Fabrication and delivery properties of soy Kunitz trypsin inhibitor nanoparticles

Abstract

Soy Kunitz trypsin inhibitor nanoparticles (KTIP) were prepared successfully by heating KTI at 80 °C in the presence of sodium sulfite. This treatment not only inactivated most trypsin inhibitor activity (TIA) of KTI (about 90.63%), but also resulting in the formation of KTI nanoparticles (about 85.92 nm) with good monodispersity (PDI: 0.18), which were confirmed by dynamic light scattering (DLS) and small-angle X-ray scattering (SAXS) determination. SDS-PAGE analysis revealed that the formation of limited aggregates (KTIP) was mediated by an –SH/–SS– exchange reaction. The delivery capacity of KTIP for curcumin as model bioactives was evaluated. The results indicated that the co-assembly of KTIP and curcumin actually greatly enhanced the dispersibility, stability and bioaccessibility of curcumin in aqueous solution. Moreover, in vitro anti-proliferative activity on tumor cells assay showed that nanoparticulate curcumin was more effective than free curcumin in solution by controlling the tumor cell growth with time. These findings suggest that KTIP could be developed as a novel nano-delivery vehicle for hydrophobic bioactives and for use in functional foods and pharmaceutics.

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Introduction

Protein-based nanoparticles have gained increasing interest as delivery systems for drugs and nutraceuticals in the past few decades.¹ To date, drug-loaded nanoparticles have been synthesized successfully from various proteins, including both water-soluble (bovine or human serum albumin, β-lactoglobulin) and insoluble proteins (zein, gliadin, barley protein etc.).^2–4 These nanoscaled systems exhibited various advantages, such as improved solubility, controlled release property and enhanced bioavailability of encapsulated nutraceuticals.^2,4 Additionally, they exhibit low toxicity due to superior biocompatibility and nutritional value.⁵ Despite these advantages, the application of protein-based nanoparticles has been impeded by several drawbacks. First, the loading content (LC) of protein nanoparticles is generally low compared to that of synthetic polymeric nanoparticles.⁶ The second disadvantage lies in the toxicity of chemical cross-linkers.⁵ Cross-linking is a necessary procedure for the preparation of nanoparticles with an amount of water-soluble proteins, which dissociate readily into their monomeric forms upon the removal of antisolvents.²

To overcome the above-mentioned drawbacks of protein-based nanoparticles, two strategies were proposed in this study. The first strategy is heat treatment for protein, because heating can result in structural unfolding and denaturation of proteins, thus increasing their surface hydrophobicity for binding hydrophobic bioactives. The second strategy is to select a disulfide bonds or cysteine-rich protein to prepare nanoparticles via intermolecular disulfide bonds hence avoiding use chemical cross-linkers.

In the last decade, American Bioscience, Inc. has developed a unique albumin-based nanoparticle technology (nab-technology) that is ideal for encapsulating lipophilic drugs into nanoparticles.⁷ Abraxane® (nab-paclitaxel; paclitaxel-albumin nanoparticle) with an approximate diameter of 130 nm is the first FDA approved nanotechnology based chemotherapeutic that has shown significant benefit in treatment of metastatic breast cancer. The market approval of Abraxane® can be viewed as a landmark not just for albumin-based drug delivery technology but also for nanomedicine.^7,8 The near future may also hold the emergence of new commercial protein nanocarrier-based products. However, at the present stage, a better fundamental understanding of the mechanisms of action of these vehicles and of the protein–drug interactions at the molecular level will provide a basis for their further optimization to ensure design of ideal protein nanocarriers and open more exciting opportunities for their use in the area of bioactives and drug delivery.⁸

Kunitz trypsin inhibitor (KTI), the major protein in soy whey protein, is a small (20.1 kDa) globular protein with two disulfide bridges (Cys39–Cys86 and Cys136–Cys145), which are solvent-exposed and critical for its inhibitory function as well as the resistance to thermal and chemical denaturation.⁹ These disulfide bonds also play an essential role in the allergenic potential of KTI.¹⁰ The crystal structure of KTI complexed with porcine trypsin has been reported¹¹ and subsequently refined¹² (PDB ID: 1AVU and 1BA7, Fig. S1†). KTI has long been known as an antinutrient in humans consuming soy proteins although the denatured inhibitors are highly nutritious, rich in sulfur amino acids and well-balanced proteins according to a dual tracer approach to measuring (DIAAS) proposed by FAO in 2014.¹³ For this reason, extensive efforts have been made to devise processing conditions for inactivating or removing trypsin inhibitors from legumes. These approaches are based largely on the heat treatment. However, heat treatments do not completely inactive all inhibitors.⁹ Therefore, alternative strategies were developed by using various reducing agents since the cleavage of disulfide bonds seems to be responsible for the inactivation of trypsin inhibitors (TIs) in soy flour.^14,15 Friedman et al.¹⁴ found that treatment of raw soy flour at 75 °C with 0.03 M sodium sulfite for 1 h completely inactivated TIs leaving no sulfite residues in the soy proteins and hence the improvement of soy flour nutrition. It is suggested that TIs in soy flour are inactivated by sulfhydryl–disulfide interchange during the first inactivation phase and by heat during the second phase.¹⁵ Nowadays, some protease inhibitors-rich protein, such as potato protein (protease inhibitor representing 50% of total protein), has been developed into commercial products such as Solanic® due to its good nutritional value and functional properties.¹⁶

The use of heat combined with sulfite to inactivate protease inhibitors usually results in protein denaturation and aggregation, which likely involves –SH/–SS– exchange reaction and can affect the native state and stability of the protease inhibitors. Consequently, a better understanding of their behavior during heating is essential for the control of their size and properties. However, the aggregation behavior of KTI, the characteristic of KTI aggregates as well as the delivery capacity of KTI aggregates for hydrophobic bioactives remain unknown. Therefore, the objectives of this work were as follows: (1) to prepare KTI nanoparticles through limited aggregation behavior and to analyze the formation mechanism of KTI aggregates during inactivation treatment by heating in the presence of sodium sulfite; (2) to evaluate the delivery capacity of KTIP for curcumin as model bioactives.

