Enzymatic hydrolysis by Alcalase of red bean protein to develop corn starch/red bean protein hydrolysate complex for reduced glycemic response in vitro

Abstract

BACKGROUND

Legume protein hydrolysates have garnered significant attention because of their multifunctional properties and potential applications in agriculture, food, and health sectors. In this study, red bean protein (RP) was extracted and subjected to Alcalase hydrolysis to obtain red bean protein hydrolysate (RPH). Subsequently, the protein hydrolysate was incorporated into corn starch (CS) to prepare corn starch/red bean protein hydrolysate complex (CS‐RPH), aiming to control the sugar release during gastrointestinal digestion of native starch.

RESULTS

RPH showed higher ABTS free radical scavenging activity, whereas FRAP activity experienced a significant drop compared to crude protein. In addition, foaming ability of RPH was remarkably improved and the protein fractions were shown to have molecular weight in the range of 15–20 kDa, in reference to 50–70 kDa of crude counterparts, as illustrated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis. Furthermore, RPH exhibited little cytotoxicity at a dose below 2000 μg mL⁻¹, despite minimal anti‐inflammatory efficacy against pro‐inflammatory cytokines nitric oxide and interleukin‐6. For the development of CS‐RPH, 2% RPH was added to 10% gelatinized CS and further subjected to in vitro gastrointestinal digestion for evaluating the sugar release. The results showed that RPH reduced 4.82% sugar release of CS, highlighting its potential in health‐related application.

CONCLUSION

Alcalase hydrolysis successfully converted red bean protein into bioactive hydrolysates with improved functional properties. Incorporation of red bean protein hydrolysate into corn starch effectively reduced sugar release during simulated gastrointestinal digestion, demonstrating its potential as a functional ingredient for developing low‐glycemic foods and managing postprandial glucose response. © 2026 Society of Chemical Industry.

INTRODUCTION

Diabetes mellitus is a chronic metabolic condition marked by hyperglycemia and is becoming recognized as a significant public health problem globally. ¹ In recent decades, the incidence of diabetes mellitus has markedly increased in both industrialized and developing nations. In 2019, the global prevalence of diabetes was 9.3% (463 million individuals), and is projected to increase to 10.2% (578 million individuals) by 2030 and 10.9% (700 million individuals) by 2045. ² Approximately 2.3 billion individuals, both children and adults, are classified as overweight worldwide, with the financial burden of obesity surpassing US$700 billion annually, resulting in various metabolic disorders in humans.^{3, 4} A nutritious plant‐based diet is more sustainable and correlates with a reduced incidence of obesity, type 2 diabetes, cardiovascular disease and some malignancies. ⁵ Diabetic patients must follow a stringent diet, particularly restricting starchy meals because monosaccharides derived from starch during digestion are the primary contributors to elevated postprandial blood glucose levels. ⁶ Consequently, the study and the production of low glycemic index dietary products are crucial for facilitating glycemic management and are garnering heightened interest. Due to the widespread occurrence of illnesses associated with glycemic disorders, there is an urgent necessity to investigate and create dietary interventions to mitigate the adverse effects of starch.

A potential method involves employing protein as a physical barrier on the starch surface, which can inhibit amylase access and hence diminish starch digestibility. ⁷ The existence of starch–protein interactions contributes to the decreased digestion of starch and the resulting glycemic reactions.^{8, 9} Ultimately, protein can impede the digestion of starch‐laden meals by blocking starch‐hydrolyzing enzymes. ¹⁰ Numerous in vitro studies have shown that including bean protein into the diet might reduce blood sugar levels in individuals. These studies indicate that pea proteins and their hydrolysates are promising dietary components because of their health advantages, including hypoglycemic properties.^{11, 12} Red kidney bean protein has been shown to significantly reduce the rate and extent of starch digestion by forming a physical barrier and interacting with starch through hydrogen bonds and hydrophobic interactions, which modifies the starch structure and inhibits α‐amylase activity in a dose‐dependent manner. ¹³ Similarly, the protein matrix in legume cotyledon cells, such as those in navy beans, acts as a secondary barrier alongside the cell wall, restricting enzyme access to starch and thereby reducing its digestibility. ¹⁴ Furthermore, proteins and their hydrolysates can modulate starch digestibility by altering the microstructure and thermal stability of starch, which can be leveraged to design low‐glycemic foods. ¹⁵ The interaction between proteins and starch affects not only the digestibility, but also the technological and nutritional properties of food because proteins can decrease gelatinization and hydrolysis rates. ¹⁶ These findings highlight the potential of using protein‐based barriers to create functional foods with controlled starch digestibility, which can help manage postprandial glucose levels and reduce the risk of diet‐related chronic diseases.^{17, 18}

Among various bean commodities, adzuki beans or red bean (Phaseolus angularis) are a leguminous plant indigenous to East Asia, extensively utilized in culinary practices and traditional medicine. Red beans are regarded as a promising functional food source as a result of their high protein content (15–30% dry weight) and rich nutritional composition, including dietary fiber, vitamins, minerals and bioactive compounds such as flavonoids and polyphenols.^{19, 20} These nutritional and bioactive constituents contribute to antioxidant, anti‐inflammatory, cardioprotective and glycemic regulatory benefits. Red beans are a significant dietary source with elevated protein levels and exceptional nutritional attributes. ²⁰ They contain high quality protein and advantageous biological substances, such as flavonoids and polyphenols, which have antioxidant and anti‐inflammatory characteristics. ²⁰ Specifically, enzymatic hydrolysates of red bean protein can provide potential bioactives, whereas the peptides responsible for this activity remain inactive inside the original protein structure. ²¹ Moreover, red bean protein hydrolysate possesses the capacity to lower blood glucose levels via modulating carbohydrate metabolism. ²²

The present study aimed to extract protein hydrolysate from red beans and integrate it with corn starch to create a low glycemic index starch complex. Crude protein extracted from red beans was subjected to Alcalase hydrolysis and the protein hydrolysate was then assessed for its technical features, namely foaming capacity, and its biological activities, including antioxidant and anti‐inflammatory effects, prior to its utilization as a raw material for the manufacture of a corn starch/red bean protein hydrolysate complex. The powdered complex was further assessed for various physical properties, spectral analysis, thermal characteristics and in vitro gastrointestinal digestion.

