Droplet-stabilized oil-in-water emulsions protect unsaturated lipids

Subscriber access provided by University of Kansas Libraries

Food and Beverage Chemistry/Biochemistry

Droplet-stabilized oil-in-water emulsions protect unsaturated lipids from oxidation Sewuese S Okubanjo, Simon M Loveday, Aiqian M. Ye, Peter J. Wilde, and Harjinder Singh J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.8b02871 • Publication Date (Web): 04 Jan 2019 Downloaded from http://pubs.acs.org on January 5, 2019

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 44

Journal of Agricultural and Food Chemistry

1

1 2 3

Droplet-stabilized oil-in-water emulsions protect unsaturated lipids from oxidation

4 5

Sewuese S. Okubanjo1, Simon M. Loveday1,2*ǂ, Aiqian Ye1, Peter J. Wilde3 , Harjinder Singh1

6 7

1. Riddet Institute, Massey University, Private Bag 11222, Palmerston North 4442, New Zealand

8 9 10 11 12

2. Food and Bio-based Products Group, AgResearch Limited, Tennent Drive, Private Bag 11008, Palmerston North 4442, New Zealand

13 14 15 16 17 18 19 20

ǂ Formerly: Massey Institute of Food Science & Technology, Massey University, Palmerston North 4442, New Zealand.

3. Quadram Institute Bioscience, Norwich Research Park, Norwich, Norfolk, NR4 7UA, UK

* Phone +64 6 351 8615 Email [email protected]

ACS Paragon Plus Environment


Page 2 of 44

2 21

Abstract

22

Droplet-stabilized emulsions use fine protein-coated lipid droplets - the shell - to emulsify

23

larger droplets of a second lipid - the core. This study investigated the oxidation resistance

24

of polyunsaturated fatty acid oil (PUFA oil) within droplet-stabilized emulsions, using shell

25

lipids with a range of melting points: olive oil (low melting), trimyristin (high-melting) and

26

palmolein oil (intermediate melting point). Oxidation of PUFA oil was accelerated with a

27

fluorescent lamp in the presence of ferrous iron (100 μM) for nine days, and PUFA

28

oxidation was monitored via conjugated dienes, lipid hydroperoxides and hexanal levels.

29

Oxidation was slower in droplet-stabilized emulsions than in conventional emulsions or

30

control emulsions of the same composition as droplet-stabilized emulsions but different

31

structure, and trimyristin gave the greatest oxidation resistance. Results suggest the

32

structured interface of droplet-stabilized emulsions limits contact between pro-oxidants

33

and oxidation-sensitive bioactives encapsulated within, and this anti-oxidative effect is

34

greatly enhanced with solid surface lipid.

35 36

Keywords: Droplet-stabilized emulsion; surface lipid; shell emulsion; core lipid; oxidation;

37

long-chain polyunsaturated fatty acids

38 39 40


Page 3 of 44


3 41

INTRODUCTION

42

As the spotlight on functional foods continues to grow, the challenges associated with

43

incorporating bioactive compounds into food systems require practical and cost-effective

44

solutions. Emulsions have become very important delivery vehicles for lipophilic

45

bioactives, both to enhance their bioavailability and protect them from degradation.

46

Lipids rich in polyunsaturated fatty acids (PUFA) are beneficial for health1, 2 but they are

47

highly susceptible to oxidation. Strategies such as the use of antioxidants and

48

microencapsulation are currently employed to protect these lipids. Many studies have

49

focused on incorporating these lipids in different emulsion systems to inhibit oxidation and

50

thus maximize their benefits. Many of these studies have been carried out with oil-in-

51

water emulsions3-5.

52

Lipid oxidation is a complex process which leads to the breakdown of lipids into products

53

that result in undesirable sensory properties of lipids. The mechanism of oxidation in bulk

54

oils and emulsions differs6. Oxidation in emulsions is mostly initiated by pro-oxidants

55

located in the aqueous phase or at the interface7,

56

characteristics and composition of the interface of an emulsion have significant impacts

57

on its oxidative stability.

58

The oxidative stability of conventional emulsions is limited by physical instability under

59

various processing conditions and characteristic porous thin interfacial layer

60

enables diffusion of pro-oxidants and subsequent reaction with the lipid phase. Interfacial

61

engineering of emulsion systems can be explored to overcome the limitations of

62

conventional emulsions because the interface is the gateway between pro-oxidants and

63

the lipid phase, creating less permeable interfaces can limit oxidation by minimizing

64

interactions between pro-oxidants and unsaturated lipids 13, 14.


8;

therefore, the structure,

9-12

which


Page 4 of 44

4 65

Despite the many studies that seek to improve the oxidative stability of PUFA lipids by

66

incorporating them into structured emulsions, several limitations such as porous

67

structure, physical instability and high cost of production exist with various emulsion

68

systems15-17 . These limitations must be overcome to achieve the goal of efficient

69

protection, delivery and enhanced bioavailability of PUFAs.

70

Ye et al.

18

reported a patented19 novel emulsion design whereby core oil droplets are

71

stabilized by protein-coated nano-droplets. The reported structure has the potential to

72

effectively protect bioactive compounds such as PUFA-rich lipids. Therefore it could be

73

of great benefit for the food industry if the potential of this novel emulsion to protect

74

lipophilic bioactives such as PUFA-rich lipid from degradation is studied and established.

75

Previous work by Ye et al. used only hexadecane in the lipid phases, whereas this study

76

explores the formation of similar structures using several food-derived lipids, and

77

elucidates the effect of lipid type on oxidative stability. Making these structures with food

78

grade oils is more challenging as the interfacial tension in triacylglycerol oils is much lower

79

than that of hexadecane used in the previous study, presenting a greater barrier for

80

protein to adsorb and stabilize the droplets.

81

This study investigated the oxidation resistance of ‘droplet-stabilized’ emulsions by

82

examining the impact of the interfacial structure and composition on the oxidative stability

83

of PUFA-rich oil incorporated within.

84


Page 5 of 44


5 85

MATERIALS AND METHODS

86

Materials

87

Milk Protein concentrate (MPC-70) was kindly donated by Fonterra Cooperative Ltd,

88

Auckland, NZ. Palmolein oil was purchased from Davis Trading (Palmerston North New

89

Zealand). Trimyristin (Dynasan 114) was kindly donated by IOI Oleo GmbH, Germany.

90

High linoleic acid (72.54% of total fatty acids) refined safflower oil was purchased from

91

Pure Nature Ingredients, Auckland New Zealand. Cumene hydroperoxide, olive oil (low

92

acidity) and hexanal standards were purchased from Sigma-Aldrich. All other reagents

93

used were of analytical grade or higher purity. Olive oil (low acidity- liquid at ambient

94

temperature), palmolein oil (melting range 20-30°C) and trimyristin (melting range 55-

95

58°C) were chosen for this study because of their varying melting points and low

96

susceptibility to oxidation. High linoleic acid safflower oil was selected because of its high

97

susceptibility to oxidation. Peroxide value of safflower oil and trimyristin as specified by

98

supplier was 3 Meq O2/kg. Initial conjugated dienes (CD), lipid hydroperoxides (LHP) and

99

hexanal (hex) values of olive, palmolein and safflower oils as analyzed was less than 30

100

mM for CD and LHP and less than 2 µM for hexanal.

