Mechanism of Formation and Stabilization of Nanoparticles Produced

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Mechanism of Formation and Stabilization of Nanoparticles Produced by Heating Electrostatic Complexes of WPI-Dextran Conjugate and Chondroitin Sulfate Qingyuan Dai, Xiuling Zhu, Jingyang Yu, Eric Karangwa, Shuqin Xia, Xiaoming Zhang, and Chengsheng Jia J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.6b01213 • Publication Date (Web): 22 Jun 2016 Downloaded from http://pubs.acs.org on June 22, 2016

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Journal of Agricultural and Food Chemistry

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Mechanism of Formation and Stabilization of Nanoparticles Produced by

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Heating Electrostatic Complexes of WPI−Dextran Conjugate and Chondroitin

3

Sulfate

4

Qingyuan Dai,†,

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Xiaoming Zhang,∗,† and Chengsheng Jia†

6

†

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Technology, Jiangnan University, Lihu Road 1800, Wuxi, Jiangsu 214122, People’s

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Republic of China

9

‡

10

‡

Xiuling Zhu,

‡

Jingyang Yu,† Eric Karangwa,† Shuqin Xia,†

State Key Laboratory of Food Science and Technology, School of Food Science and

College of Biological and Chemical Engineering, Anhui Polytechnic University,

Beijing Middle Road, Wuhu, Anhui 241000, People’s Republic of China

∗

To whom correspondence should be addressed. E-mail: [email protected] (X. Zhang). Phone: +86 510 85197217. Fax: +86 510 85884496.

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ACS Paragon Plus Environment


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ABSTRACT. Protein conformational changes were demonstrated in biopolymer

12

nanoparticles, and molecular forces were studied to elucidate the formation and

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stabilization mechanism of biopolymer nanoparticles. The biopolymer nanoparticles

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were prepared by heating electrostatic complexes of whey protein isolate (WPI)−

15

dextran conjugate (WD) and chondroitin sulfate (ChS) above the denaturation

16

temperature and near the isoelectric point of WPI. The internal characteristics of

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biopolymer nanoparticles were analyzed by spectroscopic techniques. Results showed

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that grafted dextran significantly (p < 0.05) prevented the formation of large

19

aggregates of WD dispersion during heat treatment. However, heat treatment slightly

20

induced the hydrophobicity changes of the microenvironment around fluorophores of

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WD. ChS electrostatic interaction with WD changed the fluorescence intensity of WD

22

regardless of heat treatment. Far-UV circular dichroism (CD) and attenuated total

23

reflectance Fourier transform infrared (ATR-FTIR) spectroscopies confirmed that

24

glycosylation and ionic polysaccharide did not significantly cause protein

25

conformational changes in WDC during heat treatment. In addition, hydrophobic

26

bonds were the major molecular force for the formation and stabilization of

27

biopolymer nanoparticles. However, hydrogen bonds slightly influenced their

28

formation and stabilization. Ionic bonds only promoted the formation of biopolymer

29

nanoparticles, while disulfide bonds partly contributed to their stability. This work

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will be beneficial to understand protein conformational changes and molecular forces

31

in biopolymer nanoparticles, and to prepare the stable biopolymer nanoparticles from

32

heating electrostatic complexes of native or glycosylated protein and polysaccharide.

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KEYWORDS: stabilization, nanoparticle, whey protein isolate, dextran, conjugate,

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chondroitin sulfate, electrostatic complex

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INTRODUCTION

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Biopolymer nanoparticles, as a delivery vehicle for hydrophobic bioactive

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compounds, have attracted great attention due to their remarkable nonantigenicity,

38

biocompatibility, biodegradability, and abundant renewable properties.

39

stability of biopolymer nanoparticles, especially under different physiological

40

conditions, is essential to prevent microstructural destruction before reaching certain

41

target sites after uptake. 3-5 Biopolymer nanoparticles have been prepared using native

42

or glycosylated protein and ionic polysaccharide by complex coacervation or

43

heat-induced gelation methods. 3-8 Many researchers have focused on the optimization

44

of heat-induced nanoparticles formulation from native or glycosylated protein and

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ionic polysaccharide as well as their applications. Nevertheless, much less attention

46

has been paid to the protein conformational changes in stable biopolymer

47

nanoparticles and the formation and stabilization mechanism of biopolymer

48

nanoparticles from the viewpoint of molecular forces, which were prepared by heating

49

electrostatic complexes of glycosylated protein and ionic polysaccharide.

1-5

The

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Whey protein isolate (WPI)–dextran conjugate (WD) and chondroitin sulfate (ChS)

51

were selected as model of glycosylated protein and ionic polysaccharide, respectively.

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WPI, a by-product of cheese or casein manufacturing, has been widely used as an

53

ingredient in food products for its good nutritional quality and remarkable functional

54

properties, such as emulsification, foaming ability and gelation. WPI mainly consists

55

of several globular proteins, including β-lactoglobulin (β-lg), α-lactalbumin (α-la),

56

bovine serum albumin (BSA), and immunoglobulins (IGs). 9 β-Lg is one of the major

4


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components of WPI and determines functional properties of WPI. The stability of

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complex coacervates or heat-induced nanoparticles formed by WPI and ionic

59

polysaccharide significantly decreased at the specific pH and/or at higher salt

60

concentrations, leading to precipitation or dissociation.

61

nonenzymatic glycosylation, is a series of complex reactions between free amino

62

groups of protein and reducing carbonyl groups of polysaccharide, which usually

63

occurs during thermal process in food systems. It has been reported that Maillard

64

reaction can significantly improve the solubility, thermal stability, and emulsification

65

properties of the original proteins.

66

viscosity, high solubility, and no gelation, was selected as a source of polysaccharide

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for Maillard reaction to avoid the complication during the formation of electrostatic

68

complexes between negatively and positively charged biopolymers. Dextran

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covalently conjugated to protein can provide steric hindrance against protein thermal

70

aggregation.

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disaccharide unit containing β-1,4-linked glucuronic acid and β-1,3-N-acetyl

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galactosamine, and sulfated at either the 4 or 6 position of the galactosamine residue.

73

13

74

biodegradability, and targetability.

75

charges and can be used as a vehicle of bioactive compound in delivery system with

76

positively charged substances by electrostatic interactions.

