Physical and Oxidative Stability of Emulsions Stabilized with Fractionated Potato Protein Hydrolysates Obtained from Starch Production Side Stream

Citation: Yesiltas, B.; García-Moreno, P.J.; Mikkelsen, R.K.; Echers, S.G.; Hansen, D.K.; Greve-Poulsen, M.; Hyldig, G.; Hansen, E.B.; Jacobsen, C. Physical and Oxidative Stability of Emulsions Stabilized with Fractionated Potato Protein Hydrolysates Obtained from Starch Production Side Stream. Antioxidants 2023,12, 1622. https://doi.org/ 10.3390/antiox12081622 Academic Editor: Joaquin Velasco Received: 30 June 2023 Revised: 4 August 2023 Accepted: 12 August 2023 Published: 16 August 2023 Copyright: © 2023 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/). antioxidants Article Physical and Oxidative Stability of Emulsions Stabilized with Fractionated Potato Protein Hydrolysates Obtained from Starch Production Side Stream Betül Yesiltas 1,* , Pedro J. García-Moreno 2, Rasmus K. Mikkelsen 1, Simon Gregersen Echers 3, Dennis K. Hansen 4, Mathias Greve-Poulsen 5, Grethe Hyldig 1, Egon B. Hansen 1and Charlotte Jacobsen 1,* 1National Food Institute, Technical University of Denmark, 2800 Kgs. Lyngby, Denmark; [email protected] (R.K.M.); [email protected] (G.H.); [email protected] (E.B.H.) 2Department of Chemical Engineering, University of Granada, 18012 Granada, Spain; [email protected] 3Department of Chemistry and Bioscience, Aalborg University, 2450 Copenhagen, Denmark; [email protected] 4Lihme Protein Solutions, 2750 Ballerup, Denmark; [email protected] 5KMC Amba, 7330 Brande, Denmark; [email protected] *Correspondence: [email protected] (B.Y.); [email protected] (C.J.) Abstract: This work studies the emulsifying and antioxidant properties of potato protein hydrolysates (PPHs) fractions obtained through enzymatic hydrolysis of potato protein using trypsin followed by ultrafiltration. Unfractionated (PPH1) and fractionated (PPH2 as >10 kDa, PPH3 as 10–5 kDa, PPH4 as 5–0.8 kDa, and PPH5 as <0.8 kDa) protein hydrolysates were evaluated. Pendant drop tensiometry and dilatational rheology were applied for determining the ability of PPHs to reduce interfacial tension and affect the viscoelasticity of the interfacial films at the oil–water interface. Peptides >10 kDa showed the highest ability to decrease oil–water interfacial tension. All PPH fractions predominantly provided elastic, weak, and easily stretchable interfaces. PPH2 provided a more rigid interfacial layer than the other hydrolysates. Radical scavenging and metal chelating activities of PPHs were also tested and the highest activities were provided by the unfractionated hydrolysate and the fractions with peptides >5 kDa. Furthermore, the ability of PPHs to form physically and oxidatively stable 5% fish oil-in-water emulsions (pH 7) was investigated during 8-day storage at 20 ◦ C. Our results generally show that the fractions with peptides >5 kDa provided the highest physicochemical stability, followed by the fraction with peptides between 5 and 0.8 kDa. Lastly, promising sensory results with mostly mild attributes were obtained even at protein concentration levels that are higher than needed to obtain functional properties. The more prominent attributes (e.g., bitterness and astringency) were within an acceptable range for PPH3 and PPH4. Keywords: protein hydrolysate; ultrafiltration; emulsifying peptides; oil-in-water emulsions; antioxidant activity; sensory profile 1. Introduction Following the growing demand for sustainable food ingredients sourced from natural and sustainable origins, utilization of industrial side streams gained traction as they are often abundant in nutrients and contain bioactive and techno-functional compounds [ 1 ]. By harnessing these side streams, the food industry can address the demand while adhering to clean label criteria and considering consumer acceptance [ 2 ]. By re-incorporating side streams into food production, the industry can optimize resource utilization, minimize waste, and foster a more sustainable future, where the entire lifecycle of food is maximized. As an emerging application of alternative and natural food ingredients, plant-based protein hydrolysates were investigated for their functional properties in model oil-in-water emulsion systems to gather insight into their properties in real food emulsions [ 3 – 8 ]. Oxidatively prone oils, such as fish oil rich in omega-3 polyunsaturated fatty acids, are commonly used Antioxidants 2023,12, 1622. https://doi.org/10.3390/antiox12081622 https://www.mdpi.com/journal/antioxidants Antioxidants 2023,12, 1622 2 of 18 in these model systems to increase the rate of oxidation due to the bis-allylic carbons in their molecular structure. Peptides may potentially provide protection against lipid oxidation due to a combination of both surface activity and antioxidant properties. The former is attributed to their amphiphilic character and ability to interact with/adsorb the interface between oil and water in emulsions [ 9 , 10 ]. The antioxidant activity of peptides was suggested to relate to the content of specific amino acids that have the ability to scavenge free radicals (e.g., Cys, Met, and