Article pubs.acs.org/JAFC
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† †
State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Lihu Road 1800, Wuxi, Jiangsu 214122, People’s Republic of China ‡ College of Biological and Chemical Engineering, Anhui Polytechnic University, Beijing Middle Road, Wuhu, Anhui 241000, People’s Republic of China ABSTRACT: Protein conformational changes were demonstrated in biopolymer nanoparticles, and molecular forces were studied to elucidate the formation and stabilization mechanism of biopolymer nanoparticles. The biopolymer nanoparticles were prepared by heating electrostatic complexes of whey protein isolate (WPI)−dextran conjugate (WD) and chondroitin sulfate (ChS) above the denaturation temperature and near the isoelectric point of WPI. The internal characteristics of biopolymer nanoparticles were analyzed by several spectroscopic techniques. Results showed that grafted dextran significantly (p < 0.05) prevented the formation of large aggregates of WD dispersion during heat treatment. However, heat treatment slightly induced the hydrophobicity changes of the microenvironment around fluorophores of WD. ChS electrostatic interaction with WD changed the fluorescence intensity of WD regardless of heat treatment. Far-UV circular dichroism (CD) and attenuated total reflectance Fourier transform infrared (ATR-FTIR) spectroscopies confirmed that glycosylation and ionic polysaccharide did not significantly cause protein conformational changes in WD and ChS (WDC) during heat treatment. In addition, hydrophobic bonds were the major molecular force for the formation and stabilization of biopolymer nanoparticles. However, hydrogen bonds slightly influenced their formation and stabilization. Ionic bonds only promoted the formation of biopolymer nanoparticles, while disulfide bonds partly contributed to their stability. This work will be beneficial to understand protein conformational changes and molecular forces in biopolymer nanoparticles, and to prepare the stable biopolymer nanoparticles from heating electrostatic complexes of native or glycosylated protein and polysaccharide. KEYWORDS: stabilization, nanoparticle, whey protein isolate, dextran, conjugate, chondroitin sulfate, electrostatic complex
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INTRODUCTION Biopolymer nanoparticles, as a delivery vehicle for hydrophobic bioactive compounds, have attracted great attention due to their remarkable nonantigenicity, biocompatibility, biodegradability, and abundant renewable properties.1−5 The stability of biopolymer nanoparticles, especially under different physiological conditions, is essential to prevent microstructural destruction before reaching certain target sites after uptake.3−5 Biopolymer nanoparticles have been prepared using native or glycosylated protein and ionic polysaccharide by complex coacervation or heat-induced gelation methods.3−8 Many researchers have focused on the optimization of heat-induced nanoparticle formulation from native or glycosylated protein and ionic polysaccharide as well as their applications. Nevertheless, much less attention has been paid to the protein conformational changes in stable biopolymer nanoparticles and the formation and stabilization mechanism of biopolymer nanoparticles from the viewpoint of molecular forces, which were prepared by heating electrostatic complexes of glycosylated protein and ionic polysaccharide. Whey protein isolate (WPI)−dextran conjugate (WD) and chondroitin sulfate (ChS) were selected as models of glycosylated protein and ionic polysaccharide, respectively. WPI, a byproduct of cheese or casein manufacturing, has been © 2016 American Chemical Society
widely used as an ingredient in food products for its good nutritional quality and remarkable functional properties, such as emulsification, foaming ability, and gelation. WPI mainly consists of several globular proteins, including β-lactoglobulin (β-lg), α-lactalbumin (α-la), bovine serum albumin (BSA), and immunoglobulins (IGs).9 β-Lg is one of the major components of WPI and determines functional properties of WPI. The stability of complex coacervates or heat-induced nanoparticles formed by WPI and ionic polysaccharide significantly decreased at the specific pH and/or at higher salt concentrations, leading to precipitation or dissociation.10,11 The Maillard reaction, a nonenzymatic glycosylation, is a series of complex reactions between free amino groups of protein and reducing carbonyl groups of polysaccharide, which usually occurs during thermal processes in food systems. It has been reported that the Maillard reaction can significantly improve the solubility, thermal stability, and emulsification properties of the original proteins.9,12 Dextran, a neutral polysaccharide with low viscosity, high solubility, and no gelation, was selected as a Received: Revised: Accepted: Published: 5539
March 15, 2016 May 3, 2016 June 21, 2016 June 22, 2016 DOI: 10.1021/acs.jafc.6b01213 J. Agric. Food Chem. 2016, 64, 5539−5548
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Journal of Agricultural and Food Chemistry
China). ChS consisted of 95.4% sodium ChS and 4.6% protein. Hydrochloric acid (HCl), sodium hydroxide (NaOH), o-phthalaldehyde (OPA), sodium chloride (NaCl), urea, and dithiothreitol (DTT) were purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). All materials were used without any further purification. All aqueous solutions were prepared with deionized water. Preparation of Stable Biopolymer Nanoparticles from WD and ChS. The stable biopolymer nanoparticles were prepared by heating electrostatic complexes of WD and ChS according to the methods described in our previous paper.4 Briefly, WPI and dextran were dissolved in 10 mM sodium phosphate buffer solution (PBS) (pH 6.5) with 0.02% (w/v) sodium azide, and adjusted to 7.5 and 22.5% (w/w), respectively, and to pH 6.5 using 1.0 M HCl or 1.0 M NaOH. After storage at 4 °C overnight for the complete hydration, the mixed solutions were incubated in a water bath for 48 h at 60 °C. When the Maillard reaction of the mixed solutions was finished, the reacted solutions were immediately cooled in an ice−water bath. The degree of glycosylation (DG) of WD was 9.7%, which was determined by the o-phthalaldehyde (OPA) assay from the loss of free amino groups of WPI. pH 6.5 and 60 °C were used