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. 2024 Feb 3;33(8):1847–1857. doi: 10.1007/s10068-023-01490-z

Hydrolysates with emulsifying properties prepared from protein wastes using microbial protease

D Padmapriya 1, C Shanthi 1,✉
PMCID: PMC11091031  PMID: 38752117

Abstract

Plant-based protein hydrolysates have found applications in food industry for emulsification, foaming, and increasing shelf life of food products. The objective of this study is to isolate protease-secreting bacteria hydrolyzing protein waste, and subjecting the resultant hydrolysates for the characterization for application in the food industry. Peanut cake hydrolysates were prepared using proteases from two microorganisms selected for the purpose, viz., Aneurinibacillus migulanus, VITPM11 and Aneurinibacillus aneurinilyticus, VITPS07. The cleavage specificity of the proteases from VITPM11 and VITPS07 were found to be like plasmin and elastase respectively. The cleaving sites of proteases for peanut proteins were predicted using expasy tool. The protease of VITPM11 had maximal activity of 325.8 ± 0.1 U/mL in peanut-cake media. The degree of hydrolysis (32.03 ± 0.89%), solubility (88.5 ± 1.18%), emulsion stability index (89.76 ± 2.80) and foaming stability (68.67 ± 1.53%) properties of VITPM11 protease correlated well with results from bioinformatic studies.

Supplementary Information

The online version contains supplementary material available at 10.1007/s10068-023-01490-z.

Keywords: Peanut cake wastes, Protein hydrolysate, Protein-based emulsion, Aneurinibacillus migulanus, Emulsion stability

Introduction

Bioactive compounds from agricultural and industrial by-products are emerging as an economical and eco-friendly option for use in many industries in the biotechnology age. Huge amounts of protein-rich agricultural wastes could be used as source materials for the production of bioactive hydrolysates using microbial proteases. (Santos Aguilar and Sato, 2018). Various plant and animal-based proteins are mainly used in the production of food protein hydrolysates. Nowadays, plant-based proteins and their hydrolysates are used as protein supplements since animal proteins are limited by the increasing cost,reduced availability and increasing concern relating to biodiversity loss. The production of protein hydrolysates from plant proteins is also gaining importance as more and more people, especially in Europe, are moving to a vegan diet. Protein hydrolysates have several health benefits and play interesting roles in various physiological processes (Zhang et al., 2021).

Proteins and protein hydrolysates are also used as important building blocks in the food matrix with increasing awareness about their nutritional and functional benefits. The foaming and emulsifying properties in food products such as margarine, mousses, whipped toppings, dressings, beverages, and ice cream are improved using protein-based substitutes aided by technological innovations (Muñoz-Arrieta et al., 2021). Emulsions constitute two immiscible phases that necessitate the use of emulsifiers to attain food stability without phase separation. The need for effective natural substitutes as food additives is fast growing due to increasing consumer demand for sustainable and clean-label food components (Dey et al., 2021).

The hydrolysates prepared from a protein-rich cake, a by-product of the oilseed extraction process, have found applications as emulsifiers, stabilizers, sweeteners, odor and aromatic flavor enhancers in food formulations. However, the charge characteristics, size, amphiphilic nature and surface hydrophobicity of the protein hydrolysates would have a pronounced influence on their use in various applications. All these properties of hydrolysates would in turn depend largely on the process employed for hydrolysis (Paglarini et al., 2020).

Hydrolysate production was established during the late nineteenth century, and the processes have been constantly evolving. Chemical hydrolysis of proteins was initially employed,but the use of the greener enzymatic methods are gaining importance in recent times. Enzymatic hydrolysis being perceived as safer compared to chemical methods, can be used to produce biologically active peptides with improved functional properties (Görgüç et al., 2020). Proteases derived from microbes (proteinase K, flavourzyme, protamex, alcalase, serine-protease, metalloprotease), plants (bromelain, papain, ficin) and animals (trypsin, chymotrypsin, neutrase) are commercially available. Hydrolysates derived from the same protein using different proteases show varied peptide profiles, solubility, antioxidant activity, foaming and emulsifying characteristics. Microbial fermentation has been standardized to produce commercial protein hydrolysates (Coelho et al., 2019) with specific characteristics. The main objective of the present study was to screen protease-producing bacterial strains for hydrolyzing proteins in peanut cake waste to yield hydrolysates with good emulsification and foaming properties.