101

102

Emulsion preparation

103

Droplet-stabilized emulsions were prepared by a two-step process. Firstly, solutions

104

consisting of 5% (w/w) milk protein concentrate (MPC) were made by stirring MPC

105

powder (5 g) into Milli-Q water (95 g) at room temperature for 1 h. The MPC solution was

106

heated to 65 ºC and appropriate quantities of lipid (palmolein oil, olive oil, trimyristin)

107

which is referred to as ‘surface lipid’ were added, i.e. 80% w/w MPC solution (consisting

108

of 5% w/w MPC + 95% w/w water) and 20% w/w surface lipid. The mixtures were held at



Page 6 of 44

6 109

65°C for 3-5 min and coarse emulsions made using a high-speed mixer (Labserv, D-130)

110

at 6000 rpm for 3 min. The coarse emulsion was passed through a two-stage

111

homogenizer (Homolab2, FBF Italia) at a first stage pressure of 400 bar and second stage

112

pressure of 50 bar to produce a fine emulsion which is referred to as ‘shell emulsion’. Hot

113

water was passed through the homogenizer prior to passing the coarse emulsion to pre-

114

heat it. In the second step the shell emulsion was added to appropriate quantities of

115

PUFA-rich lipid (safflower oil), which is referred to as ‘core lipid,’ and potassium

116

phosphate buffer (10mM K2HPO4 and KH2PO4, pH 6.8-7.0), i.e. 10% w/w shell droplets,

117

20% w/w core lipid and 70% w/w buffer. To form droplet-stabilized emulsions, the mixture

118

was heated to 65 °C and processed with a high-speed mixer (D-130, Labserv) at 6000

119

rpm for 5 min.

120

Droplet-stabilized emulsions comprising trimyristin surface lipid were processed at two

121

different conditions to obtain shell droplet adsorption in liquid or solid state. To obtain

122

droplet-stabilized emulsions with trimyristin in liquid or super-cooled state, shell emulsion,

123

core and buffer mixture was heated to 65 °C prior to final homogenization. To produce

124

droplet-stabilized emulsion with trimyristin in the solid state, the shell emulsion was first

125

cooled to 5 °C and then heated to 25°C prior to final homogenization with core lipid and

126

buffer.

127

Protein-stabilized safflower oil-in-water conventional control emulsions were processed

128

by mixing appropriate quantities of 0.4% w/v MPC solution and safflower oil with a high-

129

speed mixer as a reference emulsion (R). Control emulsions made up of the same

130

composition as droplet-stabilized emulsions were processed by adding shell emulsions

131

(10% w/w) to appropriate quantities of reference emulsion. The reference and control

132

emulsions were processed at the same temperature as droplet-stabilized emulsions.

133

Table 1 summarizes and shows a description of the emulsion formulations. The reference

134

emulsion was a protein-stabilized conventional emulsion while the control emulsion was


Page 7 of 44


7 135

a conventional emulsion whose composition was matched with that of droplet-stabilized

136

emulsions by addition of shell emulsion.

137 138

Particle size characterization

139

The surface area mean (Sauter mean diameter, d3,2) and volume weighted mean

140

diameter (d4,3) of emulsions were measured using a static light scattering instrument

141

(Mastersizer 2000, Malvern instruments, Worcestershire, UK). The emulsions were

142

dispersed in reverse osmosis-filtered water and the ratio of refractive index of emulsion

143

droplets (1.47) to the aqueous medium (1.33) was 1.105. The measurements were taken

144

in triplicate.

145

146

Confocal Laser Scanning Microscopy

147

The structure and morphology of emulsions was studied using a confocal laser scanning

148

microscope (DM6000B ST5, Leica Microsystems, Wetzlar, Germany). Emulsions were

149

stained according to the method described by Gallier et al. 20. Lipid droplets were stained

150

with Nile red solution dissolved in acetone (1 mg/ml) while proteins were stained with fast

151

green solution dissolved in milli-Q water (1 mg/ml). Staining involved mixing 500 µl of

152

emulsion with 10 µl and 30 µl of Nile red and fast green solutions respectively, then 25 µl

153

of the stained emulsion was deposited on concave microscope slides and fixed with 50

154

µl of agarose solution (0.5%w/w-dissolved in milli-Q water at 80°C). Slides were covered

155

with coverslips and observed under the microscope with the 63X oil immersion objective

156

lens (HCX PL APO lambda blue 63.0 x 1.40 oil UV, Leica Microsystems, Wetzlar,

157

Germany).

158

Nile red was excited at 488 nm with the argon laser and the emitted fluorescence

159

collected between 494-605 nm. Fast green was excited at 633 nm with the helium-neon

160

laser and emission collected between 638-750 nm. The samples were scanned either ACS Paragon Plus Environment


Page 8 of 44

8 161

between lines or sequentially between frames, and images were captured with LAS AF

162

software (version 2.7.3.9723) at room temperature. Images were analysed using ImageJ

163

software (Image J, National Institute of Health, Bethesda, MD, USA).

164 165

Accelerated Lipid Oxidation

166

Oxidation of safflower oil in droplet-stabilized and control emulsions was accelerated in

167

sealed 12-well cell plates (1mL emulsion per well) and 20mL headspace glass vials (95%

168

headspace) exposed to fluorescent lamp (8500-9000 lux) in the presence of ferrous iron

169

(100 μM- FeSO4.7H2O in HCl ) for 9 days at 20°C. The distance from lamp to top of plates

170

and vials was approximately 23 cm and 22 cm respectively. Sodium azide (0.02% w/v)

171

was added to the emulsions to prevent microbial growth.

172 173

Lipid Oxidation Measurements

174

Samples were collected on days 1, 3, 5, 7 and 9, and stored at -20°C. Safflower oxidation

175

was monitored via primary (conjugated dienes and lipid hydroperoxides) and secondary

176

(hexanal) oxidation products. The emulsions were analysed within 1-2 weeks of freezing.

177 178

Conjugated Dienes and Hydroperoxides

179

Extraction of lipids from emulsions was carried out according to the method described by

180

Hu et al.

181

propanol (3:1 v/v) solution. The mixture was vortexed at high speed three times for 10 s

182

each and then centrifuged (Heraeus Fresco 17, Thermo Scientific) at 2000 g for 5 min.

183

The supernatant was collected and used for conjugated dienes and lipid hydroperoxide

184

measurements.

185

Conjugated dienes were determined by the methods described by Wrolstad et al. 22, and

186

Estévez et al. 23 with slight modifications. An aliquot of emulsion was mixed with extraction

21.

Firstly, 0.3 mL of thawed emulsion was mixed with 1.5 mL of isooctane/2-


Page 9 of 44


9 187

solvent and centrifuged as previously described, then 0.2 mL of the supernatant was

188

diluted to 5 ml with isooctane. The solution was thoroughly mixed and absorbance

189

measured at 234 nm in a quartz cuvette of 1 cm path length using a spectrophotometer

190

(Genesys 10S, Thermo Scientific). Iso-octane was used as blank. The concentration of

191

conjugated dienes was calculated as follows: 𝐴234

192

𝐶𝐶𝐷 = (𝜀 ×

ɭ)

(Eq. 1)

193

Where

194

CCD = Conjugated dienes concentration in mmol/ml (molar concentration)

195

A234 = Absorbance of lipid at 234 nm

196

ɛ = Molar absorptivity of linoleic acid hydroperoxide (2.525 X 104 M-1.cm-1)

197

ɭ = Path length of cuvette in cm

198

And the conjugated dienes concentration (CD value) expressed as µM by taking into

199

consideration the volume of isooctane used for dilution.