to obtain the maximal production of Schiff base, which was the initial product of the Maillard reaction.9 The optimal concentrations of WPI and dextran under macromolecular crowding conditions were obtained under 7.5 and 22.5% (w/w), respectively.4 The incubated time of 48 h could achieve an appropriate DG of WD to prepare the stable biopolymer nanoparticles.4 ChS (1%, w/v) stock solution was obtained after dissolving ChS in deionized water and gently stirring for 2 h at room temperature. The WD stock solution and ChS stock solution were mixed (denoted as WDC). The concentrations of WPI, dextran, and ChS in WDC solution were adjusted to 0.2, 0.55, and 0.008% (w/v), respectively. After stirring for 2 h, the mixed solutions were adjusted to pH 5.2 [near the isoelectric point (pI) of WPI] with 0.1 M HCl, and heated at 85 °C for 15 min. The nanoparticle dispersions were immediately cooled for 10 min in an ice−water bath. Our previous studies showed that the secondary aggregation of heated WD (HWD) dispersion would occur at pH 4.0, and the biopolymer nanoparticles from heated WDC (HWDC) dispersion had Z-average mean diameter around 150 nm with polydispersity index (PDI) 0.08 in the pH range 1.0 to 8.0 regardless of 0.2 M NaCl. Additionally, ChS, WPI, and WD with 9.7% DG were assembled into the spherical shape and smooth surface biopolymer nanoparticles with dextran conjugated to WPI/ ChS shell and WPI/ChS core during heat treatment.4 In this work, protein conformational changes were demonstrated in biopolymer nanoparticles, and molecular forces were studied to elucidate the formation and stabilization mechanism of biopolymer nanoparticles, prepared by heating electrostatic complexes of WD with 9.7% DG and ChS. The stable biopolymer nanoparticle dispersions were kept at 4 °C before analysis. WPI and WD solutions were prepared and treated under the same conditions described above. All experiments were performed in triplicate. Intrinsic Fluorescence Emission Spectroscopy. The intrinsic fluorescence emission spectra were determined at room temperature (25 °C) using a fluorescence spectrophotometer (F-7000, Hitachi Co., Ltd., Japan). The protein concentration in each sample was diluted to 0.2 mg/mL in sodium phosphate−citric acid buffer (10 mM, pH 5.2). The emission spectra were separately recorded from 285 to 450 nm and 300 to 450 nm at the excitation wavelength of 280 and 295 nm both with a slit width of 2.5 nm, respectively. The corresponding sample without WPI was used as a control to correct the fluorescence background. Synchronous Fluorescence Spectroscopy. Synchronous fluorescence spectrometry has been widely used in multicomponent analysis to distinguish the microenvironment changes around different fluorescent groups.23 Synchronous fluorescence measurements were performed at room temperature (25 °C) using a fluorescence spectrophotometer (F-7000, Hitachi Co., Ltd., Japan). To obtain the microenvironment changes around individual tyrosine (Tyr) and tryptophan (Trp) residues in proteins, the synchronous fluorescence spectra of the same samples as intrinsic fluorescence experiments were recorded from 240 to 360 nm at fixed 15 and 60 nm intervals between
source of polysaccharide for the Maillard reaction to avoid complications during the formation of electrostatic complexes between negatively and positively charged biopolymers. Dextran covalently conjugated to protein can provide steric hindrance against protein thermal aggregation.9 ChS, a linear glycosaminoglycan, is composed of a polymerized disaccharide unit containing β-1,4-linked glucuronic acid and β-1,3-N-acetyl galactosamine, and sulfated at either the 4 or 6 position of the galactosamine residue.13 Additionally, ChS has many interesting properties, including biocompatibility, biodegradability, and targetability.14 Therefore, ChS chains have lots of anionic charges and can be used as a vehicle of bioactive compounds in delivery system with positively charged substances by electrostatic interactions. Functional properties of proteins are closely related to their structures, and protein structures are dependent on hydrophobic bonds, ionic bonds, van der Waals forces, hydrogen bonds, and disulfide bonds.15−19 Intrinsic and synchronous fluorescence spectroscopies have been used to investigate the structure, interactions, and dynamics of proteins in solution due to their high sensitivity, simplicity, and rapidity.20−23 Circular dichroism (CD) spectroscopy has been widely used to evaluate the protein conformation in solution. However, Fourier transform infrared (FTIR) spectroscopy is an excellent technique to determine the protein conformation in solutions, thin films (dry or hydrated), solids (spray-dried or lyophilized powders), or suspensions of precipitates.24−27 Compared to the traditional transmission FTIR, thin-film attenuated total reflectance FTIR (ATR-FTIR) spectroscopy is highly sensitive due to the absence of major water peak in the hydrated thinfilm sample.28,29 The contributions of different molecular forces involved in protein gels or biopolymer nanoparticles can be determined by the solubility of protein gels or particle size of biopolymer nanoparticles in various chemical reagents, which differ from each other by their functional ability to cleave specific bonds: ionic bonds (NaSCN, Na2SO4, CH3COONa, NaCl), hydrogen and hydrophobic bonds [urea, guanidine hydrochloride (GuHCl)], and disulfide bonds [β-mercaptoethanol (β-ME or 2-ME), dithiothreitol (DTT), N-ethylmaleimide (NEM)].18,29−32 The objective of the present study was to evaluate protein conformational changes in biopolymer nanoparticles, and the contributions of different molecule forces on the formation and stabilization of biopolymer nanoparticles, prepared by heating electrostatic complexes of WD and ChS. The biopolymer nanoparticles were characterized by dynamic light scattering, intrinsic fluorescence spectroscopy, synchronous fluorescence spectroscopy, CD spectroscopy, and ATR-FTIR spectroscopy. Finally, protein conformational changes were demonstrated in biopolymer nanoparticles and the mechanism of formation and stabilization of biopolymer nanoparticles was elucidated from the viewpoint of molecular forces, which will facilitate the preparation of stable biopolymer nanoparticles by heat-induced method using native or glycosylated protein and ionic polysaccharide.