Materials and methods

Materials

Deoiled peanut cakes were obtained from an oil mill near Vellore, India. Peptide substrates, viz., plasmin (D-valyl-L-leucyl-L-lysine-pNA dihydrochloride (V0882)), elastase (N-succinyl-L-alanyl-L-alanyl-L-alanine-pNA (S4760)), trypsin (Nα-benzoyl-DL-arginine-pNA hydrochloride (B4875)) and chymotrypsin (N-succinyl-L-phenylalanine-pNA (S2628)), Inhibitors viz., iodoacetate (407,719), pepstatin A (P5318) and PMSF (phenylmethylsulfonyl fluoride (P7626)) and Protein substrate, azocasein (A2765) were procured from Sigma-Aldrich, India. Protein marker (MBT092) was procured from Himedia, India. All other chemicals used in this study were of analytical grade.

Isolation and identification of protease-producing bacteria

Soil samples were collected near a fish market in Vellore, Tamil Nadu, and India. One gram of soil was soaked in 10 mL of sterile distilled water, serially diluted and spread on a 1% skim milk agar plate, the formation of clear zones around colonies was observed. The morphological characteristics of the isolates with clear zones were studied and maintained as pure cultures. Genomic DNA of the isolated bacteria was extracted from 24 h cultures using HiPuraA bacterial DNA isolation and purification kit (Himedia, India (MB508)). The isolated DNA was amplified using a Cistron master mix kit (Cistron Biolab, India (CBL-113)). Partial sequencing was carried out using primers 27F (5′-AGAGTTTGATCCTGGCTCAG-3′) and 1492R (5′-ACGGCTACCTTGTTACGACTT-3′). Gene sequencing was performed using the Sanger sequencing method (Elumalai et al., 2020). The homology and similarity were analyzed by MEGA 11 software using the available sequences present in the NCBI data bank. A BLAST search was used to construct the phylogenetic tree using the neighbor-joining method.

Optimization of culture medium for protease production

Peanut cake was chosen as substrate media for protease production by isolated bacteria. The protein content of the substrate was determined (Bradford, 1976). To standardize maximum protease production, four different concentrations of substrate (0.25, 0.5, 1 and 2 g per 100 mL) were prepared. The protease activity of 1 mL of culture supernatant was checked at regular intervals from 12 to 36 h using azocasein as a substrate (Tomarelli et al., 1949). 200 µL of cell free culture supernatant was incubated with 1% azocasein substrate solution at 40 °C for 30 min. Then the reaction mixture was centrifuged at 11,000 rpm for 10 min. 1 mL of supernatant was mixed with equal volume of 1N NaOH and optical density was measured at 440 nm. A change of 0.01 optical density per minute is equivalent to one unit of protease activity. The proteases produced by the strains VITPM11 and VITPS07 were characterized. The molecular weight of the proteases was determined using a 10% native-PAGE gel. Commercial inhibitors were used to identify the catalytic type of the proteases. Each of the enzymes (100 µL) was incubated with 100 µL of 10 mM inhibitors for 1 h and loaded onto a 7% polyacrylamide gel containing 1% gelatin. Protease specificity was determined using synthetic substrates. p-Nitroanilide-linked chromogenic synthetic peptide substrates were prepared using 25 mM Tris–HCl buffer at pH 8. The reaction was carried out by adding an equal volume of substrate (3 mM) and proteases (VITPM11 and VITPS07) separately. The reaction mixture was incubated at 40 °C for 1 h. After hydrolysis, the p-nitroanilide released during the reaction was measured in terms of absorbance at 410 nm (Shakilanishi and Shanthi, 2017).

Preparation of peanut protein hydrolysate

The peanut protein isolate (PPI) was produced using the method of (Jing et al., 2019). PPI (5% w/v) was dissolved in 25 mM sodium phosphate buffer at pH 7. The hydrolysis was done with 2000 units of protease for 1–5 h. The samples were collected every hour for 5 h. The enzymes were inactivated by heating the hydrolysate at 90 °C for 15 min. The hydrolysates prepared using VITPM11 and VITPS07 were designated as PPHM and PPHA, respectively.

Substrate specificity study on peanut proteins

The sequences of peanut proteins (Ara h 1, Ara h 2, Ara h 3, and Ara h 6) were obtained from the UniProtKB database. The cleavage sites, peptide sequence, molecular weight and gravy index of the peptides were predicted for plasmin- and elastase-type proteases using the Expasy Peptide cutter tool (https://web.expasy.org/peptide_cutter/). The emulsifying nature of the peptide sequence was analyzed using the peptide length and gravy index according to the hydrophobicity scale (Kyte and Doolittle, 1982).

Characterization of protein hydrolysates

Degree of Hydrolysis (DH)

The DH was determined using the TNBS (trinitrobenzene sulfonic acid) method ((Adler-Nissen, 1979), and the free amino groups were calculated using leucine as a standard (DH=hhtotal×100). The protein hydrolysate was subjected to 10% SDS-PAGE.