Try) or chelate metals (e.g., His, Glu, and Asp) [11,12]. Despite a modest protein content (1–2%), the magnitude of proteins obtained from the starch industry (~240,000 t/year) makes potatoes a highly relevant source of plant-based protein [ 13 ]. The identification and release of abundant proteins/peptides from plant-based sources gained significant attention from the food industry in recent years [ 9 , 14 ]. To meet the increasing demand for sustainable protein-based ingredients from alternative sources, the development of approaches combining advanced mass spectrometry-based proteomics and bioinformatics tools allowed for more time and cost-efficient research [ 15 , 16 ]. Using this approach, we previously identified several potent emulsifier peptides from abundant potato proteins [ 14 , 17 ]. While recent years showed tremendous progress in the fundamental understanding of peptide emulsification, most studies contributing to this were naturally performed on simple model systems and with individually isolated/synthesized peptides [ 9 , 10 , 18 , 19 ]. However, the use of purified peptides is not economically viable in the food industry, and thus, it is essential to further investigate and understand how hydrolysates from alternative protein sources may affect the physicochemical stability of oil-in-water emulsions. For instance, trypsin was determined the best candidate for targeted release of validated emulsifier peptides from potato protein and a tryptic hydrolysate was found to show superior emulsifying properties compared to the native protein and a range of hydrolysates obtained through conventional trial-and-error hydrolysis [20]. Previous studies reported that the size of the peptides contained in whey protein hydrolysates affects the emulsification ability; therefore, the degree of hydrolysis should be controlled [ 21 – 23 ]. In another study, sweet potato protein hydrolysates were obtained by Alcalase under high hydrostatic pressure, which significantly increased the degree of hydrolysis and the <3 kDa fraction content, thereby improving antioxidant activity [ 7 ]. Indeed, the size fractionation of the hydrolysates is an important factor to investigate both emulsifying and antioxidant activities in emulsion systems. Thus, this study aimed to investigate the emulsifying and antioxidant activities of a potato protein hydrolysate obtained by trypsin and its fractions obtained by ultrafiltration. First, the ability of the unfractionated hydrolysate and fractionated PPHs to reduce interfacial tension, as well as their viscoelasticity of the interfacial layer, was evaluated. Moreover, the radical scavenging and metal chelating activities of the PPHs were measured. In addition, we investigated the ability of PPHs to stabilize low-fat oil-in-water emulsions and control lipid oxidation by following emulsions’ physical and oxidative stability during 8 days of storage. Finally, the sensory properties of the PPHs were evaluated in a solution with a trained panel to map their sensory profile. 2. Materials and Methods 2.1. Materials Potato protein hydrolysate (PPH1) was provided by KMC AmbA (Brande, Denmark). PPH1 was produced by the hydrolysis of denatured potato protein (5 w/v%) in water using 0.05 w/v% Trypsin (Pancreatic Trypsin Novo, 6.0 S, 6.0 AU/g) (Novozymes A/S, Bagsværd, Denmark) at pH 8, 50 ◦ C for 24 h. The ratio between protein powder and Trypsin was 100:1 (w:w) . Hydrolysis was carried out in a free-fall pH mode and the change in the pH was tracked every hour. When the pH decreased to 6.6 after 7 h, it was adjusted to 7 and the final pH was noted as 7.45 after 24 h. The enzymatic hydrolysis was followed by enzyme inhibition, centrifugation (2759 × gfor 15 min) (Hermle Z 513 K, Wehingen, Germany), and freezing of the supernatant (PPH1) at − 18 ◦ C. The degree of hydrolysis of PPH1 was measured and reported as 3.2%. PPH1 was further fractionated into >10 kDa (PPH2), Antioxidants 2023,12, 1622 3 of 18 5–10 kDa (PPH3), 0.8–5 kDa (PPH4), and <0.8 kDa (PPH5) at Lihme Protein Solutions using sequential ultrafiltration with the following membranes: 10 kDa (K02-E010-05-S, 1.25 m 2 , Repligen, Waltham, MA, USA), 5 kDa (K02-E005-05-S, 1.25 m 2 , Repligen, Waltham, MA, USA), and 0.8 kDa (NFG-2B, 1812F, 0.37 m 2 , Synder, Vacaville, CA, USA). The pump used was KMPi with 8 L/min crossflow and 1 Bar differential pressure (Repligen, Waltham, MA, USA). The protein content of the unfractionated (PPH1) and fractionated (PPH2-PPH5) potato protein hydrolysates was determined by DUMAS method and the results are 2.37, 3.08, 1.28, 1.14, and 0.27 w/v% for PPH1, PPH2, PPH3, PPH4, and PPH5, respectively. Medium-chain triglycerides (MCT) oil (WITARIX ® MCT 60/40) was provided by OI Oleo GmbH (Hamburg, Germany), which was used in interfacial tension and dilatational rheology measurements. Cod liver oil was provided by Vesteraalen’s (Nordland, Norway), which was used in emulsions that were subjected to storage experiments. The peroxide value (PV) of the fish oil was 0.23 ± 0.00 meq. O 2 /kg oil. The fatty acid composition (%, w/w) of the fish oil was as follows: C14:0 (3.7), C16:0 (9.0), C16:1n-7 (9.6), C18:0 (1.9), C18:1n-9 (15.8), C18:1n-7 (4.5), C18:2n-6 (2.4), C18:3n-3 (1.0), C20:1n-9 (13.7), C20:5n-3 (8.4), C22:1n-11 (5.6), and C22:6n-3 (10.7), which was determined using GC analysis of fatty acid methyl esters (FAME) (AOCS official methods Ce2-66 and 1b-89). The tocopherol content of the fish oil was also analyzed and found as follows: α -tocopherol, 195 ± 3 µ g/g oil; β -tocopherol, 5 ± 0 µ g/g oil; γ -tocopherol, 116 ± 2 µ g/g oil; and δ -tocopherol, 43 ±1µg/g oil (AOCS Official Method Ce 8–89). 