Coating oil droplets with rice proteins to control the release rate of encapsulated beta-carotene during in vitro digestion

Abstract

Food protein-based delivery systems have unique advantages. Herein, a facile one-step fabrication of oil droplets with a tunable release rate of encapsulated lipophilic compounds during in vitro digestion is reported. Modified rice proteins (MRPs) were deposited on oil droplets with a thickness controllable by the final pH during titration of an alkaline solution of MRPs blended with oil, resulting from significant changes of secondary structures and solubility of MRPs at pH 6.0–7.0. Oil droplets were smaller than 400 nm, and the tunable conformation of MRPs and therefore resistance against peptic and pancreatic digestions resulted in the release rate of encapsulated β-carotene controllable by shell structures formed during titration. The irregular aggregates at pH 6.2 resulted in the step release of encapsulated β-carotene, releasing ca. 25% and 100%, respectively, at the simulated gastric and intestinal conditions. Whereas, the release of β-carotene was limited at the simulated gastric conditions but followed the zero-order kinetics at the simulated intestinal conditions, with a higher release rate for oil droplets with thinner shells produced at a higher pH between 6.4 and 7.0. Therefore, novel emulsion-based delivery systems can be fabricated with MRPs to control release profiles of lipophilic compounds during digestion.

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Introduction

Interest in delivery systems has been growing over the past years because numerous bioactives such as vitamins, drugs, polyphenols, and carotenoids have low bioavailability and are rapidly cleared or inactivated by blood components and detoxifying systems.^1,2 Particularly, lipophilic bioactives have been studied extensively because of their limited solubility in water, extensive first-pass metabolism, and physical, chemical, and biological instabilities,³ and emulsions commonly studied because the oil phase can be utilized to dissolve lipophilic bioactives. Properly-designed delivery systems can effectively protect the payloads against physical, chemical, and biochemical degradation mechanisms and achieve targeted delivery in cells through endocytotic pathways.⁴ For oral delivery systems, the release rate of payloads can be tuned by engineering particle structures and adopting materials based on physical, chemical, and biochemical conditions in the gastrointestinal tract. From the safety, sustainability, and consumer acceptance perspectives, oral delivery systems based on generally recognized as safe (GRAS) ingredients are advantageous when compared to many systems based on synthetic polymers.

Various polysaccharides and proteins have been studied to deliver drugs,⁵ genes,^6,7 cosmetics,³ and nutrients.⁸ However, food-sourced proteins are relatively under-studied, despite their sustainability, biocompatibility, and broad ranges of physicochemical properties. Food proteins can be modified for their functional properties using conditions practiced in food production. For example, rice proteins are abundant and inexpensive⁹ and have numerous health-related functionalities,¹⁰ including the potential hypoallergenicity which is advantageous to soy and dairy proteins for certain groups of individuals.⁹ The high glutelin content (ca. 80%) and therefore insolubility in water and other conventional solvents¹¹ however have led to very few studies about biomaterials fabricated with rice proteins.

Recently, a technology was invented in our laboratories to modify rice protein solubility. Subsequent steps of suspension in an alkaline solution at pH 12.5, incubation at −20 °C for 24 h, and freeze-milling resulted in the modified rice proteins (MRPs) being soluble at pH 7.0.¹² MRPs remained marginally soluble at acidic pH, and a sharp change of solubility between pH 6.0 and 7.0 was observed in our preliminary experiments. This unique pH-dependent solubility is desirable for many applications, for example enteric delivery systems that are currently based on modified celluloses and synthetic polymers.¹³ Because alkaline treatments are used to produce food products such as tortilla flour (masa) and hominy corn,¹⁴ MRPs can be used as GRAS polymers to engineer delivery systems with unique properties.

The overall goal of the present study was to fabricate emulsions with MRPs coating oil droplets dissolving lipophilic bioactives. Because MRPs have a limited emulsifying activity,¹⁵ a recent method based on anti-solvent precipitation of zein (alcohol-soluble corn protein) on oil droplets¹⁶ was adapted to produce emulsions based on the pH-dependent solubility characteristics of MRPs, by titrating a mixture of soybean oil and an alkaline solution of MRPs to pH 6.2–7.0. Our approach eliminates the need of both other emulsifying agents and organic solvents, which are desirable in practical applications. Specifically, the first objective was to characterize solubility and structures of MRPs as a function of pH. The second objective was to fabricate and characterize oil droplets with a MRP shell and a soybean oil core. The third objective was to characterize release profiles of β-carotene, studied as a model lipophilic bioactive, loaded in emulsions for potential oral delivery applications.