MATERIALS AND METHODS

Materials and chemicals

Red beans or Adzuki beans (P. angularis) of organic grade were purchased from C'LaVie (Ho Chi Minh City, Vietnam) with uniform size characteristic red‐brown color and showed no sign of mold or damages. Alcalase 2.5 L (activity of 2.5 AU g⁻¹, optimal at 50–60 °C and pH 7–8) were supplied by Novozymes (Bagsvaerd, Denmark). Native pure corn starch (100% purity) was collected from Jade Leaf (Thailand). It had a fine structure and uniform color and showed no lumps, unusual odor, or signs of mold. The chemicals and biological reagents, including 2,2′‐azinobis‐(3‐ethylbenzothiazoline‐6‐sulfonic acid) (ABTS), 2,4,6‐tripyridyl‐s‐triazine (TPTZ), Trolox, Coomassie Brilliant Blue G‐250, bovine serum albumin (BSA), α‐amylase (from porcine pancreas), pepsin (from porcine gastric mucosa), trypsin (from porcine pancreas), pancreatic lipase (from porcine pancreas), bile salts, lipopolysaccharide, corticosterone acetate and Dulbecco's modified Eagle's medium, were purchased from Sigma‐Aldrich (Singapore). Fetal bovine serum was obtained from Gibco (Thermo Fisher Scientific, Walthem, MA, USA). Other chemicals were of analytical grade.

Preparation of protein fraction from red bean

The protein recovery process was adapted with modifications from the method described by Austin et al. ²³ Initially, the beans were dehulled, cleaned and ground into a fine powder, which was placed in a filter bag. The fat content was extracted using hexane in a Soxhlet system for 16 h, followed by drying to eliminate any residual hexane. The defatted powder was then mixed with water at a 1:5 (w/v) ratio, adjusted to pH 9.5 using 1 m NaOH and stirred at 100 rpm at 40 °C for 60 min to extract the protein. The mixture was centrifuged at 7280 × g for 15 min at 4 °C in a TG16 high speed centrifuge (Changsha Yingtai Instrument Co., Ltd, Changsha City, China) to remove solid residues. The supernatant was adjusted to pH 4.5 with 1 m HCl and centrifuged again under the same conditions. The resulting protein precipitate was reconstituted in water at a 1:5 ratio, adjusted to pH 6.5 with 1 m NaOH, then frozen, freeze‐dried at –60 °C using the Alpha 1‐2 LDplus lyophilizer (Martin Christ Gefriertrocknungsanlagen GmbH, Osterode am Harz, Germany) and ground into crude protein powder.

Preparation of red bean protein hydrolysate and its complex with corn starch

The procedure for preparing bean protein hydrolysate was modified from the process established by Yang et al. ²⁴ The crude red bean protein (RP), prepared as described in the section ‘Preparation of protein fraction from red bean’ was quantified and transferred to a centrifuge tube, then dissolved in a 48:1:1 ratio of water, Alcalase enzyme and protein. The mixture was vigorously vortexed and incubated at 40 °C for 30 min to facilitate hydrolysis. The enzyme was inactivated by heating the mixture to 80 °C for 5 min. Ultimately, it was chilled and centrifuged at 2850 × g for 10 min to isolate the residue, resulting in the extraction of the bean protein hydrolysate (RPH). Corn starch (CS) was further incorporated into protein hydrolysate at a concentration of 10%, after which the mixture was gelatinized at 90 °C. The combination was subjected to freeze‐drying, followed by fine grinding to produce corn starch/red bean protein hydrolysate complex (CS‐RPH).

Characterization of red bean and protein hydrolysate

Proximate analysis

Moisture, crude protein, total lipid and ash content were determined by oven drying to constant mass (AOAC 935.29), the micro‐Kjeldahl method (AOAC 950.48), Soxhlet extraction (AOAC 963.15) and dry ashing (AOAC 923.03), respectively. Total carbohydrate content was calculated by difference following AOAC protocols. ²⁵

Antioxidant activities: ABTS assay

The ABTS radical scavenging activity was determined according to the method described by Nguyen et al. ²⁶ Briefly, 0.15 mL of the diluted sample was transferred into a test tube, followed by the addition of 2.85 mL of diluted ABTS reagent (adjusted to an absorbance of 1.1 ± 0.02 at 734 nm). The mixture was thoroughly mixed and incubated in the dark for 30 min. After incubation, the absorbance was measured at 734 nm using methanol as the blank. The percentage inhibition of ABTS radicals was calculated using:

$$\text{ABTS inhibition}\left(\%\right)=\frac{A_{\mathit{\text{control}}}-A_{\mathit{\text{sample}}}}{A_{\mathit{\text{control}}}}\times 100$$ (1)

where A _control is the absorbance of the ABTS reagent without sample and A _sample is the absorbance in the presence of the sample. Antioxidant activity was quantified using a Trolox standard calibration curve and expressed as mg Trolox equivalent per liter (mg TE L^–1).

Antioxidant activities: ferric reducing antioxidant power (FRAP) assay

The FRAP assay was performed following the method described by Nguyen et al. ²⁶ Briefly, 0.15 mL of the diluted sample was mixed with 2.85 mL of freshly prepared FRAP reagent, consisting of 0.3 m acetate buffer (pH 3.6), 10 mm TPTZ prepared in 40 mm HCl and 20 mm FeCl₃ at a volumetric ratio of 100:1:1. The reaction mixture was incubated in the dark for 30 min, and the absorbance was subsequently measured at 593 nm. The results were expressed as mg Trolox equivalent per liter (mg TE L^–1).