9

9, 12

10, 11

Maillard reaction, a

Dextran, a neutral polysaccharide with low

ChS, a linear glycosaminoglycan, is comprised of a polymerized

Additionally, ChS has many interesting properties, including biocompatibility, 14

Therefore, ChS chains have lots of anionic

77

Functional properties of proteins are closely related to their structures, and protein

78

structures are dependent on hydrophobic bonds, ionic bonds, van der Waals forces,

5



15-19

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hydrogen bonds and disulfide bonds.

80

spectroscopies have been used to investigate the structure, interactions, and dynamics

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of proteins in solution due to its high sensitivity, simplicity, and rapidity. 20-23 Circular

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dichroism (CD) spectroscopy has been widely used to evaluate the protein

83

conformation in solution. However, Fourier transform infrared (FTIR) spectroscopy is

84

an excellent technique to determine the protein conformation in solutions, thin films

85

(dry or hydrated), solids (spray-dried or lyophilized powders), or suspensions of

86

precipitates.

87

total reflectance FTIR (ATR−FTIR) spectroscopy is highly sensitive due to the

88

absence of major water peak in the hydrated thin-film sample.

89

of different molecular forces involved in protein gels or biopolymer nanoparticles can

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be determined by the solubility of protein gels or particle size of biopolymer

91

nanoparticles in various chemical reagents, which differ each other by their functional

92

ability to cleave specific bonds: ionic bonds (NaSCN, Na2SO4, CH3COONa, NaCl),

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hydrogen and hydrophobic bonds [urea, guanidine hydrochloride (GuHCl)], and

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disulfide bonds [β-mercaptoethanol (β-ME or 2-ME), dithiothreitol (DTT),

95

N-ethylmaleimide (NEM)]. 18, 29-32

24-27

Intrinsic and synchronous fluorescence

Compared to the traditional transmission FTIR, thin-film attenuated

28, 29

The contributions

96

The objective of the present study was to evaluate protein conformational changes

97

in biopolymer nanoparticles, and the contributions of different molecule forces on the

98

formation and stabilization of biopolymer nanoparticles, prepared by heating

99

electrostatic complexes of WD and ChS. The biopolymer nanoparticles were

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characterized by dynamic light scattering, intrinsic fluorescence spectroscopy,

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synchronous

fluorescence

spectroscopy,

CD

spectroscopy

102

spectroscopy. Finally, protein conformational changes were demonstrated in

103

biopolymer nanoparticles and the mechanism of formation and stabilization of

104

biopolymer nanoparticles was elucidated from the viewpoint of molecular forces,

105

which will facilitate the preparation of stable biopolymer nanoparticles by

106

heat-induced method using native or glycosylated protein and ionic polysaccharide.

107

MATERIALS AND METHODS

108

Materials and Reagents. WPI was obtained from Hilmar Ingredients (Hilmar,

109

California). The total solid, protein, and ash in the dry power were 95.6, 88.7, and

110

2.7%, respectively. Dextran with molecular mass of 40 kDa was purchased from

111

Sinopharm Chemical Reagent Co., Ltd (Shanghai, China). Chondroitin sulfate (ChS)

112

was kindly provided by Shandong Yibao Biologics Co., Ltd (Yanzhou, China). ChS

113

consisted of 95.4% sodium ChS and 4.6% protein. Hydrochloric acid (HCl), sodium

114

hydroxide (NaOH), o-phthalaldehyde (OPA), sodium chloride (NaCl), urea, and

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dithiothreitol (DTT) were purchased from Sinopharm Chemical Reagent Co., Ltd

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(Shanghai, China). All materials were used without any further purification. All

117

aqueous solutions were prepared with deionized water.

118

Preparation of Stable Biopolymer Nanoparticles from WD and ChS. The stable

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biopolymer nanoparticles were prepared by heating electrostatic complexes of WD

120

and ChS according to the methods described in our previous paper. 4 Briefly, WPI and

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dextran were dissolved in 10 mM sodium phosphate buffer solution (PBS) (pH 6.5)

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with 0.02% (w/v) sodium azide, and adjusted to 7.5, 22.5% (w/w), respectively, and to

7


and

ATR-FTIR


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pH 6.5 using 1.0 M HCl or 1.0 M NaOH. After storage at 4 °C overnight for the

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complete hydration, the mixed solutions were incubated in a water bath for 48 h at

125

60°C. When Maillard reaction of the mixed solutions was finished, the reacted

126

solutions were immediately cooled in an ice−water bath. The degree of glycosylation

127

(DG) of WD was 9.7 %, which was determined by the o-phthalaldehyde (OPA) assay

128

from the loss of free amino groups of WPI. pH 6.5 and 60 °C were used to obtain the

129

maximal production of Schiff base, which was the initial product of Maillard reaction.

130

9

131

conditions were obtained under 7.5 and 22.5% (w/w), respectively.

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time of 48 h could be acquired an appropriate DG of WD to prepare the stable

133

biopolymer nanoparticles.

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dissolving ChS in deionized water and gently stirring for 2 h at room temperature.

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The WD stock solution and ChS stock solution were mixed (denoted as WDC). The

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concentrations of WPI, dextran, and ChS in WDC solution were adjusted to 0.2, 0.55,

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and 0.008% (w/v), respectively. After stirring for 2 h, the mixed solutions were

138

adjusted to pH 5.2 [near the isoelectric point (pI) of WPI] with 0.1 M HCl, and heated

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at 85 °C for 15 min. The nanoparticle dispersions were immediately cooled for 10 min

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in an ice−water bath. Our previous studies showed that the secondary aggregation of

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heated WD (HWD) dispersion would occur at pH 4.0, and the biopolymer

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nanoparticles from heated WDC (HWDC) dispersion had Z-average mean diameter

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around 150 nm with polydispersity index (PDI) 0.08 in the pH range 1.0 to 8.0

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regardless of 0.2 M NaCl. Additionally, ChS, WPI and WD with 9.7% DG were

The optimal concentrations of WPI and dextran under macromolecular crowding

4

4

The incubated

ChS (1%, w/v) stock solution was obtained after

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assembled into the spherical shape and smooth surface biopolymer nanoparticles with

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dextran conjugated to WPI/ChS shell and WPI/ChS core during heat treatment.