Experimental section

Materials

Rice protein isolate powder containing 90.14 wt% protein, as determined by the Kjeldahl method with a conversion factor of 5.95,¹⁷ was purchased from Jingnong Biotechnology Ltd. (Yichun, Jiangxi, China). β-Carotene, pepsin from porcine stomach mucosa, and pancreatin from porcine pancreas were purchased from Sigma-Aldrich Corp. (St. Louis, MO, USA). All other chemicals were of an analytical grade and were used without further purification.

Preparation of MRPs

MRPs were prepared according to our previous study.¹² In brief, rice protein powder was suspended in distilled water (1 : 30, w/v) containing 0.03 M NaOH, incubated overnight at −20 °C, and directly milled using an impact mill (model XFB-500, Zhongcheng Mechanical Co., Changsha, China). The milled suspension after warming to room temperature (25 °C) was adjusted to pH 7.0 and centrifuged at 7000g for 10 min. The supernatant was transferred and desalted by dialysis against distilled water using a membrane with a molecular-weight-cut-off of 1000 Da, followed by freeze-drying to prepare lyophilized powder as the MRPs.

Solubility of MRPs

MRPs were hydrated at 1% w/v in distilled water by mixing on a magnetic stir plate at room temperature (25 °C) for 30 min, followed by adjusting to pH 2.0–10.0 using 0.1 M HCl. After stirring for another 30 min, and centrifugation at 4000g for 30 min,¹⁸ the supernatant was collected and determined for the nitrogen content using the Kjeldahl method that was converted to the protein content using a factor of 5.95.¹⁷ The solubility of MRPs was then expressed as the percentage of protein mass in the supernatant with respect to the total protein mass in the corresponding sample after centrifugation.

Secondary structures of MRPs studied with far-ultraviolet circular dichroism spectroscopy (far-UV CD)

Far-UV CD experiments were conducted using a model MOS-450 spectrometer (BioLogic Science Instruments, Ltd., Claix, France) with a 0.1 cm quartz cell at a wavelength range of 190–250 nm. MRPs were dissolved at 50 μg mL⁻¹ in 10 mM phosphate-buffered saline (PBS) adjusted to pH 6.0–7.0 with 0.2 unit increments using 0.1 M HCl. After centrifugation at 10 000g for 10 min, the supernatants were used to acquire far-UV CD spectra from averages of 3 scans collected at a step size of 0.1 nm and an average time of 2 s. Percentages of different categories of secondary structures of MRPs were determined using the CDSSTR software available from Colorado State University (Fort Collins, CO, USA).

Surface hydrophobicity (H₀) of MRPs

H₀ of MRPs at pH 6.0–7.0 was determined using 1-anilino-8-naphthalene-sulfonate (ANS) as a fluorescence probe.¹⁹ MRP samples prepared as above were diluted to 0.0015–0.015% w/v proteins in 10 mM PBS adjusted to the same pH. Four milliliters of a protein solution was thoroughly mixed with 10 μL of a freshly prepared ANS solution (8.0 mM in distilled water). The fluorescence intensity of each sample was determined at excitation and emission wavelengths of 390 and 484 nm, respectively, using an F-7000 spectrofluorometer (Hitachi Co., Tokyo, Japan). The initial slope of a fluorescence intensity versus protein concentration (% w/v) curve was used as an index of H₀. The fluorescence spectra of the samples with 0.015% w/v protein were also collected at an excitation wavelength of 390 nm and an emission wavelength range of 400–600 nm at a bandwidth of 5 nm.

Preparation of emulsions

Stock aqueous solutions were prepared with 1% w/v MRPs in distilled water and adjusted to pH 9.0 with 0.1 M NaOH. Soybean oil was mixed with the protein stock solution at a volume ratio of 1 : 10 using a mixer (model T18BS25, IKA, Wilmington, NC, USA) at gear 5 for 60 s to obtain a pre-emulsion that was constantly stirred on a magnetic stirring plate to suspend oil drops. To coat oil droplets with MRPs, 0.01 M HCl was added drop-wise using a peristaltic pump (model BT-100, Jiapeng Tec. Co., Shanghai, China) at a constant flow rate of ca. 10 μL s⁻¹. The titration experiments were performed at room temperature (25 °C) in 20–30 min to a final pH of 6.2–7.0 to produce coarse emulsions. Thereafter, the coarse emulsions were centrifuged at 4000g for 10 min (model Avanti J26-XP, Beckman Coulter Co., Brea, CA, USA) to remove free oil and large droplets (in the cream) and aggregated MRPs (in the precipitate) to obtain the serum phase as the final emulsions for characterization of properties. The final emulsions were ultra-centrifuged at 20 000g for 30 min using the same centrifuge to harvest oil droplets that were freeze-dried to prepare oil droplet powders. The serum after ultra-centrifugation had little proteins based on the Bradford assay.

