Fabrication and Characterization of Quinoa Protein Nanoparticles

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Fabrication and Characterization of Quinoa Protein Nanoparticles Stabilized Food-grade Pickering Emulsions with Ultrasound Treatment: Interfacial Adsorption/Arrangement Properties Xin-Sheng Qin, Zhigang Luo, and Xichun Peng J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.8b00225 • Publication Date (Web): 17 Apr 2018 Downloaded from http://pubs.acs.org on April 17, 2018

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Journal of Agricultural and Food Chemistry

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ACS Paragon Plus Environment


Fabrication and Characterization of Quinoa Protein Nanoparticles Stabilized Food-grade Pickering Emulsions with Ultrasound Treatment: Interfacial Adsorption/Arrangement Properties

Xin-Sheng Qin,† Zhi-Gang Luo,*,†,‡ and Xi-Chun Peng§

†

School of Food Science and Engineering, South China University of Technology, Guangzhou

510640, China ‡

Guangdong Province Key Laboratory for Green Processing of Natural Products and Product

Safety, South China University of Technology, Guangzhou 510640, China §

Department of Food Science and Engineering, College of Science and Engineering, Jinan

University, Guangzhou 510632, China

1


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ABSTRACT: The natural quinoa protein isolate (QPI) was largely reflected in the

2

nanoparticle form at pH 7.0 (about 401 nm), and the ultrasound at 20 min progressively

3

improved the contact angle (wettability) and surface hydrophobicity of the nanoparticles.

4

Ultrasound process also modified the type of intra-particle interaction, and the internal forces

5

of sonicated particles were largely maintained by both disulfide bonds and hydrophobic

6

interaction forces. In emulsion system, the ultrasound progressively increased the

7

emulsification efficiency of the QPI nanoparticles, particularly at high protein concentration

8

(c>1%, w/v), and higher emulsion stability against coalescence. As compared with the natural

9

QPI stabilized emulsions, the 20 min-sonicated emulsions exhibited higher packing and

10

adsorption of at the protein interface. The microstructure of emulsions is occurred bridging

11

flocculation of droplets at low c (≤1%, w/v), while the amount of protein particles could be

12

high enough to cover the droplets surface at high c (>1%, w/v) with hexagonal array model

13

arrangement. Thus, these results illustrated that both natural and sonicated QPI nanoparticles

14

could be performed as effective food-grade stabilizer for Pickering emulsion, however, the

15

sonicated QPI nanoparticles exhibited much better emulsifying and interfacial properties.

16

KEYWORDS: quinoa protein nanoparticles, food-grade, Pickering emulsions, ultrasonic

17

treatment, interfacial property

18 19 20 21 2



22

INTRODUCTION

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Emulsion droplets can be stabilized by either forming physical screen via surface-active

24

colloidal particles, or reducing interfacial tension through surfactants, or by forming spatial

25

interfacial films via hydrocolloids.1 Stabilization of emulsions realized by the first sort is

26

defined as Pickering emulsions.2 To show as a Pickering emulsion stabilizer, surface-active

27

particles could maintain steady in water and oil systems, and have a suitable wettability

28

(contact angle).3 There is increasing interest in Pickering emulsions, because of their

29

superiority of “surfactant-free”, favorable restores to flocculation, functionality in

30

environmental responsive emulsions of high internal phase emulsions.4 In recent years,

31

synthetic or inorganic micro/nano-particles, such as carbon nanotubes, SiO2 and laponite clay,

32

seem to be most promising to apply for Pickering emulsion stabilizer.5 However, the usage of

33

these stabilizer are restricted by their non-compatibility or non-biodegradability. Hence, the

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developments of biodegradable food-grade particles based Pickering emulsion are of

35

importance in foods and pharmaceutics fields, e.g., controlled release of some embedded

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material, and reinforcement of stability.6 The food-grade Pickering stabilizer could be

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probably classified into three types: (i) protein-type, e.g., preheated soy protein, whey protein,