Soluble protein content

The Bradford test, with a reagent consisting of 100 mg of Coomassie Brilliant Blue G 250, 50 mL of 95% ethanol and 100 mL of concentrated phosphoric acid, was employed to quantify the total soluble protein concentration. ²⁷ Following the measurement of absorbance at 595 nm, the protein concentration was determined using a BSA standard curve and represented in milligrams of BSA per liter (mg BSA L^–1).

N‐amine content and degree of hydrolysis

The degree of hydrolysis was assessed by titration following the method described by Tong et al. ²⁸ using a mixed indicator solution, comprising five volumes of 0.04% bromothymol blue and four volumes of 0.5% phenolphthalein (in alcohol). Briefly, aliquots (5 mL) were titrated with a 0.05 n NaOH solution until a light blue color appeared. Subsequently, 5 mL of neutral formalin was introduced and allowed to react for 5 min. The sample was further titrated with 0.05 n NaOH until the color changed from yellow to purple. The degree of hydrolysis of the protein solution was calculated based on the percentage difference in free N‐amine content of the protein solution before and after hydrolysis.

Fluorescence emission spectra

The protein's tertiary structure was examined utilizing fluorescence emission spectra obtained with a FluoroMax Plus fluorescence spectrophotometer (Horiba, Kyoto, Japan). The sample was diluted in a phosphate buffer solution (10 mm, pH 7.0) to a concentration of 0.05 mg mL⁻¹. The excitation wavelength was established at 280 nm, with excitation and emission slit widths of 5 nm. The emission wavelength range was 300 to 450 nm, with a scanning speed of 10 nm s⁻¹.

Zeta potential

The zeta potential of proteins was determined using the SZ‐100 particle size and zeta potential analyzer (Horiba), using water as solvent at 25 °C and a 2.8‐V electrode.

Foaming ability

The foaming capacity of protein and protein hydrolysate was assessed utilizing the technique of Yang et al., ²⁹ with some modifications. Initially, protein aliquots (10 mL) were dispensed into glass cylinders, followed by adding 30 mL of 0.05 m Tris‐HCl buffer. The mixture was well agitated to achieve homogeneity in a high‐speed homogenizer at 16 000 rpm for 2 min at 4 °C. The foaming capacity was determined by comparing the increased foam volume post‐homogenization to the original volume of the protein solution.

Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS‐PAGE)

SDS‐PAGE was conducted to examine the protein content of hydrolysate samples with a 10% SDS‐PAGE gel. Before electrophoresis, the samples were denatured by combining them with the 3X loading buffer in a 4:1 ratio, incubating at 100 °C for 3 min and chilling for 2 min, and repeating this denaturation procedure three times. A 20‐μL aliquot of the denatured protein solution was applied to a 4% stacking gel (comprising 30% acrylamide, Tris‐HCl buffer at pH 6.8, water, 10% SDS, 10% APS and TEMED) and subjected to separation at 80 V. The proteins were further separated in a 10% resolving gel, consisting of 30% acrylamide, Tris‐HCl buffer at pH 8.8, water, 10% SDS, 10% ammonium persulfate (APS) and N,N,N′,N′‐tetramethylethylenediamine (TEMED, with a current of 120 V. Subsequent to electrophoresis, the gel was subjected to staining with 0.23% Coomassie Blue R‐250 for 30 min and subsequently destained using a solution of 10% methanol and 10% acetic acid until clarity was achieved. The gel was photographed with the BioDoc‐It UVP imaging equipment (Analytik Jena AG, Jena. Germany). Protein bands showed as blue‐hued bands and were analyzed in comparison with protein standards in the range 15–250 kDa.

Cytotoxicity and anti‐inflammatory activity

The cytotoxicity and anti‐inflammatory properties of the extracts were assessed using the methodology outlined by Shan et al. ³⁰ For cell culture, RAW 264.7 cells (ATCC, TIB‐71) were grown in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum and 1% penicillin‐streptomycin, and maintained at 37 °C in a humidified incubator with 5% CO₂. The cytotoxicity of the extracts in serum‐free media was evaluated with the EZ‐Cytox test kit (Daeil Lab Service Co Ltd, Seoul, Korea) in which RAW 264.7 cells were inoculated at a density of 1 × 10⁵ per well in a 96‐well plate. Following 24 h of incubation, cells were subjected to treatment with different doses of the extracts. Thereafter, 10 μL of EZ‐Cytox was included and the mixture was incubated for an additional 2 h. Absorbance was quantified at 450 nm and cell viability was determined. The anti‐inflammatory activity was assessed by evaluating the inhibitory effect on pro‐inflammatory cytokines nitric oxide (NO) and interleukin‐6 (IL‐6). Cells were plated at a density of 1 × 10⁵ per well in 96‐well plates and incubated for 12 h, thereafter stimulated with 50 μL of 10 μg mL⁻¹ lipopolysaccharide for 4 h. Subsequently, 100 μL of the extracts at varying concentrations were introduced into the wells and incubated for 24 h. Corticosterone acetate (10 μg mL⁻¹ in dimethyl sulfoxide) served as the positive control.

Characterization of CS‐RPH

Moisture content and color attributes

Moisture content was determined as described in the section on ‘Proximate analysis’. The color characteristics in the CIE Lab* color space were measured using a Minolta CR‐400 portable colorimeter (Konica Minolta, Tokyo, Japan), and chroma (C*) and hue angle (h°) were calculated using:

$$C^{*}=\sqrt{a^{*2}+b^{*2}}$$ (2)

$$h^{°}=\mathit{\text{atan}}\frac{b^{*}}{a^{*}}$$ (3)

Solubility and wettability

The solubility and wettability was assessed according to the methodology outlined by Cano‐Chauca et al. ³¹ with some modifications. For the determination of solubility, 0.2 g of powder was precisely measured and dissolved in 10 mL of distilled water by agitation using a ZX3 vortex mixer (Velp, Usmate, Italy) at 200 rpm for 15 s to guarantee homogeneous dispersion. The sample underwent centrifugation at 2850 × g for 10 min and the supernatant was dried at 105 °C until a constant weight. For wettability, 0.2 g of powder was gradually sprinkled to 200 mL of distilled water in a 250‐mL beaker, permitting the powder to absorb without agitation. The time required for the powder particles to settle and fully dissolve was recorded as wettability.

