Characterization of protein extracted from the Omani seaweed- Hypnea bryoides

2.1.1 Sample collection

H. bryoides samples were collected by Salalah Diving Services Company (Salalah, Sultanate of Oman) from the Sadh region (Dhofar Province, Sultanate of Oman). The coordinates of the geographical sampling locations were: 1782.96’04°”N 5565.3’07°”E, 1652.18’94°”N 5447.89’80°”E, 1615.19’95°”N 5485.18’81°”E, and 1607.90’96°”N 5460.32’75°”E. The samples were collected during the period 20 September - 20 November 2021. The seaweed specimens were identified in the field based on their morphological characteristics.

2.1.2 Sample handling

After collection, seaweed samples were cleaned of any foreign materials (e.g., other seaweed plants, sand, etc.) and thoroughly washed with seawater. The samples were then placed in a cool box filled with clean seawater and transported to the facility of Salalah Diving Services Company. At the facility, the samples were thoroughly rinsed with fresh water, spread on a clean plastic sheet, and left to dry under the sun for 3 to 5 days, with daily flipping. The sun-dried seaweeds were then packed in a polyethylene bag and shipped to the Food Chemistry Lab (College of Agricultural and Marine Sciences, Sultan Qaboos University, Oman).

2.2.1 Extraction method

Prior to protein extraction, the sun-dried sample was depigmented following the procedure reported by Al-Alawi et al. (2011). Forty grams (40 g) of milled seaweed sample were mixed with 600 mL of an acetone-methanol solution (1:1) in a 1000 mL bottle. The mixture was homogenized for 5 min using an ultra-mixer (Ultra TURRAX T25 Digital Dispenser, IKA, Germany) at 10,000 rpm. Next, the mixture was stirred for 1 h at room temperature, then centrifuged for 10 min at 4000 g. The supernatant was decanted, and the sediment was re-treated with a fresh methanol-acetone solution. The mixture was then stirred and centrifuged using the same procedure described above. The extraction was repeated three times, after which the sediment was used for protein extraction.

The protein extraction process was performed as described by Mæhre et al. (2016), with slight modifications. The depigmented sample was dispersed in 0.3 M NaOH (1:10, w/v), stirred at room temperature for 1 h, and then centrifuged at 5000 g for 10 min at 4 °C. After centrifugation, the supernatant was collected, and the precipitate was re-dispersed and treated as previously described. The supernatants from both steps were combined and dialyzed using dialysis membranes (MWCO 14,000 Da, Sigma-Aldrich, USA) against deionized water for three days at room temperature, with daily water replacement. The dialyzed samples were lyophilized using a Labconco Triad freeze dryer (Labconco Corporation, USA), packed into plastic containers, and stored at 4°C until further analysis.

2.2.2 Protein concentration and purity

The crude protein concentration and purity were determined using the Bicinchoninic Acid (BCA) assay Kit method (Sigma-Aldrich, USA) according to the manufacturer’s instructions. A calibration curve was constructed using bovine serum albumin at concentrations from 0 to 1 mg/mL.

2.3.1 Fourier-transform infrared (FTIR) analysis

The analysis was performed following the procedure described by Al-Alawi et al. (2011). A FTIR spectrometer (Agilent Cary 670, Malaysia), equipped with a single-bounce diamond attenuated total reflectance (ATR) cell (GladiATR, PIKE Technologies, USA), was used to analyze the extracted protein samples. Spectra were recorded at a resolution of 4 cm⁻¹, averaging 32 scans per sample.

2.3.2 Molecular weight measurement and protein profiling

The molecular weight of the protein extract was determined using size exclusion chromatography, following the method described by Sun et al. (2010), with slight modifications. The analysis was performed on a Nexera 2X system (Shimadzu, Japan), operated by LabSolutions software (version 5.82 SP1), and equipped with an autosampler (SIL-30AC), quaternary pump (LC-30AD), a diode array detector (SPD-M30A), and a column compartment (CTO-20AC). Separation was achieved using an Ultrahydrogel linear column (300 mm × 7.8 mm, Waters, USA) maintained at 25°C.