Measurement of extrinsic fluorescence

The fluorescence intensities of PPI, PPHM and PPHA were determined (Pi et al., 2021). The sample (1 mL) was mixed with 2 mL of 20 mM phosphate buffer at pH 7.0 and 30 µL of 10 mM ANS (8-anilino-1-napthalenesulfonic acid). Using the excitation wavelength of 280 nm, the relative fluorescent emission was recorded from 380 to 600 nm by fluorescence spectrophotometer.

Measurement of particle size

Particle size was analyzed based on laser diffraction using a Malvern Instrument (Mastersizer 2000 E Version 5.61). The sample (PPI, PPHM, and PPHA) was suitably diluted and the average particle size was measured in terms of surface-weighted mean D3,2=Σnidi3∑nidi2 and volume-weighted mean D4,3=Σnidi4∑nidi3 (Hu et al., 2014).

Assessment of protein solubility

The protein solubility of PPI, PPHM, and PPHA at varying pH values was determined (Nyo and Nguyen, 2019). Twenty milliliters of each protein sample were taken, and pH was adjusted ranging from 2–12 using 1 N solutions of HCl and NaOH. The prepared solutions were continuously stirred for 30 min at room temperature and centrifuged at 10,000 rpm for 10 min. The protein content of the supernatant in each case was estimated (Lowry et al., 1951).

Determination of emulsifying properties

The emulsion activity index (EAI) and emulsion stability index (ESI) were determined using the standard protocol with slight modifications (Lopes-da-Silva and Monteiro, 2019). Sunflower oil (5 mL) was added to 15 mL of the protein sample and sonicated for 10 min. After sonication, 50 µL of the freshly prepared emulsion was diluted with 5 mL of 0.1% SDS, and absorbance was measured immediately and after 10 min at 500 nm using a UV spectrophotometer. The emulsions were prepared using PPI, PPHM and PPHA, and checked for their stability. The emulsion stability was assessed based on the time taken for phase separation observed under an optical microscope.

EAIm2/g=2×2.303×dilutionfactor×A0ϕ×C×10000
ESImin=A0A0-A10×t

The absorbance A0 and A10 are taken at 0 and 10 min, ɸ is the oil fraction, c is the protein weight (g/mL) and time interval (t). Different ranges of pH (2–10), NaCl concentrations (10–200 mM), and temperature (30–90 °C) were used to assess the stability of the emulsion. Freshly prepared emulsions were stored for 15 days at room temperature. Sodium azide (2 mM) was added to the samples during storage to prevent bacterial growth. The ESI, optical microscope microstructure of the droplet and visual images of the treated emulsions were analyzed.

Foaming properties

The foam capacity (FC) and foam stability (FS) of the hydrolysates were also determined (Jain et al., 2015). 25 mL of the sample (PPI, PPHM, and PPHA) was sonicated using an ultrasonicator for 5 min and transferred to a 100 mL measuring cylinder, and the volume of samples along with foam was determined immediately and after 10 min.

Statistical analysis

All graphs were generated using Graph Pad Prism 5. Data were calculated using Microsoft Excel and expressed as mean ± SD. Statistical significance was set at p < 0.05. All experiments were performed in triplicates.

Results and discussions

Isolation and morphological characterization of protease-producing bacteria

Two isolates with clear zones in skim milk agar plates after 24 h of incubation were subcultured and maintained in 80% glycerol at − 20 °C. Both isolates were gram-positive rods forming motile, aerobic, spore-forming, cream-colored colonies.. The isolates were identified as Aneurinibacillus migulanus (VITPM11) and Aneurinibacillus aneurinilyticus (VITPS07) based on the 16S rRNA nucleotide sequencing method. Molecular characterization revealed that the 16S rRNA nucleotide sequence had 97.95% (A. migulanus VITPM11) and 97.13% (A. aneurinilyticus VITPS07) similarity with gene sequences in the NCBI (Accession ID: OQ204387 and KX529699). Both nucleotide sequences of the strains were submitted to GenBank, and were assigned accession numbers MK571210 and MK566203 respectively. A. migulanus, which is classified under the family of Paenibacillaceae (Alenezi et al., 2016) was used to produce biosurfactants that have the property to reduce both interfacial and surface tension, and also improve stability, taste, texture and shelf-life of food products (Sellami et al., 2016). In an earlier study by Mathews et al., Aneurinibacillus aneurinilyticus was used to produce cellulase, peroxidase and laccase for the degradation of cellulose, hemicellulose and lignin (Mathews et al., 2014). In the present study, proteases from A. migulanus and A. aneurinilyticus were used to prepare hydrolysates of peanut cake, an agro-based protein waste.