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this work, protein conformational changes were demonstrated in biopolymer

148

nanoparticles, and molecular forces were studied to elucidate the formation and

149

stabilization mechanism of biopolymer nanoparticles, prepared by heating

150

electrostatic complexes of WD with 9.7% DG and ChS. The stable biopolymer

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nanoparticle dispersions were kept at 4 °C before analysis. WPI and WD solutions

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were prepared and treated under the same conditions described above. All

153

experiments were performed in triplicate.

154

Intrinsic Fluorescence Emission Spectroscopy. The intrinsic fluorescence emission

155

spectra were determined at room temperature (25 °C) using a fluorescence

156

spectrophotometer (F-7000, Hitachi Co., Ltd, Japan). The protein concentration in

157

each sample was diluted to 0.2 mg/mL in sodium phosphate-citric acid buffer (10 mM,

158

pH 5.2). The emission spectra were separately recorded from 285 to 450 nm and 300

159

to 450 nm at the excitation wavelength of 280 and 295 nm both with a slit width of

160

2.5 nm, respectively. The corresponding sample without WPI was used as a control to

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correct the fluorescence background.

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Synchronous Fluorescence Spectroscopy. Synchronous fluorescence spectrometry

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has been widely used in multicomponent analysis to distinguish the microenvironment

164

changes around different fluorescent groups.

165

measurements were performed at room temperature (25 °C) using a fluorescence

166

spectrophotometer (F-7000, Hitachi Co., Ltd, Japan). To obtain the microenvironment

23

4

In

Synchronous fluorescence

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changes around individual tyrosine (Tyr) and tryptophan (Trp) residues in proteins,

168

the synchronous fluorescence spectra of the same samples as intrinsic fluorescence

169

experiments were recorded from 240 to 360 nm at fixed 15 and 60 nm interval

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between the excitation and emission wavelength both with a slit width of 2.5 nm,

171

respectively. The fluorescence intensity of each sample blank was subtracted from

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that of corresponding sample to obtain net fluorescence intensity of each protein

173

sample.

174

Far-UV CD Spectroscopy. Far-UV CD spectroscopy of each sample was carried out

175

using a MOS-450 CD Spectropolarimeter (Biologic, Claix, France). The spectra were

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scanned from 190 to 250 nm with a 1mm path length quartz cuvette at 25 °C. The

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protein concentration in all samples was diluted to 0.2 mg/mL and adjusted to pH 5.2.

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The protein spectrum was corrected by subtracting the spectrum of a protein-free

179

solution. The molar ellipticities of protein samples were calculated as [θ]

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(deg·cm2·dmol-1) = (100 × X × M)/(L × C), where X is the signal (millidegrees)

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obtained by the CD spectrometer, M is the average molecule weight of amino acid

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residues in protein (assumed to be 115 for WPI), C is the protein concentration

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(mg/mL) of the sample, and L is the cell path length (cm).

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structures, including α-helix, β-sheet, β-turn and random coil, were analyzed by the

185

spectra

186

(http://dichroweb.cryst.bbk.ac.uk/html/process.shtml).

187

ATR-FTIR Spectroscopy. Infrared spectra were obtained at room temperature (25 °C)

188

using a FTIR spectrophotometer (Nicolet iS10, Thermo Electron Corp., Madison,

and

calculated

using

10


12

Four secondary

DICHROWEB

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Wisconsin) equipped with an Ever-Glo MIR source, a KBr beam splitter and a

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deuterated triglycine sulphate (DTGS) detector. The spectra data were collected in the

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range of 650-4000 cm-1 at a 4 cm-1 resolution and a zero filling factor of 1 using a

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Happ–Genzel apodization and Mertz phase correction. Sixteen scans were

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accumulated to obtain a reasonable signal-to-noise ratio. An aliquot of each sample

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(50µL) was placed on the aluminum foil. After 24 h of storage at room temperature,

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the dried film of each sample on the foil was formed, and then positioned directly on a

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single reflection diamond attenuated total reflectance (ATR) crystal. The ATR crystal

197

was washed with deionized water and dried with lens paper to avoid contamination

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between samples. All samples were measured under identical conditions. To ensure no

199

interference from non-protein constituents, each spectrum was obtained by subtracting

200

the corresponding background spectrum from the sample spectrum, using the Nicolet

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Omnic software (version 8.3, Thermo Electron Corp., Madison, Wisconsin). Protein

202

secondary structure is most reliably indicated by the amide I band (1600-1700 cm-1).

203

28

204

two points, and smoothed by 9-point Savitzky-Golay filter method. Second derivative

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spectrum, obtained using a third degree polynomial function with a 5-point

206

Savitsky–Golay smoothing function, was used to identify the positions of overlapping

207

components of the amide I band. The positions were then confirmed by Fourier self

208

deconvolution with a full bandwidth at half height (FWHH) of 13.0 cm-1 and a

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resolution enhancement factor (K) of 2.4. Finally, the FTIR deconvolution spectra

210

were curve-fitted by Gaussian-Lorentzian function with PeakFit software (Version

After the amide I band of the resulting different spectrum was baseline corrected by

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4.12, SeaSolve Software Inc., Framingham, Massachusetts). Quantitative estimation

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of protein secondary structure was performed by calculating the corresponding band

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percentage in the amide I band region according to the following wavenumber ranges:

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1620-1645 cm-1, β-sheet; 1645-1652 cm-1, random coil; 1652-1662 cm-1, α-helix;

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1662-1690 cm-1, β-turn. 24

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Determination of Molecular Forces for Formation and Stabilization of

217

Biopolymer Nanoparticles. The contributions of different molecular forces on the

218

formation of biopolymer nanoparticles were determined by preparing their dispersions

219

in the presence of various dissociating reagents. WD and ChS stock solutions were

220

diluted with addition of individual dissociating solution to the above-mentioned

221

concentrations. Meanwhile, the dissociating reagents in the resulting dispersions were

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adjusted to the final concentrations as follows: 0.6 M NaCl (solution S1), 1.5 M urea

223

(solution S2), 8.0 M urea (solution S3) and 10 mM DTT (solution S4). All other

224

procedures were the same as described above. To determine the contributions of

225

molecular forces on the stabilization of biopolymer nanoparticles, the stable

226

biopolymer nanoparticle dispersions were diluted 10-fold with dissociating reagents,

227

and the dissociating reagents were adjusted to the final concentrations as follows: 0.6

228

M NaCl (solution S5), 0.6 M NaCl + 1.5 M urea (solution S6), 0.6 M NaCl + 8.0 M

229

urea (solution S7), and 0.6 M NaCl + 8.0 M urea + 10 mM DTT (solution S8).

