without (control) and with addition of

0.01-2.50% 1,2-dioleoyl-sn-glycerol (oil wt.) at pH 7.0 during storage at

15ºC in the dark for 9 days. Data represents means (n=3) standard


0

100

200

300

400

500

600

0 1 2 3 4 5 6 7 8 9

Hyd

rop

ero

xid

e c

on

ce

ntr

ati

on

(m

mo

l/k

g o

il)


control (w/o DG)

0.01% DG

0.10% DG

0.50% DG

1.0% DG

2.5% DG

0

5

10

15

20

25

0 1 2 3 4 5 6 7 8 9

Hexan

al

co

nc

en

trati

on

(m

mo

l/kg

oil

)


control (w/o DG)

0.01% DG

0.10% DG

0.50% DG

1.0% DG

2.5% DG

133

There is published research on the impact of diacylglycerols on the properties of

oil-in-water emulsions. A patent filed by Momura et al. (1992) reported that

diacylglycerols with melting point 200C or less and a concentration of 30-100% (w/w) of

the oil phase produced an oil-in-water emulsion with a rich fatty flavor even at a low fat

content. Kawai and Konishi (2000) described oil-in-water emulsions made with

diacylglycerols at 30% (w/w) or greater had excellent physical stability, good taste and

appearance. Shiiba et al. (2002) found that oil-in-water emulsions comprised of at least

20% (w/w) diacylglycerols and 0.5-5.0% (w/w) of the emulsifier and crystallization

inhibitors such as polyglycerol fatty acid ester, sorbitan fatty acid ester, or sucrose fatty

acid ester had excellent physical stability at low temperature. However, there is no

information available on the impact of diacylglycerols on lipid oxidation in oil-in-water

emulsions.

5.5. Conclusions

Overall, 1,2-dioleoyl-sn-glycerol (0.01 to 2.5% of oil) was much more effective at

inhibiting lipid oxidation in the oil-in-water emulsions than 1-monooleoylglycerol. The

superior ability of 1,2-dioleoyl-sn-glycerol to inhibit lipid oxidation could be partially be

due to its ability to increased the surface charge of the emulsions droplets more than1-

monooleoylglycerol since negatively charged emulsion droplets tend to oxidize faster

since they can attract prooxidant metals to the emulsion droplet surface. However, it

seems unlikely that this is the sole effect since the changes in zeta potential were very

small (less than 4 mV for 1,2-dioleoyl-sn-glycerol and less than 2 mV for 1-

monooleoylglycerol). In addition, it does not seem possible that the ability of 1,2-

134

dioleoyl-sn-glycerol to inhibit lipid oxidation would be due to its ability to alter the

interfacial properties of the emulsion droplet in that it was much less surface active than

1-monooleoylglycerol. One possibility is that diacylglycerols have been reported to form

a liquid crystal phase (Parker, 1987) which could form a physical barrier that decreases

interactions between unsaturated fatty acids in the emulsion droplet core and prooxidants

or oxygen in the aqueous phase of the emulsion. While the antioxidant mechanism of 1,2-

dioleoyl-sn-glycerol is not currently understood, it is clear that it has potential as a novel

food antioxidant since it is already an approved generally recognized as safe food

ingredient and its is a relatively inexpensive.

135

CHAPTER 6

OVERALL CONCLUSIONS

Oil-in-water emulsions are widely used in the food industry as well as the

cosmetics, pharmaceutical and medical industries as a means to encapsulate, protect and

release bioactive lipids. However, a major concern that causes both physical and

chemical deterioration of oil-in-water emulsions is lipid oxidation. Because of many

quality deteriorations resulting from lipid oxidation such as losses in important nutrients,

formation of potentially toxic reaction products (such as aldehydes and ketones),

undesirable changes in appearance and texture, and development of rancidity that

shortens product shelf life, there are numerous studies have been conducted throughout

the years in order to gain a more thorough understanding of lipid oxidation processes in

both bulk and oil-in-water emulsions. The goal is to develop methods to control these

detrimental reactions. The focus of this research was to determine the affect of the minor

components that already present in the oils on the oxidative stability of oil-in-water

emulsions.

Even though edible oils undergo refining processes to remove undesirable

components, commercial oils still contain small amounts of minor components which are

unsaponifiable compounds such as free fatty acids, sterols, carbohydrates, phenols (e.g.

tocopherols), proteins, pigments (e.g. chlorophyll, carotenoids, flavonoids, gossypol),

trace metals, and pesticides. Some of them can act as prooxidants such as trace metals,

free fatty acids and chlorophylls and some of them can act as antioxidants such as

tocopherols. Some of these minor components such as phospholipids, free fatty acids,

mono- and diacylglycerols are surface active compounds that could affect lipid oxidation

136

by altering the chemical and physical properties of the oils and oil-in-water emulsions.