Scanning electron microscopy (SEM)‐energy‐dispersive X‐ray spectroscopy (EDX)

The surface morphology was analyzed via SEM using a FESEM S4800 (Hitachi, Tokyo, Japan) at a scanning voltage of 10 kV. EDX was concurrently conducted using the EDX H‐7593 (Horiba) to examine the surface elemental composition.

Fourier transform infrared (FT‐IR) and X‐ray diffraction (XRD) spectra

Functional groups were examined by FT‐IR spectroscopy utilizing a Frontier NIR MIR system (Perkin Elmer, Shelton, CT, USA). The crystal structure was examined using XRD utilizing EMPYREAN (PANalytical, Malvern, UK) with CuKα radiation across a 2θ range of 5‐80° at a resolution of 0.02°.

Thermal properties

Thermal characteristics were assessed using the thermogravimetric analysis (TGA)–differential scanning calorimetry (DSC)–thermogravimetric analysis (DTG) technique using the LabSys Evo TG–DSC 1600 system (LabSys, East Lansing, MI, USA), within a temperature range of 30–220 °C at a heating rate of 10 °C min⁻¹.

Evaluation of sugar release during in vitro gastrointestinal digestion of corn starch/red bean protein hydrolysate complex

Digestion procedure

The simulated digestion of native corn starch and corn starch/red bean protein hydrolysate complex was performed according to the method described in the previous study with some modifications. ³² First, 0.2 g of sample was prepared in a test tube and 2 mL of distilled water was added, and then the sample was heated at 100 °C for 5 min to simulate the processing conditions for starch‐based food products. The mixture was mixed consecutively with 5 mL of simulated salivary fluid (in mg mL⁻¹: 0.03 NaNO₂, 0.02 K₂HPO₄, 0.5 NaCl, 4.2 NaHCO₃ and 0.5 α‐amylase), 20 mL of simulated gastric fluid (mg mL⁻¹: K₂HPO₄ – 2.2, NaCl – 6.2, CaCl₂.2H₂O – 6.3, NaHCO₃ – 1.2, and pepsin – 0.5) and 20 mL of simulated intestinal fluid (in mg mL⁻¹: 0.5 KCl, 0.1 KH₂PO₄, 3.57 NaHCO₃, 0.56 NaCl, 0.22 MgCl₂.6H₂O, 0.08 CaCl₂.2H₂O, 10 bile salt, 10 trypsin and 1.0 lipase). The samples were incubated at room temperature using the CRT‐350 rotator (Jeiotech, Daejeon, Korea) at 70 rpm to simulate the churning and mixing of food during the human digestion with the incubation time of 5 min for oral phase, 30 min for gastric phase and 120 min for intestinal phase.

Preparation of analytical aliquots

After incubation, digestive fluids (5 mL) were centrifuged at 5590 × g for 5 min. Then, 2 mL of the supernatant was transferred to a test tube with 0.5 mL of 10% trichloroacetic acid and incubated for 5 min before 0.5 mL of 10% NaOH solution and 2 mL of distilled water were added to neutralize. The sample was further centrifuged at the same conditions described above and the aliquots were analyzed for reducing sugar content by the 3,5‐dinitrosalicylic acid (DNS) method.

Determination of reducing sugar content

Reducing sugar content was determined by the DNS method. ³² DNS reagent was prepared by dissolving 1 g of DNS and 30 g of Rochelle salt in 100 mL of 0.1 n NaOH at 50 °C. To determine the reducing sugar content, 1 mL of DNS reagent was pipetted into a test tube containing 2 mL of sample and boiled at 95 °C for 5 min. After cooling, the absorbance of the resulting solution was measured at 540 nm.

Statistical analysis

All statistical analyses, including the normality test (Shapiro‐Wilk's test) and the homoscedasticity of variances test (Levene's test), were conducted at a 5% significance level using R, version 4.4.2 (R Foundation, Vienna, Austria). One‐way analysis of variance and post‐hoc tests, including a t‐test for comparing the quality attributes of samples described in the sections on ‘Preparation of protein fraction from red bean’ (RP and RPH) and ‘Preparation of red bean protein hydrolysate and its complex with corn starch’ (CS and CS‐RPH) and Tukey's test for samples described in the section on ‘Evaluation of sugar release during in vitro gastrointestinal digestion of corn starch/red bean protein hydrolysate complex’ (digestive phases), were also performed. All charts were drawn using OriginPro 2025 (OriginLab Corp., Northampton, MA, USA).

RESULTS AND DISCUSSION

Proximate analysis of red bean

The proximate analysis of red beans is displayed in Table 1, indicating a comparatively low moisture content of 9.56%, which corresponds to moderate water activity. This not only extends the shelf life of unprocessed red beans, but also inhibits the proliferation of spoilage bacteria, hence preserving product quality during storage. The high total protein content (25.85%) highlights the nutritional potential of red beans as a valuable plant‐based protein source for food applications. The substantial carbohydrate content (50.06%) indicates that red beans serve as a vital energy source, essential for sustaining metabolic activity. Despite being abundant in protein and carbohydrates, red beans exhibited a relatively low ash content (2.88%), reflecting their total mineral residue content. Nonetheless, with a moderate lipid content of 11.65%, this component can still augment the overall fat content in the diet, facilitating the metabolism and absorption of fat‐soluble vitamins. The nutritional profile of red beans indicates their potential utilization in food and nutrition, particularly in protein‐ and energy‐dense formulations.

Table 1.

