microencapsulation USING SUGAR.pdf

MICROENCAPSULATION OF FISH

OIL/OLIVE OIL BLENDS USING SUGAR

BEET PECTIN AS AN ENCAPSULANT

Sudheera Polavarapu

Submitted in total fulfilment of the requirements

of the degree of Masters by Research (by Thesis Only)

June 2011

Department of Agriculture and Food Systems Melbourne School of Land and Environment

The University of Melbourne Victoria 3010, Australia

II

Abstract Fish oil (FO) is rich in the polyunsaturated fatty acids, but sensitive to oxidation by many

factors including, heat, light and atmospheric oxygen. Microencapsulation of FO has been

reported to help protect FO from oxidation, especially in the presence of antioxidants. As

extra virgin olive oil (EVOO) is rich in oleic acid, and some antioxidant compounds, this

study aimed to investigate the potential of EVOO to protect FO from oxidation in the

presence of sugar beet pectin as a wall material. Ethylenediaminetetras-acetic acid (EDTA)

was also tested as a chelating agent to retard the lipid oxidation caused by the transition

metal ions present in the sugar beet pectin.

The oil-in-water emulsions were prepared and spray-dried to form microcapsules containing

fish oil alone (control) and a blend of FO/EVOO (1:1 weight ratio), at 25% (pH 3) and 50%

oil (pH 3 and pH 6) loadings. Fish oil and Fish oil- Extra virgin olive oil emulsions with

EDTA were prepared at 50% oil loading only at pH 6. The physicochemical characteristics

and oxidative stability of all FO and FO-EVOO emulsions and microcapsules were assessed.

The microcapsules showed low moisture content (<3% w/w) and water activity (~0.3). The

encapsulation efficiencies were >90% for all the formulations proving sugar beet pectin an

effective encapsulant.

Oxidative stability was assessed by testing propanal and hexanal in the head space using GC

analysis, fatty acid content, and oxidation stability using an Oxipress. It was concluded that

EVOO improved the oxidative stability of microencapsulated FO during accelerated storage

conditions (80°C, 5 bar oxygen pressure), irrespective of oil loading. However, no effect

was detected at pH 3 during storage at room temperature (~27°C). The addition of EDTA in

the formulation of the microcapsules significantly (P<0.05) increased the oxidative stability

of the microcapsules. It was also observed that microcapsules prepared from emulsions at

pH 6 were better in terms of long term storage when compared to the microcapsules

prepared from emulsions at pH 3.

III

Declaration

This is to certify that

(i) the thesis comprises only my original work towards the Masters

(ii) due acknowledgement has been made in the text to all other material used,

(iii) the thesis is 22,584 words in length as approved by the Graduate School, Faculty or

RHD Committee.

Sudheera Polavarapu

June 2011

IV

Acknowledgements

My greatest gratitude to my supervisors Dr. Said Ajlouni, Dr. Maryann Augustin and Dr.

Christine Oliver who were abundantly helpful and offered invaluable assistance, support and

guidance from the very early stages of my Masters Project till the thesis submission. I

thank them for taking time to attend monthly meetings to discuss results and for the

constructive comments to the numerous reports I submitted. Many thanks for providing new

ideas and approaches to the project. Working under their supervision has triggered a passion

for research in me. I attribute my growth as a student to all of them and that I would benefit,

for a long time to come. I am grateful to them in every possible way. Also, my special thanks

to Dr. Christine Oliver for all her help and efforts in preparing the manuscripts for

publication.

I am grateful to all staff and students in the Department of Agriculture and Food Systems,

The University of Melbourne and CFNS (CSIRO Food and Nutritional Sciences) for their

assistance in my lab work, for sharing instruments and for creating a pleasant working

atmosphere. A warm thank you to the Senior Laboratory Manager Michelle Rhee at the

University for her assistance in ordering and supplying of chemicals and for all her support

while working in the lab. Furthermore, I convey my special acknowledgement to Li Jiang

Cheng, Zhiping Shen, Said Ajlouni, Rangika Weerakkody, Helen French, Christine Oliver

and Roderick Williams for their indispensable help and for being patient while teaching me

how to operate various lab instruments. I also appreciate their time and efforts for helping

me interpret the results obtained.

I am immensely thankful to Dr. Simon Crawford, School of Botany, The University of

Melbourne, Parkville for providing his expertise with the scanning electron microscope and

for helping me obtain the SEM images for all my samples.

Lastly, my parents deserve special mention for their endless love, encouragement and

prayers without which none of this would be of any significance. I thank my sister and

mother-in-law for always being a source of undying love, support and comfort. Words fail

V

me to express my appreciation to my husband Kiran whose love and persistent confidence

have been the foundations for my strength at difficult times and my motivation to complete

my study. I will be forever be grateful to him for understanding my passions, ambitions and

commitment for the success of the project.

VI

Table of Contents

ABSTRACT II

DECLARATION III

ACKNOWLEDGEMENTS IV

TABLE OF CONTENTS VI

ABBREVIATIONS IX

LIST OF FIGURES X

LIST OF TABLES XII

Chapter 1- INTRODUCTION 1

Chapter 2 - LITERATURE REVIEW 6

2.1 Bioactive compounds in foods 6

2.1.1 Introduction 6

2.1.2 Omega 3 fatty acids 7

2.1.3 Extra Virgin Olive Oil 8

2.1.4 Interaction of bioactives 10

2.1.5 Bioavailability 11

2.2 Microencapsulation 12

2.2.1 Microencapsulation- Today’s approach 12

2.2.2 Encapsulating materials 14

2.2.3 Emulsion preparation 16

2.2.4 Microencapsulation process 17

2.2.5 Incorporation of microencapsulated bioactives into foods 19

2.2.6 Recent developments in Microencapsulation 19

2.3 Sugar beet pectin as encapsulant 21

Chapter 3 – MATERIALS AND METHODS 23

3.1 Raw material Composition 23

3.1.1 Sugar beet pectin 23

3.1.2 Fish oil & Extra virgin olive oil 23

3.1.3 Oxidation status of Fish oil and Extra virgin olive oil 24

3.2 Preparation of emulsions & microcapsules 25

VII

3.3 Spray-dried microcapsules 29

3.4 Emulsion Characteristics 30

3.4.1 Particle size 30

3.4.2 Light microscopy 30

3.4.3 Viscosity 30

3.4.4 Oxidative stability of emulsions under accelerated conditions 30

3.4.5 Zeta-potential of emulsions 31

3.5 Characterisation of the spray-dried microcapsules 31

3.5.1 Moisture content 31

3.5.2 Water activity 31

3.5.3 Particle size of reconstituted powders 32

3.5.4 Fatty acid composition analysis 32

3.5.5 Determination of free oil content 32

3.5.6 Determination of total oil content 32

3.5.7 Encapsulation efficiency 32

3.5.8 Propanal and hexanal analysis 33

3.5.9 Oxidative stability of the microcapsules under accelerated 33

storage conditions

3.5.10 Scanning electron microscopy (SEM) 33

3.6 Statistical analysis 34

Chapter 4 – RESULTS AND DISCUSSION 35

4.1 Properties of raw material 35

4.1.1 Sugar beet pectin 35

4.1.2 Fish oil & extra virgin olive oil 35

4.1.3 Oxidation status of fish oil and extra virgin olive oil 37

4.2 Effect of partial substitution of fish oil with extra virgin olive oil 38

on emulsion and microcapsule characteristics

4.3 Effect of EDTA on emulsion and microcapsule characteristics 49

4.4 Effect of pH on the emulsion and microcapsule characteristics 57

4.5 Lipid Oxidation 65

VIII

4.5.1 Oxidative stability of emulsions and microcapsulated oil during 66

storage under ambient conditions

4.5.2 Oxidative stability of emulsions and microcapsules exposed 79

to accelerated storage conditions

Chapter 5– GENERAL CONCLUSIONS AND FUTURE DIRECTIONS 86

References 88

Appendices 109

IX

Abbreviations

SBP - Sugar beet pectin

FO - Fish oil

EVOO - Extra virgin olive oil

DGS - Dried glucose syrup

EDTA - Ethylenediaminetetras-acetic acid

EPA - Eicosapentaenoic acid

DHA - Docosahexaenoic acid

ALA- Alpha-linolenic acid

ß- CD – ß cyclodextrin

PCL- Polycaprolactone

CMC – Carboxy methyl cellulose

PEG – Polyethylene glycol

PV- Peroxide value

IP- Induction period

X

LIST OF FIGURES

Figure 1 Illustration of the steps involved and the factors considered in the

production of microcapsules

14

Figure 2 Schematic structure for generalised sugar beet pectin 22

Figure 3 Standard curve obtained after assessment of the oxidative stability

using an oxipress

25

Figure 4 Process flow chart of the experimental plan for the production of spray

dried microcapsules

27

Figure 5 Emulsions and spray-dried powders 29

Figure 6 Particle size distributions of the original (fresh) emulsions, prior to

spray drying

39

Figure 7 Brightfield micrographs of (A) FO-25 (7.5% oil, wet basis) and (B)

FO-EVOO-50 (15% oil, wet basis) emulsions at pH 3

40

Figure 8 Apparent viscosities of fish oil and fish oil-extra virgin olive oil (1:1 wt

ratio) emulsions (30% TS; 2% SBP, 20.5% DGS, 7.5% total oil and

30% TS; 2% SBP, 13% DGS, 15% total oil), intended for manufacture

of spray-dried microcapsules containing 25 and 50% oil, as a function

of shear rate

41

Figure 9 Autotitration curve for FO emulsion with zeta potential measured as a

function of pH at 22 º C

43

Figure 10 Autotitration curve for FO-EVOO emulsion with zeta potential

measured as a function of pH at 22 º C

43

Figure 11 Particle size distributions of reconstituted spray-dried microcapsules at

0, 1, 2 and 3 month (A–D) stored under room temperature conditions

(~25ºC, exposed to light)

47

Figure 12 Scanning electron microscopy images of the fish oil (A, C) and fish

oil-extra virgin olive oil (B, D) spray-dried powders (25% oil loading),

prior (A, B) and after 3 month storage (C, D) under ambient conditions

(~25°C, 0.5 aw)

48

XI

Figure 13 Particle size distributions of the original (fresh) emulsions (15% oil,

wet basis), with and without EDTA, at pH 6

50

Figure 14 Brightfield micrographs of FO-EDTA (A) and FO-EVOO-EDTA (B)

emulsions with 15% oil, at pH 6

50


ratio) emulsions (15% oil, wet basis) with and without EDTA at pH 6,

as a function of shear rate

51

Figure 16 Particle size distribution of reconstituted spray-dried microcapsules

(50% oil, dry basis), with and without EDTA at pH 6 after 0, 1, 2 and 3

month (A–D) storage at room temperature (~25ºC)

56

Figure 17 Scanning electron microscopy images of the fish oil-extra virgin olive

oil spray-dried powders (50% oil, dry basis), with (A, C) and without

(B, D) EDTA, at pH 6, prior (A, B) and subsequent (C, D) to storage for

3 months at room temperature (~25ºC)

57

Figure 18 Particle size distributions of the original (fresh) fish oil and fish

oil-extra virgin olive oil (1:1 wt ratio) emulsions made at pH 3 and 6

58

Figure 19 Brightfield micrographs of (A) fish oil (pH 3) and (B) fish oil-extra

virgin olive oil (1:1 wt ratio) emulsions (pH 6) with 15% oil

59


ratio) emulsions with 15% oil at pH 3 and pH 6, as a function of shear

rate

60

Figure 21 Particle size distributions of reconstituted spray-dried fish oil and fish

oil-extra virgin olive oil microcapsules prepared from emulsions at pH

3 and 6 with 50% oil loading prior (A), and subsequent to storage for 1

(B), 2 (C) and 3 (D) month under room temperature conditions (~25 ºC,

aw 0.5, exposed to light)

6

Figure 22 Scanning electron microscopy images of fish oil spray-dried

microcapsules from emulsions prepared at pH 3 (A, C) and pH 6 (B, D)

with 50% oil loading, prior (A, B) and subsequent (C, D) to storage at

ambient conditions (~25°C, aw 0.5, 3 months)

65

XII

LIST OF TABLES

Table 1 List of types of encapsulation processes 18

Table 2 Formulation of the fish oil and fish oil-extra virgin olive oil (1:1 wt ratio)

emulsions at pH 3

28

Table 3 Formulation of the fish oil and fish oil-extra virgin olive oil (1:1 wt ratio)

emulsions with and without EDTA at pH 6

28

Table 4 Fatty acid composition (%) of the initial oils 36

Table 5 Oxidative stability of original oils during accelerated storage (80°C,

oxygen pressure of 0.5 MPa)

38

Table 6 Zeta potential values of FO and FO-EVOO emulsions measured at 22º C 42

Table 7 Properties of spray-dried powders containing microencapsulated fish oil

and fish oil-extra virgin olive oil (1:1 wt ratio) (25 % & 50 % oil loading)

at pH 3

44

Table 8 Changes in moisture content and water activity in FO and FO-EVOO

microcapsules (25 % & 50 % oil loadings) over storage time

46

Table 9 Particle size of the reconstituted spray-dried microcapsules containing

fish oil and fish oil-extra virgin olive oil (1:1 wt ratio) during storage of

the powders in stoppered flasks at room temperature (~25ºC)

46

Table 10 Properties of powders containing microencapsulated fish oil and fish

oil-extra virgin olive oil blends (1:1 wt ratio) with and without EDTA at

pH 6, 50% oil (dry basis)

53

Table 11 Changes in moisture content and water activity of FO and FO-EVOO

microcapsules (50% oil, dry basis) (with and without EDTA) at pH 6 on

storage at room temperature (~25°C)

55

Table 12 Particle size values of the reconstituted spray-dried microcapsules (50%

oil, dry basis), with and without EDTA at pH 6, on storage at room

temperature (~25ºC)

55

Table 13 Properties of powders containing microencapsulated fish oil and fish

oil-olive oil blends (1:1 wt ratio) at pH 3 and pH 6 with 50 % oil loading

61

XIII

Table 14 Changes in moisture content and water activity of fish oil and fish

oil-extra virgin olive oil (1:1 wt ratio) microcapsules at pH 3 and pH 6

with 50% oil loading over storage time at room temperature (~25°C)

62

Table 15 Particle size values of the reconstituted spray-dried microcapsules

containing fish oil and fish oil-extra virgin olive oil (1:1 wt ratio) at pH 3

and pH 6 with 50% oil loading, during storage of the powders in stoppered

flasks at room temperature (~25°C)

64

Table 16 Propanal content in microencapsulated fish oil and fish oil-extra virgin

olive oil (1:1 wt ratio) (25 % & 50 % oil loading) at pH 3 during storage at

room temperature (~25°C, 0−3 month)

68


olive oil (1:1 wt ratio) with and without EDTA (50% oil loading) at pH 6

during storage at room temperature (~25 °C, 0−3 month)

68


olive oil (1:1 wt ratio) with 50 % oil loading at pH 3 and pH 6 during

storage at room temperature (~25°C, 0−3 month)

70

Table 19 Hexanal content in microencapsulated fish oil and fish oil-extra virgin

olive oil (1:1 wt ratio) (25 % & 50 % oil loading) at pH 3 during storage at

room temperature (~25°C, 0−3 month)

71


olive oil (1:1 wt ratio) with and without EDTA (50 % oil loading) at pH 3

and pH 6 during storage at room temperature (~25°C, 0−3 month)

72


olive oil (1:1 wt ratio) with 50 % oil loading at pH 3 and pH 6 during

storage at room temperature (~25°C, 0−3 month)

72

Table 22 a Fatty acid composition of microencapsulated fish oil (25 % and 50 % oil

loading) at pH 3 during storage at room temperature (~25°C, 0−3 month)

75

Table 22 b Fatty acid composition of microencapsulated fish oil – olive oil (25 % and

50 % oil loading) at pH 3 during storage at room temperature (~25°C, 0−3

month)

76

XIV

Table 23 a Fatty acid composition of microencapsulated fish oil (50 % oil loading)

with and without EDTA at pH 6 during storage at room temperature

(~25°C, 0−3 month)

77

Table 23 b Fatty acid composition of microencapsulated fish oil – olive oil (50 %

oil loading ) with and without EDTA at pH 6 during storage at room

temperature (~25°C, 0−3 month)

78

Table 24 Oxidative stability of bulk oils (FO, EVOO, FO-EVOO (1:1 wt ratio)

during accelerated storage (80°C, oxygen pressure of 0.5 bar)

80

Table 25 Oxidative stability of emulsions and powders containing

microencapsulated fish oil and fish oil-extra virgin olive oil (1:1 wt ratio)

(25% and 50% oil loading) at pH 3 during accelerated storage (80°C,

oxygen pressure of 0.5 MPa)

81

Table 26 Oxidative stability of emulsions and powders containing

microencapsulated fish oil and fish oil-extra virgin olive oil (1:1 wt ratio)

made with and without EDTA (50% oil loading) at pH 3 during

accelerated storage (80°C, oxygen pressure of 0.5 MPa)

84

Table 27 Oxidative stability of emulsions and powders (50% oil loading)

containing fish oil and fish oil-extra virgin olive oil (1:1 wt ratio) at pH 3

and pH 6 during accelerated storage (80°C, oxygen pressure of 0.5 bar)

85

1

CHAPTER 1 - INTRODUCTION

This chapter provides the background and purpose to the research topic and discusses the

need for further research related to the area. The project hypothesis and specific aims of the

research are outlined.

Microencapsulation technology has been the central concept of this research and the two

other areas discussed in relation to it are the protection of bioactives (omega 3 fatty acids,

and polyphenols) and the application of sugar beet pectin as an encapsulant. Literature

review on the current scientific information/existing research in the above mentioned fields

was reviewed in the second chapter.

The third chapter includes all details about the raw materials (supplier details and

manufacturer’s specification), chemicals (supplier details), equipments (with the

manufacturer’s details and model numbers) used in the research project. It also describes

the procedures/method of analysis adopted for the characterization of emulsions and

microcapsules.

Chapter four describes the results obtained after analysis of the raw materials, emulsions

and microcapsules following the analytical methods described in Chapter 3 - Methods

(Section 3.1, 3.4, 3.5, 3.6). Conclusive inferences from the results obtained were also

presented.

Based on the results of the various analyses conducted and the comparison of results

performed between samples, the final conclusions of this study is summarised in fifth

chapter.

2

1.1 BACKGROUND

Consumer behaviour and attitudes toward healthy foods continue to grow especially in

developed countries. This has sparked a wide interest in foods, such as, functional foods

which contain ingredients with health promoting properties, and can offer specific health

benefits to consumers in addition to the anticipated nutritional value.

Functional foods have the potential to contribute to a healthier society and are important

components of an overall healthful lifestyle that includes a balanced diet and physical

activity. There have been mounting evidence in recent years that food choices are an

important factor in reducing the risk of developing heart disease, cancer, obesity, high

blood pressure, osteoporosis, and other unhealthy conditions (Australian Indigenous

Healthinfonet, 2008).

Fish oils are considered functional foods as they are a rich source of omega-3 fatty acids

such as eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) (Erkkila et al.,

2006). There has been mounting evidence in the recent years that omega-3 fatty acids have

positive long-term and current health benefits. Several prospective studies and clinical

trials have shown that omega-3 fatty acids can reduce the risk of developing heart disease

(Erkkila et al., 2006; GISSI-HF investigators, 2008). Various other studies have shown

that consumption of omega-3 fatty acids have been beneficial in treating neurological

disorders such as Alzheimer’s disease (Morris et al., 2003), in reducing bone loss (Griel et

al., 2007; Weiss, Barrett-Connor & von Muhlen, 2005) in lowering incidence of

depression and other related conditions (Lin & Su, 2007). However, the incorporation of

functional ingredients in a given food system and the processing and handling of such

foods are associated with nutritional challenges for their healthy delivery. The extreme

sensitivity of fish oils to oxidation can easily lead to the development of off-flavours and

cause significant loss of product quality, stability, nutritional value, and bio-availability

and the overall acceptability of the food product (Nawar, 1996; Watkins & German, 1998).