Protease activity and hydrolysate production

The extracellular proteases produced by microbes are more stable, and require only limited space to cultivate in comparison to that from plants and animals. The production of protease varies with the type of substrates used by the microorganism. Higher protease activity was observed in media that contained 192 ± 0.27 mg of protein/g of peanut cake in 100 mL 25 mM phosphate buffer at pH 7. VITPM11 secreted protease with higher activity thanVITPS07 (Table S1). The molecular weight of the proteases is around 25 and 50 kDa for VITPM11 and VITPS07 respectively (Fig. 1a–c). In the zymogram, the absence of a clear band in Fig. 1b and c confirms that the crude enzymes of VITPM11 and VITPS07 were inhibited by phenylmethylsulfonyl fluoride (lane 5) indicating that the proteases were of the serine-type. The cleavage specificity of the serine protease group was determined using synthetic peptides. The protease from VITPM11 could cleave D-valyl-L-leucyl-L-lysine-pNA dihydrochloride (plasmin specific) with 592.42 ± 0.31 U/µg activity. Plasmin protease had a preference for C-terminal hydrophilic lysine and arginine amino acids at the P1 site of substrates. The protease from VITPS07 cleaved N-succinyl-L-alanyl-L-alanyl-L-alanine-pNA (elastase specific) with 450.27 ± 0.08 U/µg activity. Serine proteases of the elastase type catalyze the cleavage of the C-terminus located on smaller hydrophobic amino acids such as alanine, glycine, and valine (Table 1) (Li et al., 2022a).

Fig. 1.

Fig. 1

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a Native-PAGE: lane M—Molecular marker; Lane A – crude protease of VITPM11; B- crude protease of VITPS07, b and c Gelatin zymography: The effect of inhibitors on protease activity; lane M—Molecular marker (Trypsin 24 kDa); lane 1 – crude protease of VITPM11 (a) and VITPS07 (b); lane 2—EDTA; lane 3—Iodoacetate; lane 4—pepstatin A and lane 5 – PMSF

Table 1.

Protease assay using synthetic peptide substrates

Peptide substrates Enzyme activity (U/µg)
VITPM11 VITPS07
D-valyl-L-leucyl-L-lysine-pNA dihydrochloride 592.42 ± 0.31 173.33 ± 0.43
N-succinyl-L-alanyl-L-alanyl-L-alanine-pNA 266.67 ± 0.02 450.27 ± 0.08
Benzoyl-DL-arginine-pNA hydrochloride 66.62 ± 0.67 80.15 ± 0.33
N-succinyl-L-phenylalanine-pNA 159.23 ± 0.12 190.43 ± 0.16
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Characterization of protein hydrolysate

Degree of hydrolysis

Peptides with low molecular weight can be quickly absorbed into the gastrointestinal and cardiac circulation systems promoting physiological-regulating functions. The peptides formed by protein hydrolysis possess altered structure and function, increased flexibility, and surface area. These peptides have better emulsification properties, as the exposed hydrophilic amino acid residues align with water and hydrophobic residues with oil (Trigui et al., 2021). Table S1 and Fig. 2a and b show that the protease produced by strain A. migulanus showed more hydrolytic activity than that from strain A. aneurinilyticus. Moreover, in correlation with the high DH value, the residual groundnut cake slurry showed more solubilization upon hydrolysis. The highest DH values of 32.03 ± 0.89 and 24.23 ± 0.29% were obtained after 5 h of hydrolysis with strains A. migulanus VITPM11 and A. aneurinilyticus VITPS07 respectively (Fig. 2c). The protease of VITPM11 cleaves at the ends of hydrophilic amino groups such as arginine and lysine, releasing amphiphilic peptides that could promote better emulsification. In the PPI lane, eight predominant protein bands of groundnut protein were observed in the range of 64–15 kDa (Fig. 2a and b). Based on an earlier study by Busu et al., on the molecular weight of peanut proteins (Busu and Amonsou, 2019), the band range around 65 kDa denotes Ara h 1, 45–25 kDa denotes Ara h 3, and 15–20 kDa denotes Ara h 2 and Ara h 6. In 1–5, h Fig. 2b Ara h1, Ara h 2, Ara h 3, and Ara h 6 proteins were completely hydrolyzed by the protease VITPM11, whereas the Ara h 6 protein was not completely hydrolyzed by the protease VITPS07. The hydrolysate obtained after 3 h of hydrolysis was chosen for further studies based on an earlier research investigation which reported that peptides ranging in size from 11 to 27 amino acids showed good emulsification properties. Allergy-causing high molecular weight proteins like Ara h 1, Ara h 2, Ara h 3, and Ara h 6 could also be made less allergic after enzymatic hydrolysis (Bonku and Yu, 2020).