Confocal laser scanning microscopy (CLSM)

MRPs and oil were stained with 100 μM Nile blue and 100 μM Nile red, respectively, before preparing emulsions as above. Nile blue was used because it binds with hydrophobic patches of proteins.²⁰ After centrifugation, the fluorescent images of fresh emulsions were acquired on an inverted LSM-710 microscope (Zeiss Co., Oberkochen, Germany) at an excitation wavelength of 630 nm and 480 nm for Nile blue and Nile red, respectively.

Fourier transform infrared spectroscopy (FTIR)

FTIR experiments were carried out using a Nicolet iS10 FTIR spectrometer (ThermoFisher Scientific Co., Marietta, OH, USA). About 2 mg of freeze-dried oil droplet powders were mixed with KBr, followed by grinding and pressing into a pellet. The absorbance intensities were measured at a 2 cm⁻¹ resolution in the wavenumber range of 4000–400 cm⁻¹. Amide I bands (at a wavenumber range of 1700–1600 cm⁻¹) of the FTIR spectra were additionally baseline-corrected using PeakFit 4.12 (SeaSolve Software Inc., San Jose, CA, USA).

Particle size and zeta-potential

The particle size determined by dynamic light scattering (DLS) and zeta-potential of MRPs and fresh emulsions were measured using a Zetasizer Nano instrument (Malvern Instruments Ltd., Malvern, UK) after appropriate dilutions. All measurements were carried out at 25 °C and averages of three readings were reported.

Encapsulation of β-carotene

To prepare oil droplets loaded with β-carotene, β-carotene was dissolved in soybean oil at a concentration of 0.6 mg mL⁻¹, followed by preparation of emulsions as described previously. The fresh emulsions were used to determine encapsulation efficiency and release properties of β-carotene. An emulsion sample was mixed with an identical volume of petroleum ether that is a poor solvent of MRPs. After ultra-sonication (model KQ3200, Ultra-sonic Instrument Co., Kunshan, Jiangsu, China) for 1 h, the mixture was centrifuged at 10 000g for 10 min, and the supernatant was collected and measured for absorbance at 450 nm. The absorbance was calibrated using the corresponding emulsion without β-carotene extracted with petroleum ether at identical conditions. The amount of β-carotene was then determined using a standard curve established with standard solutions with different amounts of β-carotene dissolved in petroleum ether. Encapsulation efficiency (EE) and loading capacity (LC) were determined according to eqn (1) and (2), respectively. Emulsions were also freeze-dried to determine protein content using the Kjeldahl method to estimate the recovery of MRPs with respect to the amount used in emulsion preparation.where the β-carotene input is the amount used in emulsion preparation.where the mass of emulsion solids was determined after freeze-drying.

Release kinetics of β-carotene during simulated digestions

To simulate gastric digestion, 50 mL of a fresh emulsion was added with 30 mL of a saline solution containing 150 mM NaCl, 5 mM KCl, and 5 mM CaCl₂ and additional 10 mL of a 100 mM CaCl₂ solution.²¹ The mixture pH was adjusted to 1.3 using 1.0 M HCl followed by incubation at 37 °C. The simulated gastric digestion was started on a magnetic stirring hot plate by adding 10 mL of a pepsin solution (30 mg L⁻¹, dissolved in 0.1 M HCl) pre-warmed to 37 °C and continued at 37 °C for 2 h. To simulate the intestinal digestion, the treatment after 2 h peptic digestion was adjusted to pH 7.0 and dissolved with 0.06 g pancreatin. The mixture pH during pancreatic digestion at 37 °C for up to 2 h was statically maintained at 7.0 with 0.01 M NaOH (pH-stat method). Samples collected at given time points in each digestion step were heated at 100 °C for 5 min to deactivate the enzyme. The released β-carotene mass was determined by the method of Parris and coauthors with slight modifications.²² The mixture after digestion and enzyme inactivation was added with 50 mL petroleum ether and stirred for 30 min at room temperature on a magnetic stir plate. The extraction mixture was centrifuged at 10 000g for 10 min, and the supernatant was measured for absorbance at 450 nm. The absorbance at 450 nm was calibrated using control emulsions without β-carotene that were prepared and digested at conditions at identical to those with β-carotene, before determining the amount of released β-carotene, as presented previously. The percentage of released β-carotene was then calculated with respect to the total amount before peptic digestion.