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MATERIALS AND METHODS
Materials and Reagents. WPI was obtained from Hilmar Ingredients (Hilmar, CA). The total solid, protein, and ash in the dry power were 95.6, 88.7, and 2.7%, respectively. Dextran with molecular mass of 40 kDa was purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). Chondroitin sulfate (ChS) was kindly provided by Shandong Yibao Biologics Co., Ltd. (Yanzhou, 5540
DOI: 10.1021/acs.jafc.6b01213 J. Agric. Food Chem. 2016, 64, 5539−5548
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Journal of Agricultural and Food Chemistry the excitation and emission wavelength both with a slit width of 2.5 nm, respectively. The fluorescence intensity of each sample blank was subtracted from that of the corresponding sample to obtain the net fluorescence intensity of each protein sample. Far-UV CD Spectroscopy. Far-UV CD spectroscopy of each sample was carried out using a MOS-450 CD spectropolarimeter (Biologic, Claix, France). The spectra were scanned from 190 to 250 nm with a 1 mm path length quartz cuvette at 25 °C. The protein concentration in all samples was diluted to 0.2 mg/mL and adjusted to pH 5.2. The protein spectrum was corrected by subtracting the spectrum of a protein-free solution. The molar ellipticities of protein samples were calculated as [θ] (deg·cm2·dmol−1) = (100 × X × M)/(L × C), where X is the signal (millidegrees) obtained by the CD spectrometer, M is the average molecule weight of amino acid residues in protein (assumed to be 115 for WPI), C is the protein concentration (mg/mL) of the sample, and L is the cell path length (cm).12 Four secondary structures, including α-helix, β-sheet, β-turn, and random coil, were analyzed by the spectra and calculated using DICHROWEB (http://dichroweb.cryst.bbk.ac.uk/html/process. shtml). ATR-FTIR Spectroscopy. Infrared spectra were obtained at room temperature (25 °C) using a FTIR spectrophotometer (Nicolet iS10, Thermo Electron Corp., Madison, WI) equipped with an Ever-Glo MIR source, a KBr beam splitter, and a deuterated triglycine sulfate (DTGS) detector. The spectral data were collected in the range of 650−4000 cm−1 at a 4 cm−1 resolution and a zero filling factor of 1 using a Happ−Genzel apodization and Mertz phase correction. Sixteen scans were accumulated to obtain a reasonable signal-to-noise ratio. An aliquot of each sample (50 μL) was placed on the aluminum foil. After 24 h of storage at room temperature, the dried film of each sample on the foil was formed and then positioned directly on a single reflection diamond attenuated total reflectance (ATR) crystal. The ATR crystal was washed with deionized water and dried with lens paper to avoid contamination between samples. All samples were measured under identical conditions. To ensure no interference from non-protein constituents, each spectrum was obtained by subtracting the corresponding background spectrum from the sample spectrum, using the Nicolet Omnic software (version 8.3, Thermo Electron Corp., Madison, WI). Protein secondary structure is most reliably indicated by the amide I band (1600−1700 cm−1).28 The amide I band of the resulting different spectrum was baseline corrected by two points and smoothed by the 9-point Savitzky−Golay filter method. The second derivative spectrum, obtained using a third degree polynomial function with a 5-point Savitsky−Golay smoothing function, was used to identify the positions of overlapping components of the amide I band. The positions were then confirmed by Fourier self-deconvolution with a full bandwidth at half-height (fwhh) of 13.0 cm−1 and a resolution enhancement factor (K) of 2.4. Finally, the FTIR deconvolution spectra were curve-fitted by Gaussian− Lorentzian function with PeakFit software (Version 4.12, SeaSolve Software Inc., Framingham, MA). Quantitative estimation of protein secondary structure was performed by calculating the corresponding band percentage in the amide I band region according to the following wavenumber ranges: 1620−1645 cm−1, β-sheet; 1645−1652 cm−1, random coil; 1652−1662 cm−1, α-helix; 1662−1690 cm−1, β-turn.24 Determination of Molecular Forces for Formation and Stabilization of Biopolymer Nanoparticles. The contributions of different molecular forces on the formation of biopolymer nanoparticles were determined by preparing their dispersions in the presence of various dissociating reagents. WD and ChS stock solutions were diluted with addition of individual dissociating solution to the above-mentioned concentrations. Meanwhile, the dissociating reagents in the resulting dispersions were adjusted to the final concentrations as follows: 0.6 M NaCl (solution S1), 1.5 M urea (solution S2), 8.0 M urea (solution S3), and 10 mM DTT (solution S4). All other procedures were the same as described above. To determine the contributions of molecular forces on the stabilization of biopolymer nanoparticles, the stable biopolymer nanoparticle dispersions were diluted 10-fold with dissociating reagents, and the dissociating reagents were adjusted to the final concentrations as follows: 0.6 M NaCl
(solution S5), 0.6 M NaCl + 1.5 M urea (solution S6), 0.6 M NaCl + 8.0 M urea (solution S7), and 0.6 M NaCl + 8.0 M urea +10 mM DTT (solution S8).30−33 The diameter changes were used to estimate the contributions of ionic bonds (difference between S1 and control or between S5 and control, respectively), hydrogen bonds (difference between S2 and control or between S6 and S5, respectively), hydrophobic interactions (difference between S3 and S2 or between S7 and S6, respectively), and disulfide bonds (difference between S4 and control or between S8 and S7, respectively) for the formation and stabilization of biopolymer nanoparticles. The particle sizes were measured after 1 h of storage. Dynamic Laser Scattering (DLS) Measurements. The Zaverage mean diameter and polydispersity index (PDI) of biopolymer nanoparticles were obtained by dynamic light scattering using a Malvern Zetasizer (Nano ZS, Malvern Instruments Ltd., Worcestershire, U.K.) equipped with 633 nm and He−Ne laser beam. Measurements were made at 25 °C and 173° scattering angle. The nanoparticle dispersions were measured by dilution with the corresponding solutions to a final protein concentration of 0.2 mg/ mL. Each dispersion was fully shaken before measuring the Z-average mean diameter to ensure a uniform suspension of particles. Statistical Analysis. Each experiment was triplicated under the same conditions. A one-way analysis of variance (ANOVA) was applied to estimate the statistical difference. Significant differences (p < 0.05) between means were determined using Duncan’s multiple range tests. Statistical analyses were evaluated with SPSS software (version 17.0, SPSS Inc., Chicago, IL).