230

The diameter changes were used to estimate the contributions of ionic bonds

231

(difference between S1 and control or between S5 and control, respectively),

232

hydrogen bonds (difference between S2 and control or between S6 and S5,

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30-33

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respectively), hydrophobic interactions (difference between S3 and S2 or between S7

234

and S6, respectively) and disulfide bonds (difference between S4 and control or

235

between S8 and S7, respectively) for the formation and stabilization of biopolymer

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nanoparticles. The particle sizes were measured after 1 h of storage.

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Dynamic Laser Scattering (DLS) Measurements. The Z-average mean diameter

238

and polydispersity index (PDI) of biopolymer nanoparticles were obtained by

239

dynamic light scattering using a Malvern Zetasizer (Nano ZS, Malvern Instruments

240

Ltd., Worcestershire, UK) equipped with 633 nm and He−Ne laser beam.

241

Measurements were made at 25 °C and 173° scattering angle. The nanoparticle

242

dispersions were measured by dilution with the corresponding solutions to a final

243

protein concentration of 0.2 mg/mL. Each dispersion was fully shaken before

244

measuring the Z-average mean diameter to ensure a uniform suspension of particles.

245

Statistical Analysis. Each experiment was triplicated under the same conditions. A

246

one-way analysis of variance (ANOVA) was applied to estimate the statistical

247

difference. Significant differences (p < 0.05) between means were determined using

248

Duncan’s multiple range tests. Statistical analyses were evaluated with SPSS software

249

(version 17.0, SPSS Inc., Chicago, Illinois).

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RESULTS AND DISCUSSION

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Biopolymer Nanoparticles Prepared using WPI after Different Treatments. The

252

Z-average mean diameters of WPI, heated WPI (HWPI), WD, HWD, WDC, and

253

HWDC dispersions are shown in Figure 1. Around pI of WPI, the solubility of WPI

254

decreased and formed smaller biopolymer particles about 266 nm with PDI 0.505.

13



255

However, heat treatment led to the formation of large particle aggregates about

256

8890nm with PDI 0.307 in HWPI dispersion. Heat treatment might promote the

257

hydrophobic interactions and repress hydrogen interactions. In addition, near the pI of

258

protein, heat denaturation altered the hydrophobicity/hydrophilicity balance of protein

259

surface, leading to aggregation via hydrophobic interactions. These results are

260

consistent with previous studies.

261

dispersions were 128 and 159 nm, respectively. These results indicated that

262

glycosylation and ionic polysaccharide significantly prevented the formation of large

263

aggregates during heat treatment. This was due to the steric hindrance from dextran

264

chains covalently conjugated to WPI molecules and ChS chains electrostatically

265

interacted with WD. Meanwhile, the diameter changes in different WPI samples

266

might be related to protein conformational changes after different treatments. These

267

results are consistent with previous studies.

268

in diameter sizes between WD and WDC regardless of heat treatment. In our previous

269

publication, we reported that HWDC dispersion was stable against pH and salt, but

270

the secondary aggregation of HWD dispersion could occur at pH 4.0. 4 Based on these

271

findings, further studies on protein conformational changes and molecular forces in

272

the stable HWDC nanoparticles were investigated in the present work.

273

Fluorescence Spectroscopic Analysis. Intrinsic Fluorescence Emission Spectroscopy.

274

Due to the absence of external reagents, the intrinsic fluorescence spectroscopy has

275

been used as a reliable method to evaluate the changes of the microenvironment

276

around fluorescent groups in proteins.

34

The diameter sizes of WDC and HWDC

22

12, 35

There was no significant difference

The intrinsic fluorescence of protein results

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from aromatic fluorophores, including phenylalanine (Phe), tyrosine (Tyr), and

278

tryptophan (Trp) residues of proteins. Due to a very low quantum yield of Phe

279

residues, the intrinsic fluorescence of many proteins is mainly attributed to Tyr and

280

Trp residues. β-Lg, α-la, and BSA contain 2, 4 and 2 Trp residues and 4, 4 and 20 Tyr

281

residues per molecule, respectively. 21, 36 At the excitation wavelength of 280 nm, both

282

Tyr and Trp residues showed fluorescence emission spectrum, but at the excitation

283

wavelength of 295 nm, only Trp residues showed fluorescence emission spectrum.

284

At the excitation wavelength of 295 nm (Figure 2B), the maximum of emission

285

wavelength (λmax) of WPI dispersion was 335 nm, whereas the λmax of WD dispersion

286

was 333 nm. These results suggested that the polarity around Trp residues in proteins

287

decreased and the hydrophobicity increased, indicating the protein conformational

288

changes. This might be attributed to dextran covalently conjugated to WPI.

289

Meanwhile, the fluorescence intensity of WD dispersion decreased compared to that

290

of WPI dispersion. This might be due to the covalent conjugation of dextran chains to

291

WPI on fluorescence quenching of protein. These results are in agreement with the

292

fluorescence characteristics of Maillard reaction products. 37 There was no significant

293

difference in λmax between WD and WDC dispersions (Figure 2B). When WD or

294

WDC dispersions were heated at 85 °C for 15 min, both λmax were shifted from 333 to

295

335 nm, and the fluorescence intensity significantly increased (Figure 2B), indicating

296

the increase of polarity and the decrease of hydrophobicity of the microenvironment

297

around Trp residues of proteins. These fluorescence changes might be related to the

298

increase of hydrophobic interactions between protein molecules during heat treatment,

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20


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which not only contributed to the changes of the microenvironment around

300

fluorophores (Figure 2B) but also promoted the increase in particle diameters of WD

301

and WDC dispersions (Figure 1). Simion et al. reported that the fluorescence intensity

302

of β-lg significantly increased with increasing temperature (25-85 °C) at the excitation

303

wavelength of 292 nm, and the λmax of β-lg exhibited a red-shift of 2-4 nm after heat

304

treatment at 75-85 °C due to the increase of the exposure of its fluorophores.