2.2. Methods 2.2.1. Characterization of the Fractionated PPHs Molecular Weight Distribution of Hydrolysate and Fractions PPHs were analyzed by size exclusion chromatography (FPLC) using an Äkta Pure chromatography system (GE Healthcare, Brondby, Denmark). A Tricorn 10/300 column was packed with Superdex 30 prep grade (GE Healthcare, Brondby, Denmark). The mobile phase used was 0.050 M potassium phosphate buffer, 0.100 M sodium chloride pH 7.5, at a flow rate of 0.500 mL/min. The absorbance was measured at the 280 nm wavelength. Interfacial Tension and Interfacial Dilatational Rheology The dilatational surface properties were analyzed using a drop tensiometer (OCA 25, Dataphysics Instruments GmbH, Filderstadt, Germany) at 20 ◦ C. A pendant droplet of the protein fractions with a surface area of 59 mm 2 was formed at the tip of the needle (diameter 1.83 mm) in MCT oil (Witarix MCT 60/40, IOI Oleo GmbH, Hamburg, Germany) contained in a quartz glass cuvette. The aqueous solutions (pH 7) were prepared at 0.1 g protein/L. The drop contour was monitored and fitted with the Young–Laplace–Gauss equation to obtain the interfacial tension. The change in interfacial tension was monitored for 1 h, followed by a frequency and amplitude sweep. The frequency sweep was performed with frequencies of 0.01, 0.0325, 0.055, 0.0775, and 0.1 Hz at an amplitude of 2.5%. The amplitude sweep was performed with amplitudes of 1, 2, 3, 4, and 5% at a frequency of 0.01 Hz. The droplet was subjected to five oscillatory cycles at every amplitude or frequency. Via oscillatory dilatational deformation experiments (compression and expansion), the complex surface dilatational modulus was calculated from the first harmonic of the Fourier spectrum of the oscillatory interfacial tension signal. The complex surface dilatational modulus is equal to E *d = E 0 d + iE 00 d , where E 0 d is the dilatational storage modulus (elastic modulus), and E 00 d is the dilatational loss modulus (viscous modulus). The loss tangent is defined as tanΦ= (E0 d/E00 d) [10]. The measurements were conducted in triplicate. In Vitro Antioxidant Activity The radical scavenging activity of predicted antioxidant peptides was measured using the DPPH radical scavenging activity method of Yang et al. [ 24 ] with some modifications. PPHs were dissolved in water and different concentrations (0.2–50 mM) of the PPH solutions were obtained. Equal parts (100 µ L) of 0.1 mM ethanolic DPPH solution and a Antioxidants 2023,12, 1622 4 of 18 peptide solution were mixed. The mixture was transferred to the microtiter plate, kept in darkness at room temperature for 30 min, and the absorbance was measured at 515 nm using a spectrophotometer. The scavenging effect was calculated as an inhibition percentage following Equation (1): DPPH inhibition (%) = (1 −(As −A0)/Ab) ×100, (1) where As is the sample absorbance (PPHs + DPPH), Ao is the sample blank absorbance (no DPPH), and Ab is the reagent blank absorbance (no PPHs). Results were calculated for 50% inhibition concentration (IC50) and presented in mg/mL. butylated hydroxy toluene (BHT) solution was included as a positive control. All measurements were performed in triplicate. The iron chelating activity of the PPHs was measured as described in Farvin et al. [ 25 ] with some modifications. PPHs were dissolved in water and different concentrations (0.2–50 mM) of PPH solutions were obtained. Concisely, 100 µ L of the antioxidant sample was transferred to the microtiter plate and 110 µ L of water was added. To start the reaction, 20 µ L of 0.5 mM ferrous chloride was added and the plate was shaken. After 3 min, 20 µ L of 2.5 mM ferrozine was added and the plate was shaken again. After 10 min in the dark at room temperature, the absorbance was measured at 562 nm. Ethylenediaminetetraacetic acid (EDTA) was employed as a positive control. The chelating capacity and IC50 values were calculated in the same way as for the DPPH inhibition described above. 2.2.2. Production and Physicochemical Stability of Oil-in-Water Emulsions Stabilized with the PPHs Emulsion Production A total of 250 g of fish oil-in-water emulsions (5 wt%) were stabilized with 0.2 wt.