Consequently, microencapsulation has been successfully used to encapsulate omega-3 fatty

acids in order to prevent oxidation and to improve stability and bioavailability (Wakil et

3

al., 2010; Sanguansri & Augustin, 2006). Microencapsulation can be defined as a process

in which tiny particles or droplets are surrounded by a coating to give small capsules. The

material inside the microcapsule is referred to as the core, internal phase, or fill, whereas

the coating is sometimes called a shell, membrane, or wall material, as referred to

henceforth. Most microcapsules have diameters between a few micrometers and a few

millimetres (Microencapsulation, 2009). Microencapsulation is one example of technology

that has the potential to meet the challenge of successfully incorporating and delivering

functional ingredients into a range of food types.

The most common technology used for encapsulation has been spray drying because it is

efficient, cost effective, has readily available equipment and produces particles of

reasonably good quality (Dobry et al., 2009). Some of the encapsulant materials used in

the microencapsulation of foods are proteins, sugars, starches, gums, lipids and cellulosic

materials. Chitosan plus maltodextrin and liposomes have been reported in encapsulation

of omega-3 rich fish oils (Klaypradit & Huang, 2008; Reineccius, 1995). A considerable

number of research studies have been conducted on the oxidative stability of fish oil,

microencapsulated with different wall materials using one of the many chemical and

physical microencapsulation processes available. Literature review on the oxidative

stability of omega-3 fatty acids showed a variation of results due to the effectiveness of

wall materials used, changes in the process parameters, storage conditions and methods

employed for assessing the oxidative stability. Thus, further studies were even conducted

with the addition of antioxidants to improve the shelf life of the fish oil microcapsules

(Velasco, Dobarganes & Marquez –ruiz, 2000; Heinzelmann et al., 2000).

Several studies suggested that SBP could be a suitable wall material for manufacturing

lipophilic food ingredients due to its excellent emulsifying properties (Drusch, 2007;

Drusch et al., 2007; Leroux et al., 2003; Martinez-Dominguez, de la Puerta, &

Ruiz-Gutierrez, 2001). This functionality associated with SBP showed that it had potential

as a wall material for encapsulating w-3 acids in our study. Recent studies conducted by

Drusch (2007) also confer that SBP could be a promising alternative to the traditional

encapsulating agents like milk proteins and gum Arabic because of its ability to form

4

stable emulsions with improved encapsulation properties. However, further research is

needed to test the oxidative stability of the microcapsules encapsulated with SBP over

prolonged storage time. The drawback of using SBP as an encapsulant material is the

presence of transition metals in high concentrations i.e. iron (1.91 ppm) and copper (0.08

ppm) (Katsuda et al., 2008). Data reported in the literature established the fact that metals

like iron (Fe) and copper (Cu) act as catalysts for the formation of highly reactive hydroxyl

radicals which initiate a chain reaction for lipid oxidation (Benedt and Shibamoto, 2007;

Goldstein et al., 1993). Hence, in the present research study, ethylenediaminetetra-acetic

acid (EDTA) was used as a chelating agent and as an antioxidant to mask the negative

effects of these metal ions. EDTA is a polyamino carboxylic acid and a colourless,

water-soluble solid. EDTA is capable of forming complexes with metal ions in a solution

and diminishing their reactivity. EDTA and its derivatives are most commonly used

because they are inexpensive and effective even when used in small amounts in the overall

formulation (EDTA, 2010; Kabara & Orth, 1997).

1.2 THESIS HYPOTHESIS

It is hypothesised that the combination of the fatty acid composition and antioxidants

(polyphenols) in EVOO would improve the oxidative stability of the microencapsulated

fish oil powders during storage, and that SBP as the material would provide a good matrix

to maintain the nutritional value in encapsulated fish oil.

1.3 THESIS PURPOSE AND AIMS

The purpose of the research project was to microencapsulate fish oil (FO) mixed with

EVOO at a 1:1 ratio and using SBP as the wall material. EVOO was added as a source of

oleic acid and polyphenols. Polyphenolic components are known to possess potent

antioxidant properties (Olive Oil, 2008). The addition of the chelating agent, EDTA, was

evaluated for its ability to retard the lipid oxidation caused by the transition metal ions

present in SBP.

5

The aim of this research study was to examine the:

• Oxidative stability of the microencapsulated fish oil (FO) in the presence of extra

virgin olive oil (EVOO).

• Effect of SBP as the wall material to prevent oxidative stability of

microencapsulated FO.

• Influence of EDTA on the oxidative stability of FO and FO-EVOO blend

emulsions and microcapsules.

• Influence of pH on the physico-chemical characteristics and oxidative stability of

FO and FO-EVOO emulsions and microcapsules

6

CHAPTER 2 - LITERATURE REVIEW

2.1. BIOACTIVE COMPOUNDS IN FOODS

2.1.1 INTRODUCTION

Bioactive compounds are biomolecules present in foods that exhibit the capacity to

modulate one or more metabolic processes, which promote better health. They indulge in

multiple metabolic activities and are beneficial for treating several diseases and target

tissues and are usually found in glycosylated, esterified, thiolyated, or hydroxylated forms.

Bioactive food components are found abundantly in most fruits and vegetables. However,

probiotics, conjugated linolenic acid, long-chain omega-3 polyunsaturated fatty acid, and

bioactive peptides are most commonly found in animal products such as milk, fermented

milk products and cold-water fish (Encyclopaedia of Food and Culture, 2003).

Numerous bioactive components from plant sources (e.g. phytosterol, carotenoids,

flavonoids, soluble fibre, polyphenols etc.) marine oils, milk and milk products (e.g.

peptides / proteins, pre and pro-biotics) are currently used in functional food formulations

(McClements et al., 2009; Engel & Schubert, 2006; Murphy, 2001; López-López et al.,

2009). However, a significant lack of understanding of bioactive ingredient’s availability,

mechanism of action and synergistic effects exists and hence, the acceptable levels of

specific bioactive ingredients to be used in different foods have to established to establish

the appropriate dosage of healthy bioactive components.

Significant research findings stated in the Encyclopaedia of Food and Culture (2003)

indicated that bioactive food components could act simultaneously at different or identical

target sites. For instance, in cardiovascular disease, the bioactive component isoflavone

may reduce circulating oxidized low-density lipoproteins in the plasma, bind cholesterol in

the intestinal tract and thereby reduces absorption of dietary cholesterol. Another example

is the lipid-lowering mechanism of dietary fibre and phytosterol/stanols. It occurs by

sequestering cholesterol in the intestinal tract and reducing cholesterol absorption. The

7

bioactive food components have thus been proven to have health-promoting characteristics

and reduce the risk of cancer, cardiovascular disease, osteoporosis, inflammation, type 2

diabetes, and other chronic degenerative diseases (Encyclopaedia of Food and Culture,

2003; Ferguson & Philpott, 2007; Pfeuffer & Schrezenmeir, 2000; Hooper & Cassidy,

2006; Shahidi, 2006).

Clearly, bioactive food components can play an important role in better health and disease

prevention. Consequently, there is a need to produce foods with optimal levels of

health-promoting bioactive food components that would assist consumers to make healthy

food choices for their diet.

2.1.2 OMEGA 3 FATTY ACIDS

Omega-3 fatty acids are considered essential fatty acids. They are essential to human

health but cannot be manufactured by the body. For this reason, omega-3 fatty acids must

be obtained from food. Also known as polyunsaturated fatty acids (PUFAs), omega-3 fatty

acids play a crucial role in brain function as well as normal growth and development (drsri,

2009).

Fish, plant, and nut oils are the primary dietary source of omega-3 fatty acids. Best food

sources containing both eicosapentaenoic acid (EPA), C20:5(n-3), and docosahexaenoic

acid (DHA), C22:6(n-3), are cold-water fatty fishes such as, salmon, mackerel, halibut,

sardines, tuna, and herring. Dietary sources of alpha-linolenic acid (ALA), C18:3(n-3) are

flaxseeds, flaxseed oil, canola (rapeseed) oil, soybeans, soybean oil, pumpkin seeds,

pumpkin seed oil, purslane, perilla seed oil, walnuts, and walnut oil. ALA is converted in

the body into EPA and DHA, which are more readily utilised in vivo (Drsri, 2009).

Extensive research indicates that omega-3 fatty acids have antiinflammatory properties and

reduce the risk associated with chronic diseases such as heart disease, cancer, and arthritis

(Geusens et al., 1994; Richardson, 2004; GISSI-prevenzione investigators, 1999). These

essential fatty acids are an important structural component in the brain and appear to be

particularly important for cognitive (brain memory and performance) and behavioural

function (Willat et al., 1998; Mellor, Langharne & Peet, 1995). Symptoms associated with

8

omega-3 fatty acid deficiency include extreme tiredness (fatigue), poor memory, dry skin,

heart problems, mood swings or depression, and poor circulation (Omega-3-fatty acids,

2010).

An appropriate balance of omega-3 and omega-6 in the diet is vital, as they work together

to promote health. Whilst omega-3 fatty acids help reduce inflammation, most omega-6

fatty acids tend to promote inflammation. An inappropriate balance of these essential fatty

acids contributes to the development of disease, while a proper balance helps maintain and

even improve health. A healthy diet should consist of roughly 2 - 4 times more omega-6

fatty acids than omega-3 fatty acids (Omega-3-fatty acids, 2010).

Omega-3 fatty acids are extremely susceptible to oxidative damage from heat, light, and

oxygen. When exposed to the air for long periods, the fatty acids in the oil become

oxidized and rancid. Rancidity not only gives the oil disagreeable off-flavour and smell,

but it also diminishes the nutritional value. More importantly, free radicals are produced in

this process and which are implicated in the development of cancer and other degenerative

diseases (Pham-Huy, He & Pham-Huy, 2008).

Rancidity arises when the oils are removed from their food package or are stored

improperly. Hence, oils rich in polyunsaturated fatty acids should be stored in dark glass,

tightly closed containers in the refrigerator or freezer.

2.1.3 EXTRA VIRGIN OLIVE OIL

Extra virgin olive oil (EVOO) comes from cold pressing of the olives, contains no more

than 0.8% acidity, and is judged to have a superior taste, delicate flavour and most

antioxidant benefits. Olive oil is composed mainly of oleic acid, C18:1(n-9), and palmitic

acid, C16:0, as well as other minor fatty acids, along with traces of squalene (up to 0.7%)

and sterols (about 0.2% phytosterol and tocosterols). The composition varies by cultivar,

region, altitude, time of harvest, and extraction process (Alarcon de la Lastra et al., 2001;

‘Olive oil’, 2008). Olive oil also contains a group of related natural products with potent

9

antioxidant properties which give extra-virgin unprocessed olive oil its bitter and pungent

taste (Tripoli et al., 2005).

Diets rich in monounsaturated fats have been shown to reduce the risk of coronary heart

disease (Keys et al., 1986; Willet et al., 1995). This is particularly noteworthy, because

olive oil is extremely rich in monounsaturated fats, most notably oleic acid (‘Olive oil’,

2008). Another health benefit of olive oil is its ability to displace omega-6 fatty acids,

while not having any impact on omega-3 fatty acids. This way, olive oil helps to build a

more healthy balance between omega-6 and omega-3 fatty acids (Haban et al., 2004).

Oleic acid is found in various animal and vegetable sources. It comprises 55-80% of olive

oil, and the saturated form of oleic acid is called stearic acid. Oleic acid shares the normal

reactions of fatty acids and alkenes. Oxidation at the double bond can split the molecule,

yielding chain ends with aldehyde, ketone or carboxylic acid groups. This reaction occurs

slowly in air, and it causes the oil in which it is present to go rancid (‘Oleic acid’, 2008).

Polyphenols are a group of chemical substances found in plants, characterized by the

presence of more than one phenol unit or building block per molecule. Polyphenols are

generally divided into hydrolyzable tannins (gallic acid esters of glucose and other sugars)

and phenylpropanoids, such as lignins, flavonoids, and condensed tannins. Notable sources

of polyphenols include berries, tea, beer, grapes/wine, olive oil, chocolate/cocoa, coffee,

walnuts, peanuts, pomegranates, yerba mate, and other fruits and vegetables. High levels of

polyphenols can generally be found in the fruit skins (‘Polyphenols’, 2008).

Based on the epidemiologic studies conducted by Arts & Hollman, (2005), it has been

observed that olive oil phenolic content, rather than its fatty acid profile is responsible for

at least some of its cardio protective benefits. There has been growing evidence that as

antioxidants, polyphenols may protect cell constituents against oxidative damage and,

therefore, limit the risk of various degenerative diseases associated with oxidative stress

(Luqman & Rizvi, 2006; Pandey, Mishra & Rizvi, 2009). Polyphenols also exert protective

effects in treating conditions like cancer, diabetes, aging, asthma and skin damages

(Pandey & Rizvi, 2009).

10

2.1.4 INTERACTIONS OF BIOACTIVE COMPOUNDS

Bioactive compounds are susceptible to variations due to changes in their structure and

physicochemical properties as a result of processing and the possible interactions with

other food components and the intestinal microflora.

Functional foods rarely contain a single bioactive component and interactions between

bioactive components in the food is expected which could either be positive or negative.

For example, Vitamin C has been reported to reduce selenium’s effectiveness against

chemically induced colon cancer (Ip, 1986). Selenium has been shown enhance the ability

of garlic to inhibit chemically induced mammary cancer in experimental animals

(Amagase, Schaffer & Milner, 1996).

There are very few research studies that can confirm or demonstrate the interaction or

effect of polyphenols with omega-3 fatty acids. A study conducted by Ren et al. (2008)

showed that homozygous sickle cell patients (HbSS) had reduced EPA and DHA in red

cells, platelets, and mononuclear cells when compared to their healthy counterparts

(HbAA). This difference in levels of omega-3 fatty acids was not because of reduced

omega 3 intake. Something else besides dietary deficiency was causing lower cell levels of

cellular omega-3 fatty acids in these patients. The study investigated, and concluded that

the lower levels of membrane EPA and DHA in blood cells of the HbSS patients could be

attributed to peroxidation resulting from a compromised antioxidant competence. Omega-3

fatty acids aid in cell membrane function and stability and polyphenols are known to

directly and indirectly prevent peroxidation of fats. In addition, the antioxidant status

affects cell membrane levels of omega-3 fatty acids in humans, and polyphenols were

clearly found to be superior dietary antioxidants to those studied in the above mentioned

research study (beta-carotene, tocopherol). Hence, there is a possibility that polyphenols

could act to protect omega-3 fatty acids in the cell membrane.

11

2.1.5 BIOAVAILABILITY OF NUTRIENTS

From a nutritional point of view nutrient bioavailability can be defined as the amount our

digestive system will be able to extract in a form that can be absorbed into the bloodstream

(Encyclopaedia of Food and Culture, 2003).

It has been established that during fat absorption, unsaturated long chain fatty acids are

esterified at a higher rate than saturated fatty acids of similar chain length. This

phenomenon has been attributed to differences in the binding affinity of fatty acids to a

cytosolic fatty acid-binding protein (Gang et al., 1980). Studies have been conducted to

evaluate the bioavailability of omega-3 fatty acids in microencapsulated fish-oil-enriched

foods compared with an equal amount of omega-3 PUFAs contained in fish oil capsules.

The results indicated that n-3 PUFA from microencapsulated fish-oil-enriched foods were

as bioavailable as n-3 PUFA in a capsule. Therefore, fortification of foods with

microencapsulated fish oil offers an effective way of increasing omega-3 PUFA intakes

and status in line with current dietary recommendations (Wallace et al., 2000; Volker,

Weng, Quaggiotto, 2005).

Singh et al. (2008) indicated that bioavailability and pharmacokinetics of polyphenolics

are governed by a number of factors; their native form (glycosylated/aglycone), the type of

sugar moiety present, and their physiochemical properties. Bioavailability of polyphenols

varies widely from one compound to another. It depends on their chemical structure, which

determines their absorption rate through the gastrointestinal tract, metabolism, and,

therefore, bioactivity (Manach, 2004). In nature, phenolic compounds occur as glucosides.

Phenolic acids like caffeic acid are easily absorbed through the gut barrier, whereas large

molecular weight polyphenols, such as proanthocyanidins are very poorly absorbed. Once

absorbed, polyphenols are conjugated to glucuronide, sulphate and methyl groups in the

gut mucosa and inner tissues. Non-conjugated polyphenols are virtually absent in plasma.

These reactions facilitate their excretion and limit their potential toxicity. The polyphenols

reaching the colon are extensively metabolised by the microflora into a range of low

molecular weight phenolic acids (Scalbert, 2002).

12

Polyphenols present as aglycones can be absorbed from the small intestine. However, most

of them are present in the form of esters, glycosides, or polymers and are not easily

absorbed in their natural form. Glycosylation influences chemical, physical, and biological

properties of the flavonoids and their absorption. It is generally accepted that the

breakdown of these conjugates to aglycones by acid hydrolysis in the stomach and by

microflora in the gut is required to produce the bioactive components that are readily

bioavailable to the body. However, relatively little is known about the ability of these

aglycone polyphenolics to reach the target cells or what the influence of further

metabolism in the body has on their spectra of biological activities. There are numerous

sites important for the metabolism of dietary polyphenols, including the gastrointestinal

tract, the liver, and various other organs such as the skin and brain (Singh et al., 2008).

The food matrix also plays a vital role in the bioaccessibility of a bioactive component in

functional foods. For example, a lipophilic food matrix is needed for bioavailability of

carotenoids whereas when phenolic compounds are in a food matrix the bioavailability of

certain polyhenols may be lowered depending on the matrix components (Parada &

Aguilera, 2007). Another study by Kean, Hamaker, & Ferruzzi, (2008) found that specific

food preparations of maize based food products could influence the bioaccessibility of

certain bioactive carotenoid species.

2.2 MICROENCAPSULATION

2.2.1 MICROENCAPSULATION-TODAY’S APPROACH

Microencapsulation has been defined as the technology of packaging solid, liquid and

gaseous materials in small capsules that release their contents at controlled rates over

prolonged periods of time (Champagne & Fustier, 2007).

Microencapsulation technologies have been used extensively in the pharmaceuticals sector

and subsisted for many decades before its application in the food industry. With the

13

identification of exciting and new microencapsulation techniques, new and innovative

approaches have been developed for the protection and targeted delivery of bioactives.

Today, microencapsulation has become a viable means that enables the use of previously

difficult-to-use ingredients (e.g. sensitivity to air/oxygen, heat, light) and the introduction

of a variety of ingredients and properties into foods that were not previously available.

Recent years have seen a widespread popularity of this technology among food ingredients

companies to achieve a variety of foods with different functions. Figure 1 illustrates the

various factors to consider in the production of microcapsules.

Food manufacturers have now become increasingly receptive to the use of encapsulated

ingredients, which have become more economically viable. Indeed, changing consumer

trends and needs have also been the significant factors for essentially driving innovation in

the field of microencapsulation.

With consumers showing a growing preference for functional food - which now accounts

for a substantial amount of the global “wellness” market - food companies are looking for

different ways to incorporate health-promoting ingredients that deliver some kind of health

benefit to the consumer. While consumers are demanding more nutritious products, they

are unwilling to compromise on taste or quality. Microencapsulation can provide solutions

to these recent demands because of its ability to mask the undesirable tastes, prevent

oxidation of ingredients and also aid in targeted nutrition in a number of processed food

products.

14

Figure 1: Illustration of the steps involved and the factors considered in the production of

microcapsules (Sanguansri and Augustin, 2006) with some modifications.

2.2.2 ENCAPSULANTING MATERIALS

Another definition of microencapsulation is a process in which tiny particles or droplets

are surrounded by a coating, sometimes referred to as a shell, wall material, membrane or

an encapsulant material (‘Microencapsulation’, 2010).