200

Lipid hydroperoxides analysis was carried out by the ferric thiocyanate method as

201

described by Hu, McClements and Decker 21 and Katsuda, McClements, Miglioranza and

202

Decker 5. Firstly, 0.2 mL of supernatant collected after extraction was mixed with 2.8 mL

203

of methanol/butanol (2:1 v/v) solution. To this solution, 15 µL of ammonium thiocyanate

204

solution (3.94 M) and 15 µL of ferrous iron solution (prepared by mixing 0.132 M of

205

BaCl2.2H2O and 0.144 M FeSO4.7H2O) was added. The solution was vortexed and

206

incubated in the dark at room temperature for 20 min (time monitored after addition of

207

ferrous iron solution to the first reaction tube). Absorbance was read at 510 nm against a

208

cumene hydroperoxide standard curve (concentration range 0-20 µM) using a



Page 10 of 44

10 209

spectrophotometer (Genesys 10S, Thermo scientific) and peroxide value was expressed

210

as µmol cumene hydroperoxide equivalent per litre of emulsion.

211 212

All the oxidation measurements were carried out under subdued light (sample preparation

213

5-6 lux; absorbance readings 6-9 lux).

214 215

Determination of Volatile Secondary Oxidation Product (Hexanal)

216

Analysis of hexanal, a major secondary breakdown product of linoleic acid, was done by

217

solid phase micro-extraction (SPME) coupled with gas chromatography-flame ionization

218

detection. Emulsion samples were placed in 20 mL vials (Phenomenex), sealed with

219

septum crimp tops (20LLX, Phenomenex) and volatiles extracted by the SPME method.

220

The SPME auto fiber assembly (AOC-5000 auto-injector, Shimadzu) consisted of a fused

221

silica fiber coated with carboxen/polydimethylsiloxane (75 µm thickness). The samples

222

were heated in a block maintained at 50 °C for 30 min and agitated at 500 rpm to enable

223

headspace equilibration. The SPME fiber needle then penetrated the vials and the fiber

224

was exposed to the headspace for 5 min.

225

After extraction of volatiles, the fiber was withdrawn, and the adsorbed volatiles injected

226

into a gas chromatograph (GC-2010, Shimadzu) port. The injection port was maintained

227

at 230 °C and enabled thermal desorption of extracted volatiles for 2 min. The GC was

228

equipped with a flame ionisation detector (FID). The detector temperature was 250 °C.

229

The components were separated with an Rtx-1701 column (30 m, 0.25 mm inner

230

diameter, 1 µm film thickness). The column oven temperature was 40 °C (1 min hold)

231

initially and then increased to 100 °C at 5 °C/min and to end temperature of 230 °C at 10

232

°C/min. The direct injection mode was used. Helium was employed as carrier gas (flow

233

rate 1.5 mL/min). Hexanal identification and quantification in samples was determined by


Page 11 of 44


11 234

comparing retention time of peaks against hexanal external standard curves. To achieve

235

this, hexanal standard was added to fresh reference emulsion in various concentrations.

236 237

Statistical Analysis

238

Oxidation results are reported as means and standard error of six data points: two

239

independent trials with triplicate oxidation measurements in each. Data was analysed by

240

two-way analysis of variance (ANOVA) and Duncan’s multiple range test.

241 242

243

Oxidation data was fitted using the following Gaussian equations. 𝑑𝑎𝑦 ― 𝑑𝑎𝑦𝑚𝑎𝑥 2

{ 𝑒𝑥𝑝{ ― ( 𝑒𝑥𝑝{ ― (

𝐶𝐷 = 𝐶𝐷0 + 𝐶𝐷𝑚𝑎𝑥𝑒𝑥𝑝 ―

244

𝐿𝐻𝑃 = 𝐿𝐻𝑃0 + 𝐿𝐻𝑃𝑚𝑎𝑥

245

𝐻𝑒𝑥 = 𝐻𝑒𝑥0 + 𝐻𝑒𝑥𝑚𝑎𝑥

(

𝑤

)}

𝑑𝑎𝑦 ― 𝑑𝑎𝑦𝑚𝑎𝑥 2 𝑤

)}

𝑑𝑎𝑦 ― 𝑑𝑎𝑦𝑚𝑎𝑥 2 𝑤

)}

(Eq. 2) (Eq. 3) (Eq. 4)

246

CD is the level of conjugated dienes (mM), CD0 is the CD level at the start of oxidation

247

trial, CDmax is the maximum CD level, day is the day number (independent variable),

248

daymax is the day number at which CDmax occurs and w determines the width of the

249

Gaussian curve (units are days). Corresponding parameters for Eq. 3 and 4 have the

250

same meaning; units of LHP are mM and units of Hex are M.

251

The areas under the curve (AUC) were derived by numerical integration between 0 and

252

9 days using Simpson’s rule. Gaussian curves fitted the data well in most cases with the

253

exception of day 9 hexanal concentrations, which were higher than predicted by the fit

254

line. To allow for this in AUC calculations for hexanal, Gaussian curves were integrated

255

up to 7 days, and added to the area of the trapezoid described by drawing a straight line

256

between the mean (measured) values at day 7 and day 9.



Page 12 of 44

12 257

Normalisation of AUC involved dividing by the maximum AUC for each oxidation product

258

then multiplying by 100.

259


Page 13 of 44


13 260

Results

261

Formation of droplet-stabilized and control emulsions

262

Figure 1 shows confocal microscopic images of droplet-stabilized and control

263

emulsions. Particle size of olive, palmolein and trimyristin shell emulsions was the

264

same (d3,2 = 0.4 µm and d4,3 = 0.5 µm) and Table 1 shows the particle size data of

265

droplet-stabilized, reference and control emulsions. The particle sizes of shell

266

emulsions with low-, intermediate- and high-melting surface lipids were not significantly

267

different (p>0.05). Particle sizes of droplet-stabilized, reference and control emulsions

268

were also similar. Confocal microscopy and particle size analysis were used to validate

269

the differences in interfacial structure between droplet-stabilized emulsions, control

270

emulsions of same composition as droplet-stabilized emulsions, and conventional

271

reference emulsion.

272

The interface of core safflower oil droplets in the reference emulsion (R) was coated

273

by a thin protein layer (Figure 1R). For the controls of same composition as droplet-

274

stabilized emulsions (CO, CP and CT) the interfacial structure was the same as that of

275

the reference emulsion but the shell droplets were dispersed in the surrounding phase

276

(Figure 1-CO, CP & CT). In CO, CP and CT emulsions there was some spontaneous

277

adsorption of shell droplets on to the core droplets, but much less than when shell

278

emulsion and core lipid were mixed together at high speed and elevated temperature

279

in the droplet-stabilized emulsion process.

280

Core safflower oil droplets in droplet-stabilized emulsions were stabilized by protein-

281

coated shell oil droplets, which are evident in Figure 1-NO, NP, NT & NT2 at the

282

interface of larger core safflower oil droplets, as the adsorption of small shell oil

283

droplets (red) coated with protein (green) is observed. This interfacial structure is



Page 14 of 44

14 284

similar to that reported by Ye et al. 24 for hexadecane oil-in-water emulsions stabilized

285

by micellar casein coated nanoemulsions.