Results

Preparation and characterization of KTI aggregates

As shown in Fig. 1A, no aggregates were observed in the KTI solution (Fig. 1A1), the KTI solution in the presence of sodium sulfite (Fig. 1A3), and the KTI solution in the presence of sodium sulfite and N-ethylmaleimide (NEM) (Fig. 1A5) at room temperature (25 °C). The soluble aggregates appeared after heating at 80 °C for 1 h in the absence (Fig. 1A2) or presence of sodium sulfite (Fig. 1A4). Interestingly, it was noticed that there were no obvious aggregates generated after heating in the presence of the thiol-blocking agent NEM in the solution (Fig. 1A6). These observations suggest that the formation of aggregates may be mediated by disulfide bonds. The morphology of KTI aggregates generated by treatment 4 is showed in Fig. 1B. As can be seen, the size of KTI aggregates was nanoscale (about 80 nm) and appeared as a globular conformation. Size distribution of KTI with various treatments is showed in Fig. S2.† As can be seen, both treatment 2 and 4 had desired monodispersity as compare to those without heating treatments (1, 3, and 5).

Fig. 1 (A) The images of KTI solutions with various treatments. The various treatments (1–6) are described in the section of ‘Preparation of KTI solutions and inactivation treatments’ in Material and methods. (B) TEM image of treatment 4. (C) Trypsin inhibitor activity (TIA) of KTI with various treatments (1–6). Data bearing different letters are significantly different (p < 0.05).

To measure the effects of various treatments on the trypsin inhibitor activity (TIA) of KTI, TIA of native KTI and KTI with various treatments were investigated (Fig. 1C). The TIA value of native KTI was 7658 ± 230 U mg⁻¹, and it can be seen that there was only 9.37% of TIA found in the aggregates induced by heating in the presence of sodium sulfite, 33.38% of TIA found in sample 6, while the aggregates only induced by heat still retained 58.81% of TIA. Both sample 3 and 5 retained almost 100% of TIA. These results indicate that the heating combined with sodium sulfite could effectively inactive TIA of KTI. In other words, the aggregates induced by heating in the presence of sodium sulfite possessed a low TIA, suggesting that the aggregates have potential in food applications because most commercially heated meals retain up to 20% of the original trypsin and chymotrypsin inhibitory activity.¹⁷

To further verify whether disulfide bond formation is involved in the aggregation of KTI, all solutions (Fig. 1A1–6) were analyzed by SDS-PAGE under non-reducing conditions (Fig. 2A), and the solution 4 and 6 in Fig. 1A were resolved by reducing SDS-PAGE (Fig. 2B). The buffer system used with SDS-PAGE under non-reducing conditions presumably disperses non-covalently linked protein aggregates into monomers, while aggregates linked though disulfide bonds remain intact. A KTI monomer and two faint protein bands of high molecular mass could be seen in all unheated samples (Fig. 2A, lane 1, 3 and 6). After heating, heterogeneous aggregates generated in the absence of sodium sulfite (Fig. 2A, lane 2) and a small amount of KTI monomer still existed. Interestingly, in the presence of sodium sulfite, all KTI monomers were converted into homogeneous aggregates (Fig. 2A, lane 4). Moreover, the homogeneous aggregates band mainly migrated to KTI monomer when it was separated by reducing SDS-PAGE (Fig. 2B, lane 4), suggesting that disulfide bonds were responsible for the intermolecular interactions of KTI induced by heating in the presence of sodium sulfite. To get further insight into the role of the thiol group of KTI in formation of aggregates, NEM was added into the KTI solution before heating to block free thiol groups. With SDS-PAGE under non-reducing conditions, no homogeneous aggregates like in lane 4 were observed after heating (Fig. 2A, lane 4), giving further proof that polymerization must result from thiol-induced disulfide exchange reactions. Nevertheless, another narrow protein band appeared at top of separation gel of lane 2, 4, and 6 in Fig. 2A, respectively, which was also probably mediated by disulfide bonds under heating treatment, because these protein bands were disappeared under reducing SDS-PAGE (Fig. 2B). In the case of sample 6 (Fig. 2A, lane 6), the protein band indicated by green arrow may be due to free thiol groups of native KTI blocked incompletely by NEM.

Fig. 2 Non-reducing (A) and reducing (B) SDS-PAGE patterns of KTI with various treatments. The samples in lanes 1–6 are corresponding to the solutions of 1–6 in Fig. 1A, respectively. The homogeneous aggregates induced by heating in the presence of sodium sulfite are indicated by red arrow (lane 4). The aggregates induced by heating on top of separation gel are indicated by green arrows (lane 2, 4, and 6). M: molecular weight marker.

Dynamic light scattering (DLS) and small-angle X-ray scattering (SAXS) were used to characterize the particle size, shape and ζ-potential of KTI with various treatments. The radius of gyration R_g and the extrapolation of intensity to an angle of zero I₍₀₎, the two most fundamental structural parameters, were calculated by Guinier approximation derived from SAXS data, ln I_(q) versus q².¹⁸ I₍₀₎ is proportional to the molecular weight of scattering particles, while R_g represents the size or molecular compactness.¹⁹ A summary including R_g, I₍₀₎, the hydrodynamic radius R_h, and ζ-potential is reported in Table 1. As expected, R_g and I₍₀₎ of heat-treated KTI were higher than those of native protein, suggesting that protein unfolding and subsequent aggregation occurred between monomers of KTI, but the extent of KTI aggregation did not increase obviously when KTI thiol was blocked with NEM. The results of DLS experiment were similar to the changes in R_g, however, in all cases, hydrodynamic radius R_h was obviously larger than R_g. The different q range used in DLS and SAXS experiments could largely account for this discrepancy. The large aggregates mainly contribute to determine R_h at low q values during DLS experiment. At the q values seen by SAXS, this contribution was strongly reduced, and thus, R_g mostly came from the contributions of monomers. Similar observations were found in our previous works for soy protein^18,19 and were also reported for β-lactoglobulin microgels using SAXS and DSL as testing methods.²⁰ It was noteworthy that the PDI values of KTI aggregates distinctly decreased compared to that of native KTI, indicating that polydisperse native KTI could form homogeneous aggregates induced by heating.

