Heteroprotein complex formation of soy protein isolate and lactoferrin: Thermodynamic formation mechanism and morphologic structure

Food Hydrocolloids 100 (2020) 105415

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Heteroprotein complex formation of soy protein isolate and lactoferrin: Thermodynamic formation mechanism and morphologic structure Jiabao Zheng a, Qing Gao a, Chuan-he Tang a, b, Ge Ge a, Mouming Zhao a, b, c, Weizheng Sun a, b, * a

School of Food Science and Engineering, South China University of Technology, Guangzhou, 510641, China Overseas Expertise Introduction Center for Discipline Innovation of Food Nutrition and Human Health (111 Center), Guangzhou, 510641, China c Beijing Advanced Innovation Center for Food Nutrition and Human Health, BeijingTechnology & Business University, Beijing, 100048, China b

A R T I C L E I N F O

A B S T R A C T

Keywords: Heteroprotein complex coacervate Electrostatic interaction Hydrogen bond Thermodynamic formation mechanism Morphologic structure

Heteroprotein complex coacervate (HPCC) is one of the most promising electrostatically driven biopolymer materials. In this paper, the formation conditions, thermodynamic formation mechanism, and morphologic structure of soy protein isolate/lactoferrin (SPI/LF) complex coacervate were investigated. The SPI/LF complex coacervates prepared under optimal conditions were electrically neutral and followed the principle of charge compensation. Electrophoresis and isothermal titration calorimetry verified that all individual SPI fractions participated in HPCC and the stoichiometry of the SPI/LF complexes was the same as their optimal mixing ratio. Moreover, SPI/LF complex coacervation was thermodynamically favoured (ΔG < 0), which resulted from en­ tropy gain (TΔS > 0) and negative enthalpy change (ΔH < 0). Not only electrostatic interactions but also hydrogen bonds participated in the complex coacervation. The SPI/LF interaction improved the heat-stability of heat-sensitive lobe in LF. Furthermore, the SPI/LF complex (pH 6.25, SPI/LF ¼ 1:3) exhibited distinct granules, whereas the uniform crosslinking structure appeared in the other SPI/LF complex (pH 6.6, SPI/LF ¼ 1:4). Atomic force microscope showed the sphere complex (pH 6.25-SPI/LF ¼ 1:3) with a diameter of 50–150 nm and the chain-like complex (pH 6.6-SPI/LF ¼ 1:4) with a length of 50–150 nm and a width of 20–80 nm.

1. Introduction Charged polymer materials have a long history in polymer science, and the complex coacervate is one of the most important electrostati­ cally driven polymer materials (Sing, 2017). Complex coacervation is a liquid-liquid phase separation (LLPS) driven by electrostatic interactions between two oppositely charged macromolecules. The earliest research in the field of biopolymer complex coacervate was reported by Bun­ genberg et al. (1929), who studied the complex coacervation of gelatin and gum arabic. Nowadays, the biopolymer-based coacervate is still an interesting research field because of its wide applications, such as food and biomaterials ingredients (Warnakulasuriya & Nickerson, 2018), encapsulants in personal care products and functional food (Eratte, Dowling, Barrow, & Adhikari, 2018), stimuli-responsive sensitive ma­ terials (Fan, Tang, Thomas, & Olsen, 2014), and even revealing under­ lying mechanisms of some diseases (Alberti, Gladfelter, & Mittag, 2019). A considerable amount of literature has been published on complex coacervates of proteins/synthetic polyelectrolytes and proteins/

polysaccharides. Recently, there has been renewed interest in hetero­ protein complex coacervate (HPCC), which is a special case of LLPS because the dense phase is composed of at least two different proteins with opposite charges. HPCC usually forms at low ionic strength and in a pH range less than unity between the isoelectric point (pI) of two pro­ teins and follows the principle of charge and size compensation (Boire et al., 2018). So far, the HPCCs based on lactoferrin (LF) are most studied. Yan et al. (2013) found that LF/β-lactoglobulin coacervate was formed at pH 5.7–6.2 and very low salt concentration, whereas different formation conditions were reported by Anema et al. (2014) due to the difference in β-lactoglobulin monomers. Then, the structure of LF/β-lactoglobulin complex coacervates was also investigated by confocal laser scanning microscopy, small angle neutron scattering, rheology, electrostatic modeling, fluorescence recovery after photo­ bleaching, and solid-state nuclear magnetic resonance (Kizilay et al., 2014; Peixoto et al., 2016). Moreover, the protein purification based on the selective coacervation between LF and the two isoforms of β-lacto­ globulin and the encapsulation for vitamin B9 based on the complex

* Corresponding author. Overseas Expertise Introduction Center for Discipline Innovation of Food Nutrition and Human Health (111 Center), School of Food Science and Engineering, South China University of Technology, Guangzhou, 510641, China. E-mail address: [email protected] (W. Sun). https://doi.org/10.1016/j.foodhyd.2019.105415 Received 17 June 2019; Received in revised form 2 September 2019; Accepted 1 October 2019 Available online 2 October 2019 0268-005X/© 2019 Elsevier Ltd. All rights reserved.