The mobile phase consisted of a mixture of 0.05 M phosphate buffer (prepared from 1 M potassium phosphate dibasic and 1 M potassium phosphate monobasic) and 0.15 M NaCl solution. The flow rate was set to 0.5 mL/min, and the injection volume was 15 µL. Prior to injection, samples were filtered through a 0.22 µm nylon syringe filter. Molecular weight standards included amyloglucosidase (97 kDa) from Aspergillus niger (Fungi), carbonic anhydrase II (30 kDa) from bovine erythrocytes, trypsin inhibitor (20 kDa) from soybean, and vitamin B12 (cyanocobalamin, 1.355 kDa).

2.3.3 Amino acid profile and protein chemical score

The amino acid profile of H. bryoides’s protein was determined using a Nexera 2X system equipped with an LCMS-2020 Single Quadrupole LC/MS detector (Shimadzu, Japan), following the hydrolysis procedure described by Al-Saidi et al. (2011). 50 mg of the dried seaweed was placed in a hydrolysis tube with 5 mL of 6 M HCl containing 1% (v/v) phenol. The tube was flushed with nitrogen gas for 30 sec and then placed in a heating block (DRi-block DB-3D, TECHNE, UK) at 110°C for 24 h; 1 mL of the resulting hydrolysate was then mixed with 9 mL of deionized water. Subsequently, 5 µL of the sample was injected into the LC-MS system. Prior to injection, the diluted hydrolysate was filtered using a 0.22 µm nylon syringe filter.

Amino acid separation was performed using gradient elution on an Intrada Amino Acid column (50 × 3 mm, 3 µm, Imtakt, Japan) maintained at 35°C, following the manufacturer’s instructions. The mobile phases consisted of acetonitrile/formic acid (100/0.1) as solvent A, and 100 mM ammonium formate solution as solvent B. The flow rate was set at 0.6 mL/min. Elution began at 14% solvent B for 3 min, followed by a gradient increase from 14% to 100% B over 7 min, then returned to 14% B within 2 min.

The protein chemical score (CS) was calculated using the following equation, according to the method adopted by FAO/WHO (1991):

2.4.1 Oil holding capacity (OHC) and water holding capacity (WHC)

The oil and water holding capacities (O/WHC) were determined according to Garcia-Vaquero (2017). The extracted protein (i.e., 0.3 g) was weighed and transferred to an empty centrifuge tube. Then, three milliliters (3 mL) of distilled water (for WHC) or sunflower oil (for OHC) was added, mixed for one minute using a vortex mixer, and centrifuged at 2200 g for 30 min. The tube containing the sediment was weighed, and the final weight was recorded after decanting the supernatant. The formula below was used to calculate W/OHC:

3.1.1 Protein yield and purity

The crude protein yield obtained from H. bryoides was 6 ± 0.35% (db) of the total seaweed biomass (Fig. 1), representing approximately 31% of the total protein initially present in the seaweed. The extracted protein exhibited a purity of 88.5 ± 0.71%. Although other Hypnea species such as H. charoides (83.1 ± 0.94%) and H. japonica (85.0 ± 0.38%) showed comparable protein purity, their recovery yields relative to the existing protein content were markedly higher than that of H. bryoides (H. charoides: 46.3 ± 0.61%; H. japonica: 45.4 ± 0.23%) (Wong & Cheung, 2000). In contrast, the red seaweed Kappaphycus alvarezii (K. alvarezii) yielded 43% of its total protein with a purity of 62.3 ± 1.62%, which was notably lower than that achieved in this study (Kumar et al., 2017).

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Fig. 1.

The extracted protein from H. bryoides (a) before and (b) after freeze drying.

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The extraction method plays a critical role in determining both the quality and yield of the resulting protein extract. Alkaline conditions are generally known to enhance protein yield, and numerous studies support this. This increase is attributed to the improved solubilization of hydrophobic (water-insoluble) proteins from seaweed matrices through enhanced ionization under alkaline conditions (Kadam et al., 2015). However, the relatively low protein yield observed in this study, despite the use of alkaline extraction conditions, can be attributed to several factors.