Hydrolysis of MRPs during simulated digestions

To characterize proteolytic properties of MRPs, 50 mL of the above neutral protein stock solution with 1% w/v MRPs was constituted to and digested at conditions detailed in the previous section. After deactivation of enzyme, 100 μL aliquots of the mixture were added with 900 μL of pH 7.0 PBS containing 50 mM NaCl and 50 mM guanidine hydrochloride. After centrifugation at 10 000g for 10 min, the supernatant was transferred and filtered through 0.45 μm polyvinylidene fluoride filters twice (Millipore, Darmstadt, Germany). The permeate was analyzed using size exclusion chromatography (SEC) at 42 °C with a TSK-G2000 SW_XL column (TOSOH Co., Tokyo, Japan) equipped on an Agilent HPLC 1260 system (Palo Alto, CA, USA). The mobile phase at a flow rate of 0.5 mL min⁻¹ was the pH 7.0 PBS containing 50 mM NaCl and 50 mM guanidine hydrochloride. The elution was monitored for absorbance at 280 nm. The total area of peaks with an elution time greater than 20 min (P_t) and that of undigested MRPs at pH 7.0 (P₀) analyzed at the same SEC conditions were used to determine the digestibility of MRPs using eqn (3).

Statistical analysis

The experiments were performed in triplicate, and values were expressed as mean ± standard deviation (SD). Analysis of variance was evaluated using the Duncan's multiple range test.

Results and discussion

Properties of MRPs as a function of pH

Solubility and surface properties of MRPs. Fig. 1 shows the solubility of MRPs at pH 2.0–10.0. The solubility was below 10% at a pH 6.0 and below but was greater than 90% at pH 7.0 and above. A slight increase of pH between 6.0 and 7.0 significantly increased the solubility of MRPs, which may be significant to fabricate materials responsive at this pH range that is relevant to physiological conditions of many biological systems.

Fig. 1 The pH-dependent solubility measured with samples containing 1% w/v MRPs. Error bars represent SD (n = 3).

The zeta-potential magnitude of MRPs was generally smaller, while surface hydrophobicity (H₀) was higher at a lower pH between 6.0 and 7.0 (Table 1). Because a zeta-potential magnitude of 30 mV or greater is typically found for stable colloidal systems,²³ the substantial loss of protein solvation below pH 7.0 likely is driven by hydrophobic attraction due to the increased exposure of hydrophobic amino acid residues. When probed by ANS, fluorescence spectra in the range of 400–600 nm (Fig. 2a) demonstrated a red-shift of the maximum emission wavelength (λ^F_max) and the attenuation of emission intensities at a decreasing pH between 6.0 and 7.0. The results indicate that MRPs undergo molecular rearrangements (fluorescence quenching) which increase the local polarity around ANS (red-shift of λ^F_max) that weakens the binding with MRPs becoming more hydrophobic at a lowered pH.²⁴

Table 1 Surface hydrophobicity (H₀), zeta-potential, and percentages of secondary structures of MRPs at pH 6.0–7.0

pH	H0 (×10^5)^a	Zeta-potential^a (mV)	Secondary structures (%)
			α-Helix	β-Sheet	β-Turn	Random coil
a Values are mean ± SD (n = 3). Different superscript letters in the same column indicate significant differences (P < 0.05).
6.0	160.56 ± 1.73^a	−34.53 ± 1.94^bc	16.42	16.58	14.15	52.84
6.2	159.12 ± 3.46^a	−33.57 ± 0.55^c	21.06	18.08	13.69	47.22
6.4	143.50 ± 3.54^b	−34.63 ± 1.17^bc	21.67	16.12	18.39	37.26
6.6	142.00 ± 6.92^ab	−37.57 ± 1.71^ab	22.16	12.83	31.94	31.87
6.8	138.23 ± 3.61^ab	−38.07 ± 1.81^a	23.54	9.91	32.15	34.39
7.0	132.78 ± 1.41^c	−39.43 ± 2.40^a	29.03	7.60	31.12	32.25

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Fig. 2 (a) Fluorescence spectra of MRP–ANS mixtures and (b) far-UV CD spectra of MRPs at pH 6.0–7.0.