A critical step in the production of microcapsules is the selection of appropriate

encapsulating material. The encapsulant material can be selected from a wide variety of

natural or synthetic polymers depending on the stability and release characteristics

expected from the final microcapsule. It has been reported that the composition as well as

Identification of high quality source of active

ingredient to be encapsulated

Development of successful formulation

for a targeted application

Selection of a format for the delivery of the

microcapsule

- Physical and chemical properties of the bioactive

- Stability of the bioactive

- Physical and chemical properties of the bioactive

- Stability of the bioactive

- Selection of appropriate encapsulant material

- Selection of suitable microencapsulation process

- Addition of antioxidants, chelating agents, emulsifiers, stabilizers and salts

- Dry product or liquid format

15

the physical and chemical properties of the shell material can influence the functionality of

the final microcapsule and the processing technologies to be used for microencapsulation

(Sanguansri & Augustin, 2006). According to Augustin & Sanguansri (2007) a good

encapsulating material should have neutral taste and odour, low viscosity, good film

forming, gelling and barrier properties. It should also protect the core from degradation

during processing and storage and also mask any undesirable taste or odour associated with

the bioactive core when added into foods.

Some of the common encapsulant wall materials have been summarised by Vasishtha

(2003) and includes the following:

• Polysaccharides/hydrocolloids, such as starch, algin/alginate, agar/agarose,

pectin/polypectate, carrageenan, and other gums.

• Proteins such as gelatin, casein, zein, soy, and albumin.

• Fats and fatty acids such as mono-, di- and triglycerides, and lauric, capric, palmitic

and stearic acid and their salts.

• Cellulosic derivatives such as methyl- and ethyl-cellulose and CMC (carboxy

methyl cellulose)

• Hydrophilic and lipophilic waxes such as shellac, PEG (polyethylene glycol),

carnauba wax and beeswax.

• Sugar derivatives.

Conjugates of casein and whey protein have been suggested as effective encapsulating

material for fish oil (Livney, 2010). Polymers such as β-cyclodextrin (β-CD) and

polycaprolactone (PCL) were used to encapsulate fish oil using aggregation and emulsion

diffusion methods and it was found that PCL protected FO better than (β-CD) (Choia et al.,

2010). The use of complex coacervates composed of gelatin or β-lactoglobulin, gum

arabic, starch, and crosslinked with glutaraldehyde have been reported to decrease

oxidation in fish oil (Lumsdon, Friedmann & Green, 2005). Other encapsulant materials

used for the encapsulation of omega-3 fatty acids were Maillard reaction products. Fish oil

here is emulsified with heated aqueous mixtures comprising of a protein source (sodium

caseinate, whey protein isolate, soy protein isolate, or skim milk powder) and

16

carbohydrates (glucose, dried glucose syrup and oligosaccharide) (Augustin, Sanguansri

and Bode, 2006).

Different types of n-octenylsuccinate-derivatised starch have also been used as encapsulant

materials for fish oils and were found to be effective encapsulant materials (Drusch &

Schwarz, 2006). Chitosan plus maltodextrin could also offer an alternative for the

encapsulation of omega-3 rich fish oils (Daniells, 2007). Liposomes have also been

employed to extend the storage stability and shelf life of encapsulated omega 3 fatty acids

(Haynes et al., 1992). Recent studies conducted by Drusch (2007) also showed a

possibility of using sugar beet pectin as an encapsulant material for omega-3 fatty acids

2.2.3 EMULSION PREPARATION

An emulsion is a term used to describe a mixture containing two or more immiscible

liquids where one liquid (the dispersed phase) is dispersed in the other (the continuous

phase). Emulsions are thermodynamically unstable and normally do not form

spontaneously but require some form of mechanical energy input (e.g.shaking, stirring,

homogenizing or spray processes) to form an emulsion (‘Emulsion’, 2009). An emulsion

is said to be stable when there is no perceptible change in the size distribution of droplets,

its state of aggregation, or its spatial arrangement over the time-scale of observation, which

may vary from hours to months depending on the material (Vega & Roos, 2006;

Dickinson, 1994). Surfactants (e.g. polysorbates, phospholipids), proteins (e.g. milk

proteins) and/or thickening agents (gums, gelatin) can be used to increase the kinetic

stability of the emulsion (Dickinson & Woskett, 1989).

The particle size of emulsions is also a highly relevant factor as it influences the stability

of an emulsion prior to manufacture into spray-dried powder as well as the encapsulation

efficiency after spray drying. Results have shown that larger particles retained more oil

than smaller ones, but at the same time there was more unencapsulated oil at the surface of

big particles than the surface of small particles (Jafari et al., 2008; Jafari, He & Bhandari,

2007; Soottitantawat et al., 2005). The presence of unencapsulated oil on the surface of the

17

powder particles is the most undesirable property of encapsulated powders because of the

detrimental effects it has on the powder quality and stability (Vega et al., 2005).

2.2.4 MICROENCAPSULATION PROCESSES

The selection of a microencapsulation process depends on the properties of the core and

the wall materials, the release mechanisms desired, process type, and desired capsule

morphology and particle size. Most of the microencapsulation processes have been adapted

from the pharmaceutical and chemical industries (Augustin & Sanguansri, 2007). A range

of physical and chemical processes are available to encapsulate bioactive ingredients as

outlined in Table 1.

A process often used for encapsulation of oil and oil-soluble cores involves the use of

biopolymer gels as entrapment matrices. Examples include oil and oil-soluble cores

encapsulated in alginate beads by extrusion to form particles, followed by drying. For

efficient particle formation of this type methods like centrifugal extrusion and microject

methods have been developed (Benita, 1996).

Amongst the encapsulation processes mentioned above spray drying is the most commonly

used technique for microencapsulation. It is one of the oldest methods of encapsulation.

The process of spray drying is efficient, cost effective, has readily available equipment and

produces particles of reasonably good quality (Gharsallaoui, 2007). Food ingredients

microencapsulated by this method include fish oils, essential oils, vitamins, colourants,

flavours and other oil-soluble bioactives.

18

Table 1 List of Types of Encapsulation Processes (Thies, 1996)

Chemical Processes Mechanical Processes

Complex coacervation Spray drying Polymer-polymer incompatibility Spray chilling Interfacial polymerization in liquid Fluidized bed media Electrostatic deposition In situ polymerization Centrifugal extrusion In-liquid drying Spinning disk or rotational suspension separation Thermal and ionic gelation in liquid Polymerization at liquid-gas media or solid-gas interface Desolvation in liquid media Pressure extrusion or into solvent extraction bath spraying

The general process of spray drying involves the dispersion and homogenization of the

substance to be encapsulated and the encapsulating material to form a suspension in water

(slurry). The suspension is then fed into a spray drier. The solution is sprayed into the

drying chamber with the help of an atomiser or spray nozzle. There are different types of

nozzles available like rotary nozzles, single fluid pressure nozzles and ultrasonic nozzles.

Appropriate choice of the kind of nozzle to be used is based on the particle size required

for a specific application. The most common applications are in the 100 to 200 micrometre

diameter range. Hot drying gases are passed as either in the co-current or counter current

direction to the atomiser in the drying chamber. This hot air converts the suspension into

solid and the solvent into vapour. The dried particles are then collected in the particle

separator which is usually a cyclone separator (‘Spray drying’, 2010). The dry powder is

often free flowing and the microcapsules are soluble in water.

Fish oil has been encapsulated by using various formulations, processing conditions and

encapsulation technologies (Gan, Cheng & Easa, 2008; Blatt et al., 2006; Heinzelmann &

Franke, 1999; Lumsdon, Friedmann & Green, 2005; Klinkesorn et al., 2005). According to

19

Sanguansri & Augustin (2006) the formulation and processing steps prior to spray drying

are considered important in the production of stable powder microcapsules. Processing

conditions such as, temperature, pH of the emulsion, total solids, drying rate and the order

of adding each of the components used during the manufacture of fish oil powders also

influence the microcapsule properties and stability.

2.2.5 INCORPORATION OF MICROENCAPSULATED BIOACTIVES INTO

FOODS

The market for functional ingredients and foods has experienced growth in recent years

due to the increased consumer awareness and promotion of healthy eating and lifestyle.

Challenges remain to ensure that functional ingredients survive and remain ‘active’ and

‘bioavailable’ after food processing and storage. Food can be used as a vehicle for the

delivery of bioactives and micronutrients at suitable levels that provide health benefits for

consumers (Day, 2009).

Rapid developments in microencapsulation technologies and delivery strategies have

resulted in increasing numbers of successful omega-3 fortified products in the market. The

omega-3 enriched products available to consumers worldwide are dietary supplements, pet

foods, dairy products, processed fish, meat and egg products, snacks, meals, infant formula

and baby foods, soups, ice tea drinks, cakes, biscuits, bakery products and beverages

(Sanguansri & Augustin, 2006).

2.2.6 RECENT DEVELOPMENTS IN MICROENCAPSULATION (NANOENCA-

-PSULATION)

Nanoscience is an emerging area of science that has the potential to generate radical new

products and processes. Concepts in nanoscience provide a sound framework for

developing an understanding of the interactions and assembly behaviour of food

components into microstructure, which influence food structure, rheology and functional

properties at the macroscopic scale. Advances in processes for producing nanostructured

20

materials coupled with appropriate formulation strategies have made possible the

production and stabilisation of nanoparticles that have potential applications in the food

and related industries (Sanguansri & Augustin, 2006).

Nanoencapsulation technology can be used to address some extraordinary needs of many

applications by producing nanosized particles and capsules. Nanocapsules in combination

with other microencapsulation methods can lead to new release characteristics (Persyn &

Oxely, 2008).

Nanotechnology and nanoscience involve the study of phenomena and materials, and the

manipulation of structures, devices and systems that exist at the nanoscale (<100 nm in

size). The properties of nanoparticles are not governed by the same physical laws as larger

particles, but by quantum mechanics. The physical and chemical properties of

nanoparticles – for example, colour, solubility, strength, chemical reactivity and toxicity -

can therefore be quite different from those of larger particles of the same substance (Miller,

2008).

The altered properties of nanoparticles have created the possibility for many new profitable

products and applications. Engineered nanoparticles have been used to develop hundreds

of products and are available on supermarket shelves – including transparent sunscreens,

light-diffracting cosmetics, penetration enhanced moisturisers, stain and odour repellent

fabrics, dirt repellent coatings, long lasting paints and furniture varnishes, and some food

products (Miller & Kinnear, 2007). It has been speculated that nanoencapsulation may gain

importance in the near future to develop designer probiotic bacterial preparations that

could be delivered to certain parts of the gastro-intestinal tract where they interact with

specific receptors (Paques & van Rijn, 2007). Biodegradable nano/microparticles of poly

(d,l-lactide-co-glycolide) (PLGA) and PLGA-based polymers have been widely explored

as carriers for controlled delivery of macromolecular therapeutics such as proteins,

peptides, vaccines, genes, antigens and some growth factors, etc (Raghavendra et al.,

2008).

21

Nanoencapsulation just like microencapsulation has many applications like protein, DNA

and RNA stabilization, small molecule delivery, extending circulatory half-life, modifying

drug transport, clear liquid formulations, stable colloid dispersions, controlled release,

targeted delivery, triggered release (Persyn & Oxely, 2008).

There has been a rise in concern in the use of nanotechnology as it involves the

manipulation of matter at the scale of atoms and molecules and hence, potentially

introduces some serious risks to human and environmental health. Toxicological literature

suggests that nanoparticles are more reactive, more mobile, and more likely to be toxic to

humans and the environment than larger particles. Preliminary scientific research has

shown that many types of nanoparticles can result in increased oxidative stress which can

result in the formation of free radicals that can lead to cancer, DNA mutation and even cell

death. Additionally, fullerenes, carbon nanoparticles, have been found to cause brain

damage in largemouth bass, a species accepted by regulatory agencies as a model for

defining ecotoxicological effects (Miller & Kinnear, 2007) thus, limiting the use of

nanotechnology extensively.

2.3 SUGAR BEET PECTIN AS WALL MATERIAL

Pectins are a family of plant-derived heteropolysaccharides comprised predominantly of a

linear chain of (1→4)-linked α-D-galacturonic acid residues. These residues may be

partially esterified with a small percentage of rhamnose units to yield branches consisting

of neutral sugars, notably galactose and arabinose (Cho, 2001). Unlike citrus pectins, beet

pectins have ferulic acid groups esterified to some of the neutral sugars in the side-chains

of the so-called “hairy” regions (Guillon and Thibault, 1990; Micard and Thibault, 1999;

Saulnier and Thibault, 1999). Apples, guavas, quince, plums, gooseberries, oranges and

other citrus fruits, contain large amounts of pectin, while soft fruits like cherries, grapes

and strawberries contain small amounts of pectin (Cho, 2001).

The amount, structure and chemical composition of pectin differs between plants, within a

plant over time and in different parts of a plant (Cho, 2001). Pectins are generally

classified as high methyl ester (HM) pectins and low methyl ester (LM) pectins based on

22

the percentage of ester groups they possess or the degree of esterification (DE). If the DE

of the pectin is greater than 50% it is called as HM pectin, while if it is less than 50% it is

called a LM pectin (IPPA International Pectin Producers Association, 2001).

Pectins are produced commercially as a white to light brown powder, mainly extracted

from citrus fruits, and is used in food as a gelling agent particularly in jams and jellies. It is

also used in fillings, sweets, as a stabilizer in fruit juices and milk drinks and as a source of

dietary fiber (‘Pectin’, 2010). In contrast to other pectins, sugar beet pectin (SBP) is more

interesting because of its unusual emulsifying properties. The emulsifying properties of

SBP have been attributed to its high protein content (up to 11%) and high level of

acetylation (up to 5%) (Leroux et al., 2003; Funami et al., 2007). SBP also contains the

known antioxidant, ferulic acid, which occurs esterified to the oxygen on C-2 of arabinose

or C-6 of galactose (Fry, 1983) (Figure 2). However, Drusch et al., (2007) and Katsuda

et al., (2008) showed that commercial SBP contains significant amounts of transition metal

ions, namely copper and iron, which promote lipid oxidation. Hence, antioxidants were

used in this research study to improve the oxidative stability of microencapsulated fish oil

with SBP as the wall material.

Figure 2 Schematic structure for generalised sugar beet pectin. Adapted from Morris et al.,

(2010) with some modifications made.

Arabinose Galactose

Methyl groups Rhamnose

Galacturonic acid

23

CHAPTER 3 - MATERIALS AND METHODS

Fish oil (FO) (Hi-DHA 25N, EPA = 5.3%, DHA =26%, Omega-3 fatty acids =35%, PV =

1.4 meq O2/kg) was obtained from Nu-Mega Ingredients, Melbourne, Australia) and

extra-virgin olive oil (EVOO) (PV= < 20 meq O2/kg) from Boundary Bend Marketing Pty

Ltd, Lara, Australia. Sugar beet pectin (SBP, GENU® pectin type BETA, of ≥60 kDa and

degree of acetylation of 23.8% according to manufacturer’s specifications) was kindly

provided by CPKelco, Melbourne, Australia. Dried glucose syrup (DGS) (with dextrose

equivalent of 26−30 according to manufacturer’s specifications) was obtained from

Manildra, NSW, Australia. FO and EVOO were stored under nitrogen in dark bottles at

4°C (up to 6 month) or at -20°C (6-24 month).

3.1. RAW MATERIAL COMPOSITION

3.1.1 SUGAR BEET PECTIN

3.1.1.1PROTEIN CONTENT

The protein content of SBP was determined in triplicate using LECO (FP-2000 LECO

Corp., Michigan, USA) method, and the protein content was calculated using 6.25 as a

conversion factor.

3.1.1.2 IRON AND COPPER CONTENT

The iron and copper content in SBP was analyzed at DTS Laboratories, Melbourne.

3.1.2 FISH OIL AND EXTRA-VIRGIN OLIVE OIL

3.1.2.1 FATTY ACID COMPOSITION

24

The fatty acid composition of the oils prior to encapsulation was determined according to

Christie (2003) with some modification (Shen, 2010). The original and extracted oils were

derivatized using transmethylesterification and subsequently analyzed using a Varian 3400

gas chromatograph (Varian Associates Inc., USA) equipped with a BPX70 column (30 m × 0.25 mm i.d., 0.25 µm film thickness).

3.1.2.2 TOTAL PHENOLICS CONTENT IN EXTRA VIRGIN OLIVE OIL

The total phenolics content was determined using the method of Hrncirik and Fritsche

(2004). Extra virgin olive oil (2.5 g) was dissolved in 5 ml hexane (EMD Chemicals, USA)

and the phenolics were extracted with 5 ml methanol (Merck Chemicals, Germany)/water

(60:40 vol/vol) for 2 min under nitrogen atmosphere using an automatic shaker. The

mixture was then centrifuged (3500 rpm,10min) for phase separation and an aliquot (0.2

ml) of the methanolic phase was diluted with water to a total volume of 5 ml, followed by

addition of 0.5 ml Folin-Ciocalteu reagent (Sigma Chemicals Co., Australia). After 3 min,

1 ml sodium carbonate solution (35%, wt/vol) was added to the reagent mixture, which

was finally mixed and diluted with water to 10 ml. The absorbance of the solution was

measured after 2 hr against a blank sample using a UV-vis spectrophotometer (UV-1700,

Shimadzu, Suzhou manufacturing co. ltd., China) at a wavelength of 725 nm. The

calibration curve was constructed using standard solutions of caffeic acid (Sigma

Chemicals Co., Australia) within the range of 0.05–0.5 mg/ml.

3.1.3 OXIDATION STATUS OF FISH OIL AND EXTRA VIRGIN OLIVE OIL

3.1.3.1 ASSESSMENT OF OXIDATIVE STABILITY USING OXIPRESS

The oxidative stability of microcapsules containing 4g oil was assessed under oxygen

pressure (0.5 MPa) at 80°C in an ML Oxipres® apparatus (DK-8270, Mikrolab Aarhus

A/S, Højbjerg, Denmark) equiped with Paralog software. The induction period (IP) was

defined as the time (h) required for the oxidation process to be initiated. The IP was

obtained directly from the Oxipres curve and represented the starting point of pressure

25

decline. The slope (-mbar hour-1) indicates the rate at which the sample is oxidized after

the initiation of oxidation. The slope was calculated by drawing a line at the IP point

parallel to the part of the curve recording the change in pressure (Figure 3).

.

FIGURE 3 Illustration of oxidation stability measurements using ML OXIPRES as

reported by Trojakova, Reblova & Pokorny, 2001).

P- Oxygen pressure (MPa); t- Reaction time (h); t1- Tangent drawn on the curve

corresponding to the induction period; t2- Tangent drawn on the curve corresponding to the

change in oxygen pressure; p- Cross-section of the two tangents; x- end of induction

period.

3.2 PREPARATION OF EMULSIONS

Encapsulant materials were primarily selected from a range of biopolymers of various

sources such as carbohydrates( Maltodextrins, Dried Glucose syrups, Alginates, Chitosan,

Starches etc), proteins (Sodium Caseinate, Whey Proteins, Gelatins etc),

lipids (Hydrogenated fats, Vegetable oils, Palm Stearin etc) and waxes (Shellac or

Carnauba Wax etc) (Smith & Charter, 2010). In our study, DGS was used as a bulking

agent along with SBP to improve stability and microencapsulation of fish oil. Sugars in

association with emulsifying bioploymers have been shown to improve encapsulation

properties. Drusch (2007) successfully produced stable fish oil microcapsules using sugar

P t1

t

Pressure (MPa)

26

beet pectin as a wall material and glucose syrup as a bulk one. The same author reported

also that a pectin content of 1- 2% was sufficient for the formation of stable emulsions for

spray drying and proposed that the maximum oil loading in microcapsules containing FO

as the core may be limited to 50 % because of the amount of non-encapsulated fat obtained

in them. Hence, formulations were developed with 2% SBP along with DGS in the

preparation of FO and FO-EVOO emulsions with 25% and 50% oil loading at 30% TS.