This research has been conducted to investigate the role of free fatty acids and mono- and

diacylglycerols which are surface active compounds that could contribute to oxidative

stability of oil-in-water emulsions.

These studies showed that free fatty acids acted as a strong prooxidants in

stripped soybean oil-in-water emulsions. Concentrations as low as 0.1% of the lipid

accelerated lipid oxidation rate as determined by shortening the lag phase of lipid

hydroperoxide and hexanal formation. The results showed that when the pH of the

emulsions is higher than the pKa of the free fatty acids, the carboxylic acid groups on the

free fatty acids are negatively charged. Since the charged free fatty acids are surface

active, they migrate to the oil-water interface of emulsion droplets where they decrease

the negative charge of the emulsion droplet. Since addition of EDTA into oil-in-water

emulsions strongly inhibited the prooxidant activity of oleic acid, this indicates that

transition metals were responsible for accelerating oxidation. Therefore, the most likely

mechanisms for the prooxidant activity of free fatty acids is through their ability to

increase the negative charge on emulsion droplet which in turn attracts cationic transition

metals to the emulsion droplet surface where they can interact with lipid thus promote

oxidation. The studies also showed that oleic acid hydroperoxides did not increase

oxidation rates suggesting that hydroperoxides on free fatty acids are not strong

prooxidants in oil-in-water emulsion.

Further studies showed that prooxidant effect of free fatty acids was dependent on

fatty acid type with lipid oxidation rates being in the order of linolenic < linoleic < oleic.

Surprisingly, an increase in the degree of unsaturation of the free fatty acids lowers the

137

ability of the free fatty acids to promote oxidation which may be due to their differences

in geometric shape. The monounsaturated is more linear than di- and triunsaturated fatty

acids which could allow them easier access to the emulsion droplet interface as evidenced

by their ability to decrease the negative charge of the emulsion droplets more than the

polyunsaturated fatty acids. A more negatively charged emulsion droplet would be

expected to be more susceptible to lipid oxidation because it would attract transition

metals which could then more readily interact with the lipid in the emulsion droplet core.

There were no significant differences in lipid oxidation rates when free fatty acid isomers

with cis or trans double bonds were compared. Overall, free fatty acids are strong

prooxidants in oil-in-water emulsions. This prooxidant activity is dependent not only on

their concentration but also on the molecular structure of the fatty acid.

Antioxidant activity of mono- and diacylglycerols was observed in non-stripped

and stripped soybean oil-in-water emulsions by monitoring lipid hydroperoxide and

headspace hexanal formation. Addition of 1-monooleoylglycerol only had a small impact

on the oxidative stability of non-stripped soybean oil-in-water emulsions but did inhibit

lipid oxidation in emulsions prepared with stripped soybean. Much stronger antioxidant

activity was observed upon the addition of 1,2-dioleoyl-sn-glycerol to both non-stripped

and stripped soybean oil-in-water emulsions. The dramatic results found in emulsions

made with stripped soybean oil showed that lipid hydroperoxide and hexanal formation

both decreased with increasing 1,2-dioleoyl-sn-glycerol concentrations, especially, with

2.5% 1,2-dioleoyl-sn-glycerol which almost completely preventing hydroperoxide and

hexanal production. Overall, these results suggest that diacylglycerols could be an

effective antioxidant in oil-in-water emulsions which possibility due to their ability to

138

form a liquid crystal phase which could form a physical barrier that decreases interactions

between unsaturated fatty acids in the emulsion droplet core and prooxidants or oxygen

in the aqueous phase of the emulsion. Although, the antioxidant mechanism of

diacylglycerols is not currently understood and need further investigation, it is clear that

it has potential as a novel food antioxidant since it is already an approved generally

recognized as safe food ingredient and its is a relatively inexpensive.

This research leads to a new knowledge for improving the oxidative stability of

the oil-in-water emulsions regarding the content of free fatty acids and mono- and

diacylglycerols as minor components in the oil itself. The food, pharmaceutical, and

cosmetic industries should try to maintain the low level of free fatty acids (less than

0.1%) in the oils before incorporate in oil-in-water emulsions in order to improve

oxidative stability of the products. On the other hand, a slight addition of diacylglycerols

upto 2.5% could dramtically improve oxidative stability of the oil-in-water emulsions.

139

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