Fig. 2.

Fig. 2

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SDS-PAGE (10%) protein pattern of peanut and cell-free hydrolysate (1 to 5 h). The hydrolysates obtained by the strain A. aneurinilyticus VITPS07 (a), A. migulanus VITPM11 (b), 1: Ara h 1; 2: Ara h 3 acidic subunits; 3: Ara h 3 basic subunits; 4: Ara h 2; and 5: Ara h 6. M: protein markers; PPI: Peanut protein isolate. c DH values were obtained using the crude protease of VITPM11 and VITPS07. d The fluorescent intensity of PPI, PPHM, and PPHA. e Analysis of particle size of PPI, PPHM, and PPHA. f Protein solubility of PPI, PPHM, and PPHA. Emulsion characterization of PPI, PPHM, and PPHA. g EAI (m2/g) and ESI (%)

As per studies by Lin et al. on the relationship of peptide size with emulsifying properties, the peptide sequence ranging from 11–27 amino acids with a molecular weight range of 1200 to 3000 Da was filtered from the cleavage data obtained using elastase and plasmin type proteases by the expasy cutter tool (Table 2). The results hint that peptides cleaved by plasmin type proteases were more in the emulsification range with the gravy index of the peptides being hydrophilic, and the hydrophobic residues present in between probably helping in promoting emulsification.

Table 2.

Prediction of peptides generated from peanut proteins using an expasy cutter tool

Peanut protein Position of the cleavage site Name of cleaving enzyme Resulting peptide sequence Peptide length [aa] Peptide mass [Da] Gravy index
Ara h 3 66 Elastase IESEGGYIETWNPNNQEFECA 21 2431 − 1.02
Ara h 3 272 Elastase GQEEENEGGNIFSGFTPEFLA 21 2272 − 0.68
Ara h 3 116 Elastase GYFGLIFPGCPSTYEEPA 18 1948 0.06
Ara h 1 428 Elastase NPQLQDLDMMLTCV 14 1621 0.15
Ara h 1 530 Elastase SSELHLLGFGINA 13 1358 0.61
Ara h 1 159 Elastase EGEQEWGTPGSEV 13 1404 − 1.42
Ara h 3 354 Elastase GSGNGIEETICTA 13 1251 − 0.05
Ara h 1 298 Elastase NTPGQFEDFFPA 12 1369 − 0.68
Ara h 3 15 Elastase LLELSFCFCFLV 12 1434 2.38
Ara h 1 16 Elastase SPLMLLLGILV 11 1169 2.44
Ara h 3 174 Elastase FWLYNDHDTDV 11 1424 − 0.85
Ara h 1 490 Plasmin EQEWEEEEEDEEEEGSNR 18 2282 − 3.09
Ara h 2 80 Plasmin DPYSPSQDPYSPSQDPDR 18 2051 − 2.18
Ara h 3 334 Plasmin DEEEEYDEDEYEYDEEDR 18 2401 − 3.19
Ara h 3 194 Plasmin SLTDTNNNDNQLDQFPR 17 1992 − 1.73
Ara h 3 146 Plasmin LQEEDQSQQQQDSHQK 16 1956 − 2.71
Ara h 1 393 Plasmin GSEEEDITNPINLR 14 1587 − 1.16
Ara h 3 233 Plasmin SLPYSPYSPQSQPR 14 1607 − 1.42
Ara h 6 68 Plasmin IMGEQEQYDSYDIR 14 1747 − 1.31
Ara h 1 52 Plasmin CLQSCQQEPDDLK 13 1507 − 1.13
Ara h 2 115 Plasmin CCNELNEFENNQR 13 1613 − 1.61
Ara h 6 91 Plasmin CCDELNEMENTQR 13 1585 − 1.46
Ara h 1 271 Plasmin IPSGFISYILNR 12 1380 0.60
Ara h 1 405 Plasmin DGEPDLSNNFGR 12 1320 − 1.55
Ara h 6 131 Plasmin ELMNLPQQCNFR 12 1493 − 0.73
Ara h 1 312 Plasmin DQSSYLQGFSR 11 1287 − 1.14
Ara h 2 131 Plasmin LQQIMENQSDR 11 1361 − 1.46
Ara h 6 107 Plasmin LQQIMENQCDR 11 1378 − 1.16
Ara h 1 490 Plasmin EQEWEEEEEDEEEEGSNR 18 2282 − 3.09
Ara h 2 80 Plasmin DPYSPSQDPYSPSQDPDR 18 2051 − 2.18
Ara h 3 334 Plasmin DEEEEYDEDEYEYDEEDR 18 2401 − 3.19
Ara h 3 194 Plasmin SLTDTNNNDNQLDQFPR 17 1992 − 1.73
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In Fig. 2a and b, the unhydrolyzed protein bands are visible in PPHA, whereas they are fully hydrolyzed in PPHM. The basic amino acids such as arginine (11%) and lysine (3.6%) content in peanut proteins are high, and thus the plasmin-type protease VITPM11 cleaved peptides could possess better emulsification.