Table 1 Structural parameters and ζ-potential of KTI with various treatments

KTI^a	Rg^b (SAXS) (nm)	Rh^c (DLS) (nm)	PDI^d	Rg/Rh	I(0)^e	ζ-Potential (mV)
a The KTI samples of 1–6 are corresponding to the solutions of 1–6 in Fig. 1A, respectively.b Overall radius of gyration of KTI with various treatments.c Hydrodynamic radius.d Polydispersity index determined by DLS.e Extrapolation of intensity to an angle of zero.
1	2.89 ± 0.11	10.36 ± 0.44	0.38	0.28	0.67 ± 0.02	−14.6
2	8.68 ± 0.23	41.96 ± 1.52	0.15	0.21	2.51 ± 0.11	−12
3	2.96 ± 0.13	11.67 ± 0.57	0.31	0.25	0.70 ± 0.04	−8.5
4	9.65 ± 0.25	42.96 ± 1.58	0.18	0.22	2.83 ± 0.12	−18.7
5	3.01 ± 0.16	11.89 ± 0.49	0.43	0.25	0.78 ± 0.03	−15.2
6	5.13 ± 0.10	19.69 ± 0.95	0.26	0.31	2.08 ± 0.09	−10.4

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The ratio of R_g to R_h is frequently used to characterize polymeric architectures. A qualitative, approximate interpretation of the shapes for the aggregates can be provided by this ratio.¹⁹ The value of R_g/R_h is expected to be 0.775 for hard-type spheres.²¹ Polymers with unfolded structure could possess higher values. In the present study, the R_g/R_h values of all samples were all well below this value. Although this might also suggest the presence of compact objects with possible structures, the estimation of R_g/R_h from the combination of DLS and SAXS data might not predict the exact shapes of the samples. However, it was still possible to predict the general trend of the particle shape from the value of R_g/R_h. The values of R_g/R_h of samples with heating decreased somewhat except the sample in the presence of NEM, indicating that disulfide bonds linked aggregates become less dense.

The ζ-potential is the electric potential in the interfacial double layer at the location of the slipping plane relative to a point in the bulk fluid away from the interface. The ζ-potential of native KTI is about −14.6 mV, and the absolute value of ζ-potential of KTI aggregates (formed in the presence of sodium sulfite) slightly increased.

Because only KTI aggregates generated by treatment 4 lost most of its TIA, but resulting in the formation of KTI aggregates in nanoscale according to DLS data and transmission electron microscopy (TEM), we designated this KTI aggregates as KTI nanoparticles (KTIP), and only KTIP relative to native KTI were characterized in the following assays.

Circular dichroism (CD) is an important and direct technique for protein structure studies. CD spectra of KTIP relative to native KTI are shown in Fig. 3. The fractional contents of secondary structure (α-helix, β-sheet, β-turn, and random coil) of samples were calculated, and they are shown in the inset of Fig. 3. As can be seen from Fig. 3A, both KTI and KTIP have a CD spectrum characteristic of the class of β-II proteins⁹ with a minimum ellipticity at about 202 nm, indicating that there was almost no obvious change of the backbone structure in KTIP. Actually, during thermal denaturation in the present of sodium sulfite and the thiol-blocking agent NEM, the ellipticity at 202 nm showed a slight decrease and a red shift (Fig. S3A†) that was indicative of melting of secondary structure with some solvent exposure. Nevertheless, upon the formation of aggregates, these changes disappeared, which may be involved in formation of intermolecular hydrogen bonds during aggregation process.

Fig. 3 Circular dichroism (CD) spectra of KTIP relative to KTI. (A) Far-UV CD spectra, (B) near-UV CD spectra. The secondary structure was calculated by CDSSTR program.

The data in Fig. 3B showed the near-UV CD spectra of KTIP relative to KTI which was indicative of changes at the level of tertiary structure. There is maximum ellipticity of KTI due to the presence of side-chain interactions, especially the aromatic residues that lie in the hydrophobic core of the protein.⁹ The CD spectra in the region 260–320 nm arise from the aromatic amino acids (Phe, Tyr and Trp). Each of the amino acids tends to have a characteristic wavelength profile (Phe: 260–270 nm, Tyr: 275–282 nm, Trp: 290–305 nm).²² In addition, the actual shape and magnitude of the near UV CD spectrum of a protein will depend on the number of each type of aromatic amino acid present, their mobility, the nature of their environment (H-bonding, polar groups and polarisability) and their spatial disposition in the protein; near neighbours (generally less than 1 nm apart) may be able to couple as excitons, although the signals are generally too weak for this to be significant.²² For KTIP, there was a progressive decrease in ellipticity especially in the region of 275–280 nm, which was the most prominent change among KTI with various treatments (Fig. S3B†), indicating that most of the tertiary structural interactions have been lost. This decrease in near-UV ellipticity of KTIP also indicated that native KTI underwent transition in conformation and changes in the hydrophobic environment of aromatic residues (Phe, Tyr and Trp).

The surface hydrophobicity of protein was previously used to identify the structural change of proteins.^19,23 Table 2 presents the surface hydrophobicity of KTI and KTIP. The fluorescence intensity at saturating concentrations of the probe (F_max) represents an overall surface hydrophobicity (protein surface hydrophobicity, PSH). The F_max, apparent dissociation constant of the protein–8-anilino-1-naphthalenesulfonic acid ammonium salt (ANS) complex (K_d), and PSH of KTIP were significantly (p < 0.05) higher than those of native KTI, respectively, indicating the formation of new hydrophobic sites on KTIP's surface and the decrease of the binding affinity of ANS to protein, consistent with previous papers.²⁴ The results are in accordance with the data of CD spectroscopic analysis (Fig. 3).

Table 2 Protein surface hydrophobicity (PSH, assessed by titration with ANS as the hydrophobic probe) of KTI and KTIP^a

Protein samples	Fmax	Kd (μM)	PSH (F/mg μM)
a Different letters (a, b) in the column indicate significant (p < 0.05) differences among samples. Fmax: the maximum fluorescence intensity (at saturating probe concentration). Kd: the apparent dissociation constant of the protein–ANS complex. PSH: protein surface hydrophobicity index (PSH = Fmax/Kd × [protein concentration], fluorescence intensity/mg μM). The protein concentration is 0.20 mg mL^−1.
KTI	151.52 ± 4.53^b	35.32 ± 1.06^b	21.45 ± 0.89^b
KTIP	666.67 ± 19.98^a	114.93 ± 4.60^a	29.00 ± 1.15^a

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Interfacial properties of KTIP

The time evolution of surface tension (γ) for KTI and KTIP at the air–water interface is shown in Fig. 4A. The surface tension values gradually decreased with adsorption time for both samples, which can be attributed to adsorption of proteins to the interface, along with the saturation, conformational changes, and continuous conformational rearrangements within the film and/or to multilayer formation.²⁵ Although both samples had similar initial surface tension values, the extent of reduction of tension for KTI is greater than that of KTIP. This observation may be related to the relatively large size of the KTIP (Table 1), and the presence of steric hindrance derived from residual protein molecules could prevent some of the aggregated molecules from reaching the interface. Moreover, first adsorbed proteins at the interface produce a barrier to reduce the adsorption of large molecules, which are anchored into the interface only by a small part of their surface.^18,26

Fig. 4 (A) Surface tension (γ) as a function of time for solutions of 0.1% KTIP relative to 0.1% KTI at the air–water interface. (B) Time-dependent dilatational elasticity (E_d) for adsorbed layers formed from the above solutions. (C and D) AFM images of mica-supported native KTI (C) and KTIP (D) monolayer in the air–water interface at a pressure of 30 mN m⁻¹.