Proteins are classified into four distinct solubility classes based on their extraction needs: glutelins (alkali-soluble), prolamins (alcohol-soluble), globulins (salt-soluble), and albumins (water-soluble) (Osborne, 1907; Schalk et al., 2017). Since this study employed only alkaline extraction conditions, it primarily targeted glutelin-type proteins while potentially leaving substantial amounts of globulins, prolamins, and albumins (Duranti et al., 2008). Most proteins in H. bryoides may belong to solubility classes other than glutelins, which would explain the modest yield despite the use of alkaline conditions that are typically effective for protein extraction. This selectivity of the extraction method suggests that a sequential extraction approach using multiple solvents could potentially recover a more comprehensive protein fraction and achieve higher overall yields (Rasheed et al., 2020).

Additionally, protein loss during the dialysis step, which used a 14,000 MW cut-off membrane, likely allowed low-molecular-weight proteins and peptides to pass through, further contributing to the reduced yield observed in this study.

Furthermore, specific proteins in macroalgal cells form complexes with non-protein compounds, like polyphenols and polysaccharides, which can complicate their extraction and purification (Harnedy & FitzGerald, 2013).

3.2.1 FTIR analysis

Fig. 2 presents the FTIR-ATR spectra of proteins extracted from H. bryoides. FTIR spectroscopy is a valuable analytical technique for identifying the secondary structure of proteins by examining characteristic vibrational bands associated with the peptide backbone (Barth, 2007; Kong & Yu, 2007).

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Fig. 2.

FTIR-ATR spectra of an H. bryoides protein and the second derivative and curve-fitting deconvolution of the amide I band.

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In the spectrum, the prominent broad peak at approximately 3278 cm⁻¹ corresponds to N–H stretching vibrations of polypeptide chains, indicating the presence of hydrogen-bonded amide groups (Barth, 2007). The peaks observed at 2961 cm⁻¹ and 2928 cm⁻¹ are attributed to asymmetric and symmetric C–H stretching vibrations associated with CH₂ and CH₃ functional groups from amino acid side chains, respectively (Goormaghtigh et al., 2006). The spectral region ranging from 1700 to 1100 cm⁻¹ encompasses the amide bands (I, II, and III), which are directly related to the protein’s secondary structure and provide the most diagnostic information for conformational analysis.

The amide I band, which mainly indicates C=O stretching vibrations of the peptide bond. (70-85%) with minor contributions from C-N stretching (10-20%), appears as a sharp, intense peak near 1638 cm⁻¹ (Kong & Yu, 2007). This frequency is characteristic of β-sheet secondary structures, which typically absorb in the 1630-1640 cm⁻¹ range, due to intermolecular hydrogen bonding between peptide chains (Miller et al., 2013; Usoltsev et al., 2020). The medium-intensity peak at 1538 cm⁻¹ corresponds to the amide II band, which arises from N–H bending vibrations (40-60%) coupled with C–N stretching vibrations (18-40%) and is generally less sensitive to secondary structure than amide I (Barth, 2007). Additionally, a smaller peak at 1241 cm⁻¹ is attributed to the amide III band, which involves a combination of C–N stretching and N–H bending vibrations, providing supplementary structural information (Goormaghtigh et al., 2006).

For precise secondary structure analysis, the amide I band was quantitatively evaluated by second-derivative spectroscopy, followed by deconvolution using curve-fitting (Kong & Yu, 2007). The deconvolution analysis revealed that the predominant peak at 1625 cm⁻¹ accounts for approximately 56.1% of the total amide I band area, indicating that β-sheet structures constitute the major secondary structural component of H. bryoides proteins. Additional minor components were identified at 1654.9 cm⁻¹ (α-helix, ∼18.2%) and 1681.7 cm⁻¹ (β-turns or antiparallel β-sheet, ∼25.7%), providing a comprehensive secondary structure profile (Usoltsev et al., 2020).

β-sheet-rich proteins demonstrate good gel-forming capabilities due to their capacity to form intermolecular networks through hydrophobic interactions and hydrogen bonding (Dickinson, 2003). This structural arrangement enhances the water-holding capacity and mechanical strength of protein gels, making them suitable for modifying food products. The extended conformation of β-sheets also provides excellent emulsification properties, as the hydrophobic regions can interact with lipid phases while hydrophilic regions remain in the aqueous phase (Foegeding et al., 2002).