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RESULTS AND DISCUSSION Biopolymer Nanoparticles Prepared Using WPI after Different Treatments. The Z-average mean diameters of WPI, heated WPI (HWPI), WD, HWD, WDC, and HWDC dispersions are shown in Figure 1. Around pI of WPI, the
Figure 1. Z-average diameter and polydispersity index (PDI) of biopolymer particles of various dispersions. The dispersions were adjusted to pH 5.2 or heated at pH 5.2 and 85 °C for 15 min. Means ± standard deviation of triplicate analysis are given. Different letters indicate a significant difference (p < 0.05).
solubility of WPI decreased and formed smaller biopolymer particles about 266 nm with PDI 0.505. However, heat treatment led to the formation of large particle aggregates about 8890 nm with PDI 0.307 in HWPI dispersion. Heat treatment might promote the hydrophobic interactions and repress hydrogen interactions. In addition, near the pI of protein, heat denaturation altered the hydrophobicity/hydrophilicity balance of protein surface, leading to aggregation via hydrophobic interactions. These results are consistent with previous studies.34 The diameter sizes of WDC and HWDC dispersions were 128 and 159 nm, respectively. These results indicated that glycosylation and ionic polysaccharide signifi5541
DOI: 10.1021/acs.jafc.6b01213 J. Agric. Food Chem. 2016, 64, 5539−5548
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Journal of Agricultural and Food Chemistry cantly prevented the formation of large aggregates of WDC dispersion during heat treatment. This was due to the steric hindrance from dextran chains covalently conjugated to WPI molecules and ChS chains electrostatically interacting with WD. Meanwhile, the diameter changes in different WPI samples might be related to protein conformational changes after different treatments. These results are consistent with previous studies.12,35 There was no significant difference in diameter sizes between WD and WDC regardless of heat treatment. In our previous publication, we reported that HWDC dispersion was stable against pH and salt, but the secondary aggregation of HWD dispersion could occur at pH 4.0.4 Based on these findings, further studies on protein conformational changes and molecular forces in the stable HWDC nanoparticles were investigated in the present work. Fluorescence Spectroscopic Analysis. Intrinsic Fluorescence Emission Spectroscopy. Due to the absence of external reagents, the intrinsic fluorescence spectroscopy has been used as a reliable method to evaluate the changes of the microenvironment around fluorescent groups in proteins.22 The intrinsic fluorescence of protein results from aromatic fluorophores, including phenylalanine (Phe), tyrosine (Tyr), and tryptophan (Trp) residues of proteins. Due to a very low quantum yield of Phe residues, the intrinsic fluorescence of many proteins is mainly attributed to Tyr and Trp residues. βLg, α-la, and BSA contain 2, 4, and 2 Trp residues and 4, 4, and 20 Tyr residues per molecule, respectively.21,36 At the excitation wavelength of 280 nm, both Tyr and Trp residues showed a fluorescence emission spectrum, but at the excitation wavelength of 295 nm, only Trp residues showed a fluorescence emission spectrum.20 At the excitation wavelength of 295 nm (Figure 2B), the maximum of emission wavelength (λmax) of WPI dispersion was 335 nm, whereas the λmax of WD dispersion was 333 nm. These results suggested that the polarity around Trp residues in proteins decreased and the hydrophobicity increased, indicating the protein conformational changes. This might be attributed to dextran covalently conjugated to WPI. Meanwhile, the fluorescence intensity of WD dispersion decreased compared to that of WPI dispersion. This might be due to the covalent conjugation of dextran chains to WPI on fluorescence quenching of protein. These results are in agreement with the fluorescence characteristics of Maillard reaction products.37 There was no significant difference in the λmax between WD and WDC dispersions (Figure 2B). When WD or WDC dispersions were heated at 85 °C for 15 min, both λmax were shifted from 333 to 335 nm, and the fluorescence intensity significantly increased (Figure 2B), indicating the increase of polarity and the decrease of hydrophobicity of the microenvironment around Trp residues of proteins. These fluorescence changes might be related to the increase of hydrophobic interactions between protein molecules during heat treatment, which not only contributed to the changes of the microenvironment around fluorophores (Figure 2B) but also promoted the increase in particle diameters of WD and WDC dispersions (Figure 1). Simion et al. reported that the fluorescence intensity of β-lg dispersion significantly increased with increasing temperature (25−85 °C) at the excitation wavelength of 292 nm, and the λmax of β-lg dispersion exhibited a red shift of 2−4 nm after heat treatment at 75−85 °C due to the increase of the exposure of its fluorophores.21 The fluorescence intensities of WDC and HWDC dispersions were higher than those of WD and HWD dispersions, respectively, suggesting that ChS reduced the fluorescence
Figure 2. Intrinsic fluorescence emission spectra of various dispersions at the excitation wavelength of 280 nm (A) and 295 nm (B). The preparation conditions of the dispersions were the same as in Figure 1. The protein concentration in each dispersion was diluted to 0.2 mg/ mL for analysis.