Interfacial rheology of adsorbed layers is considered as indicators of structural state of proteins adsorbed at the interface and macromolecule interactions.^18,27 In all cases, it can be noticed that (data not shown) the values for surface dilatational modulus (E) were very similar to those for the dilatational elasticity (E_d), and the dilatational viscosity (E_v) values were low. Thus, from a rheological point of view, these results suggested the surface layers behaved as viscoelastic (primarily elastic) during the adsorption period studied here.²⁸ The dynamic elastic modulus of interfacial layers during protein adsorption is presented in Fig. 4B. Generally, the gradual increase in E_d with adsorption time should be attributed to protein adsorption and developing intermolecular contacts at the interface. As compared with native KTI, KTIP showed higher E_d values as soon as proteins started to adsorb, which could be associated with the rapid establishment of intermolecular contacts in the adsorbed layer. Attractive interactions (hydrophobic interactions) may contribute to the development of interfacial rheology following the formation of an interfacial layer, which is one largely accepted mechanism.²⁹ The actual molecular adsorption to the interface is believed to be accompanied by protein unfolding since KTIP possessed the weakened tertiary structures, which was evidenced by the data of CD, DLS and SAXS, and the consequent formation of intermolecular interactions might be mediated by hydrophobic forces. This was evidenced by increased surface hydrophobicity for KTIP (Table 2).

To observe the interfacial architecture formed by KTIP or KTI at the interface, the topographies of the transferred protein Langmuir–Blodgett films was imaged using AFM. As shown in Fig. 4C and D, it could be observed that both protein molecules are adsorbed onto the mica substrate as solid-like state sphere conformation. In addition, KTIP displayed a larger size globular structure as compared to KTI molecules, which matched the data of SAXS and DLS (Table 1).

Fabrication and characterization of curcumin-loaded KTIP

Over the last decade, the field of nanoparticle delivery systems for nutrients and nutraceuticals with poor water solubility has been expanding, almost exponentially, and some of these technologies are now in the process of being incorporated in food products. Because KTIP has favorable interfacial properties (Fig. 4A and B) and a nanoscale globular particle conformation (Fig. 4D), it has the potential for acting as a novel nano-delivery vehicle for hydrophobic bioactives. Therefore, curcumin was chosen as a model and curcumin-loaded KTIP were prepared by antisolvent precipitation. The entrapment efficiency (EE) and LC of curcumin in KTIP are shown in Fig. 5A. From Fig. 5A, we can interesting see that, as the concentration of KTIP increased from 0.1% to 0.5%, the EE of curcumin progressively increased from 48.6% to 96.17%, while the LC of curcumin linearly decreased from 16.28% to 7.14% (i.e., decreased from about 194.40 to 76.94 μg mg⁻¹ of KTIP). When the concentration of KTIP was more than 0.3%, the EE is comparable to that (>95%) of curcumin encapsulated in β-lactoglobulin nanoparticles produced by desolvation,³⁰ but considerably higher than that of curcumin encapsulated in zein-based colloidal particles synthesized by an antisolvent precipitation (71–87%).³¹ The loading amount of curcumin is much greater than that (1.743–1.784 μg mg⁻¹ of SPI) in soybean protein isolates (SPI)–curcumin nanocomplexes³² and that (19 μg mg⁻¹ of casein) in the supernatant of curcumin encapsulated in casein nanocapsules.³³ Generally, the typical LC for hydrophobic compound-loaded protein nanoparticles is around 5%.⁵ In the present study, the LC (7.14–16.28%) was greater than this value. It should be pointed out that a number of literature studies claimed LC values >50%, but these results were arguably inaccurate because of improper measuring methods, for example, counting precipitated nutraceutical molecules as encapsulated ones or neglecting the weight of a secondary coating when calculating the LC.⁵

Fig. 5 (A) Entrapment efficiency (EE) and loading content (LC) of curcumin in KTIP solutions with different concentration. Inset: 0–6: fresh curcumin stabilized by the 0, 0.1, 0.2, 0.3, 0.4, and 0.5% KTIP, respectively. (B) Size distributions of curcumin-loaded KTIP (fresh curcumin stabilized by the 0.4% KTIP). Inset: TEM image of curcumin-loaded KTIP. (C) XRD diffractograms of curcumin powder, KTIP power and lyophilized curcumin-loaded KTIP powder. (D) Residual ratios of curcumin as a function of storage time under ambient conditions.

Because the increase of EE was very small (only 0.39%) with increasing KTIP concentration from 0.4% to 0.5%, KTIP concentration more than 0.5% has not been studied, and 0.4% of KTIP concentration was selected as the optimum concentration for preparing KTIP–curcumin complexes in the following assays. The appearances of curcumin in pure PBS and different concentration KTIP solution can be observed in Fig. 5A (inset). The free curcumin in PBS was very turbid due to its poor water solubility. However, the dispersibility of curcumin was increased with the increase of the KTIP solution concentration, and the KTIP–curcumin mixture solutions exhibited yellow and highly transparent appearances when the concentration of KTIP solution was more than 0.2%. The water dispersibility of curcumin could reach up to 127.7 μg mL⁻¹ ((0.1 mL × 4 mg mL⁻¹) × 95.78%/3 mL) when the KTIP concentration was 0.4%, which increased approximately 49 times compared with that of free curcumin in PBS (2.6 μg mL⁻¹). It is noteworthy that curcumin dispersibility also has a great increase in PBS after the antisolvent process. A similar result has been previously reported, in which the curcumin solubility in water is 3.14 μg mL⁻¹ after a similar antisolvent treatment.³¹ Nevertheless, curcumin dispersibility is increased 116 090-fold by this antisolvent process in KTIP solutions according to previously reported results (11 μg L⁻¹).³⁴ Consequently, KTIP could be utilized as a solubilizer for curcumin. The solubilization of curcumin in KTIP solutions could be attributed to the entrapment of curcumin in the hydrophobic core or binding of curcumin to surface hydrophobic patches of KTIP. Mean particle size and size distributions of curcumin-loaded KTIP are presented in Fig. 5B. As can be seen, the mean particle size of curcumin-loaded KTIP was 93.88 nm with good monodispersity (PDI = 0.123), which was evidenced by TEM (the inset in Fig. 5B). It was slightly larger than the size of KTIP (85.92 nm, Table 1).