SBP (2 g) was dispersed in distilled water at 70°C and held at that temperature for a further

15 min before the addition of DGS. For samples with EDTA in the formulation, SBP (2 g)

was dispersed in 85 % of the total distilled water required at 70°C and EDTA was added to

chelate the metal ions before the addition of DGS. The solution was subsequently cooled to

60°C. The pH was then adjusted by adding 1M NaOH solution to samples whose pH had

to be altered to 6. The amount of NaOH solution was recorded to determine the remaining

water to be added to the pre-heated solution. The solution was subsequently cooled to

60°C. FO and EVOO were pre-heated to 60°C. A FO-EVOO blend was prepared by

combining the oils at a 1:1 weight ratio. The FO and the FO-EVOO blend were dispersed

into the pre-heated mixtures of SBP and DGS at 60°C for 1-2 min using a high shear mixer

(Silverson, London, UK) at maximum speed to obtain a pre-emulsion. The pre-emulsions

were then homogenized at 60°C using a high pressure two-stage homogenizer (APV

Rannie AS Homogenizer, Denmark) at 35 and 10 MPa. All emulsions were prepared at

30% total solids. The composition of the pre-emulsion with the percentage of each

component stated in terms of ingredient weight is outlined in Table 2 & 3. The process

flow chart of the experimental plan for the production of spray dried powders has been

presented in Figure 4. FO and FO-EVOO emulsions were prepared at 25% (pH 3) and 50%

(pH 3 & pH 6 ) oil loading in powder and emulsions with EDTA were prepared at 50% oil

loading in powder only at pH 6 yielding a total of 8 different emulsions (FO-25,

FO-EVOO-25, FO-50 (pH 3), FO-EVOO-50 (pH 3), FO-EDTA-50,FO-EVOO-EDTA-50,

FO-50 (pH 6), FO-EVOO-50 (pH 6)) (Figure 5 (A)). All emulsions were prepared in two

runs with duplicate samples in each run.

27

Dispersion of SBP (2%) in distilled water/ buffer solution (pH adjusted to 6) at 70°C

SBP-DGS mixture cooled to 60°C

SBP-DGS-Oil mixture

Pre-emulsion

Homogenization (35+10 MPa)

Stable Oil Emulsion (30 % total solids)

Spray Drying

Powder Microcapsules

Figure 4 Process flow chart of the experimental plan for the production of spray-dried

microcapsules.

Addition of EDTA then DGS

Adjusted to pH 6 & Addition of FO/FO –EVOO pre-heated to 60°C

28

Table 2 Formulation of the fish oil and fish oil-extra-virgin olive oil (1:1 wt ratio)

emulsions at pH 3.

Percentage contents in each emulsion (Ingredient weight)

Constituent

FO-25

FO-50

FO-EVOO-25

FO-EVOO-50

SBP 2 2 2 2

DGS 20.5 13 20.5 13

FO 7.5 15 3.75 7.5

EVOO 0 0 3.75 7.5

Water 70 70 70 70

Note: Moisture content of SBP: 9.92 %, DGS: 6.99 %

Table 3 Formulation of the fish oil and fish oil-extra-virgin olive oil (1:1 wt ratio)

emulsions with and without EDTA at pH 6.

Percentage contents in each emulsion (Ingredient weight)

Constituent FO-EDTA-50 FO-EVOO-EDTA-50 FO-50 FO-EVOO-50

SBP 2 2 2 2

EDTA(Na

salt)

0.05 0.05 0 0

DGS 12.95 12.95 13 13

FO 15 7.5 15 7.5

EVOO 0 7.5 0 7.5

Water 70 70 70 70

Note: Moisture content of SBP: 9.92 %, DGS: 6.99 %

29

(A) Emulsions prior to spray drying (B) Spray-dried powders

Figure 5 Emulsions (A) and spray-dried powders (B)

3.3 SPRAY-DRIED MICROCAPSULES

The homogenized emulsions were converted to spray-dried microcapsules/powders (Figure

4 (B)) using a Drytec laboratory spray dryer (Tonbridge, UK). The dryer had a water

evaporation rate of 1 L h-1 with a twin fluid nozzle at 2.5 bar atomizing pressure. The feed

was heated to 60°C before atomization. The inlet and the outlet air temperatures were

180°C and 80°C, respectively. Two independent processing runs were carried out. The

process flow chart for the production of microcapsules has been illustrated in Figure 4.

3.3.1 STORAGE CONDITIONS

Aliquots (20 g) of each of the microcapsule formulations were stored in duplicate in

transparent, stoppered, oxygen-permeable 100 ml plastic containers at room temperature

(~25°C) for 0, 1, 2 and 3 months. Zero month refers to samples analysed immediately after

the production of the microcapsules without storage. At the end of each storage time the

plastic containers were covered with aluminium foil and transferred to frozen storage

(-18°C) prior to further analysis.

30

3.4 EMULSION CHARATERISTICS

3.4.1 PARTICLE SIZE

The particle size distribution of the emulsions was determined by laser light scattering

(Mastersizer 2000G, Malvern Instruments, Worcestershire, UK) using standard optical

parameters, and a refractive index of 1.456. The emulsion was dispersed in recirculating

water within the measuring cell (Hydro SM, Worcestershire, UK) until an obscuration rate

of 10-20% was reached. The particle size measurements were reported as the

surface-volume mean diameter, d3,2.

3.4.2 LIGHT MICROSCOPY

The emulsions were visualized by light microscopy (Olympus BH-2, Anax Pty. Ltd, Japan)

equipped with an attached camera. A drop of Oil-Red-Oil dye (BDH laboratory supplies,

England) was added to each emulsion (~1 ml) and visualized using a 10x objective

magnification. The images were processed using image acquisition software (anaLYSIS

getIT).

3.4.3 VISCOSITY

The viscosity of the emulsions was measured using a cup and bob geometry (Paar Physica

Rheometer, MCR 300, Anton Paar, Austria) at 25°C within 2 h of preparation. The shear

rate (γ) was increased in 30 steps from 0.2 to 291 s−1. The duration of measurement at each

shear rate was 10 s.

3.4.4 OXIDATIVE STABILITY OF EMULSIONS UNDER ACCELERATED

CONDITIONS

The oxidative stability of the emulsions (equivalent to 4 g oil in sample) was assessed by

following the procedure described in Section 3.1.3.1.

31

3.4.5 ZETA-POTENTIAL OF EMULSIONS

Measurement of zeta potential of FO and FO-EVOO emulsions as a function of pH was

performed at 22ºC with a Zetasizer Nanoseries (Nano ZS), Malvern Instruments,

Worcestershire, United Kindom). Titrations were performed by

Autotitrator-MPT-2(Malvern Instruments, Worcestershire, United Kindom). Samples were

prepared by diluting (7µl emulsion) in buffer solution (20 ml) (the pH of distilled water

was adjusted to 6 by the addition of NaOH and HCl). Diluted emulsions (15ml) were then

filled in sample tube with a magnetic stirrer. A liquid filled pH probe was then inserted

into the sample tube. Three different titrants (0.5M HCL, 0.2 M HCL and 0.5M NaOH)

were connected simultaneously to the sample tube. The system automatically selected the

appropriate concentrations of the titrant to be mixed into the sample to adjust to the

required pH. The diluted emulsion was pumped into a capillary cell by the integral

peristaltic pump and the cell was then inserted into the measurement chamber of Zetasizer.

The program was set to measure up to 100 points between the pH range 6 – 1 and to titrate

with either acid or base.

3.5 CHARACTERIZATION OF THE SPRAY-DRIED MICROCAPSULES

3.5.1 MOISTURE CONTENT

The moisture content of the spray-dried powders was determined at 80°C using a Sartorius

infrared balance (MA30, Sartorius Mechatronics, Gottingen, Germany).

3.5.2 WATER ACTIVITY (AW)

The aw of the spray-dried powders was determined at 25°C using a Aqua-Lab Water

Activity Meter (Series 3, Decagon Devices Inc, USA).

3.5.3 PARTICLE SIZE OF THE RECONSTITUTED POWDERS

32

The spray-dried powders (10% w/v) were reconstituted at 70°C for 3 h under constant

stirring, and then rested for 1 h at room temperature (~25°C). Particle size was determined

as described above (Section 3.4.1).

3.5.4 FATTY ACID COMPOSITION

Fatty acid composition of the oils extracted from the spray-dried microcapsules was

determined as described in Section 3.1.2.1.

3.5.5 DETERMINATION OF FREE OIL CONTENTS

The free (solvent-extractable) oil content of each spray-dried powder was determined

gravimetrically according to Pisecky (1997), except that petroleum ether was used in place

of carbon tetrachloride. In this method, petroleum ether (50 ml) was added to 10 g powder.

The mixture was agitated in a stoppered tube for 15 min. The mixture was filtered

(Whatman No.541) and the solvent evaporated at 60°C using a rotary evaporator. The

remaining fat residue was then dried in an oven at 105°C for 1 h.

3.5.6 DETERMINATION OF TOTAL OIL CONTENT

The total oil content of the spray-dried microcapsules was determined gravimetrically

according to the Schmid-Bondzyndki-Ratzlaff method (1988) (AS 2300.1.3).

3.5.7 ENCAPSULATION EFFICIENCY

Encapsulation efficiency was calculated after estimating the free and total fat contents in

spray-dried microcapsules using the following formula:

Encapsulation efficiency (%) = 100- (% solvent-extractable fat / % total fat) × 100

[equation1]

33

3.5.8 PROPANAL AND HEXANAL ANALYSIS

Aliquots (2 g) of the spray-dried powders were placed in 10 ml headspace vials, sealed and

equilibrated at 60ºC for 15 min in a water bath. An aliquot (1 ml) of the headspace was

then analyzed using a Varian 3400 gas chromatograph (Varian Associates,Inc., USA)

equipped with a BPX-70 fused silica capillary column (30 m × 0.25 mm i.d., 0.25 µm

film thickness) and a FID (flame ionisation detector). The injector and detector

temperatures were 250 º C and 275 º C, respectively. The oven temperature was

programmed at 60ºC and held for 5 minutes. The temperature was then increased to 175ºC

at 10ºC min-1. The temperature was then further increased to 220 ºC at 4ºC min-1.

Propanal and hexanal were identified using external standards.

3.5.9 OXIDATIVE STABILITY OF THE SPRAY-DRIED MICROCAPSULES

UNDER ACCELERATED STORAGE CONDITIONS

The oxidative stability of the spray-dried microcapsules (equivalent to 4 g oil in sample)

was assessed was assessed by following the procedure described under Section 3.1.3.1.

3.5.10 SCANNING ELECTRON MICROSCOPY (SEM)

The morphology of the spray-dried microcapsules was examined using the SEM (Philips

XL30FEG scanning electron microscope, Eindhoven, Netherlands) in the departments of

Zoology and Biology, The University of Melbourne, Australia.

3.6 STATISTICS

All analytical determinations were carried out at least in duplicate. The results were

reported as the mean ± standard deviation (SD) of these measurements. The analysis of

variance (One way ANOVA) was performed at 95% confidence level using SPSS 18

(SPSS Inc, Chicago, USA) and LSD (least significance) was used to separate the means.

34

CHAPTER 4 - RESULTS AND DISCUSSION

4.1 PROPERTIES OF RAW MATERIALS

4.1.1 SUGAR BEET PECTIN

The protein content in SBP was 4.9 ± 0.05 (%). This value is similar to that previously

reported by Drusch (2007), and Siew & Williams (2008). However, other reported values

for protein content of SBP ranged from as low as 1.95 % (Leroux, 2003) to as high as ~

10.4 % (Thibault, 1988). This large variation in protein content reflects the sensitivity of

the protein moiety in SBP to extraction and purification processes (Kirby, Dougall &

Morris, 2006).

The iron and copper contents in the SBP were determined to be 310 ppm and 10 ppm,

respectively. This corresponds to a total iron and copper content in the emulsions of

0.000114 moles/L (i.e. 6.4 ppm, comprising 6.2 ppm Fe and 0.2 ppm Cu). Katsuda et al.,

(2008) cited that to maximize the oxidative stability of commercial oils, iron and copper

contents in oil should be as low as possible, with most specifications recommending <0.1

ppm and 0.02 ppm, respectively. The high levels of iron and copper in the SBP used in this

study are likely to promote lipid oxidation, particularly if they are not complexed by

chelating agents. To investigate the effect of the metal ions on lipid oxidation, the

emulsions (at pH 6) were prepared in the presence and absence of the metal chelator,

EDTA. EDTA was added to the emulsions at a level of 0.05% w/v (i.e. 0.000171 moles

EDTA/L emulsion). This equated to an excess of EDTA to metal ions, and thus ensured

that all the metal ions were present in the emulsion as EDTA-metal ion complexes,

preventing their interaction with lipid hydroperoxides.

4.1.2 FISH OIL AND EXTRA VIRGIN OLIVE OIL

4.1.2.1 FATTY ACID COMPOSITION

35

Results of fatty acid composition of FO and EVOO was quantified using GC are shown in

Table 4. The major fatty acids found in FO were palmitic acid (C16:0) (21%), oleic acid

(C18:1) (~14%), eicosapentaenoic acid, EPA (C20:5n-3) (6.6%), and docosahexaenoic

acid, DHA (C22:6n-3) (26.5%).The fatty acid composition of FO is highly dependent on

the age, sex, spawning cycle, season, species and origin of the fish (Donmez, 2009). In this

study, the FO was derived from tuna. The EPA and DHA content of the FO were found to

be very close to that of the manufacturer’s specification. Gotoh et al., (2006) quantified

fatty acids present in big eye tuna oil, and the reported EPA and DHA percentages were

6.5% and 24% respectively.

Results of fatty acids analysis in EVOO were in agreement with those reported in the

literature. The major fatty acid in EVOO is oleic acid, which can vary from 55−83%. It

was present at a level of 80% in the EVOO used in this work. The other major fatty acid

present is palmitic acid and normally its content ranges between 7.5−20% (‘Olive oil’,

2008). It was 7% in the EVOO used in this work.

Table 4 Fatty acid composition (%) of the initial oils

Fatty Acid

a

Fatty acid (%)

Fish Oil

Extra Virgin

Olive Oil

Palmitic acid, C16:0 21.09 ± 1.94 6.99 ± 0.13

Oleic acid, C18:1 13.51 ± 1.79 79.54 ± 0.12

Eicosapentaenoic acid, C20:5n-3 6.64 ± 0.92 ND

Docosahexaenoic acid, C22:6n-3 26.54 ± 0.53 ND

Others 32.21 ± 4.12 13.47 ± 0.01 aValues are average of duplicate measurements ± SD. ND = not detected

36

4.1.2.2 TOTAL PHENOLICS CONTENT IN EXTRA VIRGIN OLIVE OIL

The total phenolics content in EVOO was 98.11 ± 0.86 (ppm). This value falls in the

expected range specified by the manufacturer of 70−120 ppm).The phenolic content in

EVOO varies largely because of a number of factors like the olive variety, time of harvest,

environmental factors, storage conditions, and refining process, for example (‘Olive oil

source’, 2010). EVOO with a phenolic content as high as 1500 ppm has been reported

(Oliveras-López et al., 2008).

4.1.3 OXIDATION STATUS OF FISH OIL AND EXTRA VIRGIN OLIVE OIL

4.1.3.1 ASSESSMENT OF OXIDATIVE STABILITY USING THE OXIPRES®

The Oxipres® results of the original FO, EVOO and their 1:1 blend are given in Table 5.

The IP for these oils under the accelerated conditions (80ºC, 0.5 bar oxygen pressure) used

in Oxipres® assessments were 11.85 ± 0.07 h, >100 h and 16.3 ± 0.14 h, respectively, for

FO, EVOO and the 1:1 mixture of FO-EVOO. The uptake of oxygen after the IP was -210

± 9.19 mbar h-1 and -119 ± 4.24 mbar h

-1, respectively, for FO and a 1:1 mixture of

FO-EVOO. EVOO was monitored for up to 105 h, during which time no detectable

oxidation was observed. Hence no oxidation rate (slope) for this sample was provided.

This confirmed that olive oil was more stable to oxidation than FO. That might be

expected based on the fatty acid composition alone as the omega-3 fatty acids in FO are

very prone to oxidation. In addition, olive oil contains phenolic compounds, which are

known to be antioxidative (Psomiadou & Tsimidou, 2002).

37

Table 5 Oxidative stability of original oils during accelerated storage (80°C, oxygen

pressure of 0.5 bar)

aSample Induction period (h) Slope (mbar h

-1)

FO 11.85 ± 0.07 -210 ± 9.19

EVOO >100 ± 0.00 −

FO-EVOO (1:1 wt ratio) 16.3 ± 0.14 -119 ± 4.24

aValues are average of duplicate measurements ± SD

4.2 EFFECT OF PARTIAL SUBSTITUTION OF FISH OIL WITH EXTRA

VIRGIN OLIVE OIL ON EMULSION AND MICROCAPSULE

CHARACTERISTICS

The average particle size (d3,2) of the different emulsions ranged between 0.41–0.43 µm

(Figure 6). These values were comparable to those reported for oil-in-water emulsions

made by emulsifying 20% w/w orange oil with 2% SBP (Leroux et al., 2003) and

homogenised with three passes at 200 bars. SBP dissolved in water alone had a (d3,2) of

2.38 ± 0.31 µm, and contributed to the particle size of the emulsions. This observation also

accounts for the minor peak present in the particle size distribution graph of the emulsions.

There was no significant difference (P > 0.05) in the average particle sizes between the

emulsions containing 25% and 50% oil loading. This indicated that the presence of 2%

SBP was sufficient emulsifier to produce physically stable fish oil-in-water emulsions

containing 15% oil (wet basis). Similarly, other studies (Drusch, 2007; Drusch et al., 2007;

Leroux et al., 2003; Nakauma et al., 2008; Siew & Williams, 2008) have reported that

1.5−2% SBP was sufficient to produce oil-in-water emulsions containing up to 18% oil

(wet basis). For example, Nakauma et al., (2008) made stable oil-in-water emulsions with

1.5% SBP and 15% w/w oil (medium chain triglyceride) homogenised with two passes at

50 MPa.

38

Solubility is an important criterion in choosing a wall material. If the wall material is

highly insoluble it may precipitate. When this occurs the wall material (pectin) is not able

to move to the oil/water interface and the emulsifying power of the pectin will be reduced.

The particle size distribution of the emulsions revealed that SBP was fully dispersed and

that there were no large aggregates of undissolved SBP (Figure 6). Visual examination of

the emulsions by light microscopy also showed the emulsions were homogeneous and no

visual particle aggregation or ‘free’ (nonencapsulated) oil was present (Figure 7).

0

1

2

3

4

5

6

7

0.01 0.1 1 10 100 1000

Particle size (µm)

Volu

me

(%)

Figure 6 Particle size distributions of the original (fresh) emulsions, prior to spray drying.

Values are the averages of triplicate measurements ± SD; � (FO-50); � (FO-25); �

(FO-EVOO-50); � (FO-EVOO-25).

Emulsion stability is an important property to be considered in the microencapsulation

process. Emulsion stability arises from steric repulsion of aggregates by a hydrated layer

(Willats, Knox & Mikkelsenc, 2006). Pectin stabilises an emulsion by creating a layer

around the particles at the interface. It has also been reported that stable emulsions have

been formed when using low molecular weight pectins (70 kDa) (Akhtar et al., 2002) and

also because of the link between charged molecules (pectin) and protein (Leroux et al.,

2003).

39

Figure 7 Brightfield micrographs of (A) FO-25 (7.5% oil, wet basis) and (B)

FO-EVOO-50 (15% oil, wet basis) emulsions at pH 3.

The apparent viscosities of the emulsions as a function of shear rate are shown in Figure 8.

All the formulations displayed shear thinning or pseudo-elastic behaviour, which is a

common feature of oil-in-water emulsions (McClements, 1999).