In food processing applications, β-sheet proteins show enhanced thermal stability compared to α-helical proteins, maintaining their functional properties under moderate heating conditions commonly used in food preparation (Damodaran, 2017). This thermal resilience makes H. bryoides proteins particularly suitable for processed food applications that require heat treatment. Furthermore, the β-sheet structure facilitates the formation of protein films and coatings with good barrier properties, which could be valuable for food packaging applications (Gennadios et al., 1994).

The predominant β-sheet structure of H. bryoides proteins also suggests potential applications in plant-based protein products, where β-sheet-rich proteins can contribute to the fibrous texture characteristic of meat analogs through controlled aggregation and alignment during processing (Dekkers et al., 2018). β-sheet proteins’ capacity to form ordered and extended structures makes them especially suitable for creating anisotropic textures that imitate muscle fiber architecture.

Comparative analysis with other seaweed proteins reveals that the β-sheet predominance in H. bryoides is relatively uncommon, as many marine algae proteins exhibit mixed secondary structures with significant α-helical content (Fleurence, 1999). This unique structural feature may enhance the distinctive functional qualities of H. bryoides proteins and their potential benefits in certain food uses.

3.5.1 Water and oil holding capacities (W/OHC)

W/OHC are valuable to the food industry due to their influence on texture and flavor (Garcia-Vaquero et al., 2017). These functional properties serve as critical indicators of protein performance in food systems, directly affecting product quality and consumer acceptance (Damodaran, 2005). The WHC describes the protein’s capability to bind to water within the food matrix through several molecular interactions, such as electrostatic forces and hydrogen bonding (Bleakley & Hayes, 2021).

The WHC of the H. bryoides protein extract was 9.61 ± 0.15 g water/g protein, demonstrating a substantial water-binding capacity, positioning this seaweed protein as a functional ingredient for moisture-sensitive applications. This finding is slightly lower than the values of WHC found for H. japonica (11.8 ± 0.05 g water/g protein) and H. charoides (10.9 ± 0.30 g water/g protein), but comparably higher than the value found in the protein extracted from K. alvarezii (2.22 ± 0.04 g water/g protein) (Wong & Cheung, 2000; Kumar et al., 2017). Furthermore, the H. bryoides WHC is close to the WHC of legume proteins (i.e., pea, chickpea, and lentil), which range between 10.5 and 13 g water/g sample, indicating comparable functionality to established plant protein sources (Ettoumi & Chibane, 2015).

The WHC of protein is important in various foods, especially viscous foods such as dough, soup, custard, and baked goods, as they are intended to absorb water, causing protein suspension, so improving the product’s thickening and viscosity (Ettoumi & Chibane, 2015). Different factors contribute to a protein’s capability to absorb water, including protein conformation, amino acid composition, surface polarity or hydrophobicity, and physical properties such as particle size and porosity (Wong & Cheung, 2000). The molecular structure of proteins creates specific hydration sites where water molecules can be bound through various mechanisms, with polar amino acid residues playing a vital role in water retention (Zayas, 1997).

OHC is an important characteristic of food ingredients that improves mouthfeel and preserves flavor, making it a key factor in lipid-rich food products. (Ettoumi & Chibane, 2015). In this study, the OHC of the H. bryoides was found to be 13.56 ± 0.26 g oil/g protein, which is relatively higher than the OHC values reported for many other seaweeds, indicating exceptional lipid-binding capacity. For example, the OHC of K. alvarezii was 1.29 ± 0.20 g oil/g of protein (Kumar et al., 2017). Moreover, other seaweeds from the genus Hypnea showed lower OHC values, such as H. charoides (0.95 ± 0.04 g oil/g protein) and H. japonica (0.82 ± 0.01 g oil/g protein) (Wong & Cheung, 2000).