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coacervate of LF and β-lactoglobulin was also mentioned (Chapeau et al., 2016; Tavares, Croguennec, Hamon, Carvalho, & Bouhallab, 2015). In addition, in the few HPCC studies concerning plant protein, Schawartz et al. (2015) structured the assemblies of napin and β-casein, and Adal et al. (2017) identified the conditions for creation of LF/pea protein isolate (PPI) complex coacervates and analyzed their structure (size and morphology) using atomic force microscopy (AFM) and small angle X-ray scattering. Previous HPCC studies have avoided using mixed proteins like PPI due to their complexity, although such mixed proteins are common in plant protein-based materials. Briefly, with the appli­ cation of the complementary and high-resolution tools, the under­ standing of the structure of HPCC is deepening. There is a huge need to establish the correlations between protein structures and the main driven forces behind the physical processes on a thermodynamic point of view. In this paper, soy protein isolate (SPI) and LF were selected owing to the sustainability, abundance, low cost, and functionality of SPI and the foundation and universality of LF in HPCC studies. SPI mainly consists of 15S, 11S, 7S, and 2S fractions according to their sedimentation co­ efficients. The 7S and 11S globulins, also named as glycinin and β-conglycinin, are the two major globulins in SPI. LF is a dumbbellshaped iron-binding glycoprotein with a molar mass of about 83 kDa and is folded in two homologous lobes (C- and N-lobes), each composed by two domains (C1, C2 and N1, N2) (Moore, Anderson, Groom, Har­ idas, & Baker, 1997). LF has a pI of 8.6–8.9 and consequently interacts strongly with different acidic proteins under specific conditions. The purposes of this study were to (1) investigate the conditions for the creation of SPI/LF complex coacervate; (2) reveal the stoichiometry, thermal properties, and thermodynamic formation mechanism of SPI/LF complex coacervate; (3) character the apparent structure of hetero­ protein complex coacervates; (4) explore the correlations between pro­ tein structures and the main driven forces on a thermodynamic point of view.

0.1 M NaOH or HCl and centrifuged at 12000 g for 40 min. The protein content of the supernatant was defined as SPI solubility. Given SPI is a mixed protein, mass concentration was used instead of molar concentration in this study. Three common methods of preparing heteroprotein complexes were performed in this study. For “pH first”, working solutions (1 g/L) were adjusted to target pH using 0.1 M NaOH or HCl. Then, LF and SPI solutions at the same pH were proportionally mixed using a magnetic stirrer to maintain a constant total biopolymer concentration. After mixing for 30 min, the turbidity, hydrodynamic radius, and ζ-potential of mixed protein solutions were measured. Biopolymer-riched phase (complex coacervate) was collected using centrifugation at 3000g for 10 min. For “low to high”, LF and SPI solu­ tions were proportionally mixed at pH 2.5 and then the mixtures were adjusted to target pH using a magnetic stirrer. For “high to low”, LF and SPI solutions at pH 10.5 were proportionally mixed and then the mix­ tures were rapidly adjusted to target pH using a magnetic stirrer. After mixing for 30 min, the turbidity of mixed protein solutions collected from “low to high” and “high to low” were recorded. All experiments were performed at 25 � C. 2.3. Turbidity measurements The turbidity of mixtures was measured at 600 nm in 1 cm path­ length glass cuvettes using a SHIMADZU UV-1800 spectrophotometer (SHIMADZU Co., Japan). Milli-Q water was used as blank (100% transmittance). The turbidity (T) was calculated as follows (eq (1)): T¼

ln

I I0

(1)

Where I is the transmitted intensity and I0 is the incident light intensity. The turbidity growth kinetic was also recorded (0–1000 s) using a kinetic function supported by SHIMADZU UV-1800 spectrophotometer (SHIMADZU Co., Japan). The interval time between two measurements was set at 0.5 s.

2. Materials and methods 2.1. Materials

2.4. Particle size and ζ-Potential measurements

The bovine LF powder was purchased from Fonterra Cooperative group (New Zealand) and contained 96% protein (Kjeldahl, N � 6.38, dry basis). SPI was extracted by alkali extraction-acid precipitation ac­ cording to Chen, Zhao, and Sun (2013). After pH was adjusted to 7.0 with 2.0 M NaOH, the neutral SPI solution was dialysis (10 kDa) at 4 � C for 48 h, then freeze-dried and stored at 20 � C until use. As determined by inductively coupled plasma optical emission spectrometer (ICP-OES), LF contained 0.103% Na, 0.003% K, < 0.0005% Mg, < 0.002% Ca, 0.013% Fe; SPI contained 0.720% Na, 0.0369% K, 0.0635% Mg, 0.0323% Ca. Defatted soybean meal used in this study was purchased from Shandong Yuwang Industrial Co., Ltd. (Shandong, China). Milli-Q water (18.2 MΩ cm) filtered by Milli-Q apparatus (Millipore Corp., Bedford, MA) was used as a solvent in all experiments. Hydrochloric acid (HCl) and sodium hydroxide (NaOH) were purchased from Sinopharm Chemical Reagent Co., Ltd.

The mean hydrodynamic radius (Rh) and ζ-Potential of mixtures were determined by a Malvern Zetasizer Nano ZS (Malvern Panalytical Ltd., UK). Rh was measured by dynamic light scattering (DLS) at 25 � C equipped with a 4 mW helium/neon laser at a wavelength output of 633 nm and aligned for backscattering at a detection angle of 173 � C. The Rh was calculated using Stokes-Einstein equation (Rh ¼ kT=ð6πηDTÞ), where k is the Boltzmann constant, T is absolute tem­ perature, and η is solvent viscosity. The ζ-potential of mixtures was measured using a laser Doppler velocimetry and phase analysis light scattering (M3-PALS0) using disposable electrophoretic mobility cells. Equilibration time was set at 120 s and results were reported as mean Zaverage diameter and mean ζ-potential for 6 measurements (all samples were performed in duplicate, and each sample was measured 3 times, at least 11 runs for each measurement). 2.5. Microscopic observation

2.2. Preparation of SPI/LF complex coacervate

The images of SPI/LF complexes were immediately observed from the bright-field of an OLYMPUS ix73 (OLYMPUS, Japan) using objec­ tives with magnifications 40 times after mixing the working solution of SPI and LF at target pH in proportion.