305

fluorescence intensities of WDC and HWDC dispersions were higher than those of

306

WD and HWD dispersions, respectively, suggesting that ChS reduced the

307

fluorescence quenching of dextran covalently conjugated to WPI. This change might

308

be attributed to the protein conformational changes in WDC induced by electrostatic

309

interactions between ChS and WD molecules, leading to the decrease of quenching

310

effect of grafted dextran chains on the Trp fluorophore. Similar emission spectra were

311

observed regardless of the excitation wavelengths. Only slight differences in

312

fluorescence intensity between the two emission spectra were observed (Figure 2A, B,

313

respectively).

21

The

314

Synchronous Fluorescence Spectroscopy. Synchronous fluorescence spectroscopy

315

further distinguished the effects of glycosylation, ionic polysaccharide, and heat

316

treatment on the individual fluorescent groups. At fixed △λ (15 and 60 nm) between

317

excitation and emission wavelength, the synchronous fluorescence spectroscopy could

318

provide more accurate information about the microenvironment around individual Tyr

319

and Trp residues in proteins, respectively.

320

changes in the polarity and hydrophobicity of the microenvironment around

23

The shifts of the λmax are related to the

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21

321

fluorescent groups in proteins, indicating conformational changes of protein.

The

322

synchronous fluorescence spectra of all samples resulted from Tyr and Trp residues at

323

△λ = 15 and △λ = 60 nm are shown in Figure 3A, 3B, respectively. As shown in

324

Figure 3A, the fluorescence intensity of WD dispersion was lower than that of WPI

325

dispersion, indicating that dextran covalently conjugated to WPI quenched the

326

fluorescence of Tyr residues. The fluorescence intensity of WD dispersion increased

327

with addition of ChS, suggesting that ChS reduced the fluorescence quenching of

328

dextran covalently conjugated to WPI. WPI, WD, and WDC dispersions had similar

329

λmax, indicating that the polarity of the microenvironment around Tyr residues was not

330

changed. After heating WD and WDC dispersions, their fluorescence intensities

331

increased and their fluorescence spectra showed a slight blue-shift of λmax compared

332

to WPI dispersion, indicating that the hydrophobicity of the microenvironment around

333

Tyr residues was slightly changed. Additionally, the fluorescence intensity of HWD

334

dispersion was lower than that of HWDC dispersion, indicating that ChS

335

electrostatically interacted with WD. Simion et al. reported that β-lg had a 2.5 nm

336

blue-shift of the λmax (△λ = 15 nm) at 80 and 85 °C for burial of Tyr residues and its

337

fluorescence intensity significantly increased, and explained that the polarity around

338

Tyr residues decreased while the hydrophobicity increased. 21

339

As shown in Figure 3B (△λ = 60 nm), both WD and WDC dispersions showed a

340

slight blue-shift in the λmax compared to WPI dispersion. Wu et al. reported that

341

β-lg−fructooligosaccharide conjugate had a slight blue shift of the λmax (△λ = 60 nm),

342

and

explained

that

glycosylation

of

β-lg

influenced

17


the

hydrophobic


38

343

microenvironment around Trp residues.

HWD and HWDC dispersions showed a

344

slight red-shift in the λmax compared to WD and WDC dispersions, respectively.

345

Simion et al. reported that β-lg exhibited a 2.5 nm red-shift of the λmax (△λ = 60 nm)

346

at 80 and 85 °C for exposure of Trp residues and its fluorescence intensity

347

significantly increased, and demonstrated that the polarity around Trp residues

348

increased while the hydrophobicity decreased.

349

synchronous fluorescence intensity of all samples at △λ = 15 nm and △λ = 60 nm.

350

Whereas the fluorescence intensity of the latter was much higher than that of the

351

former. Additionally, the fluorescent change trend of Tyr and Trp residues induced by

352

same treatments was different in the synchronous fluorescence spectroscopy (Figure

353

3A, B, respectively).

354

Far-UV CD Spectroscopic Analysis. The far-UV CD spectra of WPI, WD, HWD,

355

WDC, HWDC dispersions are shown in Figure 4A. Conformational changes in the

356

secondary structure of proteins were studied at wavelength range between 190-250

357

nm. The broad negative peak around 206 nm represented α-helix conformation. The

358

secondary structure compositions in all samples are shown in Figure 4B. WPI had an

359

average of 33.0% α-helix, 19.4% β-sheet, 19.8% β-turn, and 27.8% random coil

360

(Figure 4B). Tomczyńska-Mleko et al. demonstrated that WPI had an average 23.1%

361

α-helix, 22.9% β-sheet, 22.2% β-turn, and 31.7% random coil at pH 5.0. This

362

difference might be due to different sources of WPI and pH condition. 39 Compared to

363

WPI, contents of β-turn and random coil in WD sample slightly increased at the

364

expense of α-helix and β-sheet (Figure 4B), indicating that glycosylation did not

21

A similar trend was observed in

18


Page 18 of 43

Page 19 of 43


365

significantly change the protein secondary structure of WD obtained under

366

macromolecular crowding conditions. Similar findings have previously been reported.

367

35, 40

368

the content of α-helix slightly decreased (Figure 4B), indicating that heat treatment

369

slightly induced protein conformational changes in WD. Perez et al. demonstrated that

370

heat treatment could promote protein conformational changes.

371

structures between HWD and HWDC were slightly different, suggesting that the

372

steric hindrance from ChS electrostatically interacted with WD did not significantly

373

induce the changes in spatial structure and unfolding of glycosylated protein during

374

heat treatment. Zhang et al. demonstrated that pectin enhanced the thermal stability of

375

WPI structure, and explained that pectin could prevent secondary structural changes

376

of WPI through electrostatic interactions. 41

377

ATR-FTIR Spectroscopic Analysis. The ATR-FTIR spectra of unheated and heated

378

WPI, WD, WDC, dextran (DEX) and dextran/ChS (DC) in the region between

379

650-4000 cm-1 are shown in Figure 5A. Although amide I, II, and III bands of FTIR

380

spectrum can be used to estimate protein secondary structure, amide I band

381

(1600-1700 cm-1) is the most sensitive to protein conformational changes, and is

382

widely used in secondary structure analysis.

383

curve-fitting individual component bands in the amide I band region of WPI are

384

shown in Figure 5B. Fourier self-deconvolution method was applied to distinguish the

385

individual components in the intrinsically overlapped amide I band contours, which

386

were assigned to different secondary structure conformations.

After heat treatment, the ellipticity of WD became less negative (Figure 4A)，and

29

22

The secondary

The FTIR spectrum and the

19


24, 28

Curve fittings of


Page 20 of 43

387

the deconvolved spectra were performed using Gaussian-Lorentzian function.