Emulsions containing 15% oil (wet basis) had greater apparent viscosities than

corresponding emulsions containing 7% oil, as expected. However, emulsions containing

either FO or a 1:1 FO-EVOO mixture with the same gross formulation had different

viscosities. It is possible that the factor responsible for these differences was the presence

of phenolic compounds in the olive oil. For example, polyphenols are known to complex

with protein (Spencer et al., 1988). An interaction between polyphenols and residual

protein in SBP can alter the interfacial properties of the oil droplet or the properties of the

bulk phase and also affect viscosity of the emulsion. Other studies showed that

polyphenol-β-casein complexes alter the properties of an air/water interface and also exist

as complexes in the bulk solution (Aguie-Beghin et al., 2008).

40

0

200

400

600

800

1000

1200

1400

1600

1800

0 50 100 150 200 250 300

Shear Rate (1/s)

Vis

cosi

ty (

mP

a/s

)

Figure 8 Apparent viscosities of fish oil and fish oil-extra virgin olive oil (1:1 wt ratio)

emulsions (30% TS; 2% SBP, 20.5% DGS, 7.5% total oil and 30% TS; 2% SBP, 13%

DGS, 15% total oil), intended for manufacture of spray-dried microcapsules containing 25

and 50% oil, as a function of shear rate. Values are the average of duplicate measurements

± SD; � (SBP-DGS-FO-50); � (SBP-DGS-FO-25); � (SBP-DGS-FO-EVOO-50); �

(SBP-DGS-FO-EVOO-25).

Zeta potential measurements of the FO and FO-EVOO emulsions (2% SBP, 13% DGS,

15% total oil) intended for the manufacture of spray-dried microcapsules with 50% oil

loading, as a function of pH, were determined using an autotitrator to predict the stability

of the emulsions (Table 6). The emulsions were measured at pH values between 6 and 1.

The zeta potential of a particle can be defined as the overall charge that the particle

acquires in a particular medium. Depending on the zeta potential of the particle, the

stability of an emulsion can be explained. If the particles in a suspension have a large

negative or positive zeta potential then they will tend to repel each other and resist the

formation of aggregates. Conversely, particles of opposite charge tend to associate. The

dividing line between stable and unstable suspensions is generally taken at either +30 mV

41

or -30 mV. Particles with zeta potentials more positive than +30 mV or more negative than

-30 mV are normally considered stable (Zeta Potential Measurement Using an Autotitrator

from Malvern Instruments and the Effect Of pH, 2005). The autotitration plots revealed

that both FO and FO-EVOO emulsions were most stable at pH greater than 5 (Figure 9 &

10). Both emulsions would be less stable as pH is reduces below 5, and would be least

stable at pH ~1.5 as the iso-electric point range for the emulsions was between 1.2 and 1.5.

It has been observed that at pH less than 5 the zeta potential of the emulsions decreases

rapidly and eventually reached an iso-electric point after pH 1.5.

Table 6 Zeta potential values of FO and FO-EVOO emulsions measured at 22º C.

aSample Zeta potential (mV)

FO-50 (pH 3) -12.12 ± 0.39

FO-50 (pH 6) -39.8 ± 0.51

FO-EVOO-50 (pH 3) -13.56 ± 0.47

FO-EVOO-50 (pH 6) -41.4 ± 0.66 aValues are average of 5 measurements ± SD

After manufacture, the moisture content of the spray-dried microcapsules made with SBP

and containing 25% and 50% oil loadings ranged from 1.6–1.8%, with a water activity of

~0.27 (Table 7).The maximum moisture specification for most spray-dried powders in the

food industry was between 3–4 % (Masters, 1991). The encapsulation efficiency of the

spray-dried microcapsules made with SBP containing 25% and 50% of FO and FO-EVOO

blend was >90%. A significant difference (P < 0.05) in the encapsulation efficiencies of

the microcapsule powders containing different oil loadings was recorded (Table 7). The

microcapsule powders with 50% oil loading had lower encapsulation efficiency (~90%)

than those with 25% oil loading (encapsulation efficiency ~98%). In concurrence with the

solvent-extractable fat contents, encapsulation efficiency was not affected by the

composition of the microencapsulated oil.

42

-50

-40

-30

-20

-10

0

10

0 1 2 3 4 5 6 7

pH

Ze

ta P

ote

nti

al

(mV

)

Figure 9 Autotitration curve for FO emulsion with zeta potential measured as a function of

pH at 22 ºC.

-45

-40

-35

-30

-25

-20

-15

-10

-5

0

5

0 1 2 3 4 5 6 7

pH

Ze

ta P

ote

nti

al

(mV

)

Figure 10 Autotitration curve for FO-EVOO emulsion with zeta potential measured as a

function of pH at 22 ºC.

43

Table 7 Properties of spray-dried powders containing microencapsulated fish oil and fish

oil-extra virgin olive oil (1:1 wt ratio) (25% and 50% oil loading) at pH 3.

Property

aMicrocapsule

FO-25 FO-EVOO-25 FO-50 FO-EVOO-50

Moisture content 1.83 ± 0.02 1.73 ± 0.01 1.66 ± 0.01 1.62 ± 0.03

Water activity (aw) 0.27 ± 0.02 0.26 ± 0.01 0.27 ± 0.02 0.27 ± 0.00

Total oil (%) 24.39 ±0.06 24.38 ± 0.06 49.09 ±0.14 49.22 ± 0.12

Solvent-extractable oil (% of free oil in powder)

0.53 ± 0.01 0.52 ± 0.00 4.99 ± 0.01 5.00 ± 0.01

Encapsulation efficiency (%)

97.85 ± 0.04a 97.87 ± 0.04a 90.43 ±0.09b 90.42 ± 0.08b

Mean particle diameter, d32, prior to spray drying

0.42 ± 0.00 0.43 ± 0.00 0.41 ± 0.00 0.43 ± 0.00

aValues are averages of triplicate analysis ± SD of 2 individual runs

Means within the row followed by different superscript letters differ significantly at P = 0.05

The amount of solvent-extractable fat in the spray-dried powders with the same oil loading

but different oil composition was comparable (Table 7). The FO and FO-EVOO powders

gave ~2% solvent-extractable fat as a percentage of total fat in powder at 25% oil loading,

and ~10% solvent-extractable fat at 50% oil loading. The amount of free oil in the powders

can generally be described as the amount of oil that can be found on the surface of the

powder particles and can include oil that is within cracks in the powder particle that is

readily accessible to solvent (Buma, 1971). Several studies had been conducted to evaluate

the factors affecting the oxidative stability of spray-dried powders containing encapsulated

oil (Velasco, Dobarganes & Márquez-Ruiz, 2003; Drusch et al., 2007; Márquez-Ruiz,

Velasco & Dobarganes, 2003). However, due to the different matrix and core materials

used, and the various methods and conditions employed to monitor oil oxidation, no

consensus has yet been reached. Velasco, Dobarganes and Marquez- Ruiz (2003) stated

that lipid distribution in a microcapsule is an important factor influencing oxidation in

encapsulated oil. They discussed various studies conducted to evaluate the oxidation of

44

microencapsulated oils during storage by separate extractions of surface and encapsulated

oil fractions. They demonstrated that it could not be hypothesised that surface oil oxidation

occurs at a faster rate than encapsulated oil even though it is more exposed to oxygen and

had no protection by the matrix. Interestingly, Drusch & Berg (2008) investigated the

oxidative storage stability of microencapsulated FO prepared under various spray drying

conditions and concluded that the surface oil protects other solvent-extractable oil fractions

(e.g. fat within capillary pores of the microcapsules) against oxidation. Furthermore, these

authors revealed that solvent-extractable oil cannot be used to predict shelf-life of

microencapsulated oils.

The moisture content and aw of the spray-dried microcapsules containing 25% and 50% oil

loadings after 3 months storage at ambient conditions were measured (Table 8). A general

increase in moisture content and aw, approximating that of the external environment (aw

~0.5), was observed with increased storage duration. The highest percentage increase in

moisture content was recorded for microcapsules with 50% oil loading.

The particle size distributions and average particle size values (d3,2) of the reconstituted

spray-dried microcapsules at 0, 1, 2 and 3 month storage under room temperature

conditions were determined (Table 9, Figure 11). The particle size distribution for the

fresh powders reconstituted before storage showed that they had particle sizes similar to

those of the respective emulsions prior to spray drying. However, there was a general

increment in the particle size distribution of the reconstituted microcapsules during storage

at room temperature. The data revealed a positive relationship between the rate of

increment in particle size and storage time. Additionally, microcapsules containing 25% oil

loading generally had smaller particle sizes than those containing 50% oil loading under

same storage conditions.

45

Table 8 Changes in moisture content and water activity in FO and FO-EVOO

microcapsules (25% and 50% oil loadings) over storage time.

aMicrocapsule

Moisture content (%) Water activity (aw)

0 month 3 month 0 month 3 month

FO-25 1.83 ± 0.02 2.18 ± 0.05 0.27 ± 0.02 0.49 ± 0.00

FO-50 1.66 ± 0.01 2.06 ± 0.03 0.27 ± 0.02 0.50 ± 0.01

FO-EVOO-25 1.73 ± 0.01 1.95 ± 0.01 0.26 ± 0.01 0.47 ± 0.01

FO-EVOO-50 1.62 ± 0.03 1.93 ± 0.02 0.27 ± 0.00 0.48 ± 0.00 aValues are averages of duplicate analysis ± SD of 2 individual runs

Table 9 Particle size of the reconstituted spray-dried microcapsules containing fish oil

and fish oil-extra virgin olive oil (1:1 wt ratio) during storage of the powders in stoppered

flasks at room temperature (~25ºC).

Microcapsule

Mean particle diameter, d3,2 (µm) a


FO-25 0.50 ± 0.01a 0.50 ± 0.00b 0.92 ± 0.00a 2.91 ± 0.10b

FO-50 0.70 ± 0.00 c 0.78 ± 0.00c 2.06 ± 0.08c 3.06 ± 0.05c

FO-EVOO-25 0.50 ±0.00 a 0.50 ± 0.00a 0.90 ± 0.01a 1.62 ± 0.01a

FO-EVOO-50 0.69 ± 0.00 b 0.79 ± 0.01d 1.69 ± 0.03b 1.69 ± 0.02a aValues are averages of triplicate analysis ± SD of 2 individual runs


46

0

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I II

III IV

A B

C D

Figure 11 Particle size distributions of reconstituted spray-dried microcapsules at 0, 1, 2

and 3 month (A–D) stored under room temperature conditions (~25ºC, exposed to light).

Values are the averages of triplicate measurements ± SD; � (FO-50); � (FO-25); �

(FO-EVOO-50); � (FO-EVOO-25).

Visual inspection of the microcapsule powders revealed that the samples stored for 2 and 3

months contained clumps that were not evident in those of the fresh microcapsule powders

(0 month), or those stored for 1 month. These observations suggested the development of

cohesive interactions between particles over the period of storage time. A number of

variables are known to contribute to the time consolidation effects on food powders.

Storage temperature, exposure of powders to moisture content in air, physical and chemical

changes in the powder sample during storage time, and variations in powder bulk density

have all been found to contribute significantly to the time-consolidation effects of powders

(Teunou, 2000; Fitzpatrick 2004; Onwulata 2005).

47

The scanning electron micrographs of the spray-dried microcapsules revealed a

polydisperse particle size distribution between ~1−25 µm in diameter (Figure 12). The

microcapsules were irregular in shape with wrinkled surfaces. The extent of surface

wrinkling has been previously associated with the concentration of SBP in the feed

emulsion (Drusch, 2007). The wrinkled surface may also be attributed to the mechanical

stresses by uneven drying at different parts of the liquid emulsion droplets at the early

stages of drying (Moreau & Rosenberg, 1993). Considerable agglomeration of the particles

was observed in powders after 3 months of storage, irrespective of the oil loading or oil

composition. Small pores were also noticed on the surfaces of a portion of particularly in

the microcapsule powders that had been stored for 3 months. The large number of pores

evident in the stored microcapsule powders indicates degradation of the matrix on storage

and consequent release of the microencapsulated oil.

A B

C D

A BB

C DD

Figure 12 Scanning electron microscopy images of the fish oil (A, C) and fish oil-extra

virgin olive oil (B, D) spray-dried powders (25% oil loading), prior (A, B) and after 3

month storage (C, D) under ambient conditions (~25°C, 0.5 aw).

48

4.3 EFFECT OF EDTA ON EMULSION AND MICROCAPSULE

CHARACTERISTICS

The average particle size (d3,2) of the FO and FO-EVOO emulsions, with and without

EDTA, ranged between 0.35–0.36 µm. The particle size distributions of the emulsions

revealed a multimodal distribution of particles, with the majority of particles ranging in

size from ~0.2−1 µm in diameter, and a minor proportion ranging from 1−10 µm (Figure

13). As mentioned in (section 4.2), SBP dissolved in water had an average particle size

(d3,2) of ~ 2.5 µm and contributed to the particle size of the emulsions. A slight difference

in mean particle size distributions was observed in FO and FO-EVOO-EDTA when

compared to FO-EVOO and FO-EDTA emulsions. That indicated that the addition of

EDTA as chelating agent had very little influence on the mean particle size of emulsions.

The average particle size (d3,2) of all the emulsions were determined after 1 month of

storage at 22ºC, and ranged between 0.35-0.36 µm. That indicated that all emulsions were

stable to droplet aggregation, as evidenced by the lack of change in the average particle

size values, (d3,2) of the fresh emulsions and corresponding emulsions after 1 month

storage. The absence of droplet aggregation could be attributed to the strong repulsive

forces between droplets which were large enough to overcome the various interactions

(mainly Van der Waals forces and hydrophobic interactions) between droplets. It was also

suggested that as pH was increased from the natural pH of pectin solution (pH 3) to pH 6,

the ratio of protonated and deprotonated carboxyl groups present on pectin molecule would

decrease, thereby increasing the negative charge of the pectin molecules. These changes in

pH values increased repulsion between the emulsion droplets and stabilises the droplets

against aggregation.

Visual examination of the emulsions by brightfield microscopy showed that all emulsions

were homogeneous without any visible particle aggregation or any marked presence of

‘free’ (nonencapsulated) oil (Figure 14). Based on the results and literature findings

discussed in section 4.2, those results again established that 2% SBP was sufficient to

produce fine and stable oil-in-water emulsions containing at least 15% w/w oil.

49

0

1

2

3

4

5

6

7

8

9

10

0.01 0.1 1 10 100 1000

Particle size (µm)

Volu

me

(%)

Figure 13 Particle size distributions of the original (fresh) emulsions (15% oil, wet basis),

with and without EDTA, at pH 6. Values are the averages of triplicate measurements ± SD;

� (FO-EDTA-50); � (FO-EVOO-EDTA-50); � (FO-50); � (FO-EVOO-50).

A BA B

Figure 14 Brightfield micrographs of FO-EDTA (A) and FO-EVOO-EDTA (B) emulsions

with 15% oil, at pH 6.

All the emulsions exhibited shear thinning behaviour with the viscosities decreasing with

increase in shear rate (Figure 15). There were some slight differences in the viscosities

noticed between the emulsions. It was observed that FO and FO-EVOO-EDTA emulsions

had slightly lower viscosities when compared to FO-EVOO and FO-EDTA over the shear

50

rate range measured. The presence of high concentrations of metal ions in SBP (Drusch et

al., 2007; Katsuda et al., 2008) and phenolic compounds in EVOO (Cicerale et al., 2009;

Frankel, 2010) could be the underlying cause for the differences in the flow properties of

FO emulsions containing both EVOO and EDTA when compared to FO emulsions

containing either EVOO or EDTA. Also, the differences in interfacial properties of FO and

FO-EVOO emulsions stabilised with SBP have already been established and discussed in

relation to polyphenol-protein complexation (section 4.2).

0

100

200

300

400

500

600

700

800

900

1000

0 50 100 150 200 250 300

Shear rate (1/s)

Vis

co

sity

(m

Pa

/s)


emulsions (15% oil, wet basis) with and without EDTA at pH 6, as a function of shear rate.

Values are the average of duplicate measurements ± SD; � (FO-EDTA-50); �

(FO-EVOO-EDTA-50); � (FO-50); � (FO-EVOO-50)

The moisture content of the spray-dried powders made with SBP (with and without EDTA)

at pH 6 with 50% oil loadings ranged from 2.83−2.96 %, with an aw of ~0.3 (Table 10).

The moisture content values were typical of that for most spray-dried powders. The

maximum moisture specification for most spray-dried powders in the food industry is

between 3–4 % (Masters 1991). This is in order to achieve microbiological stability. The

moisture content and aw of the spray-dried powders made with SBP (with and without

51

EDTA) at pH 6 with 50% oil loadings was also determined after 3 months of storage

(Table 11). An increase in moisture content and aw was observed in all microcapsules and

the percent increase in the moisture content and aw was found to be highest in

microcapsules which did not contain EDTA. That increment in moisture content was

expected as the sample containers in which the powders were stored were permeable to

moisture and the aw of the atmosphere (~0.5) was greater than that of the freshly

manufactured powders.

The spray-dried powders with and without EDTA at pH 6 and 50% oil loading gave ~10%

solvent-extractable fat of total fat in powder. The effect of solvent-extractable fat on lipid

oxidation has been investigated and it has been concluded that it cannot be used to predict

the shelf-life of microencapsulated oils (Drusch & Berg, 2008; Velasco, Dobarganes &

Márquez-Ruiz, 2003). Furthermore, since the amount of solvent-extractable fat was

considerably small, it was unlikely to correlate with the oxidative stability of the

microencapsulated oils. Also, the spray-dried powders were found to be free-flowing,

indicating that the powders did not have high free fat on the surface.

The encapsulation efficiency of the spray-dried microcapsules with and without EDTA at

pH 6 with 50% oil loading was ~90%. There was no difference in the encapsulation

efficiencies of microcapsules without EDTA and microcapsules with EDTA. Thus, it could

be concluded that neither the addition of EDTA nor the oil composition in the core of the

microcapsule had any effect on either solvent-extractable fat or encapsulation efficiency.

The particle size distribution and d3,2 values of the reconstituted spray-dried microcapsules

at 0−3 month storage under room temperature conditions was determined (Figure 16 &

Table 12). A general increase in particle size was observed in all microcapsules over

storage time at room temperature. The particle size values for FO and FO-EVOO

microcapsules over storage time were similar indicating no influence of type of oil/oils in

the core of the microcapsule on particle size.

52

Table 10 Properties of powders containing microencapsulated fish oil and fish oil-extra

virgin olive oil blends (1:1 wt ratio) with and without EDTA at pH 6, 50% oil (dry basis).

Property

aMicrocapsule

FO-EDTA

-50

FO-EVOO-

EDTA-50

FO-50 FO-EVOO

-50

Moisture content

(%)

2.83 ± 0.01 2.89 ± 0.02 2.98 ± 0.03 2.96 ± 0.04


Total oil (%) 49.27 ±0.12 49.22 ±0.13 49.29 ± 0.17 49.29 ± 0.03

Solvent-extractable

oil

(% of free oil in

powder)

4.73 ± 0.00 4.73 ± 0.00 4.72 ± 0.00 4.72 ± 0.00

Encapsulation

efficiency (%)

90.41 ± 0.08 90.40 ±0.09 90.41 ± 0.12 90.42 ± 0.02

Mean particle

diameter, d3,2,

prior to spray

drying

0.36 ± 0.00 0.35 ± 0.00 0.35 ± 0.00 0.35 ± 0.00

aValues are averages of duplicate analysis ± SD of 2 individual runs

The average oil droplet size (d3,2) of all the reconstituted microcapsules at zero month was

~0.8−0.9 µm. This showed that the particle size of zero month powders were slightly

greater than the particle size of the corresponding emulsions prior to drying. This increase

in particle size may be due to changes to the interfacial structure of SBP, causing

coalescence of oil droplets and also, some pectin aggregation during the spray drying

process. According to Rees (1969), the pectin chain interactions are promoted under low

aw conditions. Kirby, MacDougall, & Morris (2006) used atomic force microscopy (AFM)

53

to assess the structure of pectin molecules and also, to characterise protein-polysaccharide

interactions in SBP. They reported that when SBP was deposited on a mica surface it

formed small aggregates. This aggregation process was believed to be because of the

adsorbed protein-protein complexes at the air-water interface as the solvent is removed

during drying. Particle size (d3,2) values for FO and FO-EVOO microcapsules with and

without EDTA over storage time were similar indicating no influence of EDTA. The

average oil droplet size range increased from 0.84−0.90 ± 0.01 µm for microcapsules

stored for 1 month, to 1.27−1.35 ± 0.04 µm after 2 months and 1.44−1.46 ± 0.01 µm after

3 months. Visual inspection of all the stored microcapsules revealed that there were a

number of lumps in 2nd and 3rd month stored powders and absolutely no lumps in either the

0 or 1st month of storage. Also, microcapsules stored for 2 months and 3 months were

increasingly difficult to reconstitute. The increase in moisture content and aw during

storage time in the microcapsules (Table 11) and the increase in particle size / aggregation

of particles could be attributed to the change in aw over storage time. When moisture is

taken up by the powders, plasticisation of the encapsulant material occurs, leading to

increased particle-particle interaction and consequent agglomeration of powder particles

into lumps. The phenomenon of stickiness, caking or crystallisation that effect the quality

of spray-dried powders have been shown to be influenced by time, temperature and

moisture and have been numerously reported to have been encountered during the

production and storage of food powders (Levine & Slade, 1986; Bhandari et al., 1997;

Ozkan, Withy & Chen, 2003).