LF and SPI dispersions were prepared by dissolving an excess LF powder or PPI powder in Milli-Q water overnight using a magnetic stirrer to ensure complete solubilization. The dispersions were centri­ fuged at 12000 g for 40 min, filtered through 0.22 μm syringe filter to remove residues. Then the protein content of these dispersions was determined by the Kjeldahl method. LF and SPI (5 g/L) stock solutions were obtained by diluting corresponding dispersions, and working so­ lutions (1 g/L) were prepared by diluting stock solutions. The protein solubility of SPI at pH 6.0–7.0 was determined by the Kjeldahl method. The pH of the SPI working solution was adjusted to target pH using

2.6. Isothermal titration calorimetry (ITC) The MicroCal PEAQ-ITC (Malvern Panalytical Ltd., UK) was used to perform the thermodynamic analysis. The LF working solution (1 g/L) and SPI stock solution (5 g/L) were adjusted to target pH (no longer change in an hour). Then, the sample cell (200 μL) and reference cell 2

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were filled with LF (1 g/L) and Milli-Q water, respectively. The injection syringe was filled with 40 μL of SPI (5 g/L). After equilibration, LF was titrated by SPI with 19 successive injections (injection volume 2 μL) at a constant temperature of 25 � C. The initial delay, interval between ti­ trations, and stir speed were set at 60 s, 150 s, and 750 rpm, respectively. The blank was obtained by titration of SPI in the sample cell containing Milli-Q water and subtracted from the sample data. After subtraction, final results were obtained by MicroCal PEAQ-ITC Analysis Software, using the “One set of sites” fitting model.

Instruments Inc., Santa Barbara, USA). The samples were diluted by Milli-Q water to 5 μg/mL of total polymer concentration at the required pH. A volume of 5 μL final samples was pipetted onto a mica disk and incubated overnight until drying. Samples were scanned using tapping mode in air, at a scan rate of 0.977 Hz. Images were obtained at 3 μm scan size and 512-pixel resolution. AFM images were first analyzed using Nanoscope Analysis software (Bruker, version 1.8). Then, the final im­ ages were converted into 8-bit images. And the length, width, and area of each particle in these images is obtained through threshold adjust­ ment, ellipse fitting, and particles analysis using ImageJ (National In­ stitutes of Health) according to Adal et al. (2017).

2.7. Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDSPAGE)

2.12. Statistical analysis

Electrophoresis was performed on a Bio-Rad Mini-PROTEAN elec­ trophoresis system (Bio-Rad Laboratories, Hercules, CA) using 4–20% gradient gel. Marker with molecular weights of 10–180 kDa was used as standard. Samples were dissolved by 4 � loading buffer with DTT (Solarbio Ltd., China), and then boiled for 5 min. The prepared solutions (containing 20 μg or 25 μg protein) were loaded into each sample lane. Lane 1 (5 μg SPI and 15 μg LF) and Lane 2 (5 μg SPI and 20 μg LF) were set as control. They had the same amount of SPI instead of total protein for easier to distinguish the difference between these two lanes due to the overlap of the main subunits of LF and SPI. The prepared solutions containing 20 μg lyophilized coacervate (pH 6.25, SPI:LF ¼ 1:3) and 25 μg coacervate (pH 6.6, SPI:LF ¼ 1:4) were loaded into lane 3 and lane 4, respectively. They have the same total protein content as their respective control group. The molecular weight and intensity of each subunit were analyzed by Azure Biosystem C280 (Azure Biosystem, USA) equipped with Azure spot 2.0 software.

Values were expressed as means � standard deviations of triplicate determinations. The significant difference was determined at the P < 0.05 level for Duncan’s multiple range test by using SPSS software (version 16.0, Chicago, USA). 3. Results and discussion 3.1. Effect of pH and SPI/LF ratio on the formation of heteroprotein complexes The effect of pH and mass ratio on the formation of SPI/LF complexes was firstly studied through “pH first”. Fig. 1A–G shows the turbidity of SPI/LF mixtures with different mass ratios at pH 5.75–7.0 and their turbidity growth kinetics (0–1000 s). Turbidity is one of the primary indicators in the research of complex coacervation, and the mixture with the highest turbidity was generally regarded as the complex coacervate in an optimal condition (Alberti et al., 2019). In current study, among the heteroprotein blends with the highest turbidity, a higher the pro­ portion of LF was observed at a higher pH due to the changes in the charges of SPI and LF. The SPI/LF mixtures in optimal conditions at pH 5.75–6.75 exhibited higher turbidity (>2). However, SPI self-aggregation can be clearly observed at pH 5.75 and 6.0, even the SPI at pH 6.25 has begun to self-aggregate (Fig. 1A–C). A cuvette containing 1 g/L SPI solution at pH 5.75 was placed in the left-most position of Fig. 1A to show this phenomenon visually. It was suggested that self-aggregation could obscure or interfere with coacervation (Yan et al., 2013). Therefore, we focused on the formation of heteroprotein com­ plexes at pH 6.0–6.75 due to their higher turbidity and less self-aggregation. Fig. 1G shows the turbidity growth of SPI/LF mixtures at optimal conditions (pH 6.0-SPI/LF ¼ 1:2.5, pH 6.25-SPI/LF ¼ 1:3, pH 6.5-SPI/ LF ¼ 1:4, and pH 6.75-SPI/LF ¼ 1:4) from 0 s to 1000 s. The SPI/LF mixtures at lower pH not only had higher final turbidity but a faster growth rate of turbidity. Moreover, the turbidity of all mixtures has a higher growth rate before 100 s. Interestingly, Anema et al. (2016) also observed the increase in turbidity of LF and caseins mixtures from 0 s to 300 s using a similar approach. They found that the turbidity developed more rapidly for κ-casein and β-casein, reaching the plateau point within 100 s, whereas it was significantly slower for α-casein, with the turbidity still increasing after 300 s. In this study, the turbidity of SPI/LF mixtures still slowly increased after 1000 s by comparing the turbidity at 30 min with that of the first 1000 s. That turbidity increased over a long time suggested SPI/LF coacervate can fuse or coalesce up to submicron and even micron size due to the potentially metastable property of liquid assemblies which can transition into glassy or gel-like states which do not have classical liquid properties (Alberti et al., 2019). However, turbidity measurements only detect a variety of assemblies but cannot differentiate their size and underlying mechanisms. Fig. 1H and I presents the hydrodynamic radius and ζ-potential of SPI/LF mix­ tures as a function of SPI/LF mass ratio at pH 6.0–6.75, respectively. The changes in hydrodynamic radius displayed a high degree of consistency with the results of turbidity. The SPI/LF mixtures with highest turbidity