quenching of dextran covalently conjugated to WPI. This change might be attributed to the protein conformational changes in WDC induced by electrostatic interactions between ChS and WD molecules, leading to the decrease of quenching effect of grafted dextran chains on the Trp fluorophore. Similar emission spectra were observed regardless of the excitation wavelengths. Only slight differences in fluorescence intensity between the two emission spectra were observed (Figures 2A and 2B, respectively). Synchronous Fluorescence Spectroscopy. Synchronous fluorescence spectroscopy further distinguished the effects of glycosylation, ionic polysaccharide, and heat treatment on the individual fluorescent groups. At fixed Δλ (15 and 60 nm) between excitation and emission wavelengths, the synchronous fluorescence spectroscopy could provide more accurate information about the microenvironment around individual Tyr and Trp residues in proteins, respectively.23 The shifts of the λmax are related to the changes in the polarity and hydrophobicity of the microenvironment around fluorescent groups in proteins, indicating conformational changes of protein.21 The synchronous fluorescence spectra of all samples resulting from Tyr and Trp residues at Δλ = 15 nm and Δλ = 60 nm are shown in Figures 3A and 3B, respectively. As shown in Figure 3A, the fluorescence intensity of WD dispersion was lower than that of WPI dispersion, indicating that dextran covalently conjugated to WPI quenched the fluorescence of Tyr residues. The fluorescence intensity of WD dispersion increased with addition of ChS, suggesting that ChS reduced the fluorescence quenching of dextran covalently conjugated to 5542
DOI: 10.1021/acs.jafc.6b01213 J. Agric. Food Chem. 2016, 64, 5539−5548
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Journal of Agricultural and Food Chemistry
the latter was much higher than that of the former. Additionally, the fluorescent change trend of Tyr and Trp residues induced by same treatments was different in the synchronous fluorescence spectroscopy (Figures 3A and 3B, respectively). Far-UV CD Spectroscopic Analysis. The far-UV CD spectra of WPI, WD, HWD, WDC, and HWDC dispersions are shown in Figure 4A. Conformational changes in the secondary
Figure 3. Synchronous fluorescence spectra of various dispersions at the Δλ = 15 nm (A) and Δλ = 60 nm (B). The dispersions were the same as in Figure 2.
WPI. WPI, WD, and WDC dispersions had similar λmax, indicating that the polarity of the microenvironment around Tyr residues was not changed. After heating WD and WDC dispersions, their fluorescence intensities increased and their fluorescence spectra showed a slight blue shift of λmax compared to WPI dispersion, indicating that the hydrophobicity of the microenvironment around Tyr residues was slightly changed. Additionally, the fluorescence intensity of HWD dispersion was lower than that of HWDC dispersion, indicating that ChS electrostatically interacted with WD. Simion et al. reported that β-lg dispersion had a 2.5 nm blue shift of the λmax (Δλ = 15 nm) at 80 and 85 °C for burial of Tyr residues and its fluorescence intensity significantly increased, and explained that the polarity around Tyr residues decreased while the hydrophobicity increased.21 As shown in Figure 3B (Δλ = 60 nm), both WD and WDC dispersions showed a slight blue shift in the λmax compared to WPI dispersion. Wu et al. reported that β-lg−fructooligosaccharide conjugate dispersion had a slight blue shift of the λmax (Δλ = 60 nm), and explained that glycosylation of β-lg influenced the hydrophobic microenvironment around Trp residues.38 HWD and HWDC dispersions showed a slight red shift in the λmax compared to WD and WDC dispersions, respectively. Simion et al. reported that β-lg dispersion exhibited a 2.5 nm red shift of the λmax (Δλ = 60 nm) at 80 and 85 °C for exposure of Trp residues and its fluorescence intensity significantly increased, and demonstrated that the polarity around Trp residues increased while the hydrophobicity decreased.21 A similar trend was observed in synchronous fluorescence intensity of the same sample at Δλ = 15 nm and Δλ = 60 nm, whereas the fluorescence intensity of
Figure 4. Far-UV CD spectra (A) and secondary structures (B) of various dispersions. The dispersions were the same as in Figure 2. Means ± standard deviation of triplicate analysis are given. Different letters indicate a significant difference (p < 0.05).