Measurement of extrinsic fluorescence

As surface hydrophobicity influences emulsification properties, changes in this property upon enzymatic hydrolysis of peanut protein were monitored using a fluorescence spectrometer. The peanut proteins contain both hydrophobic (Gly 6.0%, Ile 3.0%, Leu 6.1%, Met 0.4%, Phe 4.9%, Tyr 3.7%, and Val 3.7%) and hydrophilic (Arg 11%, Cys 0.9%, His 2.5%, Lys 3.6%, and Thr 2.8%) amino acids. During enzymatic hydrolysis, the protein is cleaved and the buried hydrophobic regions (Tyr, Trp, and Phe) are exposed (Zheng et al., 2019). ANS preferentially binds to hydrophobic regions that are exposed to unfolded proteins. The λmax of PPI was recorded at 444 nm when excited at 280 nm. The extrinsic fluorescence shows a higher wavelength shift (redshift) in PPHM and PPHA at 467 and 465 nm (λmax) respectively, indicating the exposure of hydrophobic residues to polar solvents (Fig. 2d). The red shift was higher for PPHM because a large number of buried hydrophobic amino acid residues were exposed during hydrolysis. The peptides have improved surfactant properties with both hydrophobic and hydrophilic amino acids exposed (Jain and Anal, 2016).

Measurement of particle size

The hydrolysis of proteins results in a change in the secondary structure and the formation of lower molecular sized peptides. The surface-weighted means (D[3,2]) and volume-weighted means (D[4,3]) for PPI, PPHM, and PPHA are presented in Fig. 2e. The reduction of particle size would lead to an increase in the overall surface area and hence have a major influence on emulsion stability and oxidative stability. The particle size is inversely proportional to oxidative stability. The higher particle size of proteins would lower emulsion stability (Godoi et al., 2021). Generally, a particle size < 10 µm promotes good emulsion stability over a period. It can be seen in Fig. 2e that the surface-weighted means (D[3,2]) for PPI, PPHM and PPHA were 29.806 ± 0.20, 0.313 ± 0.01 µm and 11.592 ± 0.03, respectively. The corresponding volume-weighted means (D[4,3]) were 96.26 ± 0.60, 0.90 ± 0.01 and 16.442 ± 0.18 µm. Bimodal peaks were observed in PPI and PPHM. Apparently, the unhydrolyzed protein in PPI contributed to its poor solubility. The average particle size of PPHM was very smaller when compared with PPI and PPHA, and thus helping in maintaining the stability of the emulsion. Three peaks in the PPHA hydrolysate with varying particle size ranges indicate that the sample was partially hydrolyzed. PPHA has more hydrophobic amino acid ends that probably tend to aggregate resulting in increased particle size.

Solubility

Solubility is an important functional property of the protein hydrolysate, which can have a pronounced effect on other functionalities such as emulsification and foaming. Plant proteins have limited solubility in general, which can be increased when they are hydrolyzed at suitable pH conditions. Excellent solubility was achieved at a high degree of hydrolysis, with the solubility being the lowest near its isoelectric pH (Ashaolu et al., 2022). The food industry requires highly soluble proteins in wide pH ranges of 2–12. The solubility of PPI, PPHM, and PPHA hydrolysates increased with increasing pH. The solubility of PPHM was higher than that of PPI and PPHA (Fig. 2(f)). PPHM (31.17–88.85%) contains protein hydrolysates with low molecular weight, which correlated well with the expasy cutter results and hence was more soluble in water compared to PPI (19.58–63.89%) and PPHA (25.47–78.16%). Peptides and amino acids mainly determine water dynamics and moisture absorption (Yang et al., 2017). The hydrophilic ends (arginine and lysine) present in the PPHM probably help in enhancing the solubility of PPHM protein hydrolysate and the hydrophobic ends (alanine, valine, and glycine) exposed in PPHA impart low solubility. Poorly hydrolyzed PPI has low solubility because of its complex structure.