2.8. Fourier transform infrared spectroscopy (FT-IR) The FT-IR was performed using a VERTEX 70 FTIR Spectrometer (Bruker Co. Ltd., Germany). The freeze-dried powder of LF, SPI, and LF/ SPI complexes was crushed with KBr and pressed into a transparent sheet by a tabletting machine. All the spectra were recorded with an average of 32 scans from 4000 to 400 cm 1 at a resolution of 4 cm 1. The background was taken under the same conditions as the test samples. 2.9. Differential scanning calorimetry (DSC) The DSC was performed using a DSC3 STARe System (METTLER TOLEDO, USA). Indium (In) standards were used to calibrate the energy and temperature of the equipment, and nitrogen was the purge gas. The freeze-dried proteins and complexes powder (3 mg) was weighed in standard aluminium crucibles (40 μL) using ME analytical balance (METTLER TOLEDO, USA). At least twice the amount of water was added to adequate hydration (6 h after sealed). Samples were analyzed over a temperature range from 30 to 110 � C at the rate of 5 � C/min. An empty sealed crucible was used as a reference. The transition tempera­ tures (onset, peak, and endset temperatures) and enthalpy change were analyzed by the STARe Evaluation (METTLER TOLEDO, USA). All ex­ periments were performed in triplicate. 2.10. Scanning electron microscope (SEM) The microstructure of the freeze-dried proteins and complex powder was observed with a scanning electron microscope TM3000 (Hitachi Co., Tokyo, Japan) at a scanning voltage of 15 kV. The final samples were scanned at 200–10000 magnifications. 2.11. Atomic force microscope (AFM) The morphology of SPI, LF, and their complex coacervates was investigated with a Veeco PicoForce atomic force microscopy (Veeco 3

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Fig. 1. The effect of mass mixing ratio (SPI/LF) on turbidity of SPI/LF mixture at (A) pH 5.75; (B) pH 6.0; (C) pH 6.25; (D) pH 6.5; (E) pH 6.75; (F) pH 7.0, prepared by “pH-first” procedure. (G) Changes in turbidity after mixing LF and SPI. The effect of mixture mass ratio (SPI/LF) on (H) hydrodynamic radius and (I) ζ-potential of SPI/LF mixture at pH 6.0, 6.25, 6.5; 6.6, and 6.75, prepared by “pH-first” procedure.

(pH 6.0-SPI/LF ¼ 1:2.5, pH 6.25-SPI/LF ¼ 1:3, pH 6.5-SPI/LF ¼ 1:4, and pH 6.75-SPI/LF ¼ 1:4) also had the largest hydrodynamic radius of 1987 nm, 1731 nm, 1519 nm, and 1416 nm, respectively (Fig. 1H). By contrast, the PPI/LF complexes showed a smaller hydrodynamic radius of approximately 750 nm (Adal et al., 2017). It should be pointed out that this discrepancy may result from different biopolymer concentra­ tions and formation times besides different types of biopolymers. Similar to changes in turbidity, the hydrodynamic radius still changed over time (Anema et al., 2016). However, the description of the time interval be­ tween mixing and measurement was omitted in many studies. The changes in the ζ-potential implied electrostatic driving force for complex coacervation between the negative charged SPI and the posi­ tively charged LF. As shown in Fig. 1I, the SPI/LF mixtures gradually carried more positive charges and underwent a conversion from a negative charge to a positive charge as LF increased in these mixtures. At the same mass ratio, the higher pH of the mixture, the more negative charge. Furthermore, the mixtures with maximum turbidity and hy­ drodynamic radius (pH 6.0-SPI/LF ¼ 1:2.5, pH 6.25-SPI/LF ¼ 1:3, pH 6.5-SPI/LF ¼ 1:4, and pH 6.75-SPI/LF ¼ 1:4) approached to charge neutrality as expected, and their ζ-potential were 1.14 mV, 0.05 mV, 1.19 mV, and 0.90 mV, respectively. The complex coacervation of SPI and LF strictly adheres to the charge compensation. Furthermore, in order to reduce the build-up of charge in the mixture (SPI/LF ¼ 1:4), the hydrodynamic radius (1660 nm) and ζ-potential (0.42 mV) of the mixture (pH 6.6-SPI:LF ¼ 1:4) were also provided (Fig. S1). The procedures of “low to high” and “high to low” were performed to further investigate the effect of pH on SPI/LF complex formation and pH-induced phase transitions. The turbidity of 1 g/L SPI and LF as a function of pH was recorded to distinguish between protein selfaggregation and heteroprotein complex coacervation. As shown in Fig. 2A, the turbidity of SPI alone is greater than 1 in the range of 4–6.5 owing to self-aggregation, obtained by titration with NaOH (“low to high”). By contrast, the turbidity of LF only slightly increased around its isoelectric point (pH 8.6–9.5). However, for “high to low”, the turbidity of single SPI was greater than 1 from pH 3 to 5.5, and that of LF increased at the pH 7.5–9 (Fig. 2B). That is, the turbidity curve shifted