Proximate analysis of red bean

Attribute	Value
Moisture (%)	9.56 ± 0.22
Protein (%)	25.85 ± 0.07
Lipid (%)	11.65 ± 0.80
Ash (%)	2.88 ± 0.17
Carbohydrate (%)	50.06 ± 0.85

Characterization of red bean protein hydrolysate

Antioxidant activities

Table 2 illustrates the impact of Alcalase hydrolysis on the antioxidant activity of red bean protein. The findings indicated that the ABTS free radical scavenging activity of red bean protein greatly enhanced post‐hydrolysis (521.79 mg TE L^–1) compared to crude protein (227.50 mg TE L^–1), corroborating prior research. ²¹ The observed differences in ABTS activity may originate from variations in protein composition and surface hydrophobicity of the hydrolysate. This was reported in the study of Rizzello et al., ³³ which showed that the biological activity of peptides is not only governed by amino acid composition and sequence, but also by molecular weight, with smaller peptides generally exhibiting higher antioxidant activity. Differences in genotype and cultivation conditions may influence antioxidant activity. Samaei et al. ³⁴ also suggest that this increase may be attributed to the heightened sensitivity of the ABTS technique, which is considered to selectively interact with hydroxylated aromatic molecules within the peptide sequence.

Table 2.

Some selected physical properties of red bean protein before (RP) and after (RPH) hydrolysis by Alcalase enzyme

Attribute	RP	RPH
ABTS (mg TE L–1)	227.50 ± 3.88 a	521.79 ± 35.20 b
FRAP (mg TE L–1)	61.92 ± 0.15 a	54.90 ± 1.00 b
Foaming ability (mL)	44.3 ± 0.6 a	60.3 ± 0.6 b
Protein (mg BSA L–1)	207.17 ± 3.49 a	115.91 ± 3.91 b
N‐amine (mg L–1)	136.30 ± 3.27 a	203.30 ± 10.89 a
Degree of hydrolysis (%)	–	49.10 ± 4.42

Note: Data are presented as the mean ± SD of three replicates. Data in the same row with different letters indicate statistical differences (P < 0.05).

By contrast, the FRAP test indicated lower results for the protein hydrolysate (54.90 mg TE L^–1) compared to the crude protein (61.92 mg TE L^–1). The disparity can be elucidated by the specificity of the peptidases and the structural characteristics of the crude protein, which can markedly affect the properties of the protein hydrolysate. ³⁵ Moreover, the higher FRAP value of the crude protein indicates that molecular weight may not be the primary factor influencing iron‐reducing capacity. The reduction in surface hydrophobicity of the hydrolysate may also lead to a diminished antioxidant activity as measured by FRAP. ³⁵ This outcome aligns with the research conducted by Karimi et al., ³⁶ which assessed the biological characteristics of protein hydrolysates derived from red beans and mung beans utilizing Alcalase and Flavourzyme enzymes. This study demonstrated that Alcalase hydrolysates exhibited superior ABTS free radical scavenging activity compared to Flavourzyme and crude protein, although had diminished ferric reducing capacity, indicating that antioxidant pathways may be variably influenced by the hydrolysis process that is employed.

Foaming ability

Foaming ability is an important functional property for food applications, particularly in products such as bakery, whipped and aerated systems, where protein contributes to air incorporation and foam stabilization. The enzymatic processing of proteins, such as corn gluten and soy protein, has been shown to enhance their functional properties, including foaming and emulsion stability. ³⁷ Table 2 illustrates the changes in the foaming capacity of red bean protein as a result of hydrolysis. The findings indicate that red bean protein hydrolysate exhibited a markedly superior foaming capacity compared to crude protein, with recorded values of 60.3 mL and 44.3 mL, respectively. Plant‐derived proteins often exhibit restricted foaming capabilities as a result of their dense tertiary structure and poor solubility. Hydrolysis appears to have compromised the intricate tertiary structure of crude proteins and diminished their molecular weight. These modifications may enhance the transport and adsorption of proteins at the air–water interface, thereby increasing foaming characteristics. ³⁴ This outcome aligns with the findings of the study by Wouters et al., ³⁸ which indicated that the hydrolysis of plant proteins improves foaming capacity by decreasing surface tension and increasing the flexibility of protein molecules. This alteration may lead to increased exposure of hydrophobic residues, thereby strengthening foam stability and improving the foaming efficacy of the protein hydrolysate.

Zeta potential

The zeta potential of red bean protein, measured before and after hydrolysis with Alcalase enzyme, indicated no significant change (P > 0.05) in these values in the two samples, RP (crude protein) and RPH (hydrolyzed protein), which were –0.1 mV and –0.3 mV, respectively. This suggests that the hydrolysis process did not substantially alter the surface charge of the protein and also reveals the inadequate stability of both colloidal systems. Bhattacharjee ³⁹ stated that to provide physical stability and avert aggregation of droplets in the dispersion, the zeta potential must attain a minimum of ±30 mV to generate adequate electrostatic repulsion. The low zeta potential values of both RP and RPH samples suggest that red bean protein, in either crude or hydrolyzed form, is prone to aggregation and fails to establish a stable colloidal system. Recent investigations have shown that the zeta potential of hydrolyzed proteins can be influenced by sample processing variables and medium makeup.^{40, 41}

Soluble protein content and degree of hydrolysis

Table 2 presents the findings on the changes in soluble protein concentration and hydrolysis degree of red bean protein samples before and after hydrolysis. The results indicated a considerable disparity (P < 0.05) between the two samples, with the soluble protein level of RPH (115.91 mg BSA L^–1) being much lower than that of RP (207.17 mg BSA L^–1). This low solubility of the RPH may be attributed to the incomplete hydrolysis of some proteins, which were eliminated during centrifugation, leading to a notable drop in the quantity of soluble protein in the sample. ³⁶ Moreover, the protein hydrolysis may yield water‐insoluble peptides, hence diminishing the overall quantity of soluble protein quantified in the sample. In terms of degree of hydrolysis, results indicated that Alcalase exhibited significant hydrolysis capability, with a variation in free N‐amine concentration of 49.1%. The extent of hydrolysis significantly influenced the functional characteristics and biological activities of hydrolyzed proteins, aligning with prior research on black bean protein hydrolysates. ⁴² Liu et al. ⁴³ assessed the hydrolysis degree of mung bean protein using several commercial enzymes and determined that Alcalase exhibited the best hydrolysis efficiency. This outcome reinforces the efficacy of Alcalase as a hydrolytic enzyme with respect to enhancing the functional characteristics of plant proteins.

