The Structural Modification of Jackfruit Leaf Proteins (Artocarpus heterophyllus Lam.) by High-Intensity Ultrasound Alters Their Techno-Functional Properties and Antioxidant Capacity

Featured Application

High-intensity ultrasound is a powerful tool to improve the solubility, foaming, and emulsifying properties of modified proteins, making them suitable for emulsions or foam stabilizers, with added antioxidant benefits.

Abstract

(1) Background: Jackfruit leaves (Artocarpus heterophyllus Lam.) are rich in proteins but are under-utilized in the food industry due to their poor sensory properties and low solubility. High-intensity ultrasound (HIU) can enhance protein solubility by disrupting internal interactions and altering protein structures, making them more suitable for industrial applications. (2) Methods: This study aimed to modify the structure of jackfruit leaf proteins using HIU at different powers (600, 840, and 1080 W) and treatment times (10, 15, and 20 min). This research also characterized the amino acid composition and the techno-functional and antioxidant properties of the modified proteins. (3) Results: The HIU treatments significantly improved the foaming capacity and enhanced the emulsion stability within the proteins treated at 1080 W for 15 min, which showed a monomodal size distribution profile. Additionally, the modified proteins exhibited a higher antioxidant capacity compared to the native protein. (4) Conclusions: These findings suggest that structurally modified proteins from jackfruit leaves could be directly utilized in the formulation of emulsions or as foam stabilizers, offering added benefits to consumers due to their significant antioxidant properties.

1. Introduction

In recent years, the exploration of natural resources for the development of functional ingredients has gained attention due to their potential health benefits, the promotion of the development of new products, and sustainable sourcing. Plant-derived proteins have emerged as promising candidates among these resources due to their diverse functionalities and biocompatibilities. Jackfruit (Artocarpus heterophyllus Lam.) is a tropical fruit that is native to South and Southeast Asia. Thus, the cultivar has garnered considerable interest for its succulent fruit and for its leaves, which possess valuable compounds, including proteins [1,2]. The pruning of jackfruit trees is a common activity performed to improve the production and development of the fruits. This activity can generate between 5 and 30 kg of leaves per tree, depending on the season and the geographical zone. It is particularly affected by the tree’s age, which increases in the maturity stage [3]. Proteins are one of the main valuable components of jackfruit leaves, with the protein contents ranging from 13 to 28% on a dry basis [4].

Jackfruit leaves have been traditionally used in various cultures for their antimicrobial, anti-inflammatory, and wound-healing effects. However, at the industrial scale, there is no evidence of their use, even considering that recent studies have highlighted their potential as a source of protein [2,4]. This could be related to their low solubility and techno-functional properties. This makes jackfruit leaves a promising feedstock for protein production, which could be beneficial in addressing nutritional needs [2,3]. As the demand for alternative protein sources grows, particularly in the context of food security and environmental sustainability, jackfruit leaves offer a renewable resource that can be utilized without depleting other food sources. They are rich in proteins, which contain several EAAs, potentially serving as an economical alternative to conventional protein sources [2,3].

Traditional protein extraction and processing methods often involve harsh conditions, resulting in denaturation and the loss of bioactivity [4,5]. Therefore, there is a growing interest in exploring novel techniques to modify the structure of plant proteins while preserving or improving their inherent functionalities [5,6,7]. In this sense, the enzymatic and physical extractions including ultrasound-assisted extraction (UAE) and high hydrostatic pressure have been assessed in this regard [2,5,6,7].

High-intensity ultrasound (HIU) is a non-thermal processing technology that has gained traction in the food industry for its ability to induce physical and chemical changes in biomolecules without the need for high temperatures or chemical additives [8,9]. By subjecting protein solutions to intense acoustic waves, HIU can disrupt molecular interactions, leading to the unfolding and reconfiguration of protein structures. This can result in amino acid modification, improving the proteins’ solubility, antioxidant, and emulsifying properties, thus enhancing the techno-functional properties of the modified proteins [8,9,10]. The application of HIU for the modification of plant proteins has been investigated in various botanical sources, including soy, pea, and wheat [11,12,13]. HIU offers several advantages over conventional UAE in the obtention of modified proteins. The intense acoustic waves generated by HIU promote the rapid and efficient disruption of protein structures, leading to faster protein modification [14,15]. This results in a higher degree of protein hydrolysis, producing modified proteins with increased bioactivity and functional properties [16]. Furthermore, unlike conventional UAE, HIU requires a lower energy input to achieve the desired results [8,16]. These benefits make HIU a promising technology to produce modified proteins with enhanced functional and bioactive properties [8,9,13]. To our knowledge, its potential for enhancing the properties of proteins in jackfruit leaves remains unexplored. Therefore, this study aims to obtain modified proteins from jackfruit leaf proteins by performing HIU at different powers and times and to characterize their solubility, emulsification, and antioxidant properties.

2. Materials and Methods

2.1. Vegetal Material

The jackfruit leaves were handpicked after tree pruning in the “Tierras Grandes” orchard in Zacualpan, Compostela, Nayarit, Mexico (21.248889, −105.166944) in May 2022. Then, the leaves were placed into polyethylene bags and transported to the laboratory. After washing, the