toward the titration direction. Not only self-aggregation of a single protein, but asymmetry caused by titration path also occurred in the heteroprotein complex. Yan et al. (2013) proposed these asymmetries are different in that the ascending side is steeper than the descending side for the titration from low to high pH and ascribed this to the discrepancy between aggregation and disaggregation. But similar asymmetry occurred from “low to high” or “high to low” in this study. To further understand the effect of pH on the stage of complex coacervation and physical state of the polymeric species after perform­ ing acid or base titration, the schematic diagrams of pH-induced phase transitions were established in Fig. 2C–F. The coacervation phenomena between oppositely charged proteins were described by four critical pH (pHc, pHϕ1, pHopt, and pHϕ2). The pHc (pH corresponding to the initial increase in the slope) is related to the onset of soluble complex, which is smaller and is described as the precursor of the larger coacervate. Bio­ polymers are sufficiently charged and thus do not considerably interact with each other before pHc. The pHϕ1 (pH corresponding to the sharp increase of turbidity) and pHϕ2 (pH corresponding to the endpoint of rapid turbidity decline) meant insoluble coacervate formation and disintegration, respectively. The pHopt (pH corresponding to the maximum turbidity) is the optimum pH value utilized to prepare coac­ ervate. Santos, da Costa, et al. (2018) and Santos, de Carvalho, et al. (2018) found that the pHϕ2 values are virtually unchanged as a function of the variation of the bovine serum albumin/lysozyme ratio because the dissociation of the complexes occurred by the protonation of bovine serum albumin and not by the influence of the ratio. Moreover, although pHϕ1 and pHc dependent on the ratio, they were very similar at all albumin/lysozyme ratios. Similar results were also observed in the complex coacervates of ovalbumin/lysozyme (Santos, da Costa, & Garcia-Rojas, 2018). However, all the four critical pH increased along with LF increase in SPI/LF mixtures, as shown in Fig. 2G, which presents pHc, pHϕ1, pHopt, and pHϕ2 of SPI/LF mixtures. In summary, although pHopt values obtained from “low to high” and “high to low” was influenced by titration direction, they still were close to the result of “pH first”. The protein recovery of these three complexes (6.25-SPI/LF ¼ 1:3, pH 6.6-SPI/LF ¼ 1:4, pH 7.0-SPI/LF ¼ 1:5, obtained 4

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Fig. 2. Turbidity of SPI/LF mixtures as a function of pH at different SPI/LF mass ratio, prepared by (A) “low to high” and (B) “high to low”. Schematic diagram of pHinduced transitions at SPI/LF ratio of 1:3 based on (C) “low to high” and (D) “high to low” and at SPI/LF ratio of 1:4 based on (E) “low to high” and (F) “high to low”. (G) Four critical pH values (pHc, pHϕ1, pHopt, and pHϕ2) as a function of SPI/LF mass ratio. The illustration in the top-right corner is the photo of two SPI/LF complex coacervates.

from “pH first”) were also determined, reaching 70%, 63%, and 51%, respectively. Finally, to ensure the higher yield (>60%) of the hetero­ protein complexes, SPI/LF complexes were produced at SPI/LF ratio of 1:3 (pH 6.25) and 1:4 (pH 6.6) via “pH first” method, respectively. It is worth noting that the particle size of SPI significantly increased when its pH fell to 6.25 from 6.6 due to a reduction in electrostatic repulsion (Fig. S2). Moreover, SPI solubility slightly decreased (6.1%) at pH 6.25 (Fig. S3). These results confirmed that the SPI at pH 6.25 is present in a certain degree of self-aggregation. Although self-aggregation of SPI was observed at pH 6.25, it was used to investigate the potential effects of self-aggregation on heteroprotein complex coacervation. As shown in the upper right of Fig. 2, these two coacervates showed opaque and viscous liquid or gels, similar to the observation of LF/PPI coacervate (Adal et al., 2017). Interestingly, the micrometric spherical shape of the coacervates was still observed using optical microscopy (Fig. S4).

glycinin (acidic subunit, 35 kDa, 29.0%; basic subunit, 17 kDa, 27.9%), which in line with previous findings (Nishinari, Fang, Guo, & Phillips, 2014). It was noted that there was an overlap between the α0 subunit of β-conglycinin and the subunit of LF (75–90 kDa), which hindered the quantitative analysis of SDS-PAGE (Fig. S5). Therefore, two control groups were set up to help analysis. As shown in Fig. S5B, a total of 5 μg SPI and 15 μg LF were loaded into Lane 1; and 5 μg SPI and 20 μg LF were loaded into Lane 2. Meanwhile, 20 μg SPI/LF complex (pH 6.25, SPI/LF ¼ 1:3) and 25 μg SPI/LF complex (pH 6.6, SPI/LF ¼ 1:4) were loaded to lane 3 and lane 4, respectively. In this study, the protein profiles of lane 3 and lane 4 were consistent with that of lane 1 and lane 2 (control groups), respectively. In addition, the intensity of overlapping subunit containing α0 subunit of β-conglycinin and LF subunit in Lane 2 was 7.6% higher than that in Lane 1. Similarly, the intensity of this overlapping subunit in Lane 4 was 7.7% higher than that in Lane 3. These results indicated that the stoichiometry of these heteroprotein complexes was the same as that of the control groups and all protein subunits were involved in HPCC. In addition, since the ratio of the two proteins in the condensed phase is the same as in the mixing ratio, the ratio of SPI/LF in the diluted phase was also the same as the mixing ratio (Fig. S5). It may be an effective analytical method for the heteroprotein complex containing overlapping subunits to comparing with hetero­ protein complex by adding control groups.

3.2. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDSPAGE) The stoichiometry of protein complex and the subunits participating in HPCC can be understood by SDS-PAGE. Fig. S5 shows SDS-PAGE profiles of SPI, LF, and their complexes. SPI exhibited a wide variety of polypeptide subunits of molecular weight ranging from 10 to 102 kDa, mainly consisting of β-conglycinin (α0 -subunits, 80 kDa, 12.9%; α-subunits, 75 kDa, 14.7%; β-subunits, 51 kDa, 15.5%) and 5

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3.3. Isothermal titration calorimetry (ITC)

entropically driven due to the absolute values of TΔS were much larger than that of ΔG. It is generally accepted that driving force of interactions is electrostatics in the early studies of complex coacervation. However, in recent studies, it is found that enthalpic gain resulted from water perturbation and counterion release upon coacervation (Fu & Schlenoff, 2016). Furthermore, the negative ΔH observed in the complexes of SPI and LF can be interpreted as electrostatic interactions or hydrogen bonding interactions (Hadian et al., 2016; Kayitmazer, 2017). By contrast, hydrophobic interactions generally related to endothermic due to the energy requirement of breaking hydrogen bonds (Aberkane, Jasniewski, Gaiani, Scher, & Sanchez, 2010; Henzler et al., 2010).