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water-insoluble zein and pea protein; (ii) polysaccharide-type, e.g., starch nanocrystals,

39

hydrophobically-modified starch, cellulose and chitin; (iii) miscellaneous, e.g., lipid and

40

flavonoids.3 Many particles generally need a further chemical modification to facilitate their

41

emulsifying properties, and moreover, the creating emulsions have large droplet sizes.7 For

42

example, the formation of starch particle stabilized Pickering emulsion need to 3


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hydrophobically-modify using chemical reagent such as octenyl succinic anhydride (OSA).3

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Thus, it would be an interesting problem to find an effective and cheap food-grade

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nanoparticles to stabilize the Pickering emulsion.

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Quinoa is an Andean cereal of South America that has been planting all over the world

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recently for their abundant nutritional property, mainly owing to the high content of proteins

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that ranges from 12% to 23%.8 Quinoa protein isolate (QPI) is one of the most promising and

49

readily available quinoa products due to their high content of essential amino acids. It largely

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consists of 11S globulins (20–25 kDa for basic polypeptides and 30–40 kDa for acid

51

polypeptides) and 2S albumins (8–9 kDa), which in proportion as probably 75% of the overall

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protein amount of QPI.9 Our previous work had indicated that the natural QPI was largely

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reflected in the nanoparticle form at pH 7.0. Compared with the stabilizers, the QPI

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nanoparticles have several advantages: (i) they are functional and nutritional food stuff; (ii)

55

they can perform surface hydrophobic and/or hydrophilic property; (iii) they do not demand

56

any chemical surface modification to facilitate the particle wettability and hydrophobicity; (iv)

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they are compatible with emulsification techniques (high-speed shear/ultrasound/high pressure

58

homogenizer). In conclusion, QPI nanoparticles show good surface active nature with the

59

potentiality of providing as a promising and effective stabilizer for food-grade Pickering

60

emulsions.10

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The ultrasound process is not only correlated to particle surface property, but also affects

62

the emulsion efficiency. 11,12 Ultrasound technology applied waves at a frequency >16 kHz

63

over the damage of human hearing. The acoustic cavitation phenomenon of high intensity 4



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ultrasound (10–100 W/cm2 of power, 16–100 kHz of frequency) is utilized to destroy physical

65

bonds or facilitate chemical reactions in ingredients, thereof cavitation bubbles are quickly

66

produced and fiercely collapsed.13 The bubbles combination generates excessive pressure and

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temperature that leads to turbulence and high shear stress waves in the sonicated cavitation

68

region.14 Ultrasound treatment led to decreased size but improved solubility and surface

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hydrophobicity (Ho) of the soy protein isolate aggregates solutions.15-17

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To date, no work had explored in the emulsifying nature, adsorption and arrangement at

71

the interface of QPI nanoparticles stabilized Pickering emulsion. The objective of this work

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was to elucidate the promising potential of QPI nanoparticles to act as an effective food-grade

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Pickering-type emulsion stabilizer. Firstly, we explored the effect of the ultrasound process on

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the hydrodynamic diameter, contact angle, surface hydrophobicity and interactive forces of

75

QPI nanoparticles. Secondly, the effect of protein concentration (c; 0.25–6%, w/v) and/or corn

76

oil fraction (φ; 0.2–0.7) on the interfacial properties and stability of QPI nanoparticles

77

stabilized Pickering emulsion was determined by z-average diameters (Dz), microstructure,

78

creaming stability and adsorption/arrangement of surface particles. The correlation of varied c

79

and/or φ was further studied to preferable achieve the stabilization mechanism of QPI

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nanoparticles stabilized Pickering emulsion.