structure of proteins were studied at wavelength range between 190 and 250 nm. The broad negative peak around 206 nm represented α-helix conformation. The secondary structure compositions in all samples are shown in Figure 4B. WPI had an average of 33.0% α-helix, 19.4% β-sheet, 19.8% β-turn, and 27.8% random coil (Figure 4B). Tomczyńska-Mleko et al. demonstrated that WPI had an average 23.1% α-helix, 22.9% βsheet, 22.2% β-turn, and 31.7% random coil at pH 5.0. This difference might be due to different sources of WPI and pH condition.39 Compared to WPI, contents of β-turn and random coil in WD sample slightly increased at the expense of α-helix and β-sheet (Figure 4B), indicating that glycosylation did not significantly change the protein secondary structure of WD obtained under macromolecular crowding conditions. Similar findings have previously been reported.35,40 After heat treatment, the ellipticity of WD became less negative (Figure 4A), and the content of α-helix slightly decreased (Figure 4B), indicating that heat treatment slightly induced protein conformational changes in WD. Perez et al. demonstrated that heat treatment could promote protein conformational changes.22 The secondary structures between HWD and HWDC were slightly different, suggesting that the steric 5543
DOI: 10.1021/acs.jafc.6b01213 J. Agric. Food Chem. 2016, 64, 5539−5548
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the curve-fitting individual component bands in the amide I band region of WPI are shown in Figure 5B. The Fourier selfdeconvolution method was applied to distinguish the individual components in the intrinsically overlapped amide I band contours, which were assigned to different secondary structure conformations.24,28 Curve fittings of the deconvolved spectra were performed using the Gaussian−Lorentzian function. Consequently, quantitative estimation of protein secondary structures, including α-helix, β-sheet, β-turn, and random coil, was obtained.24,25 The percentages of four secondary structures of protein in all samples are shown in Figure 5C. Although there was difference in percentages of four secondary structures of protein, calculated by far-UV CD spectroscopy and ATR-FTIR spectroscopy, the two analytical methods showed a similar change trend in same samples. The difference might be due to different water contents. It has been reported that hydration could significantly increase contents of α-helix and random coil and lower content of β-sheet in protein by FTIR spectroscopy.42 Our results demonstrated that WPI had an average of 14.4% α-helix, 41.0% β-sheet, 29.1% β-turn, and 15.5% random coil. Similar results have previously been reported using ATRFTIR spectroscopy or Fourier transform Raman spectroscopy.26,41 Although the protein conformational changes of protein in the form of suspensions or precipitates could not be determined by fluorescence spectroscopy and CD spectroscopy, they could be determined using FTIR spectroscopy. Heat treatment slightly decreased the content of β-sheet in WPI and slightly increased the contents of β-turn and random coil (Figure 5C). Protein conformational changes might be related to larger aggregates (diameter >8800 nm) in HWPI dispersion. Similar findings have previously been reported.43 Compared to WPI, the percentage of α-helix slightly decreased in WD, and further slightly decreased in HWD (Figure 5C). These results indicated that dextran covalently conjugated to WPI and heat treatment did not significantly induce protein conformational changes of WD. These results are consistent with the findings of CD spectroscopy. ChS did not significantly induce the changes of secondary structures of WD, except for slight increase of β-sheet content in WDC. There was no significant difference in the secondary structures between HWD and HWDC (Figure 5C), indicating that ChS did not significantly change protein conformations of HWD. Molecular Forces for Formation and Stabilization of Biopolymer Nanoparticles. Molecular Forces for Formation of Biopolymer Nanoparticles. The influences of different molecular forces on the formation of biopolymer nanoparticles are shown in Figure 6A. Compared to control sample, 0.6 M NaCl induced the greatest change in the particle size of biopolymer nanoparticles followed by 8.0 M urea, 1.5 M urea, and 10 mM DTT (Figure 6A). Several researchers reported that various dissociating reagents could affect protein heat stability, rheological properties, and molecular forces within protein and water molecules, and demonstrated that the formation of protein gel networks was attributed to the balance of noncovalent interactions (ionic, hydrophobic, and hydrogen bonds) and covalent disulfide bonds.19,30−32 The biopolymer nanoparticles were prepared by heating electrostatic complexes of WD and ChS in the absence or presence of 0.6 M NaCl (control, S1 in Figure 6A, respectively). The Z-average diameter and PDI of biopolymer nanoparticles changed from 156.3 nm and 0.071 to 388.0 nm and 0.439, respectively. Electrostatic shielding effects minimized electrostatic inter-