388

Consequently, quantitative estimation of protein secondary structures, including

389

α-helix, β-sheet, β-turn, and random coil, were obtained. 24, 25

390

The percentages of four secondary structures of protein in all samples are shown in

391

Figure 5C. Although there was difference in percentages of four secondary structures

392

of protein, calculated by Far-UV CD spectroscopy and ATR-FTIR spectroscopy, the

393

two analytical methods showed similar change trend in same samples. The difference

394

might be due to different water contents. It has been reported that hydration could

395

significantly increase contents of α-helix and random coil and lower content of

396

β-sheet in protein by FTIR spectroscopy. 42 Our results demonstrated that WPI had an

397

average of 14.4% α-helix, 41.0% β-sheet, 29.1% β-turn, and 15.5% random coil.

398

Similar results have previously been reported using ATR-FTIR spectroscopy or

399

Fourier transform Raman spectroscopy.

400

changes of protein in the form of suspensions or precipitates could not be determined

401

by fluorescence spectroscopy and CD spectroscopy, they could be determined using

402

FTIR spectroscopy. Heat treatment slightly decreased the content of β-sheet in WPI

403

and slightly increased the contents of β-turn and random coil (Figure 5C). Protein

404

conformational changes might be related to larger aggregates (diameter > 8800 nm) in

405

HWPI dispersion. Similar findings have previously been reported.

406

WPI, the percentage of α-helix slightly decreased in WD, and further slightly

407

decreased in HWD (Figure 5C). These results indicated that dextran covalently

408

conjugated to WPI and heat treatment did not significantly induce protein

26, 41

Although the protein conformational

20


43

Compared to

Page 21 of 43


409

conformational changes of WD. These results are consistent with the findings of CD

410

spectroscopy. ChS did not significantly induce the changes of secondary structures of

411

WD, except for slight increase of β-sheet content in WDC. There was no significant

412

difference in the secondary structures between HWD and HWDC (Figure 5C),

413

indicating that ChS did not significantly change protein conformations of HWD.

414

Molecular Forces for Formation and Stabilization of Biopolymer Nanoparticles.

415

Molecular Forces for Formation of Biopolymer Nanoparticles. The influences of

416

different molecular forces on the formation of biopolymer nanoparticles are shown in

417

Figure 6A. Compared to control sample, 0.6 M NaCl induced the greatest change in

418

the particle size of biopolymer nanoparticles followed by 8.0 M urea, 1.5 M urea, and

419

10 mM DTT (Figure 6A). Several researchers reported that various dissociating

420

reagents could affect protein heat stability, rheological properties, and molecular

421

forces within protein and water molecules, and demonstrated that the formation of

422

protein gel networks was attributed to the balance of non-covalent interactions (ionic,

423

hydrophobic, and hydrogen bonds) and covalent disulfide bonds.

424

biopolymer nanoparticles were prepared by heating electrostatic complexes of WD

425

and ChS in the absence or presence of 0.6 M NaCl (control, S1 in Figure 6A,

426

respectively). The Z-average diameter and PDI of biopolymer nanoparticles changed

427

from 156.3 nm and 0.071 to 388.0 nm and 0.439, respectively. Electrostatic shielding

428

effects minimized electrostatic interactions between glycosylated protein and ionic

429

polysaccharide molecules and relatively increased hydrophobic interactions, which

430

promoted protein aggregation during heat treatment, indicating the destruction of the

21


19, 30-32

The


Page 22 of 43

431

original equilibrium between electrostatic and hydrophobic interactions in protein

432

dispersions. These results confirmed that ionic bonds significantly influenced the

433

formation of the biopolymer nanoparticles. Jones et al. reported that there was a weak

434

electrostatic repulsion between biopolymer particles at high salt concentration,

435

leading to the large particle aggregates.

436

that neutral salts had two antagonistic effects on electrostatic and hydrophobic

437

interactions at higher concentrations. 15

6

Additionally, Melander et al. demonstrated

438

Urea (1.5 M) was used to test hydrogen bonds (which break endothermically),

439

while hydrophobic bonds (which break exothermically) and hydrogen bonds were

440

tested with 8.0 M urea.

441

biopolymer nanoparticles decreased by 24.2% in the presence of 1.5 M urea (S2 in

442

Figure 6A), indicating that the biopolymer nanoparticles had a more compact

443

structure. However, the Z-average diameter and PDI of biopolymer nanoparticles

444

significantly increased to 273.8 nm and 0.488 in the presence of 8.0 M urea (S3 in

445

Figure 6A), respectively. Similar findings have previously been reported.

446

results indicated that although 1.5 M urea could compete with the inter- and

447

intramolecular hydrogen bonds between proteins and water, hydrogen bonds could not

448

be essential for the formation of biopolymer nanoparticles. Under similar preparation

449

conditions in the presence of 8.0 M urea, the biopolymer nanoparticles had a more

450

loose structure due to the reduction of hydrophobic interactions in strength. Therefore,

451

hydrophobic interactions played a prominent role in the formation of biopolymer

452

nanoparticles.

44, 45

Compared to the control sample, the diameter size of

22


46-48

These

Page 23 of 43


DTT is often used to reduce disulfide bonds and prevent disulfide bond formation.

453 454

19

455

of 10 mM DTT (S4 in Figure 6A). Thus disulfide bonds did not significantly

456

influence the formation of biopolymer nanoparticles. Sun and Arntfield reported that

457

no significant difference was observed on storage moduli (G') with addition of 0.1-0.3

458

M β-mercaptoethanol

459

mM N-ethylmaleimide (NEM), and explained that disulfide bonds were not required

460

for gel formation. 19

The diameter size of biopolymer nanoparticles negligibly changed in the presence

(2-ME),

0.05-0.15

M dithiothreitol

(DTT),

and 10-25

461

Molecular Forces for Maintaining Stability of Biopolymer Nanoparticles. The

462

influences of different molecular forces on the stability of biopolymer nanoparticles

463

are shown in Figure 6B. The addition of 0.6 M NaCl had almost no effect on the

464

particle size of the stable biopolymer nanoparticles compared to control sample (S5,

465

control in Figure 6B, respectively). Therefore, ionic bonds were not essential for

466

maintaining the stability of biopolymer nanoparticles. Jones and McClements reported

467

that the particle diameter of biopolymer particles, formed by heating β-lg and pectin

468

complexes in the absence of 0.2 M NaCl, had almost no change after diluting their

469

dispersion in the presence of 0.2 M NaCl, indicating its good stability to salt.