ITC is an experimental technique to truly understand the in­ teractions, binding affinity, and thermodynamics of complexation and coacervation for a system of oppositely charged macroions. Fig. 3A and B shows the ITC thermogram of the heat flow as a function of time and the binding isotherm of the SPI/LF complexes obtained by the integral of the titration peaks and subtracted from the dilution values (control), respectively. The binding stoichiometry (N), binding constant (K), enthalpy change (ΔH), entropy change (TΔS), and Gibbs-free energy change calculated by fitting with the “One set of sites” model. It should be pointed out that this model is only suited to monotonic binding iso­ therms. As shown in Fig. 3A, the height of the peaks decreases since the number of SPI that did not participate in binding increased. Once all the binding sites were occupied, the peak height tended to be constant. It was noteworthy that the calculated stoichiometric ratio (0.332 and 0.254, based on the mass ratio of SPI/LF) approached to their optimal mixing ratio at pH 6.25 and pH 6.6 (Fig. 3B). Therefore, it was feasible to predict the optimal ratio of heteroprotein protein complexes at different pH by ITC. Complex coacervation is thermodynamically favoured when ΔG is negative, which usually happens when ΔH is highly negative and/or TΔS is highly positive (Kayitmazer, 2017). In this study, the ΔH, TΔS, and ΔG of the complex (SPI/LF ¼ 1:3, pH 6.25) were 0.0043 kcal/g, 5.14 kcal/g, and 5.14 kcal/g, respectively. Similarly, these parameters of the other complex (SPI/LF ¼ 1:4, pH 6.6) were 0.0045 kcal/g, 5.89 kcal/g, and 5.90 kcal/g, respectively. Thus, although the SPI/LF interactions were exothermic processes, they were still considered to be

3.4. Fourier transform infrared spectroscopy (FT-IR) FT-IR of SPI, LF, and SPI/LF complexes are illustrated in Fig. 3C. Amides I, II, and III are the most sensitive regions to the conformational changes in the protein secondary structure in FT-IR. Amide I is between the bands of 1600 and 1720 cm 1 and is formed by in-plane stretching of the C¼O bond, weakly coupled to stretching of the C–N, and in-plane bending of the N–H bond (Talari, Martinez, Movasaghi, Rehman, & Rehman, 2017). Amide II located between the bands of 1475 and 1575 cm 1 and refers to stretching of N–H bending vibration coupled to C–N stretching). Amide III is in the range of bands 1225 and 1425 cm 1, which represents the stretching of the C–N and N–H groups (Santos, da Costa, et al., 2018). In addition, the peak near the 3300 cm 1 corre­ sponds to free O–H stretching and N–H stretching with hydrogen-bonded secondary amino groups (Sukuta & Bruch, 1999). In this study, the wavenumber of amides I in SPI/LF complexes (1657 and

Fig. 3. ITC titration graphs of SPI with LF at (A) pH 6.25 and (B) pH 6.6. Upper: Differential power signal recorded in the experiment as a function of time; lower: integrated data of enthalpy versus the mass ratio of SPI to LF. (○): the integrated data; ( ): fitted curve. (C) FT-IR of SPI, LF, and SPI/LF complexe cacervates at pH 6.25 (SPI/LF ¼ 1:3), and pH 6.6 (SPI/LF ¼ 1:4). (D) Schematic diagram of heteroprotein interactions. When macroions were close enough through electrostatic interaction, a part of the water was expelled. The peptide groups of the protein backbone and the polar groups of the side chains which originally interacted with water by hydrogen bond, turned to form intramolecular and/or intermolecular hydrogen bonds. Moreover, the entropy in this system increased due to water was no longer bound to proteins. Therefore, the released water and counterions could contribute to entropy gain of SPI/LF complex coacervation. 6

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Food Hydrocolloids 100 (2020) 105415

1665 cm 1) was different from that of single SPI and LF. Slight shifts also occurred to the wavenumber of amides II and III in SPI/LF complexes. These changes in the band of Amides I, II, and III corresponding to amides were caused by the electrostatic interaction between the COO– (C¼O) cluster of one protein and the NH3þ (NH) cluster of the other protein. Furthermore, redshifts (decreased wavenumber) occurred to the bond of 3300 cm 1 in SPI/LF complexes (3293 and 3286 cm 1) compared with that of SPI (3298 and 3296 cm 1) and LF (3306 and 3322 cm 1). This result indicated that hydrogen bonding also was involved in the heteroprotein complex coacervation between SPI and LF. As shown in Fig. 3D, this result can be interpreted as when macroions were close enough through electrostatic interaction, a part of the water was expelled (Finkelstein & Ptitsyn, 2016b). The peptide groups of the protein backbone and the polar groups of the side chains which origi­ nally interacted with water by hydrogen bond, turned to form intra­ molecular and/or intermolecular hydrogen bonds. Moreover, the entropy in this system increased due to water was no longer bound to proteins (Finkelstein & Ptitsyn, 2016a). Therefore, the released water and counterions could contribute to entropy gain of SPI/LF complex coacervation. Furthermore, the driving force underlying LLPS is the exchange of macromolecule/water interactions for macro­ molecule/macromolecule and water/water interactions under condi­ tions for which this process is energetically favorable (Alberti et al., 2019).