Secondary structures. The far-UV CD spectra between 190 and 250 nm are shown in Fig. 2b for MRPs at pH 6.0–7.0. Wide negative bands represent a combination of various types of secondary structures in the samples. The ellipticity extremum, which was negatively centered at 215 nm for MRPs at pH 7.0, blue-shifted and the magnitude decreased as pH was lowered, suggesting secondary rearrangements to proteins with fewer surface charges.²⁵ An extra negative extremum appeared at 195 nm for MRPs at pH 6.0, which means that the remaining soluble proteins after precipitation had the unordered conformations.²⁵ The determined percentages of different types of secondary structures as a function of pH are listed in Table 1, showing the increases of β-sheet and random coil structures at a lowered pH at the expense of α-helix and β-turn structures. α-Helix structures play a key role in stabilization of proteins by maximizing polar–polar interactions and minimizing aploar–polar contacts,²⁶ whereas β-sheet structures are intermediates of protein aggregation due to hydrophobic attraction.²⁷ A reduction in turn structures indicates the increased rigidity and compactness of protein structures.²⁸ The rapid increase of unordered structures in pH 6.4–6.0 samples may result from the removal of ordered proteins due to precipitation. The helix-sheet transitions bury polar groups inside proteins and increase the exposure of hydrophobic groups, which agrees with the decreased magnitude of zeta-potential and increase of H₀ (Table 1). The significant secondary structural changes of MRPs within one pH unit provide the physicochemical basis for fabrication of novel pH-responsive materials.

Properties of fresh emulsions

Emulsion structures as studied with CLSM. Emulsions formed after titration to pH 6.2–7.0 were studied using CLSM, shown in Fig. 3 for images merged with fluorescence collected from two channels imaging both proteins and soybean oil, as well as insets magnifying protein structures only. At pH 7.0 when MRPs were mostly soluble (Fig. 1), no discernable protein shells and round cavities (inset with a white ring, occupied by oil droplets) and little features of protein structures were observed at the CLSM resolution. When pH decreased to 6.8, MRPs readily precipitated on the surface of oil droplets, showing visible shell structures. As pH was further reduced to 6.6, a greater amount of protein precipitation resulted in thicker shells around oil droplets. Droplets became smaller when the system pH reached 6.4. For the sample prepared at pH 6.2, irregular structures were observed.

Fig. 3 Merged or single-channel (insets) CLSM images of soybean oil–MRP–water mixtures titrated to a final pH of (a) 7.0, (b) 6.8, (c) 6.6, (d) 6.4, and (e) 6.2 to form emulsions. Soybean oil and MRPs were stained with red and green fluorescence, respectively.

The emulsion structures observed in Fig. 3 can be affected by both the properties of MRPs and emulsion stability. MRPs are less soluble at a lower pH between 6.0 and 7.0 (Fig. 1) and therefore have a higher affinity to the oil phase, resulting in the formation of a thicker shell. When decreased to pH 6.2, the low solubility of MRPs (Fig. 1) causes rapid aggregation of MPRs and therefore irregular structures of particles (Fig. 3e). Because emulsions were centrifuged at 4000g for 10 min before taking the serum phase for CLSM, micrometer-sized droplets were likely removed (Fig. S1†). Big droplets observed in Fig. 3 likely resulted from coalescence after centrifugation and before CLSM imaging. For samples prepared at a higher pH, the shell is thinner, and the chance of coalescence is higher, corresponding to the presence of some big droplets.

Dimension of oil droplets studied with DLS. Size distributions of MRPs at pH 7.0 and fresh emulsions measured by DLS are presented in Fig. 4. All emulsion samples showed two intensity peaks, with those of the pH 6.2 treatment being larger than others. The peaks with bigger structures may be assigned to oil droplets with protein shells because the preparation process did not use high shear deformation, while the smaller ones may be assigned to protein-only particles. The mean hydrodynamic diameters of droplets of all treatments were in the range of ca. 300−400 nm (Table 2). The relatively high polydispersity index of treatments agreed with two peaks in intensity distributions (Fig. 4). In the study using a protocol similar to our study by lowering the alcohol content so that zein became precipitated on oil droplets, a mean diameter of 30–40 μm was reported.²¹ The difference resulted from the centrifugation step used in the present study to remove big droplets and precipitated protein particles, as well as the hydrophobicity difference (affinity the oil phase) between zein and MRPs and solvent polarity (with and without ethanol) in two studies.