XRD was performed to study the crystallinity of curcumin after its complexation with KTIP and the results are shown in Fig. 5C. For the XRD patterns of free curcumin exhibited intense diffraction peaks between 5° and 30°, indicating its highly crystallized structure. Oppositely, the typical amorphous XRD pattern was observed for KTIP. However, the diffraction spectrum of curcumin-loaded KTIP showed complete disappearance of all the characteristic crystalline peaks of curcumin, indicating the formation of amorphous curcumin. This observation should be attributed to the inhibition of its crystallization in the nanoscale confinement and the formation of an amorphous complex with KTIP within the particle matrix.

Curcumin in aqueous solution is readily susceptible to hydrolysis or degradation, even at physiological pH.³⁵ Herein, the short-time storage stability of free curcumin and curcumin in KTIP solutions was evaluated (Fig. 5D). The result showed that free curcumin in PBS was very unstable, and only 3.1% curcumin was retained after 24 h storage under ambient conditions. As expected, the stability of curcumin in the presence of KTIP was markedly improved. Curcumin was very stable in the initial 24 h, in spite of the stability slowly decreasing with further extension of storage time. The final retained ratio of curcumin was still above 80% in KTIP solution with a concentration of 4 mg mL⁻¹. A similar improvement of storage stability has also been observed for curcumin in complexes with serum albumins³⁵ and milk proteins including α_s1-casein³⁶ and β-lactoglobulin.³⁰

During the digestion, the nanoparticulate curcumin may undergo dramatic changes in environmental conditions (pH and ionic strength), action of proteases on the proteins, and even changes due to the presence of different active surfactants.³² After sequential processes of in vitro gastric (60 min) and intestinal (120 min) digestion, the bioaccessible amount of curcumin transferred to the aqueous phase of the digests and whole digests for free curcumin and nanoparticulate curcumin is shown in Fig. 6. For free curcumin, only 52.30% of curcumin was remained in the aqueous phase after the whole digestion. In contrast, for nanoparticulate curcumin, there was no significant loss of curcumin throughout the digestion, and the remaining amount in the aqueous phase could reach up to 94.56%, which was near the value in the whole digests (97.02%). The results indicated that KTIP could markedly increase the bioaccessibility of curcumin. Intestinal absorption of hydrophobic bioactives, such as curcumin, is dependent on their solubilization in the aqueous intestinal environment via the emulsifying action of the bile salts.³⁷ The bile salts can trap lipophilic compounds in mixed micelles or vehicles, carry them through intestinal cells barriers, and transport them into the blood circulation. In addition, the bioaccessibility of bioactives is a prerequisite for their bioavailability. It assumes that the solubilized substance may have a high potential to be absorbed by the small intestine.³⁷

Fig. 6 Percentage of curcumin remaining in the aqueous phase or the whole digests of free curcumin and curcumin-loaded KTIP after the whole simulated digestion of 180 min under darkness. Means of columns marked with different letters indicated a significant difference between each other (p < 0.05).

To assess the bioactivity of nanoparticulate curcumin, in vitro anti-proliferative activity on tumor cells of curcumin (free and nanoparticulate curcumin) was investigated in different cell line by MTT assay. All the studied cell line showed a typical dose dependent anti-proliferative effect (Fig. 7). The in vitro half maximal inhibitory concentration (IC₅₀) is the quantitative measurement for the cell toxicity induced by chemotherapeutic drug. This IC₅₀ values were calculated from the obtained curves of all the studied cell line and the result demonstrated nanoparticulate curcumin has higher anti-proliferative activity than free curcumin (Table S1†). Nanoparticulate curcumin is 1.48, 1.64, 2.08, 1.62, 2.09 and 2.33 times more effective than free curcumin as observed in A549, D145, HCT-116, HepG2, K562 and MCF-7 cell line, respectively. The obtained results demonstrated comparable inhibition of cell proliferation, where nanoparticulate curcumin was more effective than free curcumin in solution by controlling the tumor cell growth.

Fig. 7 Dose dependent cytotoxicity of KTIP, free curcumin, and curcumin-loaded KTIP in A549 (A), Du145 (B), HCT-116 (C), HepG2 (D), K562 (E), and MCF-7 (F) cell lines. The extent of growth inhibition was measured at 48 h by the MTT assay. The inhibition was calculated with respect to controls. Data as mean ± SD, n = 6. (*) p < 0.05, free curcumin in solution versus curcumin-loaded KTIP. KTIP toxicity was tested at the concentrations that are required to achieve the active concentrations of curcumin, i.e. the concentrations of KTIP were 0.06, 0.12, 0.18, 0.24, 0.30, and 0.36 mg mL⁻¹ corresponding to the concentrations of curcumin of 5, 10, 15, 20, 25, and 30 μM, respectively.

Discussion

Generally, proteins as natural polymers are heterogeneous mixtures of different sizes with a wide range of molecular weights thus producing heterogeneous nanoparticle size distribution and exhibiting batch-to-batch variation.⁸ This may hinder the scaling-up process of protein nanoparticle preparation for industrial application. An interesting strategy to overcome this drawback is the recombinant protein technology. The monodispersity and precisely defined properties of recombinant proteins as well as the predictable placement of crosslinking groups, binding moieties or their programmable degradation rates make them useful for drug delivery.⁸ Monodisperse nanoparticles based on recombinant HSA³⁸ and recombinant gelatin⁷ were successfully prepared. However, preparation of recombinant proteins is high cost and high technical requirements. On the contrary, soybean whey, an underutilized by-product of the soybean-processing industry and available in huge quantities in many countries, contains abundant KTI. In the last decade, expanded bed adsorption (EBA) has been proved to be a well-suited method for large-scale recovery of protein from a dilute watery system such as potato fruit water.³⁹ Recently, we have employed this method to recover KTI from soybean whey and have obtained desired KTI with high yield and purity (data to be published). In the present study, KTIP was prepared successfully by heating KTI at 80 °C in the presence of sodium sulfite. This treatment not only inactivated most trypsin inhibitor activity (TIA) of KTI (about 90.63%, Fig. 1C), but also resulting in the formation of KTI nanoparticles (about 85.92 nm, Table 1) with good monodispersity (PDI: 0.18, Table 1).