Emulsion stability

Emulsions are widely employed in the food sector as carriers to incorporate bioactive substances. In recent studies, plant-derived protein hydrolysates have been extensively used as healthy and nutritional emulsion stabilizers. The EAI and ESI of PPI, PPHM and PPHA are depicted in Fig. 2g. The EAI represents the capability of the protein to promote dispersion of oil-phase in water. ESI dictates the stability of the emulsion by resisting creaming, flocculation, and coalescence. A highly stabilized emulsion signifies a greater ESI value (Jain and Badve, 2022). The highest EAI and ESI values were obtained for PPHM (Fig. 2g). Peptides that are smaller in size have high mobility and can easily attach to the interface to promote emulsification. Hence such smaller peptides would rapidly adsorp oil droplets rapidly during sonication. The anchoring of peptides in the oil–water interface could make a conformational change to reduce interfacial tension. Adequate interfacial stratum formation prevents droplet aggregation and coalescence (Yang et al., 2022). Proteins that promote good emulsion stability would have smaller droplets with lower coalescence. PPHM probably has many peptides that promote good emulsification, as predicted by an expasy cutter tool than PPHA and PPI.

The emulsion stability was assessed with changes in pH, ionic strength and temperature to study their utility in the food sector. The amphiphilic structure and flexibility of peptides determine the stability of protein-based emulsions. In recent studies, it has been proven that the oxidative stability of emulsions is enhanced using protein-based emulsifiers (Li et al., 2022c). Protein emulsions were further converted into edible coatings, films and emulsion gels through various food processing techniques such as the addition of copolymers and heating. The influence of pH on emulsification properties of PPI, PPHM and PPHA was analyzed (Fig. 3A). The emulsions are more unstable near the isoelectric point because the electrostatic repulsion is less than the van der Waals force and hydrophobic interactions. The ESI was found to be minimum at pH 4. The emulsion was unstable at acidic pH (2–6) for PPI, PPHM, and PPHA with the phase separation being observed [Fig. 3A(a)]. The microstructural changes in the emulsion droplet size were higher in the acidic pH range [Fig. 3A(b)]. As the hydrolysates are precipitated at acidic pH, the emulsions become stratified. The prepared emulsions were more stable at alkaline pH with the ESI values of PPI, PPHM and PPHA at pH 8 being 49.11 ± 1.91, 89.76 ± 2.80, and 79.04 ± 1.08, respectively [Fig. 3A(c)]. The emulsion droplet size of the hydrolysates was low at alkaline pH (8–10).

Fig. 3.

Fig. 3

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A pH stability of freshly prepared emulsion (PPI, PPHM, and PPHA): (a) visual image of emulsions; (b) optical microscopy image (scale bar: 10 µm); (c) Emulsion stability index (ESI). B Ionic strength stability of freshly prepared emulsion (PPI, PPHM, and PPHA): (a) visual image of emulsions; (b) optical microscopy image (scale bar: 10 µm); (c) Emulsion stability index (ESI). C Temperature stability of freshly prepared emulsion (PPI, PPHM, and PPHA): (a) visual image of emulsions; (b) optical microscopy image (scale bar: 10 µm); (c) Emulsion stability index (ESI). D Storage stability of freshly prepared emulsion for 15 days storage (PPI, PPHM, and PPHA): (a) visual image of emulsions; (b) optical microscopy image (scale bar: 10 µm); (c) Emulsion stability index (ESI)

The ionic strength stability based on visual appearance, droplet microstructure and ESI with different NaCl concentrations of emulsions are depicted in (Fig. 3B). The emulsion stability is inversely proportional to NaCl concentration indicating that ionic concentrations highly affect aggregation. At low NaCl concentrations, PPI (10 mM) and PPHA (50 mM) were stratified, whereas PPHM was more stable. At high NaCl concentrations (100–200 mM), different levels of stratification were observed in PPHM [Fig. 3B(a)]. The PPI and PPHA were more stratified. The electrostatic attraction between the droplets was greater than the repulsion, which promoted the aggregation of droplets (Ling et al., 2020). The visual appearance and droplet microstructure of the emulsions were observed under different NaCl concentrations (10–200 mM) for PPI, PPHM, and PPHA (Fig. 3B(a&b)). The phase separation and changes in droplet morphology represent the aggregation of droplets under different ionic conditions. In Fig. 3B(c), reduced ESI values for PPI (from 49.27 ± 2.09 to 12.30 ± 0.47), PPHM (from 84.19 ± 1.35 to 33.78 ± 0.99), and PPHA (from 52.59 ± 1.88 to 19.51 ± 0.67) were observed. The PPHM emulsion was more stable at the ionic concentration range of 10–50 mM.