SDS‐PAGE

SDS‐PAGE of red bean protein before and after hydrolysis using Alcalase enzyme is depicted in Figure 1. It can be seen that the RP sample exhibited a protein spectrum in the range 15–150 kDa, with predominant protein bands at around 40, 50 and 60 kDa. These proteins were classified as globulins, the primary storage protein in red beans. ⁴⁴ Following hydrolysis with Alcalase enzyme, the resulting RPH sample exhibited protein fragments with molecular weights in the range 15–20 kDa. This indicates that Alcalase possesses significant hydrolytic capacity, fragmenting proteins into smaller peptides, in alignment with the findings regarding degree of hydrolysis presented in the previous section. This finding is consistent with the study conducted by Zheng et al., ⁴² in which the enzyme Alcalase was utilized to hydrolyze black bean protein, yielding protein fractions with molecular weights between 10 and 25 kDa. The transition to reduced molecular weight protein bands indicates that the hydrolysis generated peptides with enhanced nutritional and biological functioning potential.

Figure 1.

SDS–PAGE gel electrophoresis of red bean protein before (RP) and after (RPH) hydrolysis by Alcalase enzyme. M is standard protein with molecular weight in the range 15–250 kDa.

Tertiary structure of protein

Figure 2 displays the fluorescence emission spectra of red bean protein before and after hydrolysis with Alcalase enzyme. This measurement indicates the alteration in protein conformation based on the polarity of aromatic amino acid residues, particularly tryptophan. The findings indicated that the RP sample exhibited a maximum emission wavelength (λ _max) at 379 nm in phosphate buffer, which is near the λ _max of tryptophan residues in water at 348 nm. ⁴⁵ Simultaneously, the RPH sample exhibited a λ _max shift to around 380–390 nm comparied to RP. This change signifies that the protein has experienced considerable denaturation via hydrolysis, enhancing the accessibility of tryptophan residues to a more polar medium. ⁴⁶ Regarding fluorescence intensity, RP exhibited a value of 1 800 000 units, but RPH attained only 300 000 units. The reduction can be attributed to the observation that proteins with stable tertiary structures typically exhibit high fluorescence intensity, whereas smaller and more flexible peptides are prone to collisions with one another and the solvent, resulting in de‐excitation via non‐radiative pathways, thereby diminishing emission intensity. ⁴⁵ Gómez et al. ⁴⁵ also observed a trend when assessing the structure and antioxidant activity of black‐eyed pea proteins before and during hydrolysis under varying pH settings and hydrolysis degrees. The findings demonstrated that hydrolysis decreased the fluorescence intensity of the proteins and induced a change in the maximum emission wavelength, aligning with the results acquired in the present study.

Figure 2.

Fluorescence emission spectra of red bean protein before (RP) and after (RPH) hydrolysis by Alcalase enzyme.

Cytotoxicity and anti‐inflammatory activity

Figure 3 displays the outcomes of the toxicity investigation of red bean protein before and following hydrolysis with Alcalase enzyme on RAW 264.7 cells at various treatment doses. The findings indicated that the cell survival rate consistently exceeded 70% in both samples, even at the maximum concentration of 2000 μg mL⁻¹ compared to the control sample. This indicates that red bean protein and its hydrolysate do not induce cytotoxicity at the evaluated amounts. Diao et al. ⁴⁷ demonstrated that green bean protein hydrolyzed by several enzymes had considerably increased cytotoxicity at concentrations over 400 μg mL⁻¹ compared to lower values. In this work, red bean protein hydrolysate did not substantially influence the viability of RAW 264.7 cells, even at elevated doses. With respect to anti‐inflammatory activity, both samples, however, showed no anti‐inflammatory activity against pro‐inflammatory cytokines NO and IL‐6. Various studies have shown that peptides derived from legume proteins, such as those from jack bean, chickpea and yellow field pea, can effectively modulate inflammatory pathways. The anti‐inflammatory potential of these hydrolysates is often attributed to their bioactive peptides, which can interact with and modulate various cellular signaling pathways involved in inflammation.^{48, 49} This difference may be a result of the protein composition of red beans as well as the processing conditions of Alcalase enzyme, which converted the native protein into smaller fractions with no anti‐inflammatory activity.

Figure 3.

(A) Cytotoxicity on RAW 264.7 cells and (B) anti‐inflammatory activity against IL‐6 and NO cytokines at different doses red bean protein before (RP) and after (RPH) hydrolysis by Alcalase enzyme. Different lowercase letters for each cytokine indicate the significant differences between samples (P < 0.05).

Characterization of CS‐RPH

Physical properties

The changes in the physical characteristics of corn starch as a result of the effect of hydrolyzed red bean protein are shown in Table 3. The analytical data indicated substantial variations in moisture content and solubility between the CS and CS‐RPH samples, although wettability remained relatively unchanged. The CS‐RPH sample exhibited a reduced moisture content of 2.77% compared to CS at 3.95%, suggesting that hydrolyzed red bean protein may encapsulate the starch granules, creating a protective barrier to mitigate moisture absorption. ⁵⁰ Simultaneously, CS‐RPH exhibited a much greater solubility (5.50%) compared to CS (2.65%), most likely attributable to the breakdown of the crystalline structure of starch and the augmentation of polar groups resulting from interactions with hydrolyzed protein, which enhanced water dispersibility. ²¹ The wettability of corn starch exhibited no significant difference between the two samples, with values being in the range 94.10–98.07 s. This indicates that the hydrolyzed protein layer encasing the starch did not markedly alter the surface structure or surface polarity of the starch granules, leading to an unchanged initial water absorption rate.

Table 3.