During the inactivation processing of KTI, a limited aggregation behavior was observed. On the basis of the experimental data (Fig. 1, 2 and Table 1) combined with the structural characteristic of KTI molecule (Fig. S1†), the aggregation process can be speculated as follows: the initiation step (1) two solvent-exposed disulfide bonds of KTI molecule were cleaved by SO₃²⁻, which was followed by exposure of the free SH of native KTI, causing the protein to become reactive; the propagation reaction (2) corresponds to the build-up of aggregates via –SH/–SS– exchange reactions between activated KTI molecules; the termination step (3) due to the steric hindrance effect, the aggregation was terminated although there may be free –SH. The reaction scheme accounts for the formation of aggregates in which the monomers are linearly linked, but the aggregates are not stiff rods and may even have a spherical shape (Fig. 1B and 4D). Hoffmann et al. reported that upon heating β-lactoglobulin at 65 °C, particles of a constant average size were formed and, although more native β-lactoglobulin was converted to aggregates during prolonged heating. The molecular mass distributions remained about the same, as predicted by the model of Roefs and de Kruif.⁴⁰ This model holds for β-lactoglobulin dissolved in water at close to neutral pH and heated at relatively low temperature (65 °C), and it gives a correct description of the decrease in concentration of native β-lactoglobulin and the increase in scattered intensity, as measured by in situ light scattering.⁴¹ Accordingly, the limited aggregation behavior via disulphide linkage of KTI in the present study could also be described by a mechanism similar to the same model.

Aggregates were only mediated by hydrophobic interaction, which was not observed from SDS-PAGE patterns in the present study (Fig. 2). This may be due to the importance of non-covalent interactions varied with temperature. Galani and Apenten reported that the contribution of non-covalent interactions to the overall aggregation mechanism became important only at higher temperatures (>90 °C).⁴² The use of lower temperatures (80 °C) could be a reason why the present study found SS-mediated aggregation, or could not find clear evidence for the formation of non-covalently associated aggregates.

The structure of the molten globule state (reversible when the environment changes to one in which the native state is more stable) is characterized by a partially folded conformation with retention of the secondary structural elements whereas the tertiary structure becomes much more fluid with consequent slight swelling of the protein and greater accessibility of the hydrophobic groups of the molecule.⁴³ The data of DLS, SAXS and CD (Table 1, Fig. 3 and S2†) in the present study suggested that KTIP probably bear some features of the molten globule states, indicating that KTIP may be composed of molten globule state monomers, which was cross-linked by intermolecular disulfide bonds as well as hydrophobically driven associations may occur within the aggregates.

The interfacial properties of KTIP showed it possessed a fair surface activity and a better surface stability as compared with that of KTI (Fig. 4A and B). The pattern of the interfacial architecture on the air–water interface formed by KTIP was observed by AFM (Fig. 4D), it is a nanoscale and globular conformation particle. The surface-active and nanoscale size make KTIP can act as a novel nano-delivery vehicle for hydrophobic bioactives.

Curcumin, a yellow pigment present in the spice turmeric (Curcuma longa), has been linked with antioxidant, anti-inflammatory, antiproliferative, anticancer, antidiabetic, antirheumatic, and antiviral effects, but its optimum potential is limited by its lack of solubility in aqueous solvents and poor oral bioavailability. Herein, we employed KTIP to improve solubility, stability, bioaccessibility and bioactivity of curcumin. Unlike free curcumin, it is readily dispersed in aqueous medium (Fig. 5A, inset), showing narrow size distribution (Fig. 5B). Moreover, it displayed enhanced stability in PBS by protecting encapsulated curcumin against degradation by environmental stress (e.g., light, oxygen, or heat) (Fig. 5D), as well as the bioaccessibility of curcumin in the nanocomplexes with KTIP was greatly enhanced (Fig. 6). Most importantly, nanoparticulate curcumin was comparatively more effective than free curcumin against different cancer cell lines under in vitro condition with time resulting in reduction of cell viability by inducing apoptosis (Fig. 7).

Curcumin binds to hydrophobic clusters of proteins mainly through hydrophobic interactions.^30,33 Generally, heating can result in structural unfolding and denaturation of proteins, thus increasing their surface hydrophobicity. In the present study, KTIP was generated during inactivation treatment of KTI by heating combined with sodium sulfite, its surface hydrophobicity was improved (Table 2), the enhanced hydrophobicity for KTIP was also evidenced by the UV-Vis spectrum and intrinsic fluorescence data (Fig. S4†). The improvement of the hydrophobicity would be favorable for the binding of curcumin to KTIP. Moreover, KTIP was composed of molten globule state monomers, which was much more fluid with consequent slight swelling of the protein and greater accessibility of the hydrophobic groups or cavities of the molecule.⁴³ This structural feature can facilitate the binding of curcumin to KTIP as well. These may be the reason why KTIP can greatly enhance the solubility and stability of curcumin in aqueous solution. Although further efforts are needed to evaluate the improvement of bioavailability in vivo, it is predicted that KTIP could be developed as a novel nano-delivery vehicle for hydrophobic bioactives and for use in functional foods and pharmaceutics.

Experimental section

Materials

Soybean Kunitz trypsin inhibitor, trypsin, N-ethylmaleimide (NEM), sodium sulfite, Nα-benzoyl-DL-arginine 4-nitroanilide hydrochloride (BAPNA), 8-anilino-1-naphthalenesulfonic acid ammonium salt for fluorescence (ANS), pepsin, pancreatin powder, bile extract and curcumin were purchased from Sigma (Shanghai, China). HepG2, MCF-7, HCT-116, A549, K562, and Du145 cell lines were purchased from the American Type Culture Collection (ATCC; Manassas, VA). All other chemicals used were of analytical grade.

Preparation and characterization of KTI aggregates

Preparation of KTI solutions and inactivation treatments. 10 mg mL⁻¹ KTI solutions were prepared by dispersing KTI lyophilized powder into 10 mM phosphate buffer (pH 7.0). Then KTI solutions were treated as follows: (1) unheated; (2) heated at 80 °C for 1 h; (3) addition of sodium sulfite into the KTI solution to 10 mM without heating; (4) addition of sodium sulfite into the KTI solution to 10 mM with heating at 80 °C for 1 h; (5) addition of sodium sulfite and NEM into the KTI solution to 10 mM, sequentially, without heating; (6) addition of sodium sulfite and NEM into the KTI solution to 10 mM, sequentially, with heating at 80 °C for 1 h. After various treatments, except (1) and (2), the processed samples by other treatments ((3)–(6)) were dialysed against Millipore water for 24 h in the refrigerator (4 °C). Then the dialysed samples were analyzed immediately or freeze-dried (Christ DELTA 1-24 LSC freeze-dryer, Germany) and stored at −20 °C.

Dynamic light scattering (DLS). The KTI after various treatment were diluted to 1 mg mL⁻¹ with Millipore water, and the pH was adjusted to 7.0; then the particle size (hydrodynamic radius, R_h), ζ-potential, and polydispersity index (PDI) were measured using a Nano ZS Zetasizer instrument (Malvern Instruments, Worcestershire, UK). All measurements were carried out at 25 °C, and the results are reported as averages of three readings.