The OHC is affected by capillary attraction, which physically entraps oil via both surface adsorption and internal absorption mechanisms (Sharoba et al., 2013). Hydrophobic proteins also play a major role in oil absorption through non-polar interactions with lipid molecules (Chen et al., 2019). Thus, the variation in OHC values among different seaweed species is linked to the protein content, the type of protein, amino acid composition, and the non-polar side of the amino acids that interact with fat molecules (Zielińska et al., 2015).The current OHC value is high, even higher than those reported in legumes (i.e., peas, chickpeas, and lentils) (6 to 8 g oil/g protein) (Ettoumi & Chibane, 2015). Therefore, it is suggested that H. bryoides contains more nonpolar amino acids. High OHC is crucial for bakery products and other food applications, such as formulations containing ground meat and meat substitutes (Zielińska et al., 2015). Therefore, due to its high water and oil-holding capacity, H. bryoides is a valuable source of functional protein that could have enormous applications in food manufacturing.

3.5.2 Protein solubility

Protein solubility is regarded as the most important property of proteins as a food ingredient because it affects several functional traits, such as emulsification and foaming, and is essential to other functional characteristics (Ettoumi & Chibane, 2015). The solubility behavior of proteins depends on the balance between protein-solvent and protein-protein interactions, which are strongly influenced by environmental factors like ionic strength and pH (Kinsella, 1979). The solubility of H. bryoides protein was tested at different pHs ranging from 2 to 10. The results indicated that protein solubility increased as pH rose (Fig. 4).

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Fig. 4.

Solubility (%) of H. bryoides protein at different pH values.

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Furthermore, the protein exhibited minimal solubility at pH 2; therefore, pH 2 was identified as the protein’s isoelectric point, the pH where its net charge is zero, and protein-protein interactions are maximized. Generally, in higher acidic or alkaline conditions, protein molecules acquire charges (positive or negative), leading to electrostatic repulsion and increasing protein solubility (Kumar et al., 2017). H. bryoides protein showed a significant (p ≤ 0.05) increase in solubility starting at pH 2 (40.96 ± 3.40%) to pH 6 (93.24 ± 13.80%), followed by a slight increase (p > 0.05) up to pH 10 (97.92 ± 0.90%).

Other seaweed proteins showed different patterns, reflecting species-specific variations in amino acid composition and protein structure. For example, the solubility of K. alvarezii protein was lowest at pH 4 (33.72 ± 1.23%) and highest at pH 12 (58.72 ± 1.68%) (Kumar et al., 2017). These solubility data are lower than the solubility of H. bryoides protein in our study. On the other hand, our results were close to the data reported by Garcia-Vaquero et al. (2017), where H. elongata’s protein showed maximum and minimum solubility at pH 12 (96.15 ± 0.15%) and pH 4 (22.5 ± 0.5%), respectively. It is worth mentioning that H. elongata is a brown seaweed, and we compared it to our study due to the limited number of studies on the functional properties of proteins extracted from red seaweeds specifically and from all seaweeds in general.

High protein solubility is crucial for protein extracts used in several food products like beverages, coffee whiteners, confections, dressings, and whipped toppings, as functional ingredients, where complete dissolution is essential for product clarity and stability (Garcia-Vaquero et al., 2017).

3.5.3 Emulsifying activity (EA) and emulsifying stability (ES)

The EA of H. bryoides protein at five different pHs using sunflower oil has been shown in Fig. 5. Emulsification is a complex process that involves reducing the interfacial tension between oil and water phases through protein adsorption at the interface (McClements, 2004). The lowest EA was observed at pH 2 (5.16 mL/g), then increased as the pH increased to 46.4 mL/g at pH 8. After that, no significant increase (p > 0.05) at pH 10 (54.6 mL/g) was noticed. This pH-dependent behavior reflects the relationship between protein solubility and emulsifying capacity, as proteins need to first dissolve before moving to the oil-water interface.

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Fig. 5.

Emulsifying activity % of H. bryoides protein at different pH values.

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EA was also analyzed at pH 2 to pH 10 (Fig. 6). Protein at pH 10 showed the best stability (107.76 min), followed by pH 8, 6, and 4 (97.77 min, 87.63 min, and 44.03 min, respectively) with no significant differences between pH 6 and 8 and between 8 and 10 (p > 0.05). The pH 2 had the lowest stability (18.27 min) with highly significant differences (p < 0.05) from the other pHs. Protein extracts from the brown seaweed H. elongata showed a somewhat similar trend, as the lowest activity and stability were noticed at pH 4, while the highest activity and stability were noted at pH 8 (Garcia-Vaquero et al., 2017).