Fig. 4 DLS size distributions of MRPs at pH 7.0 and fresh emulsions prepared at pH 6.2–7.0.

Table 2 Droplet dimension, polydispersibility index, and zeta-potential of emulsions^a

Emulsion pH	Hydrodynamic diameter (nm)	Polydispersity index	Zeta-potential (mV)
a Values are mean ± SD (n = 3). Different superscript letters in the same column indicate significant differences (P < 0.05).
6.2	405 ± 65^a	0.42 ± 0.04^a	−39.0 ± 1.2^b
6.4	303 ± 22^b	0.37 ± 0.01^b	−39.1 ± 1.5^b
6.6	376 ± 45^ab	0.40 ± 0.02^ab	−47.9 ± 0.8^a
6.8	370 ± 36^ab	0.41 ± 0.00^ab	−46.8 ± 1.8^a
7.0	368 ± 23^ab	0.38 ± 0.02^ab	−47.4 ± 1.8^a

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Zeta-potential of fresh emulsions. The zeta-potential of fresh emulsions was about −39 mV for treatments titrated to pH 6.2 and 6.4, which was significantly less negative than the treatments (about −47 mV) titrated to pH 6.6, 6.8, and 7.0 (Table 2). This overall trend agreed with the zeta-potential of MRPs (Table 1). It was also noted that the magnitude of negative zeta-potentials of the oil droplets increased compared to MRPs at the same pH. The difference likely resulted from the preferential orientation of hydrophobic amino acid residues of MRPs toward the oil phase and the presence of hydrophilic and charged amino acid residues toward the continuous aqueous phase.

For particles produced with water-insoluble proteins such as gliadin and zein, the hydrophobic attraction can be stronger than the electrostatic repulsion to cause particle aggregation.^29,30 Therefore amphiphilic polyelectrolytes such as sodium caseinate³¹ and gum arabic³¹ are used to adsorb on these particles to provide electrostatic and/or steric repulsions to prevent particle aggregation. Negative zeta-potentials of oil droplets in the present study (Table 2) had a higher magnitude than those prepared with zein (<20 mV),^32,33 and electrostatic repulsion appears to be sufficient to stabilize oil droplets in the present work, as presented for no significant changes in droplet size during 9-day storage at room temperature (Fig. S4†).

Interactions among emulsion components studied using FTIR

FTIR was used to study interactions among components, compared in Fig. 5a for soybean oil, MRPs, and the freeze-dried emulsion produced at pH 6.6. The spectrum of soybean oil had peaks at 1750 cm⁻¹ (stretching vibrations of carbonyl group), 1167 cm⁻¹ (deformation vibrations of alkyl chains), and 722 cm⁻¹ (rocking vibrations of alkyl chains),³⁴ whereas that of MRPs had peaks centered at 1636 cm⁻¹ (amide I; carbonyl stretching) and 1552 cm⁻¹ (amide II; secondary N–H bending).³⁵ No extra peaks were observed in the emulsion prepared at pH 6.6, indicating the absence of new covalent bonds formed between MRPs and oil during titration to prepare emulsions and the subsequent lyophilization. Compared to the spectrum of MRPs alone, the amide II band of the freeze-dried emulsion was weaker, which can result from structural alterations of MRPs after titration to form shells on oil droplets.

Fig. 5 Comparison of FTIR spectra at (a) 4000–400 cm⁻¹ for soybean oil, MRPs, and a freeze-dried emulsion prepared at pH 6.6 and (b) 1700–1600 cm⁻¹ for MRPs and freeze-dried emulsions prepared at pH 6.2–7.0.

The FTIR spectra at 1700–1600 cm⁻¹ were further examined for changes of amide I bands (Fig. 5b). MRPs had strong absorbance at 1655 cm⁻¹ that indicated the dominance of helix structures,³⁶ with the possibility of some unordered structures. MRPs also had absorbance peaks centering on 1675, 1641, and 1630 cm⁻¹ that are attributed to turn structures,³⁷ disordered structures,³⁸ and short-segment chains connecting α-helical segments,³⁷ respectively. After mixing MRPs with oil and titration to pH 7.0–6.2 to prepare emulsions, absorbance peaks at 1675 cm⁻¹, 1641 cm⁻¹ and 1630 cm⁻¹ became more apparent, demonstrating the rearrangement from ordered α-helical structures to less ordered secondary structures. Amid I bands were similar among emulsions, indicating similar physical and chemical events after titration to pH 6.2–7.0.