Thermal treatment in the food industry helps inhibiting bacterial growth by sterilization and thus effectively increasing the shelf-life of food products (Calderón-Chiu et al., 2021). In Fig. 3C, the temperature stability of the protein-based emulsions was studied under different temperature conditions (30–90 °C). The PPHM emulsion was stable up to 90 °C and there was no phase separation. The emulsions treated at high temperatures showed an increase in droplet size for PPI and PPHA [Fig. 3C(a and b)]. The ESI value for PPHM was reduced from 86.25 ± 1.14 to 82.11 ± 2.21 for 30–90 °C. The emulsions showed phase separation from 60 °C in PPI and PPHA emulsions. The reduced ESI values of PPI (from 45.15 ± 1.08 to 33.03 ± 1.23) and PPHA (from 79.04 ± 1.08 to 55.08 ± 2.50) for 30–90 °C were noticed [Fig. 3C(c)]. The phase separation and changes in droplet size are due probably to the alteration of peptides present on the surface during thermal treatment, which modifies the surface hydrophobicity promoting flocculation and aggregation (Li et al., 2022b).

Storage stability is an important property of processed foods. PPI, PPHM, and PPHA emulsion samples were prepared and stored for 15 days at 25 °C (Fig. 3D). The visual appearance, droplet microstructure, and ESI values were used as indicators to determine the change in the stability of the emulsion. Freshly prepared emulsions did not show any phase separation. After 15 days of storage, a clear phase separation of the emulsion was observed in PPI, a slight phase separation in PPHA, and no phase separation in PPHM [Fig. 3D (a)]. The emulsion droplet size increased and stratified in PPI and PPHA after 15 days of storage [Fig. 3D(b)]. This could be due to the aggregation of unhydrolyzed proteins and hydrophobic peptides. In Fig. 3D(c), ESI values reduced from 49.11 ± 1.91 to 14.31 ± 0.10 for PPI, from 89.76 ± 2.80 to 80.32 ± 2.15 for PPI, and from 79.04 ± 1.08 to 33.78 ± 0.18 for PPHA. These results indicate that emulsions prepared using PPHM are more stable during storage.

Foaming stability

The FC and FS of PPI, PPHM and PPHA are depicted in Fig. 4. Foaming ability is also an important characteristic that helps in the formulation of food products. Foams are air bubbles in a continuous phase that play essential structural and textural roles in chocolate mousses, cakes, coffee-milk mixes, meringues, tiramisu and bread. Nowadays, the foaming properties of the plant protein hydrolysates are exploited as an alternative to egg white protein in aerated food e.g., bread and cakes (Wang et al., 2021). The foaming capacity of hydrolyzed peanut protein depends on the type of protease and the hydrolysis conditions employed. To produce better foams, the air bubbles should interact with each other in the matrix. The FC and FS were found to be higher for PPHM than PPI and PPHA (Fig. 4). The exposed sites on the amphiphilic peptides in PPHM reduce the surface tension between the air/water interface, which in turn helped in foam formation and stability. FC and FS were reduced in PPI and PPHS due to the aggregation of proteins and peptides.

Fig. 4.

Fig. 4

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FC and FS of PPI, PPHM and PPHA

In India, cold-pressed peanut oil has been conventionally used in cooking, and is again gaining popularity due to its nutritional benefits. The oil cake, a protein-rich waste from peanut oil extraction process can be a safer alternative for fat emulsifiers in food formulation. Hence, a systematic scientific attempt has been made in the study for the cleaner enzymatic method of preparation of protein hydrolysates with improved functional properties. A protease produced from VITPM11 isolated from soil samples near a fish market was found to be suitable for the purpose as assessed in terms of particle size, the emulsification, and foaming characteristics. Protein hydrolysate produced from peanut cake using the enzyme could find application as a substitute for fat emulsifiers in food formulations.

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary file1 (DOCX 1865 KB) (1.8MB, docx)

Acknowledgements

The authors thank the management of the Vellore Institute of Technology, Vellore for the encouragement and support and Dr. Anand Prem Rajan, SBST, and VIT for providing a fluorescence spectrophotometer facility.

Author contributions

DP: Methodology, Material preparation, Performed the experiments, Data curation, Data analysis, Software, Validation, and Writing-Original draft. CS: Conceptualization, Experimental designing, Resources, Supervision, Writing- Review and editing and Funding acquisition.

Funding

Not Applicable.

Declarations

Conflict of interest

The authors declare that they have no conflicts of interest.

Footnotes

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

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