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Fig. 6.

Emulsifying stability (min) of H. bryoides protein at different pH values.

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Along with the role of pH, the type of oil used in the analysis affects emulsion properties due to differences in viscosity and interfacial tension (Pearce & Kinsella, 1978). Moreover, the emulsion activity of some products might be affected by protein concentration and its nature. On the other hand, stability can be affected by several factors beyond pH, such as protein conformation, net charge, viscosity, interfacial tension, and droplet size (Bleakley & Hayes, 2021). The hydrophobic surface and the flexibility of protein molecules could be manipulated by enzymatic hydrolysis, leading to protein denaturation. It is also possible to improve the hydrophilic-lipophilic balance by uncovering the buried non-polar groups for better emulsification (Phongthai et al., 2016).

However, the strength of the hydrophobic and hydrophilic interactions affects the EA more, whereas the amount and the distribution of non-polar amino acids are responsible for ES, and a high proportion of non-polar amino acids in the protein fraction favors emulsification (Kumar et al., 2017). Protein solubility is also crucial for the emulsion, as rapid migration to the water-oil interface is required (Phongthai et al., 2016). Moreover, a protein with a lower molecular weight is attributed to increasing EA, thus, better interfacial properties of these molecules at the water-oil interface (Felix et al., 2019). The higher ES observed in this study likely results from the fact that the extracted protein includes a substantial amount of nonpolar amino acids.

3.5.4 Foaming capacity (FC) and stability (FS)

FC of H. bryoides protein at various pH values has been displayed in Fig. 7. Foaming is a surface phenomenon that requires proteins to rapidly migrate to the air-water interface and form stable films that can entrap air bubbles (Damodaran, 1997). The FC of all tested samples increased with increasing pH. The lowest value of FC was observed at pH 2 (233.33 ± 28.86%) and increased at pH 4 (406.66 ± 11.54%), pH 6 (500 ± 0%), and pH 8 (543 ± 5.77%) until it reached its highest value at pH 10 (576.66 ± 5.77%). No significant differences appeared between pHs 8 and 10 (p > 0.05).

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Fig. 7.

Foaming capacity of H. bryodies protein at different pH values.

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Low foamability coincides with low protein solubility (Fig. 4), where the polypeptide chains cannot move rapidly to the interface due to limited molecular mobility. The lowest capacity value observed at pH 2 might refer to the behavior of protein molecules at their isoelectric point (zero net charge), where protein aggregation is maximized. Whereas an increase of FC as the pH rises could be due to an increased net charge of the protein, which results in the weakening of the hydrophobic interaction, thus enhancing protein flexibility. This enhanced flexibility helps proteins diffuse and quickly move toward the air-water interface, facilitating air encapsulation and foam formation (Phongthai et al., 2016; Garcia-Vaquero et al., 2017).

In general, the current sample showed a high FC at all pH levels compared to that observed in the red seaweed K. alvarezii (53.33 ± 2.31% at pH 4.0) and the brown seaweed Himanthalia elongata (71.52% at pH 10) (Kumar et al., 2017; Garcia-Vaquero et al., 2017). The FS was measured at different times for the different pH levels (Fig. 8). The results demonstrated that foam stability decreased over time due to gravitational drainage and bubble coalescence (Murray, 2007).

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Fig. 8.

Foaming stability (min) of H. bryoides protein at different pH values.

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The result showed a significant (p ≤ 0.05) drop in FS after 15 min in pH 2 and 4 and after 30 min in pH 6, 8, and 10. No further statistical differences were observed over time (p > 0.05). Although the results showed that pH 4 had a lower FC than pH 6, 8, and 10, it exhibited the highest stability in foaming behavior, and after 30 min, there were no significant differences in FS (p > 0.05). Moreover, the findings demonstrated no significant difference between pH 8 and pH 10 in FS over time (p > 0.05). pH 4 was optimal for FS, where protein solubility/denaturation is moderate. Generally, the high FC relies on the dispersing ability of a protein, but the FS is mainly affected by the degree of protein denaturation (Kumar et al., 2017).