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MATERIALS AND METHODS

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Materials. QPI (80% protein) was acquired from Yuanye Biological Technology Co., Ltd.,

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China. Corn oil was acquired from Yihai Kerry limited, China. Potassium Sorbate was

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acquired from Kemiou Chemical Reagent Co., Ltd., China. Coomassie brilliant blue G-250,

89

1-anilino-8-naphthalene-sulfonate (ANS), bovine serum albumin (BSA), DL-Dithiothreitol

90

(DTT), urea, sodium dodecyl sulfate (SDS), Nile Blue and Nile Red were acquired from

91

Solarbio Science & Technology Co., Ltd, China. All chemical reagents were of analytical

92

grade.

93

Fabrication of QPI Nanoparticle Dispersions with Ultrasound Treatment. The QPI

94

dispersions (0.25–6%, w/v) were fabricated by immediately adding different ratio QPI

95

powders in distilled water under magnetic agitation for 2 h at 25 °C. Potassium Sorbate

96

(0.01%, w/v) served as an antimicrobial reagent. The protein dispersions were deployed to pH

97

7.0 using HCl or NaOH and stored at 4 °C for proteins hydration. The dispersions were treated

98

using a 100 W ultrasonic homogenizer model KH-250DE (Kunshan Ultrasonic Instrument

99

CO., LTD., China) at 25 °C for 20 min. And the blank control samples were conducted

100

without ultrasound treatment.

101

z-average Diameter. The z-average diameters (Dz) of QPI nanoparticles were evaluated

102

employing a Nano-ZS Zetasizer analyzer (Malvern Instruments, UK) by dynamic light

103

scatting (DLS) method. The QPI nanoparticle dispersions were diluted in deionized water at a

104

proportion of 1:5 (v/v) after stored at 4 °C overnight. Sample was measured at 25 °C and 1.33

105

of refractive index in triplicate. 6



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Surface Hydrophobicity (Ho). Surface hydrophobicity of non- or sonicated QPI samples

107

was measured based on the method of Qin et al.18 with the fluorescence probe ANS. The QPI

108

sample was attenuated to the concentration with the range of 0.1–0.0005 mg/mL with 0.01 M

109

phosphate buffer (pH = 7.0). Then, 20 µL of 8.0 mM ANS solutions was added to 3 mL of

110

each attenuated sample, and avoid light reaction for 20 min. The fluorescence intensity (FI)

111

was determined at 390 nm (excitation wavelength) and 400–750 nm (emission wavelength)

112

employing F-7000 fluorescence spectrophotometer (Hitachi Ltd., Japan). The excitation and

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emission slit widths were taken to 5 nm, and the temperature was maintained at 25 °C. Ho was

114

calculated as the slope of the FI as the protein concentration (mg/mL).

115

Evaluation of Intra-particle Interaction. The intra-particle interaction was assessed by

116

comparing the turbidity of QPI dispersions (c = 0.25%, w/v) described by Zhang and others.19

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The blank and ultrasound treated suspensions were mixed in four solvents: A) distilled water

118

(DW), B) 6.0 M urea, C) 0.5% SDS (w/v), D) 30 mM DTT, or their combinations with an

119

equal bulk. The suspensions turbidity was determined at 595 nm employing a 722SP visible

120

spectrophotometer (Shanghai Lengguang Technology CO., LTD., China) with BSA as the

121

standard by the Bradford method. The Coomassie brilliant blue were determined as blank

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controls.

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Contact Angle (θ). The θ of freeze-dried QPI samples was measured using OCA 40 micro

124

analyzer (Data Physics Corporation, Germany) loaded with a high-speed video camera. The

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sonicated protein samples were formed into 2 mm slice with a hydraulic press. The slice was

126

immersed in the oil phase and placed into a glass vessel, and a glob of Milli-Q water (3 µL) 7


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was dripped on the superficies of the slice with corn oil employing a high-precision syringe.

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Then, the contact surface image was obtained with a high-speed camera.