Small-angle X-ray scattering (SAXS). SAXS experiments were carried out using a SAXSess camera (Anton-Paar, Graz, Austria) to collect information about the particle size and shape of the samples (10 mg mL⁻¹) as described in our previous work.¹⁹

SDS-PAGE. The KTI after various treatments was assessed by non-reducing or reducing SDS-PAGE. Aliquots (50 μL) protein solutions were mixed with the same volume of non-reducing or reducing sample buffer. 10 μg of proteins were loaded into each lane. SDS-PAGE was performed on a discontinuous buffered system according to the method of Laemmli⁴⁴ using 12.5% separating gel and 5% stacking gel.

Determination of inhibition activity. Trypsin inhibitor activity (TIA) of KTI with various treatments was measured by using 0.04% (w/v) BAPNA as the substrate, which was dissolved in 0.05 M, pH 8.2, Tris–HCl buffer (1% dimethyl sulfoxide, v/v; 0.02 M CaCl₂). Assay solution was zeroed, and then 2 mL of 0.01% (w/v) trypsin in 0.04 mM HCl solution was added to initiate the reaction. Trypsin inhibitor units (TIU) were calculated as the amount of inhibitor that reduced the absorbance per minute of the standard reaction by 0.01. For accuracy, the reaction was measured in the linear portion in the 40–60% inhibition range.⁴⁵

Circular dichroism (CD) spectroscopy. Far-UV and near-UV CD spectra of KTI and KTI aggregates were collected at 25 °C using a Jasco-600 spectrophotometer (Jasco Inc., Japan) with cuvettes having a 0.1 or 1 cm path length. All experiments were performed in Millipore water. For far-UV CD, the protein concentration was 0.2 mg mL⁻¹, and the spectra were registered from 190 to 260 nm at a data pitch of 1 nm. The spectra were baseline-corrected. The molar mean residue ellipticity [MWR, θ] was expressed in degrees cm² dmol⁻¹. From the CD spectra, fractional contents of the secondary structure (α-helix, β-sheet, β-turn, and random coil) of KTI and KTI aggregates were calculated according to the CDSSTR program in the CD Pro software. For near-UV CD, the protein concentration was 1.0 mg mL⁻¹. Scans were obtained in the range of 260–340 nm by taking points every 0.5 nm, with an integration time of 1 s.

Surface hydrophobicity. The protein surface hydrophobicity was determined by titration with ANS according to the method of Liu et al.⁴⁶ with modifications.²³ The aliquots (1 mL) of protein solutions (0.2 mg mL⁻¹) were placed in the cell of an F7000 fluorescence spectrophotometer (Hitachi Co., Japan), and then, aliquots (10 μL) of ANS (5 mM in 10 mM phosphate buffer, pH 7.0) were titrated to reach a final concentration of 50 μM. The molar coefficient (5000 M⁻¹ cm⁻¹ at 350 nm) was used to calculate ANS concentration. The relative fluorescence intensity (F) was measured at 390 (excitation; slit width, 5 nm) and 470 nm (emission; slit width, 5 nm). Data were elaborated using the Lineweaver–Burk equation: 1/F = 1/F_max + (K_d/L₀)(1/F_max), where L₀ is the fluorescent probe concentration (μM), F_max is the maximum fluorescence intensity (at saturating probe concentration), and K_d is the apparent dissociation constant of a supposedly monomolecular protein/ANS complex. F_max and K_d can be calculated by standard linear regression fitting procedures. The ratio F_max/K_d, corrected for protein content, represents the protein surface hydrophobicity index (PSH).

Transmission electron microscopy (TEM). TEM was used to observe the surface morphology of KTI aggregates and to further confirm particle diameter by DLS. A drop of diluted sample was deposited onto a carbon-coated copper grid, and excess of sample was removed after 5 min with a filter paper. Then, a droplet of phosphotungstic acid (1%, w/v) was put onto the grid and removed after 5 min. Observations were made with JEM-2100F transmission electron microscope operating at 200 kV (JEOL, Japan).

Dynamic surface properties

The dynamic surface tension and the dilatational rheological properties of KTI aggregates relative to KTI at the air–water interface were measured using an optical contact angle meter (OCA-20, DataPhysics Instruments GmbH, Germany) equipped with oscillating drop accessory (ODG-20), as described in our previous work.²⁷

Dynamic surface tension. A drop of sample solution (12 μL) was delivered and allowed to stand for 10 000 s to achieve adsorption at the interface. An image of the drop was continuously recorded by a CCD camera and digitalized. The surface tension (γ) was calculated through the shape analysis of a pendant drop according to the Young–Laplace equation analysis. The surface pressure is π = γ₀ − γ, where γ₀ is the surface tension of phosphate buffer (10 mM, pH 7.0) and γ is the time-dependent surface tension of the tested solutions. The average standard accuracy of the surface tension for at least three measurements with different drops was roughly 0.5 mN m⁻¹.

Dilatational rheological properties. To obtain surface dilatational parameters, sinusoidal interfacial compression and expansion were performed by decreasing and increasing the drop volume at 10% of deformation amplitude (ΔA/A) and 0.1 Hz of angular frequency (ω). The drop was subjected to repeated measurements of five sinusoidal oscillation cycles followed by a time corresponding to 50 cycles without any oscillation up to 10 000 s required to complete adsorption. Details of this experiment as described in our previous work.¹⁸ The surface viscoelastic parameters, i.e., surface dilatational modulus, E, its elastic, E_d, and viscous, E_v, components were derived from the change in surface tension (γ) (dilatational stress) resulting from a small change in surface area (dilatational strain). The surface dilatational modulus (E) is a complex quantity and composed of real and imaginary parts (E = E_d + iE_v). The real part of the dilatational modulus or storage component is the dilatational elasticity (E_d). The imaginary part of the dilatational modulus or loss component is the surface dilatational viscosity (E_v).

To observe the morphology of KTI aggregates or KTI at the interface, protein films at a pressure of 30 mN m⁻¹ are transferred at a speed of 5 mm min⁻¹ onto freshly cleaved mica plates, using the Langmuir–Blodgett technique. The structure of the transferred proteins was imaged using a MultiMode SPM atomic force microscopy (AFM) equipped with a Nanoscope IIIa Controller (Digital Instruments, Veeco, Santa Barbara, CA) and performed in tapping mode.

Since the size of KTI aggregates is nanoscale according to DLS and SAXS data and the structure of KTI aggregates is globular conformation according to TEM, KTI aggregates is termed as KTI nanoparticles (KTIP) in the following experiments.