470

Additionally, Giroux et al. demonstrated that calcium promoted the formation of

471

nanoparticles from denatured whey protein through pH-cycling treatment, but it was

472

not necessary to maintain the stability of biopolymer nanoparticles. 32

6

473

The particle sizes of biopolymer nanoparticles increased by 12.7 and 182.0% after

474

diluting their dispersion in the presence of 0.6 M NaCl and 1.5 or 8.0 M urea (S6, 7 in

23



475

Figure 6B), respectively. Different urea concentrations caused different swelling

476

degree of the biopolymer nanoparticles. 1.5 M urea concentration slightly changed the

477

diameter size of biopolymer nanoparticles by breakage of hydrogen bonds. However,

478

8.0 M urea concentration significantly changed the diameter size by breakage of both

479

hydrogen and hydrophobic bonds.

480

that hydrogen bonds had a slight contribution on maintaining the stability of

481

biopolymer nanoparticles, while hydrophobic bonds had a predominant impact in

482

stabilizing the biopolymer nanoparticles.

18, 30, 32, 33, 49

Therefore, these findings suggested

483

Due to disulfide reduction of DTT and swelling of urea, the diameter size of

484

biopolymer nanoparticles further increased by 21.2% after dispersion dilution in the

485

presence of NaCl and urea plus DTT (S8 in Figure 6B), and PDI significantly

486

increased to 0.534, suggesting disruption of biopolymer nanoparticle dispersion.

487

These results indicated that disulfide bonds could partly maintain the stability of

488

biopolymer nanoparticles. Previous studies demonstrated that disulfide bonds could

489

partly stabilize gels of heat-induced proteins in dissociating solution (0.6 M NaCl +

490

8.0 M urea + 10 mM DTT) or (0.6 M NaCl + 8.0 M urea + 0.5 M

491

2-β-mercaptoethanol). 30, 33

492

Mechanism of Formation and Stabilization of HWDC Nanoparticles. Protein

493

structure is dependent on hydrophobic bonds, ionic bonds, van der Waals forces,

494

hydrogen bonds and disulfide bonds.

495

protein conformational changes and molecular forces in biopolymer nanoparticles.

496

The diameter size of HWPI dispersion was significantly different to those of HWD

18, 19

Therefore, it is important to understand

24


Page 24 of 43

Page 25 of 43


497

and HWDC dispersions, indicating that the steric hindrance from dextran covalently

498

conjugated to WPI and ChS electrostatically interacted with WD was a vital factor for

499

the formation of biopolymer nanoparticles. The results of fluorescence spectroscopy

500

confirmed that heat treatment slightly induced the changes in the hydrophobicity of

501

the microenvironment around fluorescent groups in WD compared to WPI (Figure 2

502

and 3). ChS induced the increase in fluorescence intensity of WD dispersion

503

regardless of heat treatment (Figure 2 and 3), since ChS electrostatically interacted

504

with WD reduced the fluorescence quenching of dextran covalently conjugated to

505

WPI. There was a blue-shift in the λmax and a decrease in the fluorescence intensity of

506

WD and WDC dispersions Compared to WPI dispersion (Figure 2B and 3B),

507

indicating the microenvironment changes around Trp residues in proteins. After heat

508

treatment, WD and WDC dispersions showed a slight red-shift in the λmax and a

509

significant increase in the fluorescence intensity (Figure 2B and 3B), suggesting that

510

the initially buried Trp residues in proteins were exposed to a more hydrophilic

511

microenvironment. These results indicated that the steric hindrance from dextran

512


513

interacted with WD molecules influenced the formation and stabilization of

514

biopolymer nanoparticles. The synchronous fluorescence spectrum showed no

515

significant difference in λmax (△λ = 15 nm) of WPI and WD dispersions (Figure 3A),

516

suggesting that glycosylation did not induce the microenvironment changes around

517

Tyr residues in WD prepared under macromolecular crowding conditions. However,

518

HWD and HWDC dispersions showed a slight blue-shift in the λmax (△λ = 15 nm) and

25



519

significantly increased fluorescence intensity compared to WD and WDC dispersions,

520

indicating that heat treatment promoted Tyr residues of protein to a more hydrophobic

521

microenvironment. Additionally, the fluorescence intensity of WDC dispersion was

522

higher than that of WD dispersion regardless of heat treatment. This might be due to

523

the electrostatic interactions between ChS and WD molecules. The effects of

524

glycosylation, ionic polysaccharide, and heat treatment on the conformational changes

525

of secondary structure of protein were confirmed by far-UV CD spectroscopy and

526

ATR-FTIR spectroscopy (Figure 4 and 5, respectively). Heat treatment slightly

527

decreased the content of β-sheet structure of WPI, which might contribute to the

528

formation of large aggregates of HWPI dispersion. Protein conformational changes

529

were closely related to the diameter changes in WPI, WD, and WDC dispersions

530

regardless of heat treatment. These results suggested that heat treatment did not

531

significantly induce protein conformational changes in the stable biopolymer

532

nanoparticles with smaller diameter, due to the steric hindrance from both dextran

533


534

interacted with WD molecules. Similar findings have previously been reported. 6, 9, 12

535

Although ionic bonds promoted the electrostatic complexation between WD and

536

ionic ChS and facilitated the formation of biopolymer nanoparticles (Figure 6A), their

537

influence for maintaining the stability of biopolymer nanoparticles was negligible

538

(Figure 6B). Hydrophobic interactions played a predominant role in the formation and

539

stabilization of biopolymer nanoparticles (Figure 6). Hydrogen bonds slightly

540

influenced the formation and stabilization of biopolymer nanoparticles (Figure 6).

26


Page 26 of 43

Page 27 of 43


541

Disulfide bonds had no impact on the formation of biopolymer nanoparticles (Figure

542

6A), but partly contributed to the stabilization of biopolymer nanoparticles (Figure

543

6B). Therefore, protein conformational changes were demonstrated in biopolymer

544

nanoparticles, and the mechanism of formation and stabilization of biopolymer

545

nanoparticle was elucidated from the viewpoint of molecular forces. This could help

546

in preparation of stable biopolymer nanoparticles from native or glycosylated protein

547

and ionic polysaccharide. Additionally, hydrophobic bonds were involved in the

548

formation and stabilization of HWDC nanoparticles, suggesting that bioactive

549

compounds could be encapsulated in HWDC nanoparticles by hydrophobic

550

interactions between hydrophobic bioactive compounds and biopolymer nanoparticles.