% protein of PPHs. PPH solutions were made in distilled water and kept in a stirrer in a refrigerator in darkness overnight to allow full hydration. The pH of the PPH solutions was checked and adjusted to 7 when necessary, using 1 M NaOH and 1 M HCl. Prehomogenization was performed using ultra turrax (Ystral, Ballrechten-Dottingen, Germany) for 3 min at 16,000 rpm. Fish oil was added into the aqueous phase within the first minute of mixing. Secondary homogenization was performed using a microfluidizer (M110L Microfluidics, Newton, MA, USA) equipped with a ceramic interaction chamber (CIXC, F20Y, internal dimension 75 µ m) at 9 kpsi pressure for three passes. Sodium azide (0.05%) and 50 µ M of FeSO 4 were added to the emulsion obtained after microfluidizer homogenization and it was stirred with a spoon. The final pH of the emulsion was measured. Emulsions were stored for 8 days at 20 ◦ C in darkness. Samples were collected for physical characterization and oxidative stability analyses during storage. Physical Stability of Emulsions (1) Creaming index Creaming index was followed throughout the storage on days 0, 1, 2, 4, and 8. The creaming index was calculated based on the equation below: creaming index (%) = (b/a) ×100, where a is the total height of the emulsion in the tube, and b is the height of the clear aqueous phase at the bottom of the tube. (2) Emulsion stability Emulsions were placed in the Turbiscan cabinet on the same day they were produced and scanned over days (Formulaction, Toulouse, France). The scanning program is shown in Table S1 in the Supplementary Material. Based on the scans, the Turbiscan stability index (TSI) is calculated by the computer. This calculation is based on an integrated algorithm Antioxidants 2023,12, 1622 5 of 18 that sums up the evolution of transmittance and backscattering light at every position (h) and based on the scan-to-scan difference over total sample height (h). TSI(t)=1 Nh tmax ∑ ti=1 zmax ∑ zi=zmin |BST(ti, zi)−BST(ti−1, zi)| where N h is the number of height position, t is the time, and z min and z max are the lower and upper height limit. BST is the signal that is taken into account (BS when T < 0.2%, otherwise T). The Turbiscan stability index (TSI) values above 3 are considered unstable while values below 1 are stable according to Formulaction’s webpage. Values in between are in the beginning of destabilization, but mostly non-visual. (3) Zeta potential The zeta potential was measured using a Zetasizer Nano ZS (Malvern Instruments, Ltd., Worcestershire, UK) on day 1. Samples were prepared by taking 80 µ L emulsion and diluting in 40 mL distilled water and mixing by a vortex. A DTS-1070 disposable folded capillary cell (Malvern Instruments, Ltd., Malvern, UK) was used for loading the samples. Measurements from the same sample were carried out in triplicate using a zeta potential range of (−) 100 to (+) 50 mV, and samples were analyzed with 100 runs at 25 ◦C. (4) Droplet size The droplet size distribution of the emulsions was measured using the Mastersizer 2000 (Malvern Instruments, Ltd., Worcestershire, UK) based on the laser diffraction technique on days 0 and 8 of the storage experiment. Emulsions were diluted in recirculating water set at 3000 rpm until an obscuration of approximately 12–15% was reached. For particle and dispersant, the refractive indices of sunflower oil (1.469) and water (1.330) were used, respectively. Measurements of droplet size were carried out in duplicates or triplicates and they were given as the surface-weighted (D[3,2]) and volume-weighted (D[4,3]) mean diameters. Oxidative Stability of Emulsions Oxidative stability analyses were carried out on the samples collected on days 0, 1, 2, 5, and 8 during the storage experiment. (1) Peroxide value Lipids were extracted using the method described by Bligh and Dyer [ 26 ] using a reduced amount of chloroform/methanol (1:1, w/w). Two extractions were made from each emulsion sample. Peroxide value (PV) was determined on lipid extracts using the colorimetric ferric thiocyanate method at 500 nm according to Shantha and Decker [ 27 ] with a spectrophotometer (Shimadzu UV-1280, Holm&Halby, Brøndby, Denmark). Measurements were performed in duplicate. (2) Tocopherol consumption Tocopherols were determined using an Agilent 1100 series HPLC (Agilent Technologies, Palo Alto, CA, USA), equipped with a fluorescence detector. Lipid extracts (2 g) from the above-mentioned Bligh and Dyer extraction were weighed and evaporated under nitrogen. Then, lipids were dissolved in 10 mL n-heptane. Heptane solutions (1 mL) were taken into separate vials before injection of an aliquot on a Spherisorb S5 W column ( 250 ×4.6 mm ) (Phase Separation Ltd., Deeside, UK). Elution was performed with an isocratic mixture of n-heptane and 2-propanol (100:0.4, v:v) at a flow of 1 mL/min injection volume of 20 µ L in a column (Waters Spherisorb 3 µ m Silica, 4.6 mm I.D. × 150 mm), preceded by a guard column (Waters Spherisorb, 5 µ m Silica, 4.6 mm I.D. × 10 mm). A fluorescence detector with excitation wavelength at 290 nm and emission wavelength at 330 nm was used according to the AOCS method [ 28 ]. A tocopherol standard mix including α , β , γ , and δ -tocopherol standards was used (Calbiochem 613424). The external quantitative standard was used for calculations. Measurements were performed in duplicate. Antioxidants 2023,12, 1622 6 of 18 (3) Volatile compound formation using dynamic headspace GC-MS Four grams of emulsion, 5 mL distilled water, and 30 mg of an internal standard (4-methyl-1-pentanol) were placed in