Some selected physical properties of corn starch (CS) and corn starch/red bean protein hydrolysate complex (CS‐RPH)

	CS	CS‐RPH
Moisture (%)	3.95 ± 0.17 a	2.77 ± 0.09 b
Solubility (%)	2.65 ± 0.07 a	5.50 ± 0.57 a
Wettability (s)	94.10 ± 3.74 a	98.07 ± 3.02 a
L*	95.98 ± 0.88 a	96.04 ± 1.01 a
a*	–0.59 ± 0.05 a	‐0.03 ± 0.02 b
b*	1.16 ± 0.07 a	2.61 ± 0.22 b
C*	1.28 ± 0.10 a	2.73 ± 0.07 b
h°	118.25 ± 5.49 a	89.83 ± 2.12 b

Note: Data are presented as the mean ± SD of three replicates. Data in the same row with different letters indicate statistical differences (P < 0.05).

Regarding color attributes, the L* brightness values of the two samples exhibited no significant difference, showing that both maintained high brightness within the range of 95.98–96.04. Nonetheless, the a* index of CS (–0.59) exhibited a tendency towards blue, whereas CS‐RPH (–0.03) approached neutrality, signifying a reduction in blue following complexation. The b* index of CS–RPH (2.61) was markedly elevated compared to that of CS (1.16), signifying an augmentation in yellow. The color saturation (C*) of CS–RPH (2.73) exceeded that of CS (1.28), signifying that the color of CS‐RPH darkened. The hue angle value (h°) of CS (118.25°) exceeded that of CS‐RPH (89.83°), suggesting that the CS sample inclined towards yellow–green, whereas CS‐RPH gravitated towards yellow–orange (Figure 4 and Table 3). This alteration may originate from the interaction between corn starch and hydrolyzed red bean protein, influencing the optical characteristics of the starch system. This may result from the existence of natural color components in hydrolyzed red bean protein or alterations in surface structure that influence light reflection.

Figure 4.

(A) Corn starch and (B) corn starch/red bean protein hydrolysate complex.

SEM‐EDX

The SEM and EDX analyses of starch samples influenced by hydrolyzed red bean protein are depicted in Figure 5(A). In the CS sample, the starch granules have a distinctive morphology with smooth surfaces and sharp edges, indicative of the unaltered structure of corn starch. Conversely, the CS‐RPH sample demonstrates that the surface of starch granules becomes more textured, exhibiting fissures and intergranular connections. This alteration may result from the breakdown of red bean protein, yielding peptides that interact with the starch surface, thus modifying the structure and shape of starch granules. Prior research indicated that incorporating protein into starch results in a structure characterized by bigger and denser holes after freeze‐drying, concurrently rendering the starch structure more tenuous, hence compromising its integrity. ⁵¹ The discrepancy in results may stem from the meticulous grinding of the samples conducted post‐freeze‐drying, which compromised the integrity of the sample structure. Furthermore, the energy dispersive X‐ray spectra of the two samples revealed no elemental presence other than carbon and oxygen, indicating that the primary constituent composition consisted of carbon (C) and oxygen (O) in an approximately 1:1 ratio for both samples. The C:O ratio indicates that the primary constituents of the two samples are polysaccharides including starch.

Figure 5.

(A) SEM‐EDX, (B) FTIR, (C) XRD spectra and (D) thermal degradation of corn starch (CS) and corn starch/red bean protein hydrolysate complex (CS‐RPH).

FTIR

FTIR spectra were conducted to assess the functional group composition in the hydrolyzed corn starch/red bean protein complex and the control starch sample. Figure 5(B) illustrates the occurrence of distinct peaks within the wavelength range of 3600–3100 cm⁻¹, indicative of intermolecular hydrogen‐bonded hydroxyl groups. ⁵² Moreover, pronounced peaks at wavelengths of 2931 and 1638 cm⁻¹ correspond to the symmetric stretching vibrations of CH₂ groups in glucose units and the stretching vibrations of C=O groups in carbohydrate moieties.^{52, 53} The signal detected at the wavelength of 1407 cm⁻¹ verifies the existence of bending CH₂ functional groups and elongated C–O–C functional groups. ⁵⁴ The peak at 1155 cm⁻¹ is indicative of starch molecules capable of interacting with other components via hydrogen bonding, ⁵⁵ whereas the 1022 cm⁻¹ wavelength pertains to the short‐range structure and hydrated crystals inside starch. ⁵⁶ The signal seen at a wavelength of 930 cm⁻¹ is ascribed to the vibrational motion of the carbon framework of the α‐1,4 glycosidic bond (C–O–C). ⁵⁴ The pronounced peaks at 854 cm⁻¹ and 529 cm⁻¹ correspond to the deformation of the CH₂ functional group and the pyranose ring's skeletal structure. ⁵⁴ The intensity disparity between the two samples indicates that the CS‐RPH sample exhibits lower intensity than the CS sample at the wavelengths of 3600‐3100, 2931, 1407, 1155, 930 and 854 cm⁻¹. The incorporation of RPH improved the short‐range ordered structure of starch, most likely attributable to the minimal steric hindrance of the small molecule peptides and amino acids in RPH, which promotes hydrogen bond formation with starch hydroxyl groups during gelatinization, resulting in the reorganization of starch chains. ⁵⁷

XRD

Figure 5(C) displays the XRD spectra of the CS‐RPH and CS complexes. CS exhibits diffraction peaks at 17°, 20°, 22° and 34°, with a pronounced peak at 22°, aligning with the characteristic A‐type crystal structure of corn starch. CS‐RPH has analogous diffraction peaks at 17°, 20°, 22° and 34°, but with diminished intensity, with new diffraction peaks at 39°, 43° and 48°. The results indicate 2θ peaks at 17°, 20°, 22° and 34°, corresponding to interplanar lengths of 5.23, 4.48, 3.97 and 2.65 Å, respectively. The A and V type crystal structures exhibit pronounced diffraction peaks at 15° and 22°, categorizing them as A type crystal structures, whereas the diffraction peak at 20° corresponds to V type crystal structures composed of endogenous lipids and starch, analogous to the findings of Wang et al. ⁵³ regarding corn starch samples. The diffraction peaks at 39° and 43°, corresponding to interplanar distances of 2.32 Å and 2.07 Å, respectively, signify the amorphous structure of the samples. The diffraction peak at 48°, exhibiting modest intensity, indicates a crystalline structure with an interplanar distance of 1.88 Å. The reduction in diffraction strength at around 2θ of 20° indicates that complexation may enhance the amorphous phase of the material.