Preparation and characterization of curcumin-loaded KTIP

Preparation of curcumin-loaded KTIP. To obtain curcumin-loaded KTIP, 0.1 mL stock solution of curcumin (4 mg mL⁻¹ in ethanol) was added into 2.9 mL of KTIP solutions (1, 2, 3, 4, and 5 mg mL⁻¹) in successive titrations with magnetic stirring. The mixtures were centrifuged at 10 000g, 25 °C for 20 min to pellet the unbound curcumin, and the supernatants containing curcumin nanocomplexes were preserved in a light-resistant container at 4 °C for determination. As contrasts, KTIP without curcumin and curcumin without KTIP in the same PBS solution with homologous concentration were also prepared. The particle size and size distributions of the nanocomplexes were measured using dynamic light scattering (DLS).

Particle morphology. TEM was used to observe the surface morphology of curcumin-loaded KTIP and to further confirm particle diameter by DLS. The procedure of TEM as mentioned above.

Determination of the entrapment efficiency (EE) and loading content (LC). The EE (%) of curcumin in the curcumin-loaded KTIP was estimated as the percentage of curcumin encapsulated in the proteins by the following equation: EE (%) = 100 − (amount of free curcumin (mg)/total amount of added curcumin (mg)) × 100, where the amount of free curcumin is determined from the precipitate obtained by centrifugation. The precipitate was extracted in 5 mL of ethanol with mild stirring for 5 min under magnetically stirred conditions and then centrifuged at 10 000g for 15 min at 25 °C to remove the protein aggregates. The supernatant was subjected to spectrophotometric analysis at 426 nm with a GENESYS 10S UV-Vis spectrophotometer (Thermo Scientific, USA), and the curcumin concentration was determined using an established standard curve of curcumin (y = 0.1159x + 0.1028, R² = 0.9958). The loading content (LC) of the samples was calculated with the following equations: LC (%) = mass of encapsulated curcumin/(total mass of encapsulated curcumin + KTIP).

Stability measurement. The free curcumin and the freshly prepared nanocomplex dispersions containing sodium azide (0.002%, w/v) were settled under room temperature (25 °C). At the specified time points, samples (200 μL) were taken out and added to 1800 μL of ethanol for quantitative analysis of curcumin by spectrophotometer, as described above. The results were represented by a retained ratio of curcumin, which was calculated as the percentage of the retained curcumin at a certain time point with respect to the initial value.

Sequential in vitro gastric and intestinal digestion. An in vitro model that stimulated sequential gastric and intestinal digestion was applied to assess the effect of digestion on the in vitro bioaccessibility of free curcumin and nanoparticulate curcumin according to the method described elsewhere³² with slight modifications. Briefly, 10 mL of freshly prepared samples were well mixed with 40 mL of 0.1 mol L⁻¹ HCl (pH = 1.5), and preincubated in a shaker (at 37 °C) at a rate of 100 rpm for 10 min. If necessary, the pH of the mixtures was adjusted to 1.5 using 1 mol L⁻¹ HCl. Subsequently, 10 mg of pepsin powder was added and well mixed to start the simulated gastric digestion (0–60 min). After 60 min, the pH of the pepsin-digests was immediately adjusted to 7.0 using 4 mol L⁻¹ NaOH. Then 250 mg of bile extract was added and well dispersed in the shaker for 10 min. Last, 20 mg of pancreatin powder was added to start the intestinal digestion (60–180 min). After 180 min, 500 μL of the final digest dispersion was also collected and centrifuged at 55 000g at 4 °C for 95 min (Micro Ultracentrifuge CS15ONX, Hitachi, Japan). The aqueous fraction was collected from the centrifuge tube and then passed through a filter with 0.22 μm pores (Millipore, Billerica, MA, USA) to ensure that the curcumin in the aqueous fraction were actually in KTIP. The amounts of curcumin in KTIP and the whole digests were extracted and determined according to the method described above. The whole process was kept in the dark in order to avoid light-induced degradation.

In vitro anti-proliferative activity on tumor cells assay of curcumin-loaded KTIP. The anti-proliferative effects of curcumin-loaded KTIP were analyzed by the MTT assay.⁴⁷ Briefly, HepG2, MCF-7, HCT-116, A549, K562, and Du145 cell lines were seeded at 5000 cells per well density in 96-well plates (Corning, NY, USA). After 24 h, the cells were treated with a medium containing DMSO-dissolved or different concentration (0, 5, 10, 15, 20, 25 and 30 μM) of curcumin-loaded KTIP. Other cells were untreated (negative controls, NC) or treated only with DMSO or KTIP at the concentrations as in the dispersions with encapsulated curcumin (positive controls, PC). Cells were incubated for 48 h for assessing the toxicity of samples. A standard MTT based colorimetric assay was used to determine cell viability. Relative cell viability was expressed as the absorbance normalized by that of the control cells treated with same amounts of DMSO and KTIP as in DMSO-dissolved and encapsulated curcumin samples. The mean and standard deviation from six-well replicates were calculated. The normalized cell viability was obtained after normalizing the viability of a treatment by the viability of NC and PC.

Cell viability (%) = [(A_treated − A_PC)/(A_NC − A_PC)] × 100

where A_treated, A_PC and A_NC are the absorbance of the wells with cells treated by curcumin, the positive and the negative control, respectively.

Statistical analysis

An analysis of variance (ANOVA) of the data was performed, and a least significant difference (LSD) with a confidence interval of 95% was used to compare the means.

Conclusions

In summary, soy Kunitz trypsin inhibitor nanoparticles (KTIP, about 85.92 nm) with good monodispersity (PDI: 0.18) were fabricated successfully during inactivation treatment by heating at 80 °C and pH 7.0 for 1 h in the presence of sodium sulfite. The formation of KTIP was probably mediated by sulfhydryl–disulfide interchange, as evidenced by the SDS-PAGE patterns with or without adding the thiol-blocking agent NEM. KTIP not only retained a small amount of trypsin inhibitor activity, but also possessed favorable surface activity. Curcumin-loaded KTIP prepared by desolvation are found to encapsulate curcumin with high efficiency and relatively high loading content. The co-assembly of KTIP and curcumin actually greatly enhanced the solubility, stability, bioaccessibility and in vitro anti-proliferative activity on tumor cells of curcumin in aqueous solution. These findings suggest that KTIP could be developed as a novel nano-delivery vehicle for hydrophobic bioactives and for use in functional foods and pharmaceutics.

Acknowledgements

This research was supported by grants from the Chinese National Natural Science Foundation (serial numbers: 31130042 and 31371744), and the Project of National Key Technology Research and Development Program for the National High Technology Research and Development Program of China (863 Program: 2013AA102208).

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Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra19886d

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