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Preparation of QPI Stabilized Pickering Emulsion. The QPI Pickering emulsions were

130

fabricated after the fabrication of QPI nanoparticle dispersions (0.25–6%, w/v) with

131

ultrasound treatment. And the emulsions were conducted with variation in corn oil fraction (φ)

132

from 0.2 to 0.7, while dispersed with a T25 digital Ultra homogenizer (IKA Inc., Germany) at

133

20000 rpm for 1 min. The emulsions formed freshly were immediately applied to

134

emulsification determination, or stored for 7 days for further emulsion physical stability

135

determination.

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Droplet Size Distribution. The surface- or volume- mean diameter (D32 and D43) and

137

droplet size distribution of various QPI stabilized Pickering emulsions, were measured

138

employing a Malvern Mastersizer analyzer 2000 (Malvern Instruments, UK).13 The samples

139

were scattered in deionized water at 2500 rpm of pump speed and 0.001 absorption index. The

140

comparative refractive index of the emulsions was set as 1.095, due to the proportion of the

141

refractive index of corn oil (1.467) to that of deionized water (1.33). All determinations were

142

taken the average of three determinations.

143

Creaming Index (CI) of Emulsion Stability. CI of emulsions formed at varying cases used

144

to evaluate the stability commonly. 20 mL of each emulsion sample was transferred to a scale

145

tube (10 cm height × 2 cm diameter) and then depot at room temperature. Height of the serum

146

(Hs) and total height (Ht) of emulsions were measured before or after storage (7 days). The

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proportion of CI% was defined as ( Hs / Ht ) × 100%. 8



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Microstructure of Emulsions. Optical Microscopy. The microstructure of QPI

149

nanoparticles stabilized emulsions was measured using a BH2 light microscope analyzer

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(Olympus Co. Ltd., Japan) outfitted with a photographic camera. A 50 µL aliquot of each

151

emulsion was taken and transferred onto a glass slide, then coated with a coverslip. The

152

measurements were performed at room temperature with 500 magnification lens, as described

153

by Gao et al.20

154

Confocal Laser Scanning Microscopy (CLSM). The microstructure of protein adsorbed to

155

QPI Pickering emulsion interface was observed by CLSM using TCS-SP5 microscope

156

analyzer (Leica Microsystem Inc. Germany). Emulsions were pre-stained with Nile Blue A

157

and Nile Red during the emulsion fabrication process. Then, samples were transferred onto a

158

glass slide and coated with a coverslip. Silicone oil was used to avoid water evaporation. The

159

samples were explored by an argon/krypton laser at an excitation wavelength of 488 and 633

160

nm (HeNe laser) with a 100 magnification lens, respectively.

161

Percentage of Adsorbed Proteins (AP%). The amount of QPI proteins adsorbed to the

162

interface of the droplets was evaluated according to Isaschar-Ovdat et al.21 with some

163

modifications. A 5 mL QPI Pickering emulsion was centrifuged for 10 min at 4000 rpm

164

(Hitachi Ltd., Japan). Two layers were obtained after centrifugation: the aqueous layer on

165

bottom, the creamed oil droplet layer of the emulsion at the top of the tube. The clarified

166

aqueous phase was extracted by injector and then colated using a 0.45-mm aqueous film. Cf is

167

the protein concentration in the filtered aqueous layer with the Bradford method using BSA

9


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reagent. Co is the initial protein concentration in the protein dispersions. The AP% of the

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emulsion was calculated as follows:

170

AP% = (Co – Cf) × 100%/Co

171

Surface Particle Coverage at the Interface of Droplets. Limited coalescence process

172

occurs in Pickering emulsions where solid particles are inconvertible adsorbed at the droplets

173

interface. When the matrix is conducted with a small number of solid particles, the particles is

174

inadequate to protect the newly formed droplets. After the homogeneity, the total particle

175

amount of droplets interface is reduced because of the droplets coalescence. Since the solid

176

particles are inconvertibly anchored, the droplet coalescence course stops when the interface is

177

competent coated.22,23 Particles arrangement at the oil-water interface could be indicated from

178

the surface particle coverage (SC%), which is defined as the proportion between the droplets

179

interfacial area (Sint) and the total particle area (Seq). The total particle area can be

180

approximately calculated from their equatorial area: Seq = nparticlesπ(Dz/2)2, where nparticles was

181

the total amount of solid particles and Dz was the hydrodynamic diameter. The nparticles was

182

calculated by the adsorbed proteins (AP%) at the interface. The Sint is directly connected to

183

D32 (Sint = 6Vd/D32), where Vd is the oil capacity. Hence, the surface particle coverage SC% =

184

Seq/Sint. The SC% can be deduced from the slope of (1/D32) against (Seq/Vd).