hindrance from ChS electrostatically interacting with WD did not significantly induce the changes in spatial structure and unfolding of glycosylated protein during heat treatment. Zhang et al. demonstrated that pectin enhanced the thermal stability of WPI structure, and explained that pectin could prevent secondary structural changes of WPI through electrostatic interactions.41 ATR-FTIR Spectroscopic Analysis. The ATR-FTIR spectra of unheated and heated WPI, WD, WDC, dextran (DEX), and dextran/ChS (DC) in the region between 650 and 4000 cm−1 are shown in Figure 5A. Although amide I, II, and III bands of FTIR spectrum can be used to estimate protein secondary structure, amide I band (1600−1700 cm−1) is the most sensitive to protein conformational changes and is widely used in secondary structure analysis.29 The FTIR spectrum and
Figure 5. ATR-FTIR spectra of various samples (A), ATR-FTIR spectrum and curve-fitting individual component bands in the amide I band region of WPI (B), and protein secondary structures in various samples (C). The preparation conditions of various samples were as in Figure 1, and the samples were dried at room temperature. Means ± standard deviation of triplicate analysis are given. Different letters indicate a significant difference (p < 0.05). 5544
DOI: 10.1021/acs.jafc.6b01213 J. Agric. Food Chem. 2016, 64, 5539−5548
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Journal of Agricultural and Food Chemistry
have previously been reported.46−48 These results indicated that although 1.5 M urea could compete with the inter- and intramolecular hydrogen bonds between proteins and water, hydrogen bonds could not be essential for the formation of biopolymer nanoparticles. Under similar preparation conditions in the presence of 8.0 M urea, the biopolymer nanoparticles had a more loose structure due to the reduction of hydrophobic interactions in strength. Therefore, hydrophobic interactions played a prominent role in the formation of biopolymer nanoparticles. DTT was often used to reduce disulfide bonds and prevent disulfide bond formation.19 The diameter size of biopolymer nanoparticles negligibly changed in the presence of 10 mM DTT (S4 in Figure 6A). Thus, disulfide bonds did not significantly influence the formation of biopolymer nanoparticles. Sun and Arntfield reported that no significant difference was observed on storage moduli (G′) with addition of 0.1−0.3 M β-mercaptoethanol (2-ME), 0.05−0.15 M dithiothreitol (DTT), and 10−25 mM N-ethylmaleimide (NEM), and explained that disulfide bonds were not required for gel formation.19 Molecular Forces for Maintaining Stability of Biopolymer Nanoparticles. The influences of different molecular forces on the stability of biopolymer nanoparticles are shown in Figure 6B. The addition of 0.6 M NaCl had almost no effect on the particle size of the stable biopolymer nanoparticles compared to control sample (S5, control in Figure 6B, respectively). Therefore, ionic bonds were not essential for maintaining the stability of biopolymer nanoparticles. Jones and McClements reported that the particle diameter of biopolymer particles, formed by heating β-lg and pectin complexes in the absence of 0.2 M NaCl, had almost no change after diluting their dispersion in the presence of 0.2 M NaCl, indicating its good stability to salt.6 Additionally, Giroux et al. demonstrated that calcium promoted the formation of nanoparticles from denatured whey protein through pH-cycling treatment, but it was not necessary to maintain the stability of biopolymer nanoparticles.32 The particle sizes of biopolymer nanoparticles increased by 12.7 and 182.0% after diluting their dispersion in the presence of 0.6 M NaCl and 1.5 or 8.0 M urea (S6, 7 in Figure 6B), respectively. Different urea concentrations caused different swelling degree of the biopolymer nanoparticles. 1.5 M urea concentration slightly changed the diameter size of biopolymer nanoparticles by breakage of hydrogen bonds. However, 8.0 M urea concentration significantly changed the diameter size by breakage of both hydrogen and hydrophobic bonds.18,30,32,33,49 Therefore, these findings suggested that hydrogen bonds had a slight contribution in maintaining the stability of biopolymer nanoparticles, while hydrophobic bonds had a predominant impact in stabilizing the biopolymer nanoparticles. Due to disulfide reduction of DTT and swelling of urea, the diameter size of biopolymer nanoparticles further increased by 21.2% after dispersion dilution in the presence of NaCl and urea plus DTT (S8 in Figure 6B), and PDI significantly increased to 0.534, suggesting disruption of biopolymer nanoparticle dispersion. These results indicated that disulfide bonds could partly maintain the stability of biopolymer nanoparticles. Previous studies demonstrated that disulfide bonds could partly stabilize gels of heat-induced proteins by dissociating solutions [(0.6 M NaCl + 8.0 M urea + 10 mM DTT) or (0.6 M NaCl + 8.0 M urea + 0.5 M 2-βmercaptoethanol)].30,33
Figure 6. Contributions of molecular forces for the formation and stabilization of biopolymer nanoparticles. Biopolymer nanoparticle dispersions were prepared by heating mixed solutions of WD and ChS in the presence of various dissociating reagents [0.6 M NaCl (S1), 1.5 M urea (S2), 8.0 M urea (S3), 10 mM DTT (S4)] at pH 5.2 and 85 °C for 15 min (A). Biopolymer nanoparticle dispersions were diluted in various dissociating solutions [0.6 M NaCl (S5), 0.6 M NaCl and 1.5 M urea (S6), 0.6 M NaCl and 8.0 M urea (S7), 0.6 M NaCl and 8.0 M urea plus 10 mM DTT (S8)] after they were prepared by heating mixed solutions of WD and ChS in the absence of any dissociating reagents at pH 5.2 and 85 °C for 15 min (B). Means ± standard deviation of triplicate analysis are given. Different letters indicate a significant difference (p < 0.05).