lobes of dumbbell-shaped LF. The difference in iron saturation between N and C lobes and more compact structure of the C lobe in the iron-saturated protein may result in their different thermal sensitivities (Bokkhim, Tran, Bansal, Grondahl, & Bhandari, 2014; F. G.; Liu, Sun, Wang, Yuan, & Gao, 2015). Notably, SPI/LF complexes still showed two peaks. The appearance of SPI significantly (P < 0.05) increased the denaturation temperature of the first peak of LF, and the more SPI in complexes, the higher the denaturation temperature. In addition, ΔH of two peaks corresponding to SPI/LF complexes is smaller than that of LF. These demonstrated that the interaction between SPI and LF could improve the thermostability of heat-sensitive lobe in LF, and it is not uncommon to improve the thermal stability of proteins via HPCC (Santos, da Costa, et al., 2018). 3.6. Scanning electron microscope (SEM) In previously published studies, two forms of SPI were observed by SEM. The first one was a spherical shape with a wrinkled surface; the other one was a sheet-like structure with a smooth surface. After sum­ ming up these papers, it was found that the spherical structures appeared in commercial SPI obtained by spray drying, and the smooth flaky structures were observed in the freeze-dried SPI (Liu et al., 2016; Qi, Venkateshan, Mo, Zhang, & Sun, 2011; Zhao et al., 2015). Even other plant proteins had similar patterns (Zhao, Liu, Zhang, Zhu, & Ao, 2019). As shown in Fig. 5A and B, both SPI and LF exhibited a flaky structure, and the sheet-like structure of SPI was thinner than that of LF. Inter­ estingly, a large number of gel-like crosslinked particles were observed in the SPI/LF complexes through a series of magnification (Fig. 5C and D), demonstrating strong interactions between SPI and LF. Even in the freeze-dried complexes, these gel-like crosslinked structures can be clearly observed. Similar cross-linking structures were also present in the supramolecular assembly of apo α-lactalbumin and lysozyme, which resulted from the coalescence between microspheres assembled by these proteins (Nigen, Croguennec, Madec, & Bouhallab, 2007). Furthermore, the SPI/LF complex (pH 6.25, SPI/LF ¼ 1:3) exhibited distinct granules, whereas the uniform crosslinking structure was observed in the other SPI/LF complex (pH 6.6, SPI/LF ¼ 1:4). This may be closely related to the self-aggregated SPI in the SPI/LF complex (pH 6.25, SPI/LF ¼ 1:3).

3.5. Differential scanning calorimetry (DSC) Fig. 4 shows the DSC thermograms of SPI, LF, and their complexes. DSC characteristics, such as ΔH, To, Tp, and Te, are summarized in Table 1. As shown in Fig. 4A, the heat flow curve of SPI displayed classical two peaks, and the first peak was related to the denaturation of β-conglycinin at 76.7 � C and the second peak corresponding to the denaturation of glycinin at 95.0 � C (Tang, Choi, & Ma, 2007). LF also showed two denaturation peaks, the first peak was at a lower temper­ ature of 62.5 � C followed by a second peak at a higher temperature of 89.3 � C (Fig. 4B). Moreover, the absolute value of ΔH corresponding to the first peak ( 9.78 J/g) is significantly (P < 0.05) greater than that of the second peak ( 2.50 J/g). These peaks were attributed to the two

Fig. 4. DSC graphs of (A) SPI, (B) LF, (C) SPI/LF complex coacervates at pH 6.25 (1:3) and (D) pH 6.6 (1:4). 7

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Table 1 Thermal properties of SPI, LF, and SPI/LF complexes. Samples

Peak

SPI LF SPI/LF 1:3–6.25 SPI/LF 1:4–6.6 SPI LF SPI/LF 1:3–6.25 SPI/LF 1:4–6.6

Peak1

Peak2

ΔH (J/g) 1.58 � 0.13 9.78 � 0.15 5.08 � 0.30 6.88 � 0.05 7.94 � 0.79 2.50 � 0.23 1.69 � 0.04 2.46 � 0.08

To (� C)

Tp (� C)

Te (� C)

72.31 � 0.23 52.30 � 0.12 57.98 � 0.64 55.58 � 0.10 89.05 � 0.27 84.67 � 0.28 83.09 � 0.23 82.29 � 0.15

76.67 � 0.17 62.45 � 0.29 68.51 � 0.38 65.37 � 0.19 95.01 � 0.48 89.30 � 0.30 87.88 � 0.33 87.79 � 0.19

80.65 � 0.16 68.35 � 0.43 75.52 � 0.57 74.02 � 0.11 100.1 � 0.71 92.73 � 0.28 92.74 � 0.38 92.46 � 0.10

Fig. 5. SEM micrographs of (A) SPI observed at �500 and �2000; (B) LF observed at �200 and �2000; (C) SPI/LF complex (pH 6.25, SPI/LF ¼ 1:3) observed at �200, �2000, and �10000; and (D) SPI/LF complex (pH 6.6, SPI/LF ¼ 1:4) observed at �200, �2000, and �10000.

In general, this large crosslinked gel-like and agglomerated structure result from the conglutination of many spherical structures, and these spheres are originally formed via LLPS, which does not have to result in liquid, freely fusing assemblies; instead, coacervates can undergo further gel-transitions. In addition, nucleation and growth (NG) and spinodal decomposition (SD) are two classical mechanisms of LLPS (Pathak, Priyadarshini, Rawat, & Bohidar, 2017). The solid-like network and/or complex 3D connected network resulted from protein-rich continuous phase reaches glass transition conditions may imply that LLPS was related to SD (Boire et al., 2019). Moreover, the particle size of the SPI/LF complex coacervates in this study is larger than that of LF/caseins coacervates previous reported by Anema et al. (2016), who suggested that LF/as-casein coacervates coalesced through a nucleation and growth process. Given the restricted size and growth of coacervates occurred via a nucleation barrier, SPI/LF complex coacervation was more likely to come from SD (Falahati & Haji-Akbari, 2019).