a purge bottle. The purge bottle was subjected to a water bath at 45 ◦ C for 30 min under purging with nitrogen (150 mL/min) and the volatile compounds were trapped in Tenax GR tubes. Afterward, the tubes were put into an Automatic Thermal Desorber (ATD-350, Perkin Elmer, Norwalk, CN), which was connected to gas chromatography (GC Agilent 6890 N, Palo Alto, CA, USA; Column: DB-1701, 30 m ×0.25 mm ×1.0 µm ). The oven program had an initial temperature of 45 ◦ C for 5 min, increasing with 1.5 ◦ C/min until 55 ◦ C, with 2.5 ◦ C/min until 90 ◦ C, and with 12.0 ◦ C/min until 220 ◦ C, where the temperature was kept for 4 min. A mass spectrometer (MS Agilent 5973 , Agilent Technologies, USA; electron ionization mode, 70 eV; mass to charge ratio scan between 30 and 250) was used to separate the individual volatile compounds and these compounds were identified by MS library searches (Wiley 138 K, John Wiley and Sons, Hewlett-Packard, New York, NY, USA). The following standards were selected for quantification: 2-ethyl furan, 1-penten-3-one, pentanal, 1-penten-3-ol, 1-pentanol, hexanal, (E)-2-hexenal, heptanal, (E)-2-heptenal, (Z)-4-heptenal, octanal, (E,E)-2,4-hexadienal, (E,E)-2,4-heptadienal , (E)-2-nonenal, and (E,E)-2,4-decadienal. An ethanolic solution of the standards was prepared and diluted into nine levels (0.1–250 µ g/mL) for the calibration curve. Each dilution (30 mg) was added into an emulsion produced in the same way as in one of the emulsions (PPH2), which allows a similar release for standards, as it was for volatile compounds formed in all emulsions. Analysis was run in triplicate. 2.2.3. Sensory Analysis of the PPHs The sensory evaluation was performed by a tested and trained objective sensory panel at DTU Food. The panel was specifically trained in objective descriptive analysis and the sensory evaluations were performed in a sensory lab with separated booths under normal daylight and at ambient temperature according to the standards and guidelines for the design and construction of sensory lab to ISO 13300-25492 (2006); ISO 3972 (2011) and ISO 3972 (2011)/Cor 1 (2012); 8586 (2012); and ISO 8589, 2007; NMKL Procedure No. 6, 2023 (Table S3, Supplementary Material). The first sessions were used to develop a vocabulary to describe the sensory characteristics of the samples. The next three sessions were used to train the sensory panel in measuring and scoring the intensity of the selected sensory attributes. For the training, a set of samples with the different attributes and intensities of the attributes was used to ensure that all experimental conditions were represented. The scale was an unstructured 15 cm line scale with an anchor point at 1.5 cm and 13.5 cm. The samples were poured in plastic cups with lids and stored at 20 ◦ C for one hour before serving. All samples were served in three replicates and in random order with three digital numbers and the assessors were served neutral flat bread and water to clean their mouths between samples. The final vocabulary was defined in four categories as listed here: appearance: transparent, color (light/dark); odor: sweet, floral-like, sourish; flavor: sweet, earthy, potato, salty, green, cold tea, umami, bitter; texture: sticky, astringency, and viscous. 2.2.4. Statistical Analysis Mean value and standard deviations were introduced to Statgraphics 18 (Statistical Graphics Corp., Rockville, MD, USA) for the data analysis. Multiple sample comparison was performed to identify the significant differences between samples at certain sampling days and between sampling days for each sample during storage using Tukey as a post hoc test at p< 0.05 significance level. Antioxidants 2023,12, 1622 7 of 18 3. Results and Discussion 3.1. Characterization of the Unfractionated Hydrolysate and Their Fractions Obtained by Ultrafiltration 3.1.1. Molecular Weight Distribution The retention volume of the peptides by size exclusion increases from PPH2, PPH3, PPH4, to PPH 5, owing to decreasing peptide sizes (Figure 1). This is in accordance with the decreasing molecular weight cut-off of the membranes that they were retained on during the sequential filtration (10 kDa, 5 kDa, and 0.8 kDa permeate, respectively). PPH1, unfractionated PPH, spans the complete range seen in the fractionated samples. Larger molecules, with a retention volume of less than 11 mL, were exclusively retained in the PPH2 sample. Antioxidants 2023, 12, x FOR PEER REVIEW 7 of 19 2.2.4. Statistical Analysis Mean value and standard deviations were introduced to Statgraphics 18 (Statistical Graphics Corp., Rockville, MD, USA) for the data analysis. Multiple sample comparison was performed to identify the significant differences between samples at certain sampling days and between sampling days for each sample during storage using Tukey as a post hoc test at p < 0.05 significance level. 