Mechanism of Formation and Stabilization of Nanoparticles Produced

Jun 22, 2016 - 1800, Wuxi, Jiangsu 214122, People,s Republic of China. ‡. College of Biological and Chemical Engineering, Anhui Polytechnic Universi...
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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

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Sulfate

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Qingyuan Dai,†,

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Xiaoming Zhang,∗,† and Chengsheng Jia†

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

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Republic of China

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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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ABSTRACT. Protein conformational changes were demonstrated in biopolymer

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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)−

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

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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

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

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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

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

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

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glycosylation and ionic polysaccharide did not significantly cause protein

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

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

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biopolymer nanoparticles. However, hydrogen bonds slightly influenced their

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

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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

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

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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,

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biocompatibility, biodegradability, and abundant renewable properties.

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stability of biopolymer nanoparticles, especially under different physiological

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conditions, is essential to prevent microstructural destruction before reaching certain

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

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or glycosylated protein and ionic polysaccharide by complex coacervation or

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

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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

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has been paid to the protein conformational changes in stable biopolymer

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nanoparticles and the formation and stabilization mechanism of biopolymer

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

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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)

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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

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

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

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

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

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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

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

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concentrations, leading to precipitation or dissociation.

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

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

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

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

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properties of the original proteins.

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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

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complexes between negatively and positively charged biopolymers. Dextran

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covalently conjugated to protein can provide steric hindrance against protein thermal

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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.

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biodegradability, and targetability.

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

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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

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

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

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hydrogen bonds and disulfide bonds.

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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

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

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

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

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precipitates.

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

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

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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

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

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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),

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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

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

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

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formation and stabilization of biopolymer nanoparticles, prepared by heating

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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

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spectroscopy. Finally, protein conformational changes were demonstrated in

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biopolymer nanoparticles and the mechanism of formation and stabilization of

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biopolymer nanoparticles was elucidated from the viewpoint of molecular forces,

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which will facilitate the preparation of stable biopolymer nanoparticles by

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

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MATERIALS AND METHODS

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

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

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

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

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

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

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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

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aqueous solutions were prepared with deionized water.

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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

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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

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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

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

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

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

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

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

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9

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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

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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

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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

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

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stabilization mechanism of biopolymer nanoparticles, prepared by heating

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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

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experiments were performed in triplicate.

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Intrinsic Fluorescence Emission Spectroscopy. The intrinsic fluorescence emission

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

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

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

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

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

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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

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changes around different fluorescent groups.

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

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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,

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the synchronous fluorescence spectra of the same samples as intrinsic fluorescence

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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,

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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

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sample.

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

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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

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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

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spectra

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

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

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

and

calculated

using

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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

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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

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

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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

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

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28

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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

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

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

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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

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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

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Biopolymer Nanoparticles. The contributions of different molecular forces on the

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formation of biopolymer nanoparticles were determined by preparing their dispersions

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

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

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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

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

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

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molecular forces on the stabilization of biopolymer nanoparticles, the stable

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

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

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

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

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

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

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

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

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

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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

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

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

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

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

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

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

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

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

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

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

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

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(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

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

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

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

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

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

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

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

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

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consistent with previous studies.

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

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glycosylation and ionic polysaccharide significantly prevented the formation of large

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

264

chains covalently conjugated to WPI molecules and ChS chains electrostatically

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

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

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results are consistent with previous studies.

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

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

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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

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the stable HWDC nanoparticles were investigated in the present work.

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Fluorescence Spectroscopic Analysis. Intrinsic Fluorescence Emission Spectroscopy.

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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.

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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

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

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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

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

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

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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

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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.

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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

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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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which not only contributed to the changes of the microenvironment around

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

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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

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

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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

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

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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

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provide more accurate information about the microenvironment around individual Tyr

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and Trp residues in proteins, respectively.

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changes in the polarity and hydrophobicity of the microenvironment around

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The shifts of the λmax are related to the

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fluorescent groups in proteins, indicating conformational changes of protein.

The

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

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△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

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the

hydrophobic

Journal of Agricultural and Food Chemistry

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

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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

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24, 28

Curve fittings of

Journal of Agricultural and Food Chemistry

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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

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43

Compared to

Page 21 of 43

Journal of Agricultural and Food Chemistry

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

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19, 30-32

The

Journal of Agricultural and Food Chemistry

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

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46-48

These

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Journal of Agricultural and Food Chemistry

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

ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry

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

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Page 25 of 43

Journal of Agricultural and Food Chemistry

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

chains covalently conjugated to WPI molecules and ChS chains electrostatically

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

ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry

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

chains covalently conjugated to WPI molecules and ChS chains electrostatically

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).

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Page 27 of 43

Journal of Agricultural and Food Chemistry

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

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Journal of Agricultural and Food Chemistry

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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biochemical characteristics of high-pressure- and heat-induced gels from blue whiting

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(35) Perusko, M.; Al-Hanish, A.; Cirkovic Velickovic, T.; Stanic-Vucinic, D.

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Macromolecular crowding conditions enhance glycation and oxidation of whey

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interactions of α-Lactalbumin: X. Effect of acylation of tyrosyl and lysyl side chains

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on molecular conformations. J. Biol. Chem. 1971, 246, 1909-1921.

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(38) Wu, X.; Liu, M.; Xia, L.; Wu, H.; Liu, Z.; Xu, X. Conjugation of functional

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

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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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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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Fourier-transform infrared spectroscopic investigation of protein stability in the

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

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Equilibrium denaturation of recombinant human FK binding protein in urea.

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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

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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).

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Figure 1

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Figure 2

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Figure 3

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Figure 4

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Figure 5

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Figure 6

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Table of Contents (TOC)

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