SEM of the microcapsules indicated that the stored powders had an increase in

agglomeration in the powder particles when compared to the SEM images of the fresh

powders (Figure 17). The microcapsules were regular in shape with wrinkled surface and

no cracks or pores. This surface wrinkling has been associated to be caused by the

mechanical stress caused during spray drying process and has been discussed in section

4.2. Most images had microcapsules as discrete units with no incomplete or damaged shell.

54

Table 11 Changes in moisture content and water activity of FO and FO-EVOO

microcapsules (50% oil, dry basis) (with and without EDTA) at pH 6 on storage at room

temperature (~25°C).

aMicrocapsule Moisture content (%) Water activity (aw)


FO-EDTA-50 2.83 ± 0.01 3.25 ± 0.01 0.30 ± 0.01 0.49 ± 0.01

FO-EVOO-EDTA-50 2.89 ± 0.02 3.01 ± 0.06 0.30 ± 0.00 0.48 ± 0.00

FO-50 2.98 ± 0.03 3.53 ± 0.06 0.29 ± 0.00 0.49 ± 0.01

FO-EVOO-50 2.96 ± 0.04 3.19 ± 0.12 0.29 ± 0.00 0.49 ± 0.00 aValues are averages of duplicate analysis ± SD of 2 individual runs

Table 12 Particle size values of the reconstituted spray-dried microcapsules (50% oil, dry

basis), with and without EDTA at pH 6, on storage at room temperature (~25ºC).

Microcapsule

Mean particle diameter, d3,2, (µm)a


FO-EDTA-50 0.87 ± 0.01b 0.88 ± 0.01a 1.30 ± 0.03ab 1.46 ± 0.03a

FO-EVOO-EDTA-50 0.84 ± 0.00a 0.89 ± 0.00a 1.35 ± 0.04b 1.44 ± 0.03a

FO-50 0.89 ±0.00b 0.89 ± 0.00a 1.28 ± 0.06a 1.44 ± 0.04a

FO-EVOO-50 0.87 ± 0.01c 0.90 ± 0.02a 1.27 ± 0.04a 1.46 ± 0.03a aValues are averages of triplicate analysis ± SD of 2 individual runs


55

0

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olu

me

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olu

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0.01 0.1 1 10 100 1000

Particle size (µm)

Vo

lum

e (

%)

A B

C D

Figure 16 Particle size distribution of reconstituted spray-dried microcapsules (50% oil,

dry basis), with and without EDTA at pH 6 after 0, 1, 2 and 3 month (A–D) storage at

room temperature (~25ºC). Values are the averages of triplicate measurements ± SD; �

(FO-EDTA-50); � (FO-EVOO-EDTA-50); � (FO-50); � (FO-EVOO-50).

56

A B

C D

A B

C

A B

C DD

Figure 17 Scanning electron microscopy images of the fish oil-extra virgin olive oil

spray-dried powders (50% oil, dry basis), with (A, C) and without (B, D) EDTA, at pH 6,

prior (A, B) and subsequent (C, D) to storage for 3 months at room temperature (~25ºC).

4.4 EFFECT OF pH ON EMULSION AND MICROCAPSULE

CHARACTERISTICS

The mean particle diameters (d3,2) of FO and FO-EVOO emulsions at pH 3 with 50% oil

loading were 0.41 ± 0.00 µm and 0.43 ± 0.00 µm, respectively. In comparison, both the FO

and FO-EVOO emulsions at pH 6 with 50% oil loading had a slightly smaller mean

particle diameter of 0.35 ± 0.0 µm. The difference in particle size observed for emulsions

at pH 3 and pH 6 indicated a slight influence of pH on the emulsion particle size. It has

already been discussed in section 4.2 that 2% SBP was sufficient to prepare stable FO

emulsions with 50% oil loading. The small difference in mean particle diameters of the

57

emulsions made at pH 3 and pH 6 suggests that there was sufficient emulsifier present to

prepare finely-dispersed emulsions and any change in conformation / charge of the SBP as

pH is raised from pH 3 to 6 did not have a marked effect on the emulsifying capacity of

SBP.

The particle size distributions of FO and FO-EVOO emulsions at pH 3 and pH 6 showed a

non-Gaussian, largely uni-modal distribution (Figure 18). It was observed that there was a

shoulder on the major peak in the distribution curve in all the 4 emulsions, corresponding

to particles having a mean diameter between 1−10 µm, and this was more prominent in the

emulsions prepared at pH 3. This shoulder could be because of the presence of small

aggregates caused by the interaction between particles. When the pH of a solution is

altered the proteins present in the mixture may experience some conformational changes

and as they approach their isoelectric point they precipitate (Vaclavik & Christian, 2008).

This precipitation may cause the proteins to loose their emulsifying capacity as the

precipitated particles collide, stick and break apart until a stable mean particle size is

reached and in the due process causes some aggregation.

0

1

2

3

4

5

6

7

8

9

0.01 0.1 1 10 100 1000

Particle size (µm)

Vo

lum

e (

%)

Figure 18 Particle size distributions of the original (fresh) fish oil and fish oil-extra virgin

olive oil (1:1 wt ratio) emulsions made at pH 3 and 6. Values are the averages of triplicate

measurements ± SD; � (FO-50 (pH 3)); � (FO-EVOO-50 (pH 3)); � (FO-50 (pH 6)); �

(FO-EVOO-50 (pH 6)).

58

The brightfield micrographs of the emulsions are shown in Figure 19. The micrographs of

pH 3 and pH 6 emulsions were comparable and appeared to have no large aggregates of

undissolved SBP. Additionally, all the emulsions were homogeneous with no indication of

any unencapsulated oil.

Figure 19 Brightfield micrographs of (A) fish oil (pH 3) and (B) fish oil-extra virgin olive

oil (1:1 wt ratio) emulsions (pH 6) with 15% oil.

In general, all the emulsions exhibited shear thinning behaviour with the viscosities

decreasing with increase in shear rate (Figure 20). FO and FO-EVOO emulsions having

15% oil (wet basis) that were adjusted to pH 6 were less viscous than the FO and

FO-EVOO emulsions at pH 3, indicating the influence of pH. The zeta potential data

showed that the SBP-stabilised oil droplets had an isoelectric point of ~!1.5. Thus the

droplets were more negatively charged at pH 6 (ξ-potential = -41.5 ± 2.12 mV) than pH 3

(ξ-potential = -14.0 ± 1.41 mV). As the pH is increased to pH 6, the carboxyl groups on the

pectin molecule (as well as protein) lose H+ and therefore the negative charge on the SBP

will increase, resulting in increased electrostatic repulsion. At pH 3, there is less charge

(smaller zeta potential) and therefore the tendency for the particles to aggregate is

increased. The increased interaction between particles lead to an increase in viscosity of

the emulsions made at pH 3.

59

0

200

400

600

800

1000

1200

1400

1600

1800

0 50 100 150 200 250 300

Shear rate (1/s)

Vis

co

sity

(m

Pa

/s)


emulsions with 15% oil at pH 3 and pH 6, as a function of shear rate. Values are the

average of duplicate measurements ± SD; � (FO-50 (pH 3)); � (FO-EVOO-50 (pH 3));

� (FO-50 (pH 6)); � (FO-EVOO-50 (pH 6)).

The moisture content of the FO and FO-EVOO powders at pH 6 with 50% oil loading was

2.98 ± 0.03% and 2.96 ± 0.04%, respectively at 0 month storage (Table 13). The moisture

content of the FO and FO-EVOO powders at pH 3 with 50% oil loading was 1.66 ±

0.01% and 1.62 ± 0.03%, respectively. A water activity of ~0.3 was found for all the

powders. The higher moisture content in pH 6 microcapsules was directly related to the

smaller particle size diameter. Smaller particle size provided the larger surface area and

facilitated entrapping more water molecules between particles.

Moisture content and aw of FO and FO-EVOO powders made at pH 3 and pH 6 with 50%

oil loading was determined after 3 months of storage under ambient conditions to assess

their physical stability. Both properties increased with increased storage duration. It was

observed that the percent increase in moisture content and aw was greater in microcapsules

made at pH 3 (Table 14) and as pointed out previously (section 4.3), that was probably due

to absorption of moisture from the surrounding environment (aw ~0.5) as the samples were

stored in (plastic) containers that were permeable to moisture. The moisture content

60

measurements were in agreement with the results of aw, with the aw increasing with the

increase in moisture content.

FO and FO-EVOO microcapsules prepared at pH 3 and pH 6 with 5 % oil loading both

had ~10% solvent-extractable fat and encapsulation efficiencies of ~90% (Table 13). Thus,

it can be concluded that neither pH nor the type of oil/oils in the core of microcapsule had

any effect on either solvent-extractable fat or encapsulation efficiency.

It can be collectively concluded that there were no differences among the microcapsules at

pH 3 and pH 6 when total oil percentage, extractable oil percentage and encapsulation

efficiency was compared.

Table 13 Properties of powders containing microencapsulated fish oil and fish oil-olive oil

(1:1 wt ratio) at pH 3 and pH 6 with 50% oil loading.

Property

aMicrocapsule

FO-50

(pH 3)

FO-EVOO-50

(pH 3)

FO-50

(pH 6)

FO-EVOO-50

(pH 6)

Moisture content (%) 1.66 ± 0.01 1.62 ± 0.03 2.98 ± 0.03 2.96 ± 0.04


Total oil (%) 49.09 ± 0.14 49.22 ± 0.12 49.29 ±

0.17

49.29 ± 0.03

Solvent-extractable oil (%) 4.99 ± 0.01 5.00 ± 0.01 4.72 ± 0.00 4.72 ± 0.00

Encapsulation efficiency

(%)

90.43 ± 0.09 90.42 ± 0.08 90.41 ±

0.12

90.42 ± 0.02

Mean particle diameter, d3,2, prior to spray drying (µm)

0.41 ± 0.00 0.43 ± 0.00 0.35 ± 0.00 0.35 ± 0.00

aValues are averages of duplicate analysis ± SD of 2 individual runs

61

Table 14 Changes in moisture content and water activity of fish oil and fish oil-extra

virgin olive oil (1:1 wt ratio) microcapsules at pH 3 and pH 6 with 50% oil loading over

storage time at room temperature (~25°C).

aMicrocapsule

Moisture content (%) Water activity (aw)


FO-50 (pH 3) 1.66 ± 0.01 2.06 ± 0.03 0.27 ± 0.02 0.5 ± 0.01

FO-EVOO-50 (pH 3) 1.62 ± 0.03 1.93 ± 0.02 0.27 ± 0.0 0.48 ± 0.0

FO-50 (pH 6) 2.98 ± 0.03 3.53 ± 0.06 0.29 ± 0.0 0.49 ± 0.01

FO-EVOO-50 (pH 6) 2.96 ± 0.04 3.19 ± 0.12 0.29 ± 0.0 0.49 ± 0.0 aValues are averages of duplicate analysis ± SD of 2 individual runs

The particle size distribution of the reconstituted spray-dried microcapsules at 0, 1, 2 and 3

month storage under room temperature conditions was determined (Figure 21 & Table

15). A general increase in particle size was observed in all microcapsules over storage time

at room temperature. The increase in particle size for FO and FO-EVOO microcapsules

over storage time were similar, indicating only a minor influence of the type of oil/oils in

the core of the microcapsule. The particle size of all zero month powders at pH 3 and pH 6

with 50% oil loading were slightly greater than the particle size of the corresponding

emulsions prior to spray drying. As stated previously in section 4.3, this could be attributed

to the destabilisation of emulsion droplets during the spray drying process as well the

formation of pectin aggregates.

A significant (P < 0.05) increase in particle size of all the microcapsules at pH 6 with 50%

oil loading was observed after spray drying. However, the increase in particle size of the

microcapsules over storage was comparatively small in comparison with the particle size

of the FO and FO-EVOO microcapsules at pH 3 with 50% oil loading during storage.

62

Figure 21 Particle size distributions of reconstituted spray-dried fish oil and fish oil-extra

virgin olive oil microcapsules prepared from emulsions at pH 3 and 6 with 50% oil loading

prior (A), and subsequent to storage for 1 (B), 2 (C) and 3 (D) month under room

temperature conditions (~25 ºC, aw 0.5, exposed to light). Values are the averages of

triplicate measurements ± SD; � (FO-50 (pH 3)); � (FO-EVOO-50 (pH 3)); � (FO-50

(pH 6)); � (FO-EVOO-50 (pH 6))

0

1

2

3

4

5

6

7

0.01 0.1 1 10 100 1000

Particle size (µm)

Vo

lum

e (%

)

0

1

2

3

4

5

6

7

0.01 0.1 1 10 100 1000

Particle size (µm)

Vo

lum

e (%

)

0

1

2

3

4

5

6

7

0.01 0.1 1 10 100 1000

Particle size (µm)

Vo

lum

e (%

)

0

1

2

3

4

5

6

7

0.01 0.1 1 10 100 1000

Particle size (µm)

Vo

lum

e (%

)

A B

C D

63

Table 15 Particle size values of the reconstituted spray-dried microcapsules containing

fish oil and fish oil-extra virgin olive oil (1:1 wt ratio) at pH 3 and pH 6 with 50% oil

loading, during storage of the powders in stoppered flasks at room temperature (~25ºC).

aValues are averages of triplicate analysis ± SD of 2 individual runs


SEM of FO and FO-EVOO microcapsules at pH 3 and pH 6 revealed comparable surface

topology. The microcapsules had a wrinkled surface (Figure 22). As discussed previously

in section 4.2 and 4.3 the wrinkles have been associated with the SBP (Drusch, 2007) but

may also be due to particle shrinkage during the early stages of the drying process (Moreau

& Rosenberg, 1993). Given the surface of the microcapsules is continuous and devoid of

any cracks or pores, it is likely that the microcapsules would protect the encapsulated oil

against rapid oxidation.

The SEM images of the FO and FO-EVOO microcapsules after storage at room

temperature for 3 months show an increased agglomeration of the powder particles. This

phenomenon of lumping can be attributed to the change in moisture content and aw that

occurred in the microcapsules over storage time (Table 14).

aMicrocapsule

Mean particle diameter, d3,2, (µm)


FO-50 (pH 3) 0.70 ± 0.00c 0.78 ± 0.00c 2.06 ±0.08c 3.06 ± 0.05c

FO-EVOO-50 (pH 3) 0.69 ± 0.00b 0.79 ± 0.01d 1.69 ± 0.03b 1.69 ± 0.02a

FO-50 (pH 6) 0.89 ± 0.00b 0.89 ± 0.00a 1.28 ± 0.06a 1.44 ± 0.04a

FO-EVOO-50 (pH 6) 0.87 ± 0.01c 0.90 ± 0.02a 1.27 ± 0.04a 1.46 ± 0.03a

64

A

C

AA

A

A

C

B

D

A

C

AA

A

A

C

B

D

Figure 22 Scanning electron microscopy images of fish oil spray-dried microcapsules

from emulsions prepared at pH 3 (A, C) and pH 6 (B, D) with 50% oil loading, prior (A,

B) and subsequent (C, D) to storage at ambient conditions (~25°C, aw 0.5, 3 months).

4.5 LIPID OXIDATION

Lipid oxidation is a major issue when dealing with oils and powders encapsulated with oils

because of the thermal and oxidative reactions that take place during storage. The primary

products of lipid oxidation are the hydroperoxides which are colourless, tasteless and

odourless. These hydroperoxides are unstable and can further degrade into various low

molecular weight compounds with distinctive colour, odour and flavour characteristics.

These low molecular weight compounds include alkanes, alkenes, alcohols, ketones,

aldehydes, acids and esters. Some of these compounds impart off-flavor/taste to the

powders and drastically decrease the shelf-life of the product (Tamime, 2009).

65

Many secondary products have been identified from the radical and photosensitized

oxidations of polyunsaturated lipids. These secondary products mainly consist of

oxygenated monomeric materials including epoxy-hydroperoxides, oxo-hydroperoxides,

hydroperoxy epidioxides, dihydroperoxides, hydroperoxy bis-epidioxides, and

hydroperoxy bicycloendoperoxides. Some higher molecular weight dimeric compounds

have also been identified from autoxidized methyl linoleate and linolenate. Decomposition

of these oxidation products form a wide range of carbonyl compounds, hydrocarbons and

furans, for example, that contribute to the flavour deterioration of foods (Frankel, 1987).

4.5.1 OXIDATIVE STABILITY OF EMULSIONS AND MICROCAPSULATED

OIL DURING STORAGE UNDER AMBIENT CONDITIONS

4.5.1.1 PROPANAL HEADSPACE ANALYSIS

Propanal and hexanal are the main volatiles formed by oxidative decomposition of

omega-3 fatty acids and omega-6 fatty acids, respectively (Frankel et al., 1994). Hence, the

propanal and hexanal contents measured at different storage times in the microcapsule

powders can be used as indicators of oxidative stability.

Propanal was detected in all the fresh (0 month) FO and FO-EVOO microcapsules made

with 25% and 50% oil loading at pH 3 (Table 16) indicating a possibility of lipid oxidation

taking place either during the emulsion preparation or spray drying process. An increase in

the propanal content in all powders was positively correlated with the increase in storage

time. The amount of propanal detected in the headspace of all the fresh (0 month)

microcapsules were similar to amounts detected by Drusch et al., 2007; Serfert, Drusch &

Schwarz, 2009 for SBP-, caseinate-, n- octenylsuccinate starch and gum arabic stabilised

FO-in-water emulsions, but only in the absence of added antioxidants. The highest content

of propanal in the microcapsule powders was recorded after 3 months of storage in FO-50

(32.35 ± 0.92 µg/g powder) and FO-EVOO-50 (28.70 ± 4.53 µg/g powder) powders made

at pH 3. Significant differences in propanal content (P < 0.05) amongst all the samples

stored for up to 2 months were noted. The propanal content in microcapsule powders

66

stored 0−2 months was higher in microcapsule powders containing 50% oil loading than

powders containing 25% oil loading. This is expected as a more robust interface is

anticipated for the 25% microcapsules, resulting in reduced porosity causing less oxygen

diffusion and hence, less oxidation on storage. The results also showed that the FO-EVOO

powders generally had slightly higher propanal concentrations compared to FO powders

alone, irrespective of oil loading, at most times during storage. However, there were no

significant differences (P > 0.05) in the propanal content in the FO and FO-EVOO samples

at 3 months. The continuous increment of propanal in all powders during storage indicated

the steady oxidation and degradation of omega-3 fatty acids. These observations revealed

that the addition of EVOO was not able to protect FO from oxidation. One of the reasons

for EVOO not being able to increase the oxidative stability of microencapsulated FO could

be the presence of transition metal ions (e.g. Fe, Cu) present in SBP and also, the low pH

environment that led to rapid oxidation.