3. Results and Discussion 3.1. Characterization of the Unfractionated Hydrolysate and Their Fractions Obtained by Ultrafiltration 3.1.1. Molecular Weight Distribution The retention volume of the peptides by size exclusion increases from PPH2, PPH3, PPH4, to PPH 5, owing to decreasing peptide sizes (Figure 1). This is in accordance with the decreasing molecular weight cut-off of the membranes that they were retained on during the sequential filtration (10 kDa, 5 kDa, and 0.8 kDa permeate, respectively). PPH1, unfractionated PPH, spans the complete range seen in the fractionated samples. Larger molecules, with a retention volume of less than 11 mL, were exclusively retained in the PPH2 sample. Figure 1. Fast protein liquid chromatography results from potato protein hydrolysates (PPHs). The absorbance was measured at a 280 nm wavelength. PPH1 to PPH5 denotes: unfractionated PPH, >10 kDa, 5–10 kDa, 0.8–5 kDa, and <0.8 kDa, respectively. In contrast, lower-mass peptides (i.e., a retention volume in the range 20–25 mL) seem to be distributed across the different fractions, indicating that lower-mass peptides reside in the retentate post ultracentrifugation. To facilitate more efficient fractionation by size, samples may be passed through the filters with multiple passes by washing in a diafiltration-like manner. However, this would come at the expense of sample dilution and impose substantial additional costs if adapting the process to full-scale production. 3.1.2. Interfacial Tension and Interfacial Dilatational Rheology Emulsifying peptides may reduce interfacial tension by (i) diffusing from the bulk aqueous phase to the interface, (ii) adsorbing at the oil/water interface, and (iii) rearranging their structure at the interface to project their hydrophobic residues to the oil phase [16]. Figure 2a shows the reduction in the interfacial tension of the potato protein 0 50 100 150 200 250 300 0 5 10 15 20 25 30 35 Absorbance at 280 nm (mAU) Elution volume (ml) PPH1 PPH2 PPH3 PPH4 PPH5 Figure 1. Fast protein liquid chromatography results from potato protein hydrolysates (PPHs). The absorbance was measured at a 280 nm wavelength. PPH1 to PPH5 denotes: unfractionated PPH, >10 kDa, 5–10 kDa, 0.8–5 kDa, and <0.8 kDa, respectively. In contrast, lower-mass peptides (i.e., a retention volume in the range 20–25 mL) seem to be distributed across the different fractions, indicating that lower-mass peptides reside in the retentate post ultracentrifugation. To facilitate more efficient fractionation by size, samples may be passed through the filters with multiple passes by washing in a diafiltration-like manner. However, this would come at the expense of sample dilution and impose substantial additional costs if adapting the process to full-scale production. 3.1.2. Interfacial Tension and Interfacial Dilatational Rheology Emulsifying peptides may reduce interfacial tension by (i) diffusing from the bulk aqueous phase to the interface, (ii) adsorbing at the oil/water interface, and (iii) rearranging their structure at the interface to project their hydrophobic residues to the oil phase [ 16 ]. Figure 2a shows the reduction in the interfacial tension of the potato protein hydrolysates studied (PPH1) and their fractions obtained by ultrafiltration (PPH2-PPH5). It was observed that the bulk hydrolysate (PPH1) notoriously reduced the interfacial tension during the first 10 min, leveling off at a value of 20.9 ± 0.4 mN/m after 60 min. An identical trend to that observed for the bulk hydrolysate was also found for the hydrolysate fraction PPH4 ( 5–0.8 kDa ). On the other hand, the hydrolysate fractions PPH3 (10–5 kDa) and PPH5 (<0.8 kDa) reduced interfacial tension to a significantly (p< 0.05) lower extent when compared to PPH1, reaching a value of 22.9 ± 0.3 mN/m after 60 min. Interestingly, the hydrolysate fraction PPH2 (>10 kDa) led to an initial significantly (p< 0.05) faster decrease in interfacial tension when compared to PPH1, leveling off after 25 min and reaching a value of 17.7 ±0.4 mN/m after 60 min. This finding is in agreement with Antioxidants 2023,12, 1622 8 of 18 previous studies reporting that limited protein hydrolysis (degree of hydrolysis, DH, of 3–6%), which results in larger peptides, favored the emulsifying properties of the obtained hydrolysates [ 20 – 23 , 29 ]. In fact, PPH2 presents larger peptides than the rest of the fractions with higher potential to present large hydrophobic patches within their structure. These large hydrophobic patches are key to the diffusion and later adsorption of the peptides at the oil/water interface [ 14 , 29 ]. In any case, it should be noted that the interfacial tension values obtained for the potato protein hydrolysate, or its fractions obtained by ultrafiltration were higher than those previously reported for potato emulsifying peptides [ 10 , 14 ]. This finding is ascribed to the complex nature of a hydrolysate and its fractions, where not all peptides present have amphiphilic properties. Interfacial dilatational rheology allows the characterization of interfacial layers, providing information on their viscoelasticity, structure, and intermolecular connections [ 30 ]. The complex dilatational modulus (E *d = E 0 d + iE 00 d ) determines the resistance of the interfacial layer to external perturbations of its equilibrium state [ 31 ]. The elastic part (E 0 d ) of the complex dilatational modulus indicates the recoverable