and G), whereas LF was smaller spherical particles, with a length of 5–30 nm (average length is approximately 10.7 nm) and a width of 3–20 nm (average length is approximately 6.6 nm). Adal et al. (2017) observed uniform spherical particles of LF, with a mean radius of 6.2 nm. The slightly higher average size resulted from the stacked pro­ teins on mica sheets (Fig. 6A and B). Moreover, that these two different complexes strictly abided by charge compensation implied that the biopolymers involved in coacervation could rearrange itself via chain conformational changes to alter the charge distribution (i.e., change in the charge anisotropy) and thus respond to the environmental condi­ tions (Moschakis & Biliaderis, 2017). A significant difference in apparent morphology of two SPI/LF complexes was observed via AFM, although they both strictly followed the electrostatic compensation mechanism and had similar thermodynamic formation mechanisms. The SPI/LF complex (pH 6.25, SPI/LF ¼ 1:3) showed larger agglomerates and even some aggregates larger than 300 nm in diameter were observed. By contrast, the other SPI/LF complex (pH 6.6, SPI/LF ¼ 1:4) exhibited cluster and chain structure formed by flocculation of small coacervate units. Similar chain structure was observed in the PPI/LF complex (Adal et al., 2017). Fig. 6I and J displays the length, width and area distribution of these particles collected from AFM micrographs. The particles length in both complexes focused on 50–150 nm; and their width (pH 6.25, SPI/LF ¼ 1:3 and pH 6.6, SPI/LF ¼ 1:4) focused on

3.7. Atomic force microscope (AFM) Fig. 6 shows the AFM micrographs of SPI, LF, and SPI/LF complexes and corresponding particle size. SPI exhibited spheroidicity with a length of 25–75 nm (average length was approximately 45.9 nm) and a width of 20–60 nm (average width was approximately 29.0 nm) (Fig. 6A 8

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Fig. 6. AFM micrographs of (A) SPI observed at 3 μm � 3 μm; (B) LF observed at 1 μm �1 μm; (C) SPI/LF complex (pH 6.25, SPI/LF ¼ 1:3) observed at 3 μm � 3 μm; (D) SPI/LF complex (pH 6.6, SPI/LF ¼ 1:4) observed at 3 μm � 3 μm. Cartoon schematic diagrams of SPI/LF complexes prepared at (E) pH 6.25-SPI/LF ¼ 1:3 and (F) pH 6.6-SPI/LF ¼ 1:4. Bubble diagrams corresponding to respective AFM micrographs and describing the size distribution and shape of (G) SPI, (H) LF, and SPI/LF complexes prepared at (I) pH 6.25 SPI/LF ¼ 1:3 and (J) pH 6.6 SPI/LF ¼ 1:4. The size of each bubble is proportional to the size of the protein or complex.

50–150 nm and 20–80 nm, respectively. Moreover, the particles in the complex (pH 6.25, SPI/LF ¼ 1:3) were more closed to the dotted line represented for the spheres. These results demonstrated the hetero­ rprotein complex (pH 6.25, SPI/LF ¼ 1:3) was approximate to the spheres, whereas the other complex (pH 6.25, SPI/LF ¼ 1:3) was approximate to the chains. This difference should be related to the participation of SPI self-aggregates in SPI/LF complex coacervation. More positively charged biopolymers and residual positively charged soluble complexes were needed to fuse with SPI self-aggregates, which had large size and abundant surface charge. Hence, the build-up of charge may lead the coacervates contained SPI self-aggregates to form larger ellipsoids. Two cartoon schematics were provided to distinguish these complexes (Fig. 6E and F). It should be pointed out that the size of SPI/LF complexes observed from AFM was significantly lower than that of DLS. This discrepancy may be explained by the high dilution of AFM samples because of complex coacervation was biopolymer concentration-dependent (Moschakis et al., 2017). In addition, the flocculated chain structure of SPI/LF complex (pH 6.6, SPI/LF ¼ 1:4) also confirmed that SPI/LF complex coacervation might be generated from SD, due to NG usually appeared as hydrodynamically driven coa­ lescence (Pathak et al., 2017).

by significant entropy gain. When SPI and LF were close enough by electrostatic interaction, a portion of the water and counterions was released. The peptide groups in protein backbones and the polar groups in side chains originally interacted with water via hydrogen bonding turned to build intramolecular and/or intermolecular hydrogen bonds, and thus increasing the entropy. Therefore, the energy advantaged HPCC was not only as a result of the protein/protein interaction, but also the interaction of protein/water and water/water. Interestingly, the SPI/LF interaction improved the thermostability of heat-sensitive lobe in LF. A large number of gel-like crosslinked particles and agglomerated structures originated from the conglutination of many spherical struc­ tures originally formed via LLPS (i.e., gel-transition) were observed by SEM. This gel-like network and/or complex 3D connected network resulted from protein-rich continuous phase reaches gel-transition may imply that LLPS was related to SD (Boire et al., 2019). Furthermore, the sphere complex (pH 6.25-SPI/LF ¼ 1:3) with a diameter of 50–150 nm and the chain-like complex (pH 6.6-SPI/LF ¼ 1:4) with a length of 20–50 nm and a width of 10–40 nm were observed by AFM. Their different apparent structure may result from the SPI self-aggregation. The biopolymers involved in coacervation could rearrange itself via chain conformational changes to alter the charge distribution (i.e., change in the charge anisotropy) and thus respond to the environmental conditions. Unfortunately, this study is limited by the complexity of SPI. However, we found that all the major individual proteins in SPI participated in HPCC. Therefore, the high-resolution tools, such as solid nuclear magnetic resonance and modeling, can be performed to explore the HPCC between these individual SPI fractions and LF in further research.

4. Conclusion SPI/LF complex coacervate can be generated via electrostatic inter­ action, and stoichiometry of two optimal complex coacervates (pH 6.25SPI/LF ¼ 1:3 and pH 6.6-SPI/LF ¼ 1:4) was identical to their mixing ratio. SPI/LF complex coacervation was exothermic and accompanied 9

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Declaration of competing interest

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