The defined bands located at 1687 cm⁻¹ and 1618 cm⁻¹ of freeze-dried emulsions (Fig. 5b) suggest the formation of inter- and intra-molecular antiparallel pleated β-sheet structures, respectively.^39,40 β-Sheet structures are correlated with protein folding and aggregation,²⁷ and the increase of β-sheet structures agreed with the lowered solubility of MRPs at a lower pH (Fig. 1). Thereafter, secondary structural changes of MRPs after titration resulted in the deposition of MRPs on oil droplets and shell formation, as presented previously. The formation of intermolecular β-sheet structures also corresponds to the lowed solubility of MRPs due to aggregation and precipitation. Most proteins from cereals and legumes precipitate around their isoelectric point (pI),^41–43 and the precipitation triggers subtle conformational alterations of proteins.¹ The rapid solubility decrease of MRPs from pH 7.0 to 6.0 enables the fabrication of materials such as emulsions above the MRP pI at pH 4.5–5.0 (ref. 11) using pH as a variable.

Properties of encapsulating β-carotene

The encapsulation efficiency and loading capacity of β-carotene loaded in emulsions prepared at pH 6.2–7.0 are listed in Table 3. Both encapsulation efficiency and loading capacity of β-carotene increased for emulsions prepared at a lower pH, except no significant difference between the pH 6.2 and 6.4 treatments. Because MRPs are more soluble (Fig. 1) and shells on oil droplets are thinner at a higher pH (Fig. 2), oil droplets may be subjected to flocculation, coalescence, and demulsification to form larger aggregates that were removed after centrifugation (Fig. S1†), which lowers both encapsulation efficiency and loading capacity. This speculation was further supported by the about 90% recovery of MRPs (Table 3).

Table 3 Properties of encapsulating β-carotene in fresh emulsions^a

pH after titration	Encapsulation efficiency (%)	Loading capacity (mg g^−1)	MRP recovery (%)
a Values are mean ± SD (n = 3). Different superscript letters in the same column indicate significant differences (P < 0.05).
6.2	13.38 ± 1.70^a	0.41 ± 0.05^a	87.1 ± 0.3^b
6.4	14.40 ± 0.96^a	0.42 ± 0.03^a	91.0 ± 0.7^a
6.6	7.18 ± 0.97^b	0.29 ± 0.04^b	91.6 ± 0.5^a
6.8	8.88 ± 0.57^b	0.33 ± 0.02^b	92.3 ± 0.7^a
7.0	3.94 ± 0.93^c	0.20 ± 0.03^c	89.3 ± 0.8^ab

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Release kinetics of β-carotene during simulated digestions

Fig. 6 shows the kinetics of β-carotene release from fresh emulsions during the simulated 2 h gastric digestion and subsequent 2 h intestinal digestion. A burst release was observed in the first 10 min of the simulated gastric digestion, followed by a negligible release rate in the next 110 min (Fig. 6a). This observation correlated well with the digestion profile of MRPs that were susceptible to peptic digestion in the first 10 min followed by insignificant further hydrolysis (Fig. 7). Because MRPs are insoluble at gastric acidity (Fig. 1), droplets became aggregated when subjected to the simulated gastric treatment (shown in Fig. S2† for the pH 6.6 treatment). The measured β-carotene in the supernatant after extraction with petroleum ether and centrifugation likely resulted from the release of oil phase during peptic digestion of MRP shell on oil droplets. With the exception of the emulsion prepared at pH 6.2, thicker shells of emulsions formed at a lower pH generally agreed with the better retention of β-carotene (Fig. 6a), and the low percentage of release (<10%) suggests a lower extent of peptic digestion of MRPs on oil droplets than the neutral MRP solution (Fig. 7b). This likely results from conformations of MRPs on oil droplets being different from those in the aqueous phase, as evidenced by the zeta-potential (Tables 1 vs. 2) and FTIR data (Fig. 5), which changes the susceptibility to pepsin digestion. For the emulsion prepared at pH 6.2, the loosely aggregated structures were observed before the simulated digestion (Fig. 3e), but big droplets were also found after treatment by pepsin (Fig. S3†) which was different from the emulsion prepared at pH 6.6 (Fig. S2†). It is possible that the emulsion prepared at pH 6.2 had loosely aggregated MRPs and the initial pepsin digestion resulted in the release of a higher extent of the oil phase than other pH treatments.