286

Impact of droplet-stabilized emulsion structure on oxidative stability of core

287

safflower oil

288

Figure 2 shows the evolution of conjugated dienes, hydroperoxides and hexanal in

289

droplet-stabilized, reference and control emulsions.

290

The oxidation of core safflower oil steadily increased with >1 day of exposure to light.

291

The highest conjugated dienes and hydroperoxide levels were observed after 5 days

292

exposure, which was followed by a steady decrease, indicating decomposition of

293

oxidation products. Primary oxidation products such as lipid hydroperoxides are usually

294

stable at low temperatures and in the absence of metals but in the presence of metals

295

they decompose very fast

296

conjugated dienes and lipid hydroperoxides, some studies on oxidative stability of

297

emulsions have also shown similar trends 26, 27.

298

Hexanal increased due to the breakdown of primary oxidation products and then

299

decreased, which was unexpected. Hexanal may have reacted or bound with other

300

constituents (e.g proteins), rendering it non-volatile, which would lead to decreased

301

quantification on days 7 and 9 28. The fact that hexanal does not return to zero suggests

302

that binding/reaction sites have been saturated. A few published studies have shown a

303

similar trend 21, 29 and the kinetics of secondary product development are known to differ

304

for different oils

305

is competitive adsorption of other volatiles produced during oxidation which may have

306

limited hexanal adsorption on SPME fibre.

25.

25

so it is not surprising that we observed a rapid decrease in

Another probable explanation for the observed decrease in hexanal

307 308

There were significant differences (p < 0.05) in core safflower oil oxidation between

309

samples after 1, 3, 5, 7 and 9 days exposure. Oxidation of safflower oil in dropletACS Paragon Plus Environment

Page 15 of 44


15 310

stabilized emulsions was slower and/or less extensive than in reference emulsion and

311

composition-matched controls, for both the primary and secondary oxidation products.

312

Table 2 shows the parameters used to fit oxidation time courses in Figure 2, and the total

313

amounts of oxidation products are expressed as area under the curve (AUC) values in

314

Table 3. Table 2 shows the highest conjugated dienes (CDmax), lipid hydroperoxides

315

(LHPmax) and hexanal (Hexmax) values for each emulsion. CDmax, LHPmax and Hexmax were

316

highest in reference and controls (R, CO, CP & CT) than droplet stabilized emulsions and

317

lowest in droplet-stabilized emulsions of trimyristin surface lipid (NT & NT2) than olive

318

and palmolein oil surface lipids (NO & NP).

319 320

According to AUC values, oxidation was reduced by up to 55% by the droplet-stabilized

321

emulsion structure. All droplet-stabilized emulsions had lower values of all oxidation

322

products compared with reference and controls, which showed small differences among

323

themselves. It was clear that trimyristin was the most effective surface lipid, followed by

324

olive oil and palm oil.

325

Formation of hexanal in controls of same composition as droplet-stabilized emulsion was

326

slower than in reference emulsion (R) and this indicates that the addition of shell emulsion

327

delayed oxidation slightly, which may be due to spontaneous adsorption of protein-coated

328

shell droplets to some core droplets (Figure 3D).

329

Figure 3 and Figure 4 show the structural changes in droplet-stabilized, reference and

330

control emulsions during oxidation. Aggregation was quantified by measuring the particle

331

size distribution and microscopic structure was examined as oxidation products peaked

332

(5 days) and decreased (7 days). For 5 days exposure, core droplet aggregation was

333

observed in reference emulsion (Figure 3B). Shell droplets in control emulsions of same

334

composition as droplet-stabilized emulsion flocculated and formed a network (Figure 3E)

335

and core droplet aggregation was also observed. We did not observe structural changes



Page 16 of 44

16 336

in droplet-stabilized emulsions after 5 days exposure as the ‘droplet-stabilized structure’

337

was still evident at the interface of most core droplets (fig. 4). The changes observed at

338

7 days were same as observed at 5 days, but more aggregation was observed in each

339

case.

340

compromised interfacial structure integrity in some cases (fig. 4 I & L).

341

Figure 5 shows particle size changes for droplet-stabilized, reference and control

342

emulsions during oxidation. The observed particle size changes correspond to the

343

changes in structure because particle size of reference and control emulsions increased

344

significantly during oxidation (p < 0.05). There were no significant changes in particle size

345

of droplet-stabilized emulsions after 5 days, but after 7 days there was a significant

346

increase in droplet size.

For droplet-stabilized emulsions, shell droplet aggregation appeared to have

347 348

Effect of surface lipid type on oxidative stability of droplet-stabilized safflower oil

349

emulsion

350

Safflower oil oxidation levels in droplet-stabilized emulsions of intermediate-melting

351

(palmolein oil) surface lipid (NP) were higher than droplet-stabilized emulsions of low-

352

(NO) and high-melting surface lipids (NT & NT2). Differences between NT and NT2

353

were subtle: for the NT emulsion AUC values were lower for CD and LHP and higher

354

for hexanal, compared with NT2.

355

Higher levels of safflower oil oxidation in droplet-stabilized emulsion of medium-melting

356

surface lipid (palmolein oil) were unexpected. However, structural changes observed

357

during oxidation provide an explanation for these results. While droplet-stabilized

358

emulsions made up of low- (NO) and high-melting surface lipids (NT and NT2)

359

remained relatively stable during oxidation, that of medium melting surface lipid was

360

less stable as observed with changes in structure (Figure 4) and droplet size (Figure

361

5). More uniform coating of core safflower oil droplets was observed with proteinACS Paragon Plus Environment

Page 17 of 44


17 362

coated olive and trimyristin shell droplets than with palmolein oil shell droplets (Figure

363

4). After five days’ oxidation we observed a shift in the particle size distribution of

364

palmolein oil droplet-stabilized emulsions to larger sizes (Figure 5B and H).

365

Impact of surface lipid state on oxidative stability of droplet-stabilized emulsion

366

A significant difference (p < 0.05) in safflower oxidation levels between droplet stabilized

367

emulsions of high melting surface lipid processed above its melting temperature (NT) and

368

that processed below melting temperature (NT2) was only found after 9 days exposure

369

for conjugated dienes, hydroperoxides and hexanal analysis (Figure 2).

370 371

Our hypothesis was that for emulsions processed above trimyristin’s melting temperature,

372

trimyristin shell droplets adsorbed to safflower oil-water interface in a liquid state, while

373

for those processed below trimyristin’s melting temperature, shell droplets adsorbed in a

374

solid state. Even though the emulsions processed above melting temperature

375

subsequently cooled and remained in a supercooled state30 , the expectation was that

376

droplet-stabilized emulsions processed below melting temperature (adsorption in solid

377

state) would provide better oxidative stability than that processed above melting

378

temperature (adsorption in liquid state). For NT2 we expected that the solid matrix would

379

entrap the core lipid immediately after processing and delay oxidation, but we found no

380

significant difference (p > 0.05) in oxidation levels between these formulations except

381

after 7 d exposure. The emulsion processed above melting temperature (NT) appeared

382

to inhibit oxidation better than that processed below melting temperature (NT2) (Table 2)

383

but oxidation peak time for NT2 was longer than NT for formation of primary oxidation

384

products (CD and LHP) and shorter for secondary oxidation product (hexanal). This

385

suggests a distinct delay in oxidation with NT2 emulsions (Table 3).