No propanal was detected in all the fresh (0 month) FO and FO-EVOO powders made with

and without EDTA (50% oil loading) at pH 6 (Table 17). However, a general increase in

propanal content was observed with increase in storage time. The microcapsules with

EDTA had less propanal content compared to the microcapsules without EDTA indicating

that EDTA exerted a significant (P < 0.05) protective effect on the microcapsules with

regard to their oxidative stability.

The results also showed that the FO-EVOO powders had propanal contents significantly (P

< 0.05) lower than FO powders alone indicating that the antioxidant compounds in EVOO

were effective at pH 6. For example, addition of EVOO at 50% oil loading decreased

propanal content by 2.13 fold compared to microcapsules without EDTA after 3 months of

storage (Table 17).

67

Table 16 Propanal content in microencapsulated fish oil and fish oil-extra virgin olive oil

(1:1 wt ratio) (25% and 50% oil loading) at pH 3 during storage at room temperature (~25

°C, 0−3 month) .

aMicrocapsule

Propanal content (µg g-1

powder)


FO-25 1.49 ± 0.09ab 2.63 ± 0.25a 8.50 ± 0.14a 25.85 ± 2.62 a

FO-50 1.66 ± 0.03b 4.55 ± 0.21b 19.15 ± 1.63 b 28.70 ± 4.53 a

FO-EVOO-25 1.41 ± 0.02a 3.97 ± 0.24b 9.28 ± 0.16 a 24.60 ± 3.25 a

FO-EVOO-50 1.58 ± 0.03ab 6.35 ±0.12c 20.30 ± 0.14 b 32.35 ± 0.92 a aValues are average of duplicate analysis ± SD of 2 individual runs

Means within columns followed by different superscript letters differ significantly at 5% level


(1:1 wt ratio) with and without EDTA (50% oil loading) at pH 6 during storage at room

temperature (~25 °C, 0−3 month).

aValues are average of duplicate analysis ± SD of 2 individual runs


Data reported in literature established the fact that metals like Fe and Cu act as catalysts for

the formation of highly reactive hydroxyl radicals which initiate a chain reaction for lipid

oxidation (Benedt & Shibamoto 2007; Goldstein, Meyerstein & Czapski 1993). The

significant (P < 0.05) differences in the propanal content between microcapsules with and

without EDTA confirmed EDTA’s efficiency in inhibiting oxidation and also that metal

aMicrocapsule


powder)


FO-EDTA-50 0a 1.90 ± 0.42a 1.95± 1.01a 18.90 ± 0.00 b

FO-EVOO-EDTA-50 0 a 1.06 ± 0.08a 1.19 ± 0.08a 7.68 ± 0.40 a

FO-50 0 a 2.37 ± 0.01a 9.15 ± 0.92 b 27.55 ± 0.07 c

FO-EVOO-50 0 a 1.88 ±0.21a 7.50 ± 0.85 b 8.80 ± 0.71 a

68

ion-induced oxidation reactions was one of the major reasons for the oxidative

deterioration of the FO and FO-EVOO microcapsules during storage. Jacobsen (2010)

performed some studies to retard lipid oxidation in foods enriched with omega-3-fatty

acids. It was found that the addition of EDTA (5 mg/kg milk) to skimmed milk

significantly retarded the oxidation when the emulsions were enriched with 1.5% FO.

Data gained from those studies also indicated that EDTA was effective in preventing

oxidation in omega-3 fatty acid-enriched mayonnaises and salad dressings when compared

to fitness bars. The article also illustrated that the effectiveness of EDTA might also

depend on the initial level of lipid hydroperoxides in the FO or the emulsion.

At pH 6, improved oxidative stability can be expected as the solubility of iron in water

increases with decreasing pH (Donnelly et al., 1998; Graf et al., 1984; Mancuso et

al.,1999). Therefore, the high propanal contents of FO and FO-EVOO microcapsules at pH

3 as compared to FO and FO-EVOO microcapsules at pH 6 could be attributed to the

increased solubility of iron. Indeed, some amount of propanal was found in FO and

FO-EVOO microcapsules at pH 3 prior to storage (zero month), whereas no detectable

propanal was observed in FO and FO-EVOO microcapsules at pH 6 at zero month. The

propanal content of FO and FO-EVOO with 50% oil loading at pH 6 were significantly

lower than the FO and FO-EVOO samples with 50% oil loading at pH 3 (P < 0.05),

irrespective of the storage duration (Table 18). This also showed that there was an effect of

pH on the oxidative stability of the powders. The results revealed that the FO-EVOO

powders at pH 6 had propanal contents (8.8 ± 0.71 µg/g powder) significantly (P < 0.05)

lower than FO powders (27.55 ± 0.07 µg/g powder) alone after 3 months of storage.

In addition to its influence on the solubilisation of transition metals, pH also affects oxygen

solubility and mobility, and the rate of non-enzymatic browning reaction in foods . There

have been studies where FO-enriched mayonnaise showed increased oxidation with

decreasing pH during storage and also that metal ions significantly promoted oxidation

(Tong et al., 2000; Jacobsen, Timm & Meyer, 2001). In another study, polyoxyethylene 10

lauryl ether was used as an emulsifier in making model emulsions and it was found that

iron showed a highly increased pro-oxidative effect at pH 3 compared to that at pH 7

69

because of the increased solubility of iron at low pH (Cho et al., 2003). However, in the

current study the data indicated that the effect of pH was not significant (P > 0.05) in the

case of FO without added EVOO. Consequently, it could be concluded that the antioxidant

effect of EVOO was readily available at pH 6.


(1:1 wt ratio) with 50% oil loading at pH 3 and pH 6 during storage at room temperature

(~25°C, 0−3 month) .



4.5.1.2 HEXANAL HEADSPACE ANALYSIS

There was no detectable hexanal for 0−2 months storage in either FO or FO-EVOO

microcapsules (25% and 50% oil loading) at pH 3 (Table 19) and FO or FO-EVOO

microcapsules (50% oil loading) with EDTA at pH 6 (Table 20). FO and FO-EVOO

microcapsules (50% oil loading) without EDTA at pH 6 had some amount of detectable

hexanal in the 2nd and 3rd month and no detectable hexanal prior to 2 months (Table 21).

The later development of hexanal compared to propanal may be expected as hexanal and

propanal are secondary oxidation products of omega-6 (e.g. C18:2) and omega-3 fatty

acids (e.g. DHA, EPA, C18:3), respectively. It is well known that omega-3 fatty acids are

more prone to oxidation than omega-6 fatty acids. It has, however, been reported that

hexanal cannot be used as an adequate marker for the beginning of oxidation in case of

aMicrocapsule


powder)


FO-50 (pH 3) 1.66 ± 0.03b 4.55 ± 0.21b 19.15 ± 1.63b 28.70 ± 4.53bc

FO-EVOO-50 (pH 3) 1.58 ±0.03ab 6.35 ±0.12c 20.30 ± 0.14b 32.35 ± 0.92c

FO-50 (pH 6) 0a 2.37 ± 0.01a 9.15 ± 0.92a 27.55 ± 0.07b

FO-EVOO-50 (pH 6) 0a 1.88 ±0.21a 7.50 ± 0.85a 8.80 ± 0.71a

70

virgin olive oil, although it has been successful with refined vegetable oils (Snyder et al.,

1988; Warner et al., 1988).

Table 19 Hexanal content in microencapsulated fish oil and fish oil-extra virgin olive oil

(1:1 wt ratio) at pH 3 during storage at room temperature (~25°C, 0−3 month)

aMicrocapsule Hexanal content (µg g

-1 powder) after 3 months storage

FO-25 1.4 ± 0.21a

FO-50 1.6 ± 0.36a

FO-EVOO-25 2.1 ± 0.42a

FO-EVOO-50 1.8 ± 0.71a aValues are average of duplicate analysis ± SD of 2 individual runs


No significant difference (P > 0.05) in the hexanal content between FO and FO-EVOO

microcapsules (25% and 50% oil loading) at pH 3 microcapsules after 3 months of storage

was observed. FO and FO-EVOO microcapsules with EDTA had significantly less hexanal

content compared to the samples without EDTA (P < 0.05). It was also observed that

FO-EVOO microcapsules with and without EDTA had hexanal contents significantly

lower than FO microcapsules with and without EDTA (P < 0.05). Also, the hexanal

content of FO and FO-EVOO with 50% oil loading at pH 6 were significantly lower than

the FO and FO-EVOO samples with 50% oil loading at pH 3 (P < 0.05).. That was in

agreement with the propanal results and confirmed that pH was also an influencing factor

in the oxidation of the microencapsulated oil.

Objectionable rancid odour was noted in all microcapsule powders on sniffing the samples

stored in containers for 3 months at ambient conditions .There are a wide range of volatile

compounds present in FO and EVOO and these volatiles bring out different tastes and

odours. Each of these compounds have different thresholds of perception. It is anticipated

that propanal and hexanal give rise to a rancid odour.

71


(1:1 wt ratio) with and without EDTA (50% oil loading) at pH 6 during storage at room

temperature (~25°C, 0−3 month) .




(1:1 wt ratio) with 50% oil loading at pH 3 and pH 6 during storage at room temperature

(~25°C, 0−3 month).



4.5.1.3 EVALUATION OF LIPID OXIDATION BY DETERMINING THE

CHANGES IN THE FATTY ACIDS COMPOSITION OVER STORAGE TIME

The major fatty acids present in the oils (FO and EVOO) and all encapsulated oils were

palmitic acid (C16:0), oleic acid (C18:1), EPA (C20:5) and DHA (C22:6). Hence, the

aMicrocapsule

Hexanal content (µg g-1

powder)


FO-EDTA-50 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 1.26 ± 0.02c

FO-EVOO-EDTA-50 0.00 ± 0.00a 0.00 ± 0.00a 0 .00± 0.00a 0.18 ± 0.04a

FO-50 0.00 ± 0.00a 0.00 ± 0.00a 0.12 ± 0.01a 0.51 ± 0.01b

FO-EVOO-50 0.00 ± 0.00a 0.00 ± 0.00a 0.41 ± 0.28a 0.44 ± 0.06b

aMicrocapsule

Hexanal content (µg g-1

powder)


FO-50 (pH 3) 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 1.60 ± 0.36a

FO-EVOO-50 (pH 3) 0.00 ± 0.00a 0.00 ± 0.00a 0 .00± 0.00a 1.80 ± 0.71a

FO-50 (pH 6) 0.00 ± 0.00a 0.00 ± 0.00a 0.12 ± 0.01a 0.51 ± 0.01b

FO-EVOO-50 (pH 6) 0.00 ± 0.00a 0.00 ± 0.00a 0.41 ± 0.28a 0.44 ± 0.06b

72

percentage composition of each of the major fatty acids mentioned above was determined

[Table 22 (a & b), Table 23 (a & b) and Table 24 (a & b)] after extraction of the oil

from the microcapsules. The ratio of EPA to C16:0 and DHA to C16:0 were also provided.

Other minor fatty acids present in each of the oils were calculated as a percentage and

represented as ‘others’ in the tables. The data has been expressed as the unsaturated fatty

acids to C16:0 because the absolute value of C16:0 did not change, only its % relative to

the unsaturated fatty acid changed as oxidation proceeded.

As expected, EPA and DHA, being polyunsaturated fatty acids had significant changes in

their composition over the storage period (P > 0.05). The amounts of EPA (C20:5n-3) and

DHA (C22:6n-3) in the oil extracted from all fresh powders containing microencapsulated

FO were slightly lower than that of the pure FO. The extreme susceptibility of

polyunsaturated fatty acids to oxygen, light and temperature is widely known (Garg et al.,

2006; Kolanowski & Laufenberg, 2006) and hence, the oxidation of the omega-3-fatty

acids in the oil must have occurred during the preparation of emulsions and/or the spray

drying process. The monounsaturated fatty acid composition was comparable. As

expected, the polyunsaturated omega-3 fatty acids EPA and DHA in all microcapsules

decreased over the storage period. In contrast, there was no significant change (P > 0.05)

in the oleic acid content with storage time. This result might be explained in terms of the

structure of the fatty acids. Generally, the greater the number of double bonds in the fatty

acid, the more unstable the fatty acid and the greater the ease of its oxidation (e.g. by

oxygen, light) (Olive oil source, 2010).

The percentage decrease in EPA and DHA was found to be greater in microcapsules

having 50% oil loading when compared to microcapsules with 25% oil loading, at pH 3

(Tables 22, a & b). The highest stability of EPA and DHA was found to be in FO

microcapsules (25% oil loading) with the EPA decreasing from 6.4% at 0 month to 6.2% at

the 3rd month, and DHA from 23.6% at 0 month to 21.8% at the 3rd month. FO-EVOO-50

microcapsules and FO-50 microcapsules were found to be the least stable after 3 months of

storage. The EPA content decreased from 3.3% to 2.7% in FO-EVOO-50 and from 6.7%

to 5.6% in FO-50, and DHA from 12% to 9.3% in FO-EVOO-50 and from 24% to 18% in

73

FO-50 microcapsules. A percentage decrease of 16.31% and 22.84% in EPA and DHA

was observed in FO-EVOO-50 microcapsules over 3 months of storage whereas, a

percentage decrease of 16.49% EPA and 25.55% DHA was recorded in FO-50

microcapsules indicating that EVOO had a slight antioxidative effect on controlling lipid

oxidation of microencapsulated FO. On the contrary, FO-EVOO-25 had a greater

percentage decrease of EPA and DHA, 12.04% and 17.75%, respectively, when compared

to 2.82% EPA and 7.83% DHA in FO-25. That indicated that the concentration of

antioxidative components present in EVOO was not sufficient to protect the FO from

oxidation or other mechanisms come into play which offset or (negated) the antioxidant

effects of EVOO components.

FO and FO-EVOO microcapsules without EDTA at pH 6 in the formulation were found to

be least stable when compared to the microcapsules with EDTA (Table 23, a & b). That

suggested that the metal ions present in SBP were one of the underlying factors responsible

for lipid oxidation in FO and FO-EVOO microcapsules prepared at pH 3. FO-EVOO

microcapsules were less oxidised than FO microcapsules made with 50% oil loading at pH

6. EPA and DHA contents decreased by 0.33% and 1.99%, respectively, in FO-EVOO-50

microcapsules and 0.74% for EPA and 4.46% for DHA in FO-50 microcapsules.

Consequently, it could be concluded that EVOO was as effective in preventing lipid

oxidation in FO microcapsules as EDTA during long term storage. That might be

attributed to the high antioxidant content and radical scavenging activity of EVOO.

FO and FO-EVOO microcapsules (50% oil loading) at pH 3 were found to be less stable

when compared to FO and FO-EVOO microcapsules (50% oil loading) at pH 6 [Table 22

(a & b) and Table 23 (a & b)]. EPA and DHA contents decreased by 16.49% and

25.55%, respectively, in FO-50 microcapsules and EPA by 16.31% and DHA by 22.84%

in FO-EVOO-50 microcapsules at pH 3. EPA and DHA contents decreased by 12.71% and

19.16%, respectively in FO-50 microcapsules and 10.41% and 15.89%, respectively, in

FO-EVOO-50 microcapsules at pH 6. This shows that FO microcapsules were less stable

than FO-EVOO microcapsule at both pH 3 and pH 6.

74

Table 22 a Fatty acid composition (%) of microencapsulated fish oil (25% and 50% oil loading) at pH 3 during storage at room

temperature (~25°C, 0−3 months).

Fatty acid

Microcapsule

FO-25 FO-50

0 month 1 month 2 month 3 month 0 month 1 month 2 month 3 month

C16:0 19.78 ± 1.25 21.14 ± 0.13 22.16 ± 0.01 23.05 ± 0.29 20.78 ± 0.15 21.53 ± 0.49 23.58 ± 0.08 25.34 ± 0.08

C18:1 14.07 ± 0.86 15.07 ± 0.39 14.72 ± 0.13 16.11 ± 0.17 14.55 ± 0.12 14.69±0.65 16.03 ± 0.05 15.31 ± 0.04

C20:5n-3 6.39 ± 0.14 6.36 ± 0.17 6.42 ± 0.05 6.21 ± 0.07 6.67 ± 0.29 6.14 ± 0.12 5.40 ± 0.01 5.57 ± 0.11

C22:6n-3 23.63 ± 0.17 22.55 ± 0.92 21.66 ± 0.22 21.78 ± 0.08 24.23 ± 1.24 21.07 ± 0.26 18.35 ± 0.11 18.04 ± 0.12

Others 37.13 ± 0.67 35.05 ± 0.57 33.77 ± 0.42 36.56 ± 0.61 33.77 ± 0.57 36.56 ± 0.21 24 ± 0.24 24.46 ± 0.19

DHA:EPA 3.7± 0.11 3.54 ± 0.05 3.4 ±0.00 3.5 ± 0.01 3.63 ± 0.03 3.44 ± 0.02 3.34 ± 0.01 3.24 ± 0.04

EPA:C16 0.32 ± 0.01 0.30 ± 0.01 0.29 ± 0.00 0.27 ± 0.00 0.32 ± 0.01 0.29 ± 0.01 0.23 ± 0.00 0.22 ± 0.00

DHA:C16 1.20 ± 0.08 1.07 ± 0.05 0.98 ± 0.01 0.94 ± 0.01 1.17 ± 0.05 0.98 ± 0.01 0.78 ± 0.01 0.71 ± 0.01

Values are average of duplicate measurements ± SD

75

Table 22 b Fatty acid composition (%) of microencapsulated fish oil–extra virgin olive oil (1:1 wt ratio) (25% and 50% oil loading) at

pH 3 during storage at room temperature (~25°C, 0−3 months).

Fatty acid

Microcapsule

FO-EVOO-25 FO-EVOO-50


C16:0 14.71 ± 0.08 15.02 ± 0.04 14.87± 0.21 15.72 ± 0.01 14.57± 0.17 15.01± 0.05 14.94 ± 0.05 15.52 ± 0.16

C18:1 46.05 ± 0.27 46.89 ± 0.09 46.59 ± 0.37 48.52 ± 0.07 45.41 ± 0.15 46.18 ± 0.10 46.98 ± 0.06 48.62 ± 0.30

C20:5n-3 3.24 ± 0.03 3.15 ± 0.01 3.07 ± 0.03 2.85 ± 0.10 3.25 ± 0.06 3.09 ± 0.07 2.92 ± 0.01 2.72 ± 0.03

C22:6n-3 12.00 ± 0.23 11.30 ± 0.13 11.02 ± 0.03 9.87 ± 0.03 12.04 ± 4.4 10.74 ± 0.32 10.26 ± 0.03 9.29 ± 0.17

Others 24.00 ± 0.61 24.4 ± 0.26 24.73 ± 0.11 24.91 ± 0.01 24.73 ± 0.45 24.91 ± 0.34 24.91 ± 0.15 23.84 ± 0.34

DHA:EPA 3.70± 0.04 3.61 ± 0.01 3.6 ± 0.01 3.46 ± 0.13 3.71 ± 0.06 3.48 ± 0.04 3.50 ± 0.00 3.42 ± 0.03

EPA:C16 0.22 ± 0.00 0.21 ± 0.00 0.21 ± 0.00 0.18 ± 0.00 0.22 ± 0.01 0.21 ± 0.00 0.19 ± 0.00 0.18 ± 0.00

DHA:C16 0.82 ± 0.01 0.75 ± 0.01 0.74 ± 0.00 0.63 ± 0.00 0.81 ± 0.05 0.72 ± 0.02 0.69 ± 0.00 0.60 ± 0.02


76

Table 23 a Fatty acid composition (%) of microencapsulated fish oil (50% oil loading) with and without EDTA at pH 6, during

storage at room temperature (~25°C, 0−3 months).