Fig. 6 Release kinetics of β-carotene during (a) 2 h simulated gastric digestion and (b) subsequent 2 h simulated intestinal digestion of fresh emulsions. Error bars represent SD (n = 3).

Fig. 7 Digestion profile of MRPs: (a) size-exclusion chromatograms showing MRPs after peptic digestion for up to 120 min and the subsequent pancreatic digestion for up to 120 min, and (b) digestability of MRPs calculated from the chromatograms. The elution times of protein standards with molecular weights of 77 (transferrin), 16 (apomyoglobin) and 12 (cytochrome c) kDa are marked with vertical arrow lines labeled a, b, and c, respectively.

During the subsequent simulated intestinal digestion, the emulsion prepared at pH 6.2 quickly reached 100% release of β-carotene (Fig. 6b), which may have resulted from the loose MRPs structures becoming dissolved at pH 7.0 and subsequently releasing oil (Fig. S3†). Other treatments exhibited the zero-order release kinetics (R² > 0.9, Table 4) that had a higher release rate for the emulsion prepared at a higher pH (Fig. 6b and Table 4) and followed the same trend as observed in the simulated gastric digestion (Fig. 6a). Because MRPs are soluble at neutral intestinal acidity (Fig. 1) and MRPs can be digested to greater than 80% after 2 h pancreatic treatment (Fig. 7b), the results in Fig. 6b further support the hypothesis about differences in the digestion properties of MRPs in the aqueous phase and on oil droplets. As a result, oil droplets prepared at a higher pH had a thinner shell and were demulsified quicker during pancreatic hydrolysis of shells, corresponding to a higher release rate (Fig. 6b and Table 4).

Table 4 Parameters after linear fitting (y = kx + b) of β-carotene release kinetics during the simulated intestinal digestion of emulsions prepared at pH 6.4–7.0

Emulsion pH	k	b	R^2
7.0	0.79	4.50	0.98
6.8	0.17	3.93	0.92
6.6	0.09	4.12	0.93
6.4	0.04	1.38	0.93

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The retention of most β-carotene after gastric digestion (Fig. 6) is advantageous to many protein-based delivery systems. For example, rapid release of encapsulated compound during gastric digestion was observed for chitosan–protein complex nanoparticles,⁴⁴ soy protein microspheres,¹ and zein nanoparticles.⁴⁵ To address this challenge, shell glycosylation⁴⁶ and shell coating⁴⁷ were studied, which increases the complexity of encapsulation and reduces practicality of the technology. The retention of about 90% of β-carotene during the 2 h simulated gastric digestion and the release of over 90% of β-carotene following the zero-order kinetics show the potential application of the emulsion fabricated at pH 7.0 as enteric delivery systems. Conversely, the limited release of β-carotene from emulsions prepared at pH 6.4–6.8 after the simulated gastric and intestinal digestions indicates the possibility of delivering the encapsulated bioactives to the colon, e.g., for anti-inflammatory drugs to treat relevant bowel diseases¹³ if MRPs can be digested by enzymes secreted by colorectal microflora.

Conclusions

A one-step titration method was used to successfully fabricate emulsions utilizing significant changes in solubility and conformations of MRPs at pH 6.0–7.0. Titration resulted in precipitation of MRPs on oil droplets to form shells with the thickness controllable by the final pH of titration. The diameter of oil droplets was smaller than 400 nm even without using an emulsification unit operation applying high mechanical energy. The tunable conformational and digestion properties of MRPs on oil droplets and the shell thickness enabled the limited release of encapsulated β-carotene during the simulated gastric digestion and the controllable zero-order release rate in the subsequent simulated intestinal digestion. The presented results highlight the potential of MRP-based emulsions as a carrier of lipophilic bioactives for controlled release after oral intake.

Acknowledgements

This work was supported by the National High Technology Research Development Program of China (863 Program) (No. 2013AA102204, 2013AA102206), National Natural Science Foundation of China (No. 31201381, 31471616 and 31371874) and Special Fund for Agro-Scientific Research in the Public Interest of China (No. 201303071). T. Wang would like to thank the scholarship provided by the China Scholarship Council. Q. Zhong would like to acknowledge the support from the University of Tennessee and the USDA National Institute of Food and Agriculture Hatch Project 223984.

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Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra14943j

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