energy stored in the interface, whereas the viscous part (E 00 d ) represents the energy lost through relaxation processes [ 32 ]. The hydrolysate and all the ultrafiltration fractions formed predominantly elastic interfacial layers at the tested frequencies and amplitudes, having values of E 0 d markedly larger than E 00 d as well as low values of phase angles ( Φ≤ 26 ◦ ). This finding denotes that all the peptide-based interfacial layers evaluated showed a gel-like structure with peptide–peptide interactions. The complex dilatational modulus observed for the interfaces studied ranged from 3.5 ± 0.9 to 33.2 ±1.5 mN/m (Figure 2b,c), which is in line with previously reported values for whey and fish protein hydrolysates [ 21 , 31 ]. All the interfacial layers were amplitude-independent in the range 1–5%, denoting that they were within the linear viscoelastic regime (LVE). Moreover, the lack of change in the interfacial moduli with increasing amplitude (Figure 2b) suggests weak and easily stretchable interfaces that did not suffer a disruption in their microstructure in this amplitude range [10]. Figure 2c shows that the elasticity of all the interfaces, but particularly PPH2, PPH1, and PPH4, increased when increasing frequency. This is due to the shorter time needed to adapt to the deformation via the diffusional exchange of peptides with the aqueous phase or relaxation processes within the network [ 31 ]. Interestingly, Figure 2b,c shows that PPH2 formed the most rigid interfacial layer (e.g., highest E *d ), followed by the unfractionated hydrolysate (PPH1) and PPH3-PPH4, with PPH5 presenting the lowest E *d (Figure 2b,c). Indeed, PPH2 (>10 kDa) presented large peptides and unhydrolyzed proteins that re-arrange at the interface, facilitating intermolecular connections that result in an interface with higher elasticity and strength [ 21 ]. On the contrary, low molecular weight peptides present in PPH5 (<0.8 kDa) were not able to interconnect at the interface, leading to the weakest interface. Antioxidants 2023, 12, x FOR PEER REVIEW 8 of 19 hydrolysates studied (PPH1) and their fractions obtained by ultrafiltration (PPH2-PPH5). It was observed that the bulk hydrolysate (PPH1) notoriously reduced the interfacial tension during the first 10 min, leveling off at a value of 20.9 ± 0.4 mN/m after 60 min. An identical trend to that observed for the bulk hydrolysate was also found for the hydrolysate fraction PPH4 (5–0.8 kDa). On the other hand, the hydrolysate fractions PPH3 (10–5 kDa) and PPH5 (<0.8 kDa) reduced interfacial tension to a significantly (p < 0.05) lower extent when compared to PPH1, reaching a value of 22.9 ± 0.3 mN/m after 60 min. Interestingly, the hydrolysate fraction PPH2 (>10 kDa) led to an initial significantly (p < 0.05) faster decrease in interfacial tension when compared to PPH1, leveling off after 25 min and reaching a value of 17.7 ± 0.4 mN/m after 60 min. This finding is in agreement with previous studies reporting that limited protein hydrolysis (degree of hydrolysis, DH, of 3–6%), which results in larger peptides, favored the emulsifying properties of the obtained hydrolysates [20–23,29]. In fact, PPH2 presents larger peptides than the rest of the fractions with higher potential to present large hydrophobic patches within their structure. These large hydrophobic patches are key to the diffusion and later adsorption of the peptides at the oil/water interface [14,29]. In any case, it should be noted that the interfacial tension values obtained for the potato protein hydrolysate, or its fractions obtained by ultrafiltration were higher than those previously reported for potato emulsifying peptides [10,14]. This finding is ascribed to the complex nature of a hydrolysate and its fractions, where not all peptides present have amphiphilic properties. (a) 0 5 10 15 20 25 30 010 20 30 40 50 60 IFT, mN/m Time, min PPH2 PPH4 PPH1 PPH3 PPH5 Figure 2. Cont. Antioxidants 2023,12, 1622 9 of 18 Antioxidants 2023, 12, x FOR PEER REVIEW 9 of 19 (b) (c) Figure 2. Interfacial properties of PPHs: (a) Interfacial tension as a function of time, (b) complex surface dilatational modulus (E*) as a function of amplitude (1–5%, frequency: 0.01 Hz), and (c) complex surface dilatational modulus (E*) as a function of frequency (0.005 to 0.1 Hz, amplitude: 2.5%). Measurements were carried out for 0.1 g protein/L PPHs aqueous solutions (pH 7) at 20 °C using MCT oil. The bare MCT oil–water interfacial tension was 26 mN/m. PPH1 to PPH5 denotes: unfractionated PPH, >10 kDa, 5–10 kDa, 0.8–5 kDa, and <0.8 kDa, respectively. Interfacial dilatational rheology allows the characterization of interfacial layers, providing information on their viscoelasticity, structure, and intermolecular connections [30]. The complex dilatational modulus (E*d = E’d + iE”d) determines the resistance of the interfacial layer to external perturbations of its equilibrium state [31]. The elastic part (E’d) of the complex dilatational modulus indicates the recoverable energy stored in the interface, whereas the viscous part (E”d) represents the energy lost through relaxation processes [32]. The hydrolysate