Fatty acid

Microcapsule

FO-EDTA-50 FO-50


C16:0 19.50 ± 0.66 19.66 ± 0.35 20.13 ± 0.50 19.71 ± 0.44 19.10 ± 0.57 19.48 ± 0.65 20.37 ± 0.11 20.16 ± 0.38

C18:1 14.71 ± 0.72 13.56 ± 0.28 15.06 ± 0.85 15.36± 1.93 17.38 ± 2.29 16.95 ± 2.88 18.16 ± 2.70 18.74 ± 2.29

C20:5n-3 6.06 ± 0.24 6.44 ± 0.33 5.89 ± 0.21 5.48 ± 0.35 5.82 ± 0.06 5.64 ± 0.02 5.36 ± 0.05 5.08 ± 0.06

C22:6n-3 24.13 ± 0.18 25.05 ± 0.33 22.52 ± 0.16 20.57 ± 1.33 23.28 ± 0.15 22.30 ± 0.37 20.43 ± 0.60 18.82 ± 0.18

Others 35.52 ± 0.36 35.30± 0.27 36.65 ± 0.29 38.88 ± 1.90 34.42 ± 1.66 34.98 ± 1.85 35.84 ± 1.94 37.20 ± 2.00

DHA:EPA 3.99 ± 0.13 3.89 ± 0.04 3.83 ±0.16 3.76 ± 0.02 4.00 ± 0.06 3.95 ± 0.06 3.78 ± 0.08 3.71 ± 0.03

EPA:C16 0.31 ± 0.00 0.33 ± 0.01 0.29 ± 0.00 0.28 ± 0.02 0.31 ± 0.01 0.29 ± 0.01 0.26 ± 0.00 0.25 ± 0.00

DHA:C16 1.22 ± 0.03 1.29 ± 0.04 1.12 ± 0.04 1.04 ± 0.06 1.22 ± 0.03 1.16 ± 0.02 1.00 ± 0.02 0.93 ± 0.01


77

Table 23 b Fatty acid composition (%) of microencapsulated fish oil–extra virgin olive oil (1:1 wt ratio) (50% oil loading) with and

without EDTA at pH 6, during storage at room temperature (~25°C, 0−3 months).

Fatty acid

Microcapsule

FO-EVOO-EDTA-50 FO-EVOO-50


C16:0 15.30 ± 1.02 16.31 ± 0.25 16.39 ± 0.55 16.56 ± 0.78 15.76 ± 0.23 15.86 ± 0.13 16.21 ± 0.21 16.25 ± 0.08

C18:1 35.88 ± 2.09 40.40 ± 0.68 39.80 ± 0.37 40.70 ± 2.01 40.26 ± 1.21 41.18 ± 1.54 42.19 ± 0.99 42.10 ± 0.90

C20:5n-3 3.69 ± 0.08 3.22 ± 0.24 3.22 ± 0.14 3.19 ± 0.04 3.17 ± 0.28 3.04 ± 0.26 2.84 ± 0.15 2.84 ± 0.12

C22:6n-3 14.39 ± 0.25 12.66 ± 0.68 12.60 ± 0.03 12.11 ± 0.11 12.52 ± 0.68 11.88 ± 0.61 10.77 ± 0.08 10.53 ± 0.38

Others 30.74 ± 2.78 27.99 ± 0.51 27.99 ± 0.33 27.44 ± 2.86 28.30 ± 0.09 28.04 ± 0.54 27.99 ± 0.55 28.28 ± 0.96

DHA:EPA 3.90 ± 0.02 3.90 ± 0.09 3.92 ± 0.17 3.79 ± 0.06 3.96 ± 0.14 3.91 ± 0.14 3.80 ± 0.17 3.71 ± 0.06

EPA:C16 0.24 ± 0.02 0.24 ± 0.01 0.20 ± 0.00 0.19 ± 0.01 0.18 ± 0.02 0.18 ± 0.01 0.16 ± 0.01 0.16 ± 0.01

DHA:C16 0.92 ± 0.08 0.77 ± 0.03 0.77 ± 0.03 0.73 ± 0.03 0.79 ± 0.03 0.75 ± 0.03 0.66 ± 0.00 0.65 ± 0.02


78

4.5.2 OXIDATIVE STABILITY OF EMULSIONS AND MICROCAPSULES

EXPOSED TO ACCELERATED STORAGE CONDITIONS

Accelerated oxidation tests are generally performed to compare the oxidative stability of

samples containing different ingredients or samples held at varying storage conditions.

Such studies are used also to understand the effectiveness of various antioxidants used in

preventing oxidative deterioration in samples based on values of the induction period (IP).

Accelerated storage tests artificially hasten or speed up the oxidation process by exposing

samples to heat, oxygen, light, metal catalysts, or enzymes. Shelf-life analysis of samples

under ambient conditions should also be conducted for comparison. Accelerated oxidation

tests should be used basically as a rapid screening test. The main difficulty with

accelerated storage test results is that they are conducted under artificial conditions and we

assume that the reactions occurring under these conditions are similar to those reactions

that occur at ambient conditions (Nielsen, 2010).

Results from accelerated storage revealed that the IP of the pure oils and FO-EVOO blend

was greatest for EVOO followed by FO-EVOO and then FO (Table 24). A similar trend

was observed in their rate of oxidation. Oxipres® results for microencapsulated FO and

FO-EVOO at 25% and 50% oil loadings, with and without inclusion of EDTA, at pH 3 and

pH 6 are shown in Tables 25−−−−27. The emulsions and microencapsulated oils had a lower

IP than the pure oils (Table 24 & 25). One reason for this is the larger surface area of the

emulsified oil droplets and microencapsulated oil compared to that of the bulk oil/air

interface. The presence of small oil droplets in the emulsions and microencapsulated oil

allows rapid access of oxygen to the oil.

79

Table 24 Oxidative stability of bulk oils (FO, EVOO, FO-EVOO (1:1 wt ratio) during

accelerated storage (80°C, oxygen pressure of 0.5 bar).

aSample Induction period (h) Slope (mbar h

-1)

FO 11.85 ± 0.07 -210 ± 9.19

EVOO >100 ± 0.00 −

FO:EVOO (1:1 wt ratio) 16.3 ± 0.14 -119 ± 4.24

aValues are averages of duplicate analysis ± SD

There was no significant (P > 0.05) difference in the IP values between emulsions made

for manufacture of powders with 25% and 50% oil loading at pH 3 (Table 25). That also

suggested that there was no effect of oil composition or oil loading on the IP values of

emulsions. Similarly, FO microcapsules (25% and 50% oil loading) revealed the same IP

values (6.95 ± 0.07 h). However, the IP value increased from 7.90 h to 11.90 h when the

oil load was increased from 25% to 50% in FO-EVOO microcapsules. The results

indicated that spray-dried FO-EVOO microcapsules had greater oxidative stability than

spray-dried FO microcapsules under the accelerated storage conditions.

The enhanced oxidative stability of the FO-EVOO microcapsules could be due to the

presence of olive oil, which contains large amounts of oleic acid (55−83%) and much

smaller amounts of saturated fatty acids (14%). That interpretation was in agreement with

the findings of Parkanyiova et al., (2000) who reported that triacylglycerols containing

bound linoleic acid were oxidised several times faster than triacylglycerols containing only

oleic and saturated fatty acids. However, the Oxipres® data were in direct contrast to those

obtained for powders prepared at pH 3 and stored under ambient conditions, as determined

by propanal and hexanal analysis and fatty acid composition data. This could be attributed

to a variety of auto oxidative mechanisms that take place with respect to the change in

temperature of oxidation. It has been widely reported that the antioxidative potential of

virgin olive oil is affected by both storage and thermal treatment due to modifications in

80

oxidative/hydrolytic reactions. Other studies have shown that thermal treatment of virgin

olive oil resulted in an increase in the formation of hydroxytyrosol and tyrosol from virgin

olive oil secoiridoids (Bešter et al., 2008; Brenes et al., 2002; Lerma-Garcia et al., 2009;

Sacchi et al., 2002). Another study by Carrasco-pancorbo et al., (2005) demonstrated that

hydroxytyrosol contributed to the oxidative stability of virgin olive oil.

Table 25 Oxidative stability of emulsions and powders (25% and 50% oil loading)

containing fish oil and fish oil-extra virgin olive oil (1:1 wt ratio) at pH 3 during

accelerated storage (80°C, oxygen pressure of 0.5 bar).

Sample Induction period (h) Slope (mbar h-1

)

(A) Emulsion

FO-25 4.90 ± 0.14a -28.00 ± 4.24b

FO-50 4.60 ± 0.00a -38.00 ± 1.41a

FO-EVOO-25 5.00 ± 0.14a -21.50 ± 0.71b

FO-EVOO-50 4.65 ± 0.07a -27.00 ± 1.41b

(B) Microcapsule

FO-25 6.95 ± 0.07a -225.00 ±14.14d

FO-50 6.95 ± 0.07a -2350.50 ± 7.78a

FO-EVOO-25 7.90 ± 0.00b -612.50 ± 3.54c

FO-EVOO-50 11.95 ± 0.35c -792.00 ± 1.41b

Values are averages of duplicate analysis ± SD of 2 individual runs


As expected, the addition of antioxidant i.e EDTA, in the formulation of the microcapsules

significantly increased the IP of FO and FO-EVOO microcapsules and emulsions with

EDTA, offering better oxidative stability. The FO-EVOO emulsions and microcapsules

showed greater oxidative stability with greater IP’s as compared to FO microcapsules at

pH 6 with 50% oil loading (Table 26B). The enhanced stability of FO-EVOO

microcapsules could be attributed to the absence of polyunsaturated fatty acids.

Monounsaturated fatty acids possess only one single double bond making them more

81

stable than polyunsaturated fatty acids yet not as stable as saturated fats (‘Unsaturated fats’

2010; ‘ Extra virgin olive oil’ 2010). All emulsions (FO and FO-EVOO) made with and

without EDTA at pH 6 had relatively small slope values indicating a slow rate of oxidation

in the emulsions, whereas these values were much larger in all the microcapsules (Table

26). This difference in oxidative stability between emulsions and microcapsules may be

due to the orientation of the EDTA and the phenolic compounds present in EVOO in an

aqueous phase and in a dried state. Transition metal ions, such as iron and copper, would

be located in the aqueous phase of oil-in-water emulsions and oriented in the oil/water

interface (Frankel et al., 1994; Jacobsen et al., 2008). EDTA, omega-3 hydroperoxides and

the polar phenolics would also be present in the aqueous phase of the liquid emulsions.

Therefore, it is possible that the shared location and high level of contact between the pro-

and antioxidants in the aqueous phase of the liquid emulsions was more effective in

preventing lipid oxidation than in dried state, due to the water restricted environment in the

latter. Other factors that could possibly cause differences in lipid oxidation in emulsions

and microcapsules would be factors affecting the antioxidant activity and rate of oxidation

in both systems. The effectiveness of antioxidants in oil-in-water emulsions is dependent

on several parameters, such as their structure, polarity, location, radical scavenging and

metal-chelating attributes (Mattia et al., 2010; Paiva-Martins & Gordon, 2002,

Paiva-Martins et al., 2006). The rate and extent of lipid oxidation may also be influenced

by numerous variables like pH, type of emulsifier, oxygen availability and structure,

thickness and composition of the interface (Coupland & McClements, 1996; McClements

& Decker, 2000; Velasco et al., 2003, 2006).

The FO and FO-EVOO emulsions had greater IP values at pH 3 than FO and FO-EVOO

emulsions at pH 6 (Table 27). No significant difference (P > 0.05) in IP values was

observed between FO and FO-EVOO emulsions made with 50% oil loading at pH 3. The

FO-EVOO emulsion had a similar IP value to FO emulsion made with 50% oil loading at

pH 6. All emulsions made with 50% oil loading at pH 3 and pH 6 had lower slope values

than the microcapsulated oil, indicating a slow rate of oxidation. No significant difference

(P > 0.05) between the spray-dried FO and FO-EVOO microcapsules at pH 3 with 50% oil

loading when compared to the corresponding spray-dried FO and FO-EVOO

82

microcapsules at pH 6 (Table 27). Data revealed that the FO-EVOO microcapsules were

better in terms of oxidative stability when compared to FO microcapsules both at pH 3 and

pH 6. In case of FO-EVOO and FO emulsions, the IP values were very small and did not

vary significantly (P > 0.05) from each other irrespective of oil composition and pH. This

result indicated that microencapsulation was significantly (P < 0.05) effective in

preventing lipid oxidation than in liquid emulsions. It has been previously discussed

(section 4.5.2) as to how the effectiveness of the antioxidants may be affected by numerous

variables like structure, polarity, location (e.g. water, oil, interface) and levels of metals

(Fe & Cu) availability. It is possible that the efficacy of antioxidants present in EVOO has

been affected by one of these factors and that could have caused these differences in

microcapsules and emulsions lipid oxidation.

83

Table 26 Oxidative stability of emulsions and powders (50% oil loading) containing fish

oil and fish oil-extra virgin olive oil (1:1 wt ratio) made with and without EDTA (50% oil

loading) at pH 6 during accelerated storage (80°C, oxygen pressure of 0.5 bar).


)

(A) Emulsion

FO-EDTA-50 13.90 ± 0.14b -16.50 ± 2.12ab

FO-EVOO-EDTA-50 23.65 ± 0.49c -11.50 ± 4.95c

FO-50 3.20 ± 0.28a -24.00 ± 7.07ab

FO-EVOO-50 3.90 ± 0.28a -31.50 ± 3.54a

(B) Microcapsule

FO-EDTA-50 10.00 ± 0.28b -421.50 ± 6.36b

FO-EVOO-EDTA-50 19.65 ± 1.06d -131.50 ± 3.54d

FO-50 7.35 ± 0.21a -480.50 ± 4.95a

FO-EVOO-50 12.70 ± 0.14c -97.00 ± 16.97c



84

Table 27 Oxidative stability of emulsions and powders (50% oil loading) containing fish

oil and fish oil-extra virgin olive oil (1:1 wt ratio) at pH 3 and pH 6 during accelerated

storage (80°C, oxygen pressure of 0.5 bar).


)

(A) Emulsion

FO-50 (pH 3) 4.60 ± 0.00a -38.00 ± 1.41a

FO-EVOO-50 (pH 3) 4.65 ± 0.07a -27.00 ± 1.41b

FO-50 (pH 6) 3.20 ± 0.28a -24.00 ± 7.07ab

FO-EVOO-50 (pH 6) 3.90 ± 0.28a -31.50 ± 3.54a

(B) Microcapsule

FO-50 (pH 3) 6.95 ± 0.07a -2350.50 ± 7.78a

FO-EVOO-50 (pH 3) 11.95 ± 0.35c -792.00 ± 1.41b

FO-50 (pH 6) 7.35 ± 0.21a -480.50 ± 4.95a

FO-EVOO-50 (pH 6) 12.70 ± 0.14c -97.00 ± 16.97c



85

CHAPTER 5 - GENERAL CONCLUSIONS AND FUTURE DIRECTIONS

The experimental results obtained in the study confirmed some previous conclusions

drawn by other researchers and generated some new findings supporting initial research

objectives. The following were the main conclusions made:

• SBP was an effective encapsulant only in terms of microencapsulation efficiency of

the microcapsules and required the addition of a chelating agent, such as EDTA, to

neutralize the negative effects of transition metals that were associated with the

polysaccharide to improve the oxidative stability of the microcapsules.

• At pH 3

- EVOO did not improve the oxidative stability of microencapsulated FO during

storage at room temperature.

- In contrast, EVOO improved the oxidative stability of microencapsulated FO

during accelerated storage conditions.

• At pH 6

- FO-EVOO microcapsules had greater oxidative stability than FO microcapsules

both during storage at room temperature and under accelerated storage conditions.

• Addition of EDTA as a chelating agent significantly increased the oxidative

stability of the microcapsules.

• Microcapsules prepared from emulsions at pH 6 were better in terms of long term

oxidative storage stability than microcapsules prepared from emulsions at pH 3.

The main focus of this work was to produce physically stable powders containing

microencapsulated FO with EVOO added for its natural antioxidants and additional health

benefits. Results demonstrated that EVOO did contribute to some antioxidative effect

when microcapsules were prepared at pH 3 and 6 under accelerated storage conditions.

86

However, the same could not be said about the protective effect of EVOO on FO-EVOO

microcapsules prepared at pH 3 when stored under ambient conditions. A significant and

positive effect on oxidative stability of FO microcapsules was observed with the addition

of EDTA in the microcapsule formulation. Similarly, SBP was effective as a wall material

but, for long term stability of FO microcapsules it required EDTA (a metal-chelating

additive) to counteract the pro-oxidative effects caused by the metal ions (iron and copper)

associated with it.

The results obtained through this research project helped to improve understanding of

o the protection requirements to minimise oxidation of FO during processing of

FO microcapsules.

o how and where to add the microencapsulated omega 3 fatty acids in a product.

o the possible interactions of omega-3-fatty acids with other ingredients.

o the physical and shelf-life properties of the FO-EVOO powders.

FUTURE DIRECTIONS

This study identified the potential application of SBP as a food matrix, and tried to

protect FO with the natural antioxidants present in EVOO by optimising the process

parameters to stabilize microcapsules. However, it could be suggested that testing SBP

after depolymerising could yield good results. It is recommended also for further

studies that concentrate on natural food ingredients rich in antioxidants such as

phytosterols could be useful to protect FO from oxidation. Furthermore, the in vitro

analysis of FO-EVOO powders could be examined to examine the susceptibility of the

microcapsule to simulated gastric and intestinal digestion.

Finally, testing the delivery of omega-3 fatty acids in the form of FO-EVOO powder

into various food products (cereal products e.g. breakfast cereals, bakery products e.g.

bread or into cake mixes and fruit nut bars) could have significance nutritional value

and possible industrial applications.

87

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Effect Of pH. (2005). [online]. Available at: http://www.azom. com/details .asp? Article

ID=2840 [Retrieved 9 December 2010].

108

APPENDICES

1. Percentage protein present in the SBP sample.

The nitrogen content in SBP sample (triplicate analysis) found by LECO was:

SBP replicate 1 = 3.887 mg



Average of replicate 1 + 2 + 3= 3.9 mg

N % = N in mg/( Mass of sample (g) * 10)

= 3.9/ (0.5*10)

= 0.78

% protein = N% * factor

(Factor for general samples= 6.25

Factor for samples containing milk= 6.38)

Therefore, % protein = 0.78 *6.25

= 4.9

2. Propanal content calculation

std 0.1 µl = 1987073 peak area counts

Sample (2 g) gave 80137 peak area counts

Therefore amount of propanal equivalents is;

X µl propanal = 80137

so 80137 x 0.1/1987073

=0.00403 µl/2 g

=0.0020 µl/g sample

109

Density of propanal is 0.81 g/ml, therefore we need to convert the propanal concentration

to ml/g to have the same units, so have to multiply by 10-3

so 0.0020 x 10-3 ml (0.81 g/ml)

is 1.62 * 10-6 g

which is 1.62 µg/g of powder.

3. Hexanal content calculation

std 0.1 µl = 1996252 peak area counts

Sample (2 g) gave 75559 peak area counts

Therefore amount of hexanal equivalents is

X µl hexanal = 75559

So 75559 x 0.1/1996252

=0.0038 µl/2 g

=0.00189 µl/g sample

Density of hexanal is 0.81 g/ml, therefore we need to convert the hexanal concentration to

ml/g to have the same units, so have to multiply by 10-3

so 0.00189 x 10-3 ml (0.81 g/ml)

is 0.00000153 g

which is 1.53 µg/g of powder.

4. Total phenolic content in EVOO

The equation obtained from the calibration curve plotted for the caffeic acid is as follows:

y= 3.8617x+ 0.0759

110

Substituting the value of y (absorbance value of the extract at 725 nm) in the above

equation,

x = 0.257- 0.0759 / 3.8617

= 0.047 mg/ ml caffeic acid in 0.2 ml

Therefore, in 5.2 ml of the extract there is 0.244 mg caffeic acid.

0.244 mg caffeic acid is present in 2.5 g oil and hence, there is 97.5 mg/kg oil.

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