551

The stable HWDC nanoparticles with pH and salt resistance can be produced

552

large-scalely. Therefore, HWDC nanoparticles could be used as a promising carrier

553

system for hydrophobic nutrients in physiological conditions.

554

AUTHOR INFORMATION

555

Corresponding Author

556

Postal address: State Key Laboratory of Food Science and Technology, School of

557

Food Science and Technology, Jiangnan University, Lihu Road 1800, Wuxi, Jiangsu

558

214122, People’s Republic of China. E-mail: [email protected] (X. Zhang).

559

Tel.: +86 510 85197217. Fax: +86 510 85884496.

560

Funding

561

This research was financially supported by the National 125 Program of China

562

(2013AA102204, 2012BAD33B05, and 2011BAD23B04), the National Natural

27



563

Science Foundation of China (31471624), the Anhui Provincial Natural Science

564

Foundation (1608085MC71 and 1608085MC72), and the Natural Science Research

565

Program of Higher Education Institutions of Anhui Province (KJ2016A065 and

566

KJ2016A800).

567

Notes

568

The authors declare no competing financial interest.

569

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570

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Ozimek, L.; Kowaluk, G.; Gustaw, W.; Wesołowska-Trojanowska, M. Changes of

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secondary structure and surface tension of whey protein isolate dispersions upon pH

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and temperature. Czech J. Food Sci. Vol 2014, 32, 82-89.

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(40) Zhuo, X.-Y.; Qi, J.-R.; Yin, S.-W.; Yang, X.-Q.; Zhu, J.-H.; Huang, L.-X.

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Formation of soy protein isolate–dextran conjugates by moderate Maillard reaction in

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macromolecular crowding conditions. J. Sci. Food Agric. 2013, 93, 316-323.

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(41) Zhang, S.; Zhang, Z.; Lin, M.; Vardhanabhuti, B. Raman spectroscopic

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characterization of structural changes in heated whey protein isolate upon soluble

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complex formation with pectin at near neutral pH. J. Agric. Food Chem. 2012, 60,

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12029-12035.

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(42) Costantino, H. R.; Griebenow, K.; Mishra, P.; Langer, R.; Klibanov, A. M.

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Fourier-transform infrared spectroscopic investigation of protein stability in the

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lyophilized form. Biochim. Biophys. Acta-Protein Struct. Molec. Enzym. 1995, 1253,

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69-74.

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(43) Prabakaran, S.; Damodaran, S. Thermal unfolding of β-lactoglobulin:

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Characterization of initial unfolding events responsible for heat-induced aggregation.

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(44) Kauzmann, W. Some factors in the interpretation of protein denaturation. In

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Advances in Protein Chemistry, Anfinsen, C. B.; Anson, M. L.; Bailey, K.; Edsall, J.

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T., Eds.; Academic Press: New York and London, 1959; Vol. 14, pp 1-63.

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Equilibrium denaturation of recombinant human FK binding protein in urea.

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Stability of casein micelles cross-linked by transglutaminase. J. Dairy Sci. 2006, 89,

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proteins by urea. Proc. Natl. Acad. Sci. U. S. A. 2003, 100, 5142-5147.

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715

Figure Captions

716

Figure 1. Z-average diameter and polydispersity index (PDI) of biopolymer particles

717

of various dispersions. The dispersions were adjusted to pH 5.2 or heated at pH 5.2

718

and 85 °C for 15 min. Means ± standard deviation of triplicate analysis are given.

719

Different letters indicate a significant difference (p < 0.05).

720

Figure 2. Intrinsic fluorescence emission spectra of various dispersions at the

721

excitation wavelength of 280 nm (A) and 295 nm (B). The preparation conditions of

722

the dispersions were as in Figure 1. The protein concentration in each dispersion was

723

diluted to 0.2 mg/mL for analysis.

724

Figure 3. Synchronous fluorescence spectra of various dispersions at the △λ = 15 nm

725

(A) and △λ = 60 nm (B). The dispersions were the same as in Figure 2.

726

Figure 4. Far-UV CD spectra (A) and secondary structures (B) of various dispersions.

727

The dispersions were the same as in Figure 2. Means ± standard deviation of triplicate

728

analysis are given. Different letters indicate a significant difference (p < 0.05).

729

Figure 5. ATR-FTIR spectra of various samples (A), ATR-FTIR spectrum and

730

curve-fitting individual component bands in the amide I band region of WPI (B), and

731

protein secondary structures in various samples (C). The preparation conditions of

732

various samples were as in Figure 1 and dried at room temperature. Means ± standard

733

deviation of triplicate analysis are given. Different letters indicate a significant

734

difference (p < 0.05).

735

Figure 6. The contributions of molecular forces for the formation and stabilization of

736

biopolymer nanoparticles. Biopolymer nanoparticle dispersions were prepared by

35



737

heating mixed solutions of WD and ChS in the presence of various dissociating

738

reagents [0.6 M NaCl (S1), 1.5 M urea (S2), 8.0 M urea (S3), 10 mM DTT (S4)] at

739

pH 5.2 and 85 °C for 15 min (A). Biopolymer nanoparticle dispersions were diluted in

740

various dissociating solutions [0.6 M NaCl (S5), 0.6 M NaCl and 1.5 M urea (S6), 0.6

741

M NaCl and 8.0 M urea (S7), 0.6 M NaCl and 8.0 M urea plus 10 mM DTT (S8)]

742

after they were prepared by heating mixed solutions of WD and ChS in the absence of

743

any dissociating reagents at pH 5.2 and 85 °C for 15 min (B). Means ± standard

744

deviation of triplicate analysis are given. Different letters indicate a significant

745

difference (p < 0.05).

36


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Page 37 of 43


Figure 1

37



Figure 2

38


Page 38 of 43

Page 39 of 43


Figure 3

39



Figure 4

40


Page 40 of 43

Page 41 of 43


Figure 5

41



Figure 6

42


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Table of Contents (TOC)

43


Mechanism of Formation and Stabilization of Nanoparticles Produced

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