TGA‐DSC‐DTG

Thermal analysis was carried out to evaluate the thermal stability and thermal evolution of CS and CS‐RPH samples over a specified temperature range. The results of thermal decomposition of CS and CS‐RPH are presented in Figure 5(D) and Table 4. The TGA curve shows that the thermal decomposition of the CS sample was divided into two main stages. The first stage was observed in the temperature range of 31.21–109.79 °C with a mass loss of 6.45%, whereas the second stage occurred from 289.4 °C and ended at 346.4 °C with a mass loss of 64.64%. The DTG curve also confirmed that the peak temperatures of the two stages were at 97.43 °C and 329.80 °C, respectively. Similar results were observed in the CS‐RPH sample with two main stages of thermal decomposition. The first stage started at 40.2 °C and ended at 108.94 °C with a peak temperature of 96.89 °C, and with a mass loss of 5.93%. The second stage occurred in the temperature range of 347.5–382.5 °C characterized by a mass loss of 53.59%, with a peak temperature confirmed at 376.4 °C. The mass loss at these stages was a result of the loss of water from the molecules in the first stage and the decomposition of hydrolyzed red bean protein molecules in the second stage, respectively. ⁵⁸ The CS and CS‐RPH samples suffered slight weight loss in the temperature range of 30–110 °C indicating the disappearance of water. Thermogravimetric analysis revealed a distinct phase of rapid mass loss occurring between 289 °C and 382 °C, indicating a clear evaporation behavior. A gradual mass loss began, whereas the thermal decomposition of the complexes accelerated, leading to a rapid weight loss. This phenomenon may be a result of the depolymerization of starch macromolecules to glucose and a series of gaseous products of lower molecular weight. Furthermore, as the temperature increased, glycosylation of glucose and decomposition of some polyhydroxy groups also contributed to the weight loss. ⁵³

Table 4.

Thermal degradation of corn starch (CS) and corn starch/red bean protein hydrolysate complex (CS‐RPH)

Sample	Stage	Temperature (°C)			Δm (%)
		Onset	Peak	End
CS	Stage 1	31.21	97.43	109.79	–6.45
	Stage 2	289.40	329.80	346.40	–64.64
CS‐RPH	Stage 1	40.20	96.89	108.94	–5.93
	Stage 2	347.50	376.40	382.50	–53.59

Sugar release during in vitro gastrointestinal digestion

Figure 6 illustrates a notable disparity in the sugar release capacity between pure corn starch (CS) and the corn starch/red bean protein hydrolysate complex (CS‐RPH) after in vitro digestion. In the simulated salivary phase (SSF), CS‐RPH exhibited a higher sugar release rate compared to CS. However, in the simulated gastric phase (SGF), CS (70.67%) exhibited a higher sugar release rate compared to CS‐RPH (68.04%), most likely a result of the hydrolyzed protein creating an interaction network with starch, which limited the accessibility of digestive enzymes.^{15, 59} During the simulated intestinal phase (SIF), both samples attained peak sugar release levels; however, CS exhibited a total sugar release compared to CS‐RPH (96.78%), suggesting that the hydrolyzed red bean protein may provide a protective barrier around the starch, therefore decelerating the breakdown rate. This outcome indicates that the mixture of hydrolyzed red bean protein and corn starch may regulate glucose release, hence decelerating carbohydrate digestion in the colon. The variation in released sugar levels results from the crystal structure and the inhibition of amylase activity in the CS‐RPH complex, ⁶⁰ signifying that RPH diminishes starch digestibility, which is also influenced by the concentration of RPH inside the complex. The V‐shaped crystalline structure of starch exhibits increased resistance to α‐amylase hydrolysis, potentially reducing the rate of starch hydrolysis. ⁶¹ A study conducted by Wang et al. ⁵³ assessed the digestibility and estimated glycemic index of a maize starch/soy isoflavone combination. The findings indicated that soybean isoflavones positively influenced starch digestibility and decreased postprandial blood glucose levels.

Figure 6.

Sugar release during in vitro gastrointestinal digestion by simulated salovary fluid (SSF), simulated gastric fluid (SGF) and simulated intestinal fluid (SIF) of corn starch (CS) and corn starch/red bean protein hydrolysate complex (CS‐RPH). Different lowercase and uppercase letters indicate the significant differences between samples (P < 0.05) regarding digestive phase and powder, respectively.

CONCLUSIONS

Protein hydrolysates derived from red beans have been shown to possess significant bioactive properties, making them promising candidates for various health‐related and technological applications. The use of proteins as a physical barrier to inhibit amylase access and reduce starch digestibility is a promising approach. In the present study, the corn starch/red bean protein hydrolysate complex was shown to have control on the sugar release during in vitro gastrointestinal digestion. Consequently, it may serve as an innovative method to regulate starch digestibility and glucose release. Moreover, additional assessment of the dose‐response relationship is required to quantify the postprandial glycemic response.

FUNDING INFORMATION

This research did not receive any specific grant from funding agencies in the public, commercial or not‐for‐profit sectors.

CONFLICTS OF INTEREST

The authors declare that they have no conflicts of interest.

AUTHOR CONTRIBUTIONS

T‐T‐DN, M‐TT, T‐V‐LN, TTVT, ADD, D‐TN, Q‐TH and Q‐DN were responsible for data curation. T‐T‐DN, M‐TT, ADD, D‐TN, Q‐TH and Q‐DN were responsible for investigations. T‐T‐DN, M‐TT, T‐V‐LN, TTVT and Q‐DN were responsible for methodology and visualization. T‐T‐DN, M‐TT, T‐V‐LN, TTVT, ADD, D‐TN, Q‐TH and Q‐DN were responsible for writing the original draft. Q‐DN was responsible for conceptualization and reviewing and editing.

DATA AVAILABILITY STATEMENT

The data that support the findings of this study are available from the corresponding author upon reasonable request.