and all the ultrafiltration fractions formed predominantly elastic interfacial layers at the tested frequencies and amplitudes, having values of E’d markedly larger than E”d as well as low values of phase angles (Φ ≤ 26°). This finding denotes that all the peptide-based interfacial layers evaluated showed a gel-like structure with peptide–peptide interactions. The complex dilatational modulus observed for the interfaces studied ranged from 3.5 ± 0.9 to 33.2 ± 1.5 mN/m (Figure 2b,c), which is in line with previously reported values for whey and fish protein hydrolysates [21,31]. All the 0 5 10 15 20 25 1 2 3 4 5 E*, mN/m Amplitude, % PPH1 PPH2 PPH3 PPH4 PPH5 0 5 10 15 20 25 30 35 0.01 0.1 E*, mN/m Frequency, Hz PPH1 PPH2 PPH3 PPH4 PPH5 Figure 2. Interfacial properties of PPHs: ( a ) Interfacial tension as a function of time, ( b ) complex surface dilatational modulus (E * ) as a function of amplitude (1–5%, frequency: 0.01 Hz), and ( c ) complex surface dilatational modulus (E * ) as a function of frequency (0.005 to 0.1 Hz, amplitude: 2.5%). Measurements were carried out for 0.1 g protein/L PPHs aqueous solutions (pH 7) at 20 ◦ C using MCT oil. The bare MCT oil–water interfacial tension was 26 mN/m. PPH1 to PPH5 denotes: unfractionated PPH, >10 kDa, 5–10 kDa, 0.8–5 kDa, and <0.8 kDa, respectively. 3.1.3. In Vitro Antioxidant Activity Based on the DPPH radical scavenging activity results, PPH2 exhibited a significantly lower IC50 value compared to PPH1, PPH3, and PPH4 (Figure 3). PPH5 was found to be the least effective to scavenge radicals. On the other hand, based on the ferrozine assay, PPH3 was better at chelating metals followed by PPH2, PPH1, PPH5, and PPH4 (Figure 3). This is a different trend compared to the radical scavenging activity of the same PPHs, which could be attributed to the impact of peptide size on their ability to act as antioxidants via different mechanisms. In a previous study, peptide size was reported to affect the potency of peptides to retard lipid oxidation, where shorter peptides are generally attributed to higher scavenging and lipid peroxidation inhibition activities compared to long-chain peptides [ 16 ]. Ren et al. outlined that peptide chains containing <3 kDa peptides were more effective scavengers of hydroxyl radicals and inhibitors of lipid peroxidation when compared to chains containing >3 kDa peptides [ 33 ]. Therefore, it can be inferred that Antioxidants 2023,12, 1622 16 of 18 mitigating the bitter taste and ensuring the acceptability of food products incorporating protein hydrolysates. 4. Conclusions Emulsions produced with unfractionated and size-fractionated potato protein hydrolysates (PPHs) obtained by targeted hydrolysis with trypsin provided physiochemically stable emulsions. Different fractions showed differences in their in vitro antioxidant activity in terms of radical scavenging and metal chelating activities, where PPH2 (>10 kDa) had the best radical scavenging activity and PPH3 (5–10 kDa) had the best metal chelating activity. PPH2 (>10 kDa) had significantly stronger peptide–peptide interactions leading to the formation of more rigid and stable oil–water interfacial layers. Overall, emulsions containing PPHs higher than 5 kDa resulted in higher physical and oxidative stability during 8-day storage, indicating that the size fractionation influences techno-functional properties. This is crucial in the utilization of these ingredients as the fractions can be selected based on their best performance in the required functional property as antioxidants, emulsifiers, or both. Promising sensory results were also obtained at a higher protein concentration than needed to obtain expected functional properties. Supplementary Materials: The following supporting information can be downloaded at: https: //www.mdpi.com/article/10.3390/antiox12081622/s1, Table S1: Scanning program for the Turbiscan; Table S2: Statistical analysis for the (a) formation of hydroperoxides; (b) consumption of α -tocopherols; (c) development of 1-penten-3-ol and (d) development of (E,E)-2,4-heptadienal; Figure S1 : The development of volatile compounds during 8 days of storage; Table S3: References for sensory analysis. Author Contributions: Conceptualization, B.Y., P.J.G.-M. and C.J.; methodology, B.Y., P.J.G.-M., D.K.H., M.G.-P., G.H. and C.J.; validation, B.Y., P.J.G.-M., R.K.M. and G.H.; formal analysis, B.Y., P.J.G.-M., R.K.M., D.K.H., M.G.-P. and G.H.; investigation, B.Y., P.J.G.-M., R.K.M., D.K.H., M.G.-P. and G.H.; data curation, B.Y., P.J.G.-M., R.K.M. and G.H.; writing—original draft preparation, B.Y. and P.J.G.-M.; writing—review and editing, B.Y., P.J.G.-M., R.K.M., S.G.E., D.K.H., M.G.-P., G.H. and C.J.; visualization, B.Y., P.J.G.-M. and G.H.; supervision, B.Y. and C.J.; project administration, E.B.H. and C.J.; funding acquisition, E.B.H. and C.J. All authors have read and agreed to the published version of the manuscript. Funding: This research was funded by Innovation Fund Denmark, grant number 7045-00021B (PROVIDE Project). Data Availability Statement: Data is available on request. 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