(1)

185

Statistical Analysis. The determinations presented as the average of three samples ±

186

standard deviations. The data statistics were determined employing SPSS software (p < 0.05)

187

by ANOVA.

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RESULTS AND DISCUSSION

190

Characterization of QPI Nanoparticles by Ultrasound Treatment. The size, surface

191

hydrophobicity and wettability of particle in the natural and sonicated (at 20 min) QPI

192

dispersions at c = 0.25% (w/v) were measured by dynamic light scattering, fluorescence probe

193

and oil-in-water contact angle. Figure 1 shows the Dz of QPI dispersions changed remarkably

194

with the increasing ultrasound time. The Dz of the natural QPI nanoparticles was 401 nm,

195

while it significantly decreased to 207 nm for 20 min-sonicated sample. The results illustrated

196

that the ultrasound treatment markedly reduced particle size because of the micro-steaming

197

and turbulent forces by cavitation effect. A similar result about the effect of ultrasound on the

198

protein particles has been obtained by Gao et al. which was well consistent with the fact that

199

the ultrasound treatment usually led to rapidly production and violently sink of cavitation

200

vesicles.20,24

201

The surface characteristics of Pickering stabilizer are very important for their emulsifying

202

property. Such as organic and inorganic particles, including starch granules, microcrystalline

203

cellulose, or solid lipid nanoparticles, this particles generally need surface hydrophobic

204

modification.4,5,7 Surface hydrophobicity (Ho) also plays a vital part in the conformation,

205

function and stability of proteins. Figure 2 displays the typical intrinsic fluorescence emission

206

profiles of the natural and 20 min-sonicated QPI nanoparticles. The principle of fluorescence

207

probe ANS is as follows: the fluorescence spectra of QPI samples are very susceptive to the

208

hydrophobic Trp or Tyr residues chromophores in the polar environment.25 The Ho values are

209

calculated by the spectra and slope of FI as protein concentration in Figure 2. Ho values of the 11


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natural and sonicated QPI samples are also summarized and presented in Figure 1. The Ho of

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the 20 min-sonicated QPI dispersion displayed a remarkable increase contrasted with the

212

natural sample (Ho increased from 22 to 44). These results were attributed to the cavitation

213

phenomenon of ultrasound, which could induce unfolding of the QPI nanoparticles, lead to the

214

exposure of hydrophobic regions from the interior to the exterior of QPI nanoparticles.26

215

The wettability of the solid particles plays an important part for the fabrication of

216

Pickering-stabilized emulsions. The wettability of particles was usually determined by the

217

contact angle θ. The θ value close to 90° can favor the packing behavior of particles at the

218

droplet interface.6 For a spherical particle, the energy obtained for separating the solid

219

particles from oil/water interface (∆E), which count as follows: ∆E = πR2γ(1±cosθ)2, where γ

220

is the particle surface tension. The particles at the droplet interface can be inconvertible

221

adsorbed to oil droplets if 30° 1%, w/v), the number of QPI particles could be

276

sufficient to stabilize the droplets surface. The sonicated QPI stabilized emulsions with better

277

emulsification against flocculation might be due to the greater intra-particle hydrophobic

278

interactions and more effective adsorption at the droplets interface. A similar limitation of

279

flocculation by increasing c has been obtained for the heat induced soy glycinin nanoparticles

280

stabilized Pickering emulsion.28

281

Creaming Index (CI) of Emulsion Stability. All the QPI stabilized emulsions formed at