386



Page 18 of 44

18 387 388 389 390

Discussion

391

Lipid oxidation in emulsions is influenced by catalysts such as singlet oxygen, metal

392

ions, light, temperature and enzymes. Oxidation by light follows two possible pathways

393

or mechanisms. In type 1 mechanism, an excited sensitizer such as chlorophyll or

394

riboflavin directly abstracts hydrogen from a fatty acid and produces a free-radical, this

395

mechanism is most likely to occur under low oxygen concentrations. While in type 2

396

mechanism when oxygen is readily available, an excited sensitizer emits or transfers

397

energy absorbed to triplet oxygen (3O2) generating singlet oxygen (1O2). Singlet oxygen

398

directly reacts with the double bonds of unsaturated fatty acids 31. In our study the most

399

probable mechanism of accelerated oxidation is the type 2 mechanism. As emulsions

400

were exposed to light in the presence of ferrous ions, we envisage accelerated

401

oxidation by oxygen and ferrous ions. Singlet oxygen (1O2) reacts with core safflower

402

oil to yield lipid hydroperoxides (LHP) and then ferrous ions catalyse decomposition of

403

hydroperoxides.

404 405

Droplet surface area, droplet charge, interfacial permeability, interfacial thickness and

406

interfacial composition are some of the factors that can impact lipid oxidation in oil-in-

407

water emulsions

408

emulsions have reported about the impact of droplet surface area and charge on

409

oxidation, it is important to emphasise that oxidation is not only influenced by these

410

factors.

32.

While many studies on the oxidative stability of protein-stabilized


Page 19 of 44


19 411 412

Reports on the effect of droplet size on oxidative stability are contradictory: some

413

studies report decreased oxidative stability with increased surface area

414

studies show the reverse trend

415

and control emulsions was not so different. Even though the particle size of reference

416

emulsion (R) was slightly higher than droplet stabilized emulsions (Table 1), it is

417

unlikely that increased oxidative stability of droplet stabilized emulsions was due to

418

droplet surface area, because if this was the predominant factor, the oxidative stability

419

of R should have been better than other controls (CO, CP & CT) and droplet-stabilized

420

emulsions or vice versa.

421

The pH of emulsions was measured after 5 and 7 days under accelerated oxidation

422

conditions. All emulsions had pH values in the range 6 to 7 (where milk proteins would

423

carry a net negative charge), except R in which pH decreased to 5.5 after five days under

424

accelerated oxidation conditions. Haahr and Jacobsen

425

peroxides and volatiles at pH 3 than pH 7 in omega-3 enriched emulsions irrespective of

426

iron addition, a similar trend was also reported for mayonnaise enriched with fish oil

427

Another study reported slower formation of peroxides and volatiles at pH 3 than pH 7 in

428

salmon oil-in-water emulsions without addition of iron

429

oxidation rates at pH 7 to low solubility of iron in the continuous phase at pH 7. In our

430

study, the pH of emulsions after homogenization was the same therefore differences in

431

oxidative stability between emulsions could not have been influenced by differences in

432

pH. The observed pH decrease in R on day 5 can be attributed to the production of short

433

chain aliphatic acids during the formation of secondary oxidation products 37

34.

33

but other

In our study, the particle size of droplet-stabilized

434


35

7

reported faster formation of

36.

and attributed the increased


Page 20 of 44

20 435

The ability of proteins to bind or chelate metals has been reported to limit lipid oxidation

436

by partitioning metal ions away from the droplet interface and reducing their reactivity 38-

437

40

438

oxidation 41. We did not quantify the amount of unadsorbed proteins in our study and so

439

it is difficult to evaluate the effect, but the amounts of protein used in droplet-stabilized

440

emulsions and composition-matched controls were similar therefore it is unlikely that the

441

presence of unadsorbed proteins in the continuous phase favoured oxidative stability of

442

droplet-stabilized emulsions over composition-matched controls. Composition-matched

443

controls (CO, CP & CT) consisted of two types of droplets: shell droplets and the core

444

emulsion droplets, both stabilized by the adsorption of MPC proteins. The actual

445

structure of MPC proteins on the droplet surface is not clearly known but likely to contain

446

casein micelle and its fragments and some whey proteins. The interfacial structures of

447

MPC proteins in shell droplets and the core emulsion droplets are likely to be similar,

448

although it is possible that casein micelles adsorbed onto the shell droplets may be more

449

dissociated due to high homogenisation pressure used in their preparation. Assuming

450

there are no interactions between shell droplets and the core emulsion droplets upon

451

gentle mixing, the proportions of shell droplets in composition-matched controls and

452

droplet-stabilized emulsions would be the same. The only expected difference would be

453

the location of the droplets. In droplet stabilized emulsions, the shell droplets stabilized

454

by MPC protein act as an emulsifying agent for the core oil, because the outer regions of

455

adsorbed casein micelles are able to adsorb and stabilize the core oil droplets as the shell

456

droplets connect with each other and form a layer along the core droplet interface as

457

reported by Ye, Zhu and Singh 18 . The actual structure and conformation of protein that

458

stabilize the core droplets is not known and being investigated in our other studies.

. The presence of unadsorbed proteins in emulsions has also been shown to limit

459


Page 21 of 44


21 460

The physicochemical properties of lipids can influence the physical stability of an

461

emulsion and consequently impact oxidative stability

462

impact of surface lipids of varying physical properties and it is possible that due to the

463

physical instability (interface thinning and partial coalescence) of palmolein oil droplet-

464

stabilized emulsions during oxidation, the droplet-stabilized interface became more

465

accessible to ferrous ions and oxygen thus promoting interactions between ferrous

466

ions or oxygen and core safflower oil which resulted in higher levels of conjugated

467

dienes, lipid hydroperoxides and hexanal recorded.

468

The presence of naturally-occurring antioxidants such as tocopherols or carotenoids

469

in oils can also inhibit oxidation in emulsions

470

products and pro-oxidants such as chlorophyll. The olive oil used here was a highly-

471

refined low-acidity product without added antioxidants. Similarly, trimyristin was free

472

from added antioxidants and as a fully saturated lipid, it is unlikely to contain oxidation

473

products at significant levels. Palmolein oil did not contain added antioxidants but may

474

have contained low levels of oxidation products and/or naturally-occurring pro- and

475

antioxidants. It is clear from Figure 2 that the initial levels of oxidation products in

476

emulsions varied very little with surface lipid. Palmolein droplet-stabilized emulsions

477

generally oxidised faster than comparable emulsions with other surface lipids, and a

478

contribution from naturally-occurring pro-oxidants in palmolein cannot be ruled out.

479

Another factor that may have contributed to the differences in oxidative stability

480

between surface lipids is their differences in hydrophobicity (low levels of polar groups)

481

which could have influenced the partitioning of ferrous ion and oxygen, we did not carry

482

out any experiments to characterise lipid hydrophobicity so it is difficult to evaluate.

14,

42.

Our study also examined the

as can naturally-occurring oxidation

483



Page 22 of 44

22 484

The physical state of lipids in emulsions can influence oxidation because it affects the

485

distribution of lipid-soluble components between the phases, diffusion rate of

486

molecules through the lipid droplets 43 and may trap the lipid within a crystalline matrix

487

making it less accessible to aqueous pro-oxidants and oxygen

488

increased oxidative stability with NT and NT2 emulsions relative to other surface lipids

489

(Figure 2, Table 2, Table 3), which may be attributable to the protective effect of droplet

490

stabilization and the solid lipid matrix formed at the interface upon trimyristin

491

crystallization.

492

Looking at the structural changes of NT and NT2 during oxidation, there appear to be

493

no marked differences in structure between these two emulsions that could have given

494

rise to the observed differences in oxidation rates; however, one possible explanation

495

can be drawn from the observed differences in structure when the emulsions are

496

freshly formed. In both emulsions, crystalline structures were observed but the

497

crystalline morphology (Figure 6) in the emulsions processed below melting

498

temperature (NT2) was different from that processed above melting temperature (NT)

499

and this morphology may have resulted in a different distribution or location of

500

trimyristin shell droplets which impacted oxidative stability of core unsaturated

501

safflower oil.

502

Overall, both emulsions (NT and NT2) showed significantly increased oxidative stability

503

over reference emulsion (R), control emulsion (CT), and droplet-stabilized emulsions of

504

low (NO) and medium (NP) melting surface lipids (Table 2, Table 3). These results show

505

that the oxidation resistance of droplet-stabilized emulsions can be greatly enhanced with

506

the use of high melting surface lipids.

507

The ability of pro-oxidant metal ions or oxygen to diffuse into the lipid phase of an

508

emulsion can be limited by making the interface less accessible and so carefully


16.

We observed

Page 23 of 44


23 509

structuring the interface of emulsions may be a good strategy that can inhibit oxidation

510

in emulsions 6. The interfacial layer of an oil-in-water emulsion plays a significant role

511

in the oxidative stability of the oil phase because it is the gateway between aqueous

512

pro-oxidant metals, oxygen and the oil phase. Once metal ions and oxygen cross the

513

interface they are able to interact with the core-unsaturated lipid 12, 44

514

The formation of cohesive protein layers by cross-linking protein with transglutaminase

515

to increase oxidative stability has been reported but, these layers were still porous and

516

did not inhibit lipid oxidation 45. Alternatively, structural design of emulsion interface to

517

make it thicker and less permeable as seen with multi-layered and Pickering emulsions

518

have been reported to increase oxidative stability

519

multi-layered emulsions inhibit oxidation by metal ions differs with that of Pickering

520

emulsions while the former depends greatly on electrostatic repulsions between the

521

charged outer layer and metal ions, the latter tends to form a rigid barrier which

522

prevents metal ions from reacting with the lipid phase.

523

Our study shows that the structured interface of droplet-stabilized emulsions provides

524

a barrier to pro-oxidant metal ions and oxygen, which can almost halve oxidation

525

compared with conventional protein-stabilized emulsions (Figure 2 and Table 3).

526

Proteins are reported to stabilize emulsions by forming thick interfaces which can

527

retard oxidation

528

protein-coated lipid droplets in a somewhat aggregated network and this appears to

529

form a protective interface that may be responsible for the increased oxidative stability

530

observed. It is also possible that this protective effect was further enhanced by the

531

adsorption of any free non-adsorbed proteins to the interfaces of core oil droplets not

532

fully covered or coated by shell droplets.

21, 49.

44, 46-48.

The mechanism by which

In droplet-stabilized emulsions, the interface is covered by



Page 24 of 44

24 7

533

According to Mancuso, McClements and Decker

increased pH implies a decrease in

534

iron solubility in the continuous phase, causing it to precipitate at the droplet interface and

535

promote oxidation. If ferrous ion partitioned at the interface of emulsions and promoted

536

oxidation in controls but oxidation was less extensive in droplet-stabilized emulsions, we

537

hypothesise that the lipid droplet interface (shell droplets) in droplet-stabilized emulsions

538

limited the proximity between ferrous ion and lipid hydroperoxides, as illustrated in Figure

539

7.

540

We also hypothesise that the lipid droplet interface in droplet-stabilized emulsions may

541

have influenced partitioning of pro-oxidant metal ions away from the droplet interface

542

and limited interactions that promote oxidation.

543

In conclusion, safflower oil oxidation in droplet-stabilized emulsion was significantly

544

slower and/or less extensive than in composition-matched conventional protein-

545

stabilized emulsions. These results indicate that the structured interface of droplet-

546

stabilized emulsion limits contact between pro-oxidant metals, oxygen and lipid

547

substrate. Addition of shell droplets to conventional emulsions delayed formation of

548

secondary oxidation products which also shows the impact of protein-coated shell

549

droplet stabilization. These results confirm the oxidation resistance of droplet-stabilized

550

emulsions. Oxidative stability of safflower oil in droplet-stabilized emulsions was

551

greatly enhanced with protein-coated high-melting surface lipid, both when the

552

emulsions were processed above and below the surface lipid’s melting temperature.

553

These results show that oxidation resistance of droplet-stabilized emulsions can also

554

be improved by changing the interfacial composition.

555

This study provides critical knowledge for the functional food industry about plausible

556

applications and functional properties of droplet-stabilized emulsions.

557


Page 25 of 44


25 558 559 560 561 562 563

Acknowledgements

564 565 566 567

This research was funded by the Riddet Institute, a national Centre of Research Excellence, funded by the New Zealand Tertiary Education Commission. PW gratefully acknowledges support from the Biotechnology and Biological Sciences Research Council (UK).

568

Supporting Information

569 570

Further images of droplet-stabilized trimyristin emulsions NT and NT2. Images showing structural changes in emulsions during the oxidation trials.

571

References

572 573 574 575 576 577 578 579 580 581 582 583 584 585 586 587 588 589 590 591 592 593 594 595 596 597 598

1. Mozaffarian, D.; Wu, J. H. Y., Omega-3 Fatty Acids and Cardiovascular Disease: Effects on Risk Factors, Molecular Pathways, and Clinical Events. Journal of the American College of Cardiology 2011, 58, 2047-2067. 2. Yeong, L. H.; So Bong, J., Effects of conjugated linoleic acid on body fat reduction and physical exercise enhancement of obese male middle school students. Korean Journal of Life Science 2010, 20, 1844-1850. 3. Djordjevic, D.; McClements, D. J.; Decker, E. A., Oxidative Stability of Whey Protein-stabilized Oilin-water Emulsions at pH 3: Potential ω-3 Fatty Acid Delivery Systems (Part B). Journal of Food Science 2004, 69, C356-C362. 4. Frankel, E. N.; Satue-Gracia, T.; Meyer, A. S.; German, J. B., Oxidative stability of fish and algae oils containing long-chain polyunsaturated fatty acids in bulk and in oil-in-water emulsions. Journal of Agricultural and Food Chemistry 2002, 50, 2094-2099. 5. Katsuda, M. S.; McClements, D. J.; Miglioranza, L. H. S.; Decker, E. A., Physical and Oxidative Stability of Fish Oil-in-Water Emulsions Stabilized with β-Lactoglobulin and Pectin. Journal of Agricultural and Food Chemistry 2008, 56, 5926-5931. 6. McClements, D. J.; Decker, E. A., Lipid Oxidation in Oil-in-Water Emulsions: Impact of Molecular Environment on Chemical Reactions in Heterogeneous Food Systems. Journal of Food Science 2000, 65, 1270-1282. 7. Mancuso, J. R.; McClements, D. J.; Decker, E. A., The Effects of Surfactant Type, pH, and Chelators on the Oxidation of Salmon Oil-in-Water Emulsions. Journal of Agricultural and Food Chemistry 1999, 47, 4112-4116. 8. Yoshida, Y.; Niki, E., Oxidation of methyl linoleate in aqueous dispersions induced by copper and iron. Archives of Biochemistry and Biophysics 1992, 295, 107-114. 9. McClements, D. J., Emulsion Design to Improve the Delivery of Functional Lipophilic Components. Annual Review of Food Science and Technology 2010, 1, 241-269. 10. Berton-Carabin, C. C.; Sagis, L.; Schroën, K., Formation, Structure, and Functionality of Interfacial Layers in Food Emulsions. Annual Review of Food Science and Technology 2018, 9, 551-587.



Page 26 of 44

26 599 600 601 602 603 604 605 606 607 608 609 610 611 612 613 614 615 616 617 618 619 620 621 622 623 624 625 626 627 628 629 630 631 632 633 634 635 636 637 638 639 640 641 642 643 644 645 646 647 648 649 650 651 652

11. Lesmes, U.; Sandra, S.; Decker, E. A.; McClements, D. J., Impact of surface deposition of lactoferrin on physical and chemical stability of omega-3 rich lipid droplets stabilized by caseinate. Food Chemistry 2010, 123, 99-106. 12. Silvestre, M. P. C.; Chaiyasit, W.; Brannan, R. G.; McClements, D. J.; Decker, E. A., Ability of Surfactant Headgroup Size To Alter Lipid and Antioxidant Oxidation in Oil-in-Water Emulsions. Journal of Agricultural and Food Chemistry 2000, 48, 2057-2061. 13. Berton, C.; Ropers, M.-H.; Viau, M.; Genot, C., Contribution of the interfacial layer to the protection of emulsified lipids against oxidation. Journal of Agricultural and Food Chemistry 2011, 59, 5052-5061. 14. Berton-Carabin, C. C.; Ropers, M.-H.; Genot, C., Lipid Oxidation in Oil-in-Water Emulsions: Involvement of the Interfacial Layer. Comprehensive Reviews in Food Science and Food Safety 2014, 13, 945-977. 15. Komaiko, J.; Sastrosubroto, A.; McClements, D. J., Encapsulation of ω-3 fatty acids in nanoemulsion-based delivery systems fabricated from natural emulsifiers: Sunflower phospholipids. Food Chemistry 2016, 203, 331-339. 16. McClements, D. J.; Decker, E. A.; Weiss, J., Emulsion-Based Delivery Systems for Lipophilic Bioactive Components. Journal of Food Science 2007, 72, R109-R124. 17. Walker, R.; Decker, E. A.; McClements, D. J., Development of food-grade nanoemulsions and emulsions for delivery of omega-3 fatty acids: opportunities and obstacles in the food industry. Food & Function 2015, 6, 41-54. 18. Ye, A.; Zhu, X.; Singh, H., Oil-in-Water Emulsion System Stabilized by Protein-Coated Nanoemulsion Droplets. Langmuir 2013, 29, 14403-14410. 19. Singh, H.; Zhu, X. Q.; Ye, A. Lipid encapsulation. patent US2009/0029017, 2009. 20. Gallier, S.; Gragson, D.; Jimenez-Flores, R.; Everett, D., Using Confocal Laser Scanning Microscopy To Probe the Milk Fat Globule Membrane and Associated Proteins. Journal of Agricultural and Food Chemistry 2010, 58, 4250-4257. 21. Hu, M.; McClements, D. J.; Decker, E. A., Lipid Oxidation in Corn Oil-in-Water Emulsions Stabilized by Casein, Whey Protein Isolate, and Soy Protein Isolate. Journal of Agricultural and Food Chemistry 2003, 51, 1696-1700. 22. Wrolstad, E. R.; Acree, E. T.; Decker, E. A.; Penner, M. H.; Reid, D. S.; Schwartz, J. S.; Shoemaker, C. F.; Smith, D.; Sporns, P., Handbook of Food Analytical chemistry. John wiley & sons Inc.: New Jersey, 2005; p 515-517. 23. Estévez, M.; Kylli, P.; Puolanne, E.; Kivikari, R.; Heinonen, M., Fluorescence spectroscopy as a novel approach for the assessment of myofibrillar protein oxidation in oil-in-water emulsions. Meat Science 2008, 80, 1290-1296. 24. Ye; Zhu; Singh, Oil-in-Water Emulsion System Stabilized by Protein-Coated Nanoemulsion Droplets. Langmuir 2013, 29, 14403-14410. 25. Choe, E.; Min, D. B., Mechanisms and Factors for Edible Oil Oxidation. Comprehensive Reviews in Food Science and Food Safety 2006, 5, 169-186. 26. O’ Dwyer, S. P.; O’ Beirne, D.; Eidhin, D. N.; O’ Kennedy, B. T., Effects of sodium caseinate concentration and storage conditions on the oxidative stability of oil-in-water emulsions. Food Chemistry 2013, 138, 1145-1152. 27. Ponginebbi, L.; Nawar, W. W.; Chinachoti, P., Oxidation of linoleic acid in emulsions: Effect of substrate, emulsifier, and sugar concentration. Journal of the American Oil Chemists' Society 1999, 76, 131. 28. Shahidi, F.; Zhong, Y., Lipid Oxidation: Measurement Methods. In Bailey's Industrial Oil and Fat Products, John Wiley & Sons, Inc.: 2005. 29. Iglesias, J.; Lois, S.; Medina, I., Development of a solid-phase microextraction method for determination of volatile oxidation compounds in fish oil emulsions. Journal of Chromatography A 2007, 1163, 277-287. 30. Westesen, K.; Bunjes, H., Do nanoparticles prepared from lipids solid at room temperature always possess a solid lipid matrix? International Journal of Pharmaceutics 1995, 115, 129-131. 31. Kanofsky, R. J., Photosensitization. In Singlet Oxygen: Applications in Biosciences and Nanosciences, Nonell, S.; Flors, C., Eds. Royal Society of Chemistry: Cambridge, UK, 2016; pp 93-103. ACS Paragon Plus Environment

Page 27 of 44


27 653 654 655 656 657 658 659 660 661 662 663 664 665 666 667 668 669 670 671 672 673 674 675 676 677 678 679 680 681 682 683 684 685 686 687 688 689 690 691 692 693 694 695 696 697

32. Waraho, T.; McClements, D. J.; Decker, E. A., Mechanisms of lipid oxidation in food dispersions. Trends in Food Science & Technology 2011, 22, 3-13. 33. Lee, S. J.; Choi, S. J.; Li, Y.; Decker, E. A.; McClements, D. J., Protein-Stabilized Nanoemulsions and Emulsions: Comparison of Physicochemical Stability, Lipid Oxidation, and Lipase Digestibility. Journal of Agricultural and Food Chemistry 2011, 59, 415-427. 34. Nakaya, K.; Ushio, H.; Matsukawa, S.; Shimizu, M.; Ohshima, T., Effects of droplet size on the oxidative stability of oil-in-water emulsions. Lipids 2005, 40, 501-507. 35. Haahr, A. M.; Jacobsen, C., Emulsifier type, metal chelation and pH affect oxidative stability of n-3-enriched emulsions. European Journal of Lipid Science and Technology 2008, 110, 949-961. 36. Jacobsen, C.; Timm, M.; Meyer, A. S., Oxidation in Fish Oil Enriched Mayonnaise: Ascorbic Acid and Low pH Increase Oxidative Deterioration. Journal of Agricultural and Food Chemistry 2001, 49, 39473956. 37. Dimakou, C. P.; Kiokias, S. N.; Tsaprouni, I. V.; Oreopoulou, V., Effect of Processing and Storage Parameters on the Oxidative Deterioration of Oil-in-Water Emulsions. Food Biophysics 2007, 2, 38. 38. Caetano-Silva, M. E.; Mariutti, L. R. B.; Bragagnolo, N.; Bertoldo-Pacheco, M. T.; Netto, F. M., Whey Peptide-Iron Complexes Increase the Oxidative Stability of Oil-in-Water Emulsions in Comparison to Iron Salts. Journal of Agricultural and Food Chemistry 2018, 66, 1981-1989. 39. Guzun-Cojocaru, T.; Koev, C.; Yordanov, M.; Karbowiak, T.; Cases, E.; Cayot, P., Oxidative stability of oil-in-water emulsions containing iron chelates: Transfer of iron from chelates to milk proteins at interface. Food chemistry 2011, 125, 326-333. 40. Sugiarto, M.; Ye, A.; Taylor, M. W.; Singh, H., Milk protein-iron complexes: Inhibition of lipid oxidation in an emulsion. Dairy Science & Technology 2010, 90, 87-98. 41. Faraji, H.; McClements, D. J.; Decker, E. A., Role of Continuous Phase Protein on the Oxidative Stability of Fish Oil-in-Water Emulsions. Journal of Agricultural and Food Chemistry 2004, 52, 4558-4564. 42. McClements, D. J., Food Emulsions : Principles, Practices, and Techniques, Third Edition. Chapman and Hall/CRC: Boca Raton, UNITED KINGDOM, 2015. 43. Okuda, S.; McClements, D. J.; Decker, E. A., Impact of Lipid Physical State on the Oxidation of Methyl Linolenate in Oil-in-Water Emulsions. Journal of Agricultural and Food Chemistry 2005, 53, 96249628. 44. Kargar, M.; Spyropoulos, F.; Norton, I. T., The effect of interfacial microstructure on the lipid oxidation stability of oil-in-water emulsions. Journal of Colloid and Interface Science 2011, 357, 527-533. 45. Kellerby, S. S.; Gu, Y. S.; McClements, D. J.; Decker, E. A., Lipid Oxidation in a Menhaden Oil-inWater Emulsion Stabilized by Sodium Caseinate Cross-Linked with Transglutaminase. Journal of Agricultural and Food Chemistry 2006, 54, 10222-10227. 46. Gudipati, V.; Sandra, S.; McClements, D. J.; Decker, E. A., Oxidative Stability and in Vitro Digestibility of Fish Oil-in-Water Emulsions Containing Multilayered Membranes. Journal of Agricultural and Food Chemistry 2010, 58, 8093-8099. 47. Klinkesorn, U.; Sophanodora, P.; Chinachoti, P.; McClements, D. J.; Decker, E. A., Increasing the Oxidative Stability of Liquid and Dried Tuna Oil-in-Water Emulsions with Electrostatic Layer-by-Layer Deposition Technology. Journal of Agricultural and Food Chemistry 2005, 53, 4561-4566. 48. Ogawa, S.; Decker, E. A.; McClements, D. J., Influence of Environmental Conditions on the Stability of Oil in Water Emulsions Containing Droplets Stabilized by Lecithin−Chitosan Membranes. Journal of Agricultural and Food Chemistry 2003, 51, 5522-5527. 49. Fang, Y.; Dalgleish, D. G., Dimensions of the Adsorbed Layers in Oil-in-Water Emulsions Stabilized by Caseins. Journal of Colloid and Interface Science 1993, 156, 329-334.

698 699 700 701 702 ACS Paragon Plus Environment


Page 28 of 44

28 703

Table 1: Average particle size of droplet-stabilized and control emulsions

Emulsions

R

Volume weighted

mean diameter,

mean diameter,

D3,2 (μm)

D4,3 (µm)

None

13.2 ± 2.5c

15.3 ± 3.0c

Surface lipid

protein stabilized safflower oil-in water emulsion (reference emulsion)

CO

R with added shell emulsion

Olive oil

7.1 ± 0.3a

14.6 ± 1.1a

CP


Palmolein oil

7.1 ± 0.1a

14.0 ± 2.2a

CT


Trimyristin

6.7 ± 0.2a

15.0 ± 1.6a

NO

droplet-stabilized emulsion droplet-stabilized emulsion

Olive oil

8.9 ± 0.6b

15.2 ± 2.8b

Palmolein oil

8.6 ± 0.3b

13.3 ± 1.3b

Trimyristin

9.0 ± 1.0b

15.8 ± 4.9b

Trimyristin

9.5 ± 1.8b

21.0 ± 2.6d

NP

NT

NT2

704 705 706 707 708 709 710 711 712

Description

Surface-weighted

droplet-stabilized emulsion processed at temperature above 56°C droplet-stabilized emulsion processed at temperature below 56°C

Values are means of three measurements (3 runs per measurement) ± standard deviation. Different superscript letters (a, b, c) represent different treatments. Means with same superscript are not significantly different (p>0.05). Core lipid in all emulsions= safflower oil; R= protein-stabilized oil-in-water emulsion (reference emulsion); CO= R with added olive oil shell emulsion; CP= R with added palmolein oil shell emulsion; CT= R with added trimyristin shell emulsion; NO= droplet stabilized emulsion with olive oil shell droplets; NP= droplet stabilized emulsion with palmolen oil shell droplets; NT= droplet stabilized with trimyristin shell droplets processed above 56°C; NT2= droplet stabilized emulsion with trimyristin shell droplets processed below 56°C

713


Page 29 of 44


29 714 715

716

717

Table 2: Oxidation parameters derived from fitting equations 2, 3 and 4 to oxidation data. Values in brackets are standard errors. Conjugated Dienes (CD)

CD0 (mM)

CDmax (mM)

daymax

w

R

8.6 (1.3)a

50.0 (2.7)a

5.3 (0.1)bc

1.7 (0.1)b

CO

7.7 (1.5)a

43.0 (2.9)ab

5.2 (0.1)cd

1.7 (0.1)b

CP

8.1 (1.2)a

42.0 (2.2)ab

4.8 (0.1)d

1.9 (0.1)b

CT

5.7 (1.4)a

47.5 (2.6)ab

5.2 (0.1)c

1.8 (0.1)b

NO

4.4 (1.1)a

24.2 (1.7)c

5.7 (0.1)b

2.3 (0.2)ab

NP

6.4 (0.7)a

36.0 (1.4)c

5.2 (0.1)c

1.8 (0.1)b

NT

4.0 (0.7)a

19.0 (1.3)c

5.2 (0.1)b

2.0 (0.2)b

NT2

3.2 (1.6)a

18.3 (1.7)c

6.4 (0.2)a

3.4 (0.5)a

Superscript letters indicate significant differences (P

Droplet-stabilized oil-in-water emulsions protect unsaturated lipids

Recommend Documents