actions between glycosylated protein and ionic polysaccharide molecules and relatively increased hydrophobic interactions, which promoted protein aggregation during heat treatment, indicating the destruction of the original equilibrium between electrostatic and hydrophobic interactions in protein dispersions. These results confirmed that ionic bonds significantly influenced the formation of the biopolymer nanoparticles. Jones et al. reported that there was a weak electrostatic repulsion between biopolymer particles at high salt concentrations, leading to the large particle aggregates.6 Additionally, Melander et al. demonstrated that neutral salts had two antagonistic effects on electrostatic and hydrophobic interactions at higher concentrations.15 Urea (1.5 M) was used to test hydrogen bonds (which break endothermically), while hydrophobic bonds (which break exothermically) and hydrogen bonds were tested with 8.0 M urea.44,45 Compared to control sample, the diameter size of biopolymer nanoparticles decreased by 24.2% in the presence of 1.5 M urea (S2 in Figure 6A), indicating that the biopolymer nanoparticles had a more compact structure. However, the Zaverage diameter and PDI of biopolymer nanoparticles significantly increased to 273.8 nm and 0.488 in the presence of 8.0 M urea (S3 in Figure 6A), respectively. Similar findings 5545
DOI: 10.1021/acs.jafc.6b01213 J. Agric. Food Chem. 2016, 64, 5539−5548
Article
Journal of Agricultural and Food Chemistry Mechanism of Formation and Stabilization of HWDC Nanoparticles. Protein structure is dependent on hydrophobic bonds, ionic bonds, van der Waals forces, hydrogen bonds, and disulfide bonds.18,19 Therefore, it is important to understand protein conformational changes and molecular forces in biopolymer nanoparticles. The diameter size of HWPI dispersion was significantly different from those of HWD and HWDC dispersions, indicating that the steric hindrance from dextran covalently conjugated to WPI and ChS electrostatically interacting with WD was a vital factor for the formation of biopolymer nanoparticles. The results of fluorescence spectroscopy confirmed that heat treatment slightly induced the changes in the hydrophobicity of the microenvironment around fluorescent groups in WD compared to WPI (Figures 2 and 3, respectively). ChS induced the increase in fluorescence intensity of WD dispersion regardless of heat treatment (Figures 2 and 3, respectively), since ChS electrostatically interacting with WD reduced the fluorescence quenching of dextran covalently conjugated to WPI. There was a blue shift in the λmax and a decrease in the fluorescence intensity of WD and WDC dispersions compared to WPI dispersion (Figure 2B and 3B, respectively), indicating the microenvironment changes around Trp residues in proteins. After heat treatment, WD and WDC dispersions showed a slight red shift in the λmax and a significant increase in the fluorescence intensity (Figures 2B and 3B, respectively), suggesting that the initially buried Trp residues in proteins were exposed to a more hydrophilic microenvironment. These results indicated that the steric hindrance from dextran chains covalently conjugated to WPI molecules and ChS chains electrostatically interacting with WD molecules influenced the formation and stabilization of biopolymer nanoparticles. The synchronous fluorescence spectra of WPI and WD dispersions showed no significant difference in the λmax (Δλ = 15 nm) (Figure 3A), suggesting that glycosylation did not induce the microenvironment changes around Tyr residues in WD prepared under macromolecular crowding conditions. However, HWD and HWDC dispersions showed a slight blue shift in the λmax (Δλ = 15 nm) and significantly increased fluorescence intensity compared to WD and WDC dispersions, indicating that heat treatment promoted Tyr residues of protein to a more hydrophobic microenvironment. Additionally, the fluorescence intensity of WDC dispersion was higher than that of WD dispersion regardless of heat treatment. This might be due to the electrostatic interactions between ChS and WD molecules. The effects of glycosylation, ionic polysaccharide, and heat treatment on the conformational changes of secondary structure of protein were confirmed by far-UV CD spectroscopy and ATRFTIR spectroscopy (Figures 4 and 5, respectively). Heat treatment slightly decreased the content of β-sheet structure of WPI, which might contribute to the formation of large aggregates of HWPI dispersion. Protein conformational changes were closely related to the diameter changes in WPI, WD, and WDC dispersions regardless of heat treatment. These results suggested that heat treatment did not significantly induce protein conformational changes in the stable biopolymer nanoparticles with smaller diameter, due to the steric hindrance from both dextran chains covalently conjugated to WPI molecules and ChS chains electrostatically interacting with WD molecules. Similar findings have previously been reported.6,9,12 Although ionic bonds promoted the electrostatic complexation between WD and ionic ChS and facilitated the formation
of biopolymer nanoparticles (Figure 6A), their influence for maintaining the stability of biopolymer nanoparticles was negligible (Figure 6B). Hydrophobic interactions played a predominant role in the formation and stabilization of biopolymer nanoparticles (Figure 6). Hydrogen bonds slightly influenced the formation and stabilization of biopolymer nanoparticles (Figure 6). Disulfide bonds had no impact on the formation of biopolymer nanoparticles (Figure 6A), but partly contributed to the stabilization of biopolymer nanoparticles (Figure 6B). Therefore, protein conformational changes were demonstrated in biopolymer nanoparticles, and the mechanism of formation and stabilization of biopolymer nanoparticles was elucidated from the viewpoint of molecular forces. This could help in preparation of stable biopolymer nanoparticles from native or glycosylated protein and ionic polysaccharide. Additionally, hydrophobic bonds were involved in the formation and stabilization of HWDC nanoparticles, suggesting that bioactive compounds could be encapsulated in HWDC nanoparticles by hydrophobic interactions between hydrophobic bioactive compounds and biopolymer nanoparticles. The stable HWDC nanoparticles with pH and salt resistance can be produced on a large scale. Therefore, HWDC nanoparticles could be used as a promising carrier system for hydrophobic nutrients in physiological conditions.
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AUTHOR INFORMATION
Corresponding Author
*State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Lihu Road 1800, Wuxi, Jiangsu 214122, People’s Republic of China. E-mail:
[email protected]. Tel: +86 510 85197217. Fax: +86 510 85884496. Funding
This research was financially supported by the Program of "Collaborative Innovation Center of Food Safety and Quality Control in Jiangsu Province", the National 125 Program of China (2013AA102204), the National Natural Science Foundation of China (31471624), the Anhui Provincial Natural Science Foundation (1608085MC71 and 1608085MC72), and the Natural Science Research Program of Higher Education Institutions of Anhui Province (KJ2016A065 and KJ2016A800). Notes
The authors declare no competing financial interest.
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