282

c = 0.25–6% with different φ values of 0.2–0.7 after 0 and 7 days storage, gradually to divide

283

into two layers: the creamed layer on the upside and the serum layer at the underpart (Figure

284

5A). In these Pickering emulsions, the storage of 7 days was sufficient for creaming, and a

285

further store time did not increase the creaming index any more. Figure 5B shows the

286

creaming index of QPI nanoparticles stabilized emulsions changed with the applied c and φ

287

after 7 days storage. The observations suggested that the application of the increasing c or φ

288

could enhance the creaming stability of the QPI Pickering emulsions against oiling off. The

289

increase in c resulted in a prominent reduction in CI%, indicating the limitation of creaming

290

by improving the particle concentration. Analogical data about the effect of c on creaming

291

stability have been obtained for the Pickering emulsions stabilized by zein, chitin nanocrystal

292

particles and soy glycinin.28,32-33 These results might be due to the increasing c reduced the

293

droplet size.16 Another possible reason for the improvement of creaming stability might be 15


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attributed to the formation of a thicker tier at the surface of oil droplets at higher c.32 It was

295

also obtained that the physical creaming stability of the emulsions increased upon with

296

different φ values of 0.2–0.7 at a specific c value. The observations were in accordance with

297

Liu and Tang,28 which revealed that high φ increased the particle amount of per interfacial

298

area adsorbed at the droplets surface.

299

Percentage of Adsorbed Proteins (AP%). In the protein particles stabilized Pickering

300

emulsions, the physical creaming stability and interfacial property is not only contacted with

301

attractive and repulsive inter-droplet interactions, but also related to inter-protein interactions

302

including unadsorbed and adsorbed protein at the interface.17 To further elaborate the

303

significance of inter- protein interactions for QPI stabilized emulsions, we determined the

304

number of adsorbed proteins at the droplets interface of the natural and 20 min-sonicated QPI

305

stabilized emulsions (Figure 6). Under all the investigated conditions, the AP% data of

306

sonicated QPI stabilized emulsions were obviously larger than that of natural QPI stabilized

307

emulsions. From Figure 6, the AP% of QPI stabilized emulsions progressively improved with

308

the increasing c. At higher c values, more protein particles were adsorbed at the droplet

309

interfaces, thus formed a thicker interfacial protein layer. With the increasing φ, the AP%

310

values of 20 min-sonicated QPI stabilized emulsions were in the range of 56–80.96%. This

311

conclusion was consistent with the creaming stability, illustrating that the high φ provided

312

larger interface area for protein to adsorb.

313

Nanoparticle Packing at the Interface and Surface Particle Coverage (SC). The

314

limited coalescence phenomenon of Pickering emulsion has been successfully used to supply 16



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an deep understanding of packing, adsorption and arrangement of solid particles at the droplet

316

interface. The percentage of surface particle coverage (SC%) can be examined by the slope of

317

the reciprocal surface diameter (1/D32) against the total particle area (Seq) divided by the oil

318

capacity (Vd). The simulation of (1/D32) of the natural and 20 min-sonicated QPI stabilized

319

Pickering emulsions as a control of the (Seq/Vd) is displayed in Figure 7A. The results revealed

320

that all the tested emulsion performed a good linear relation between (1/D32) and (Seq/Vd) with

321

a coefficient factor R2 of 0.988-0.993. Similar evolutions have been found for Pickering

322

emulsions stabilized by heat pretreated soy glycinin nanoparticles, whey protein microgel

323

particles, kafirin nanoparticles and quinoa starch granules.6,7,28,31 The SC% and QPI particle

324

center-to-center distance at the droplet interface (Dc-to-c) values of the natural and 20

325

min-sonicated QPI stabilized emulsions are summarized in Table 2. It can be observed that the

326

natural and sonicated QPI nanoparticles performed a relatively low SC% about 48.67–54.43%

327

compared to theoretical hexagonal array at interface (SC% =

328

nanoparticles exhibited high coating availability and efficient coverage at the droplets

329

interface. Ultrasound treatment progressively increased the SC% from 48.67 to 54.43%, thus

330

suggesting that the sonicated QPI nanoparticles would be more effective to serve as Pickering

331

stabilizer. We can still approximately calculate the Dc-to-c at the interface between two

332

neighboring particles according to square or hexagonal array models (Figure 7B and Table

333

2).31 The 20 min-sonicated QPI nanoparticles performed strong interactive forces between

334

adsorbed proteins, which led to large inter-particle space of particles at the interface.

335

According to comparison, the hexagonal array model appears to be more suitable for the 17


√ ×100%=90.7%).31

QPI

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arrangement of 20 min-sonicated QPI nanoparticles at the droplets interface than the square

337

array model.

338

We demonstrated the QPI nanoparticles could be used as effective food-grade stabilizers

339

of O/W Pickering emulsion. The supposed scheme for the formation of QPI nanoparticles

340

stabilized food-grade Pickering emulsions with ultrasound treatment is shown in Figure 8.

341

Ultrasound treatment considerably reduced the QPI particle size, and revealed the hydrophobic

342

groups from the interior to the outside of the QPI nanoparticles. The sonicated QPI stabilized

343

emulsions exhibited better emulsification efficiency and stronger inter-particle structure, and

344

the higher stability against coalescence and coacervation. Compared with the natural QPI

345

stabilized emulsions, the 20 min-sonicated emulsions exhibited higher packing and adsorption

346

of protein particles at the droplets interface. The microstructure of QPI Pickering emulsions

347

are occurred bridging flocculation of droplets at low c (≤1%, w/v), while the number of

348

particles could be high sufficient to fully cover the droplets surface at high c (>1%, w/v) with

349

hexagonal array model arrangement. Thus, the results would likely be of major significance

350

not only for the progress of QPI stabilized food-grade Pickering emulsions, but also for

351

facilitating the bioactive ingredients (e.g. polyphenols, carotenoid) applied in food and

352

pharmaceutical delivery products with health effects.

353 354 355 356 18



357

AUTHOR INFORMATION

358

Corresponding Authors

359

* Fax: +86-20-87113848. Tel.: +86-20-87113845. E-mail: [email protected]

360

Notes

361

The authors declare no competing financial interest.

362

ACKNOWLEDGMENTS

363

Financial support from the China Postdoctoral Science Foundation (2016M590787,

364

2017T100616), Key Project of Science and Technology of Guangdong Province of China

365

(2017B090901002), and the National Natural Science Foundation of China (21376097,

366

21576098).

367 368 369 370 371 372 373 374 375 376 377 19


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intensity ultrasonic pretreatment. Ultrason. Sonochem. 2016, 31, 590–597.

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extraction. Ultrason. Sonochem. 2011, 18(4), 813–835.

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459

Figure Captions

460

Figure 1. z-average diameter (Dz, histogram), surface hydrophobicity (Ho, alignment), and

461

contact angle (θ) of natural and sonicated (20 min) QPI nanoparticles at c = 0.25%.

462

Figure 2. Typical intrinsic fluorescence emission profiles of the natural (A) and 20

463

min-sonicated (B) QPI nanoparticles at c = 0.25%.

464

Figure 3. Effect of various solvents (DW, 6.0 M urea, 0.5% SDS, 30 mM DTT) on the

465

turbidity of particles in the natural and 20 min-sonicated QPI dispersions at c = 0.25%.

466

Figure 4. (A) Droplet size distribution and representative optical microscopy images (500×

467

magnification) of QPI stabilized Pickering emulsions formed at φ = 0.5 with varying c values

468

of 0.25–6%. (B) CLSM images microstructure of the 20 min-sonicated QPI nanoparticles

469

stabilized emulsion at φ = 0.5.

470

Figure 5. (A) Visual observation of the QPI stabilized Pickering emulsions prepared at c =

471

0.25–6% with varying φ values of 0.2–0.7 after 0 and 7 days storage. (B) Creaming index (%)

472

of emulsions stabilized by the QPI nanoparticles formed at c = 0.25–6% with varying φ values

473

of 0.2–0.7 after 7 days storage.

474

Figure 6. Percentage of adsorbed protein (%) for the natural and 20 min-sonicated QPI

475

stabilized emulsions, prepared by varying c values (at a specific φ value of 0.5, histogram), or

476

at varying φ values (at a specific c value of 6%, alignment).

477

Figure 7. (A) Simulation of the average inverse surface diameter (1/D32) of the natural and 20

478

min-sonicated QPI Pickering emulsions as a function of the total particle area (Seq) normalized

24



479

by the oil volume (Vd). (B) Graphic scheme of QPI nanoparticles packing and arrangement at

480

interface in a square or hexagonal array.

481

Figure 8. Supposed mechanism for the formation of food-grade Pickering emulsions

482

stabilized by the QPI nanoparticles with ultrasound treatment.

25


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Page 27 of 37


Table 1. The D43 and D32 values of droplets in the QPI Pickering emulsions, formed at c = 0.25– 6% with varying φ values of 0.2–0.7 by ultrasound treatment. Parameter φ

D43 (µm) 0.25

1

2

6

0.25

1

2

6

0.2

79.128a

61.692b

50.253c

42.95d

11.253a

8.689b

8.187c

8.044c

0.5

121.687a

104.588b

89.302c

62.006d

85.238a

68.799b

54.167c

39.262d

0.7

127.099a

123.016b

120.964c

120.76c

89.322a

85.463b

84.353b

81.2d

20 min

0.2

53.839a

48.294b

47.495c

35.395d

8.195a

7.782b

8.375b

6.59d

-sonication

0.5

113.552a

102.461b

85.875c

55.065d

75.957a

68.974b

13.428c

10.242d

0.7

120.55a

117.169b

112.383c

111.408c

82.487a

81.532b

78.051c

77.675c

natural

c

D32 (µm)

Values represent the means ± standard error (n = 3). Different letters in the same row indicate significant differences (p < 0.05).

26



Page 28 of 37

Table 2. The SC% and particle center-to-center distance at interface (Dc-to-c) values of the natural and 20 min-sonicated QPI stabilized emulsions. QPI

Dz (nm)

SC%

Dc-to-c (nm)

Nanoparticles

Square array

Hexagonal array

4 Dz

√3Dz 6

natural

401.37

48.67

50.97

54.79

20 min-sonication

207.57

54.43

24.93

26.79

27


Page 29 of 37


Figure 1

28



Figure 2

29


Page 30 of 37

Page 31 of 37


Figure 3

120

a

A

Control 20 min-sonication

B

b

Turbidity (µ g/mL)

100

80

C c

60

D

40 d

20 DW

urea

SDS

30


DTT


Figure 4

31


Page 32 of 37

Page 33 of 37


Figure 5

32



Page 34 of 37

Creaming Index (%)

B 80

0.25% 1% 2% 6%

60

40

20

0.2

0.5 ϕ

33


0.7

Page 35 of 37


Figure 6

0.5

0.2

0.7

40

60

75 Control 20 min-sonication

80

50

100

25

0

0.25

2

1

Protein concentration (%, w/v)

34


6

120

Percentage of Adsorbed Proteins (%)

Percentage of Adsorbed Proteins (%)

100


Page 36 of 37

Figure 7

A

200 Control 20 min-sonication

150 2

-1

1/D32 (cm )

Y=0.34242X-0.14363 (R =0.9884)

100

2

Y=0.30623X+4.5794 (R =0.99255)

50

0 0

200

400 -1

Seq/Vd (cm )

35


600

Page 37 of 37


Figure 8

36


Fabrication and Characterization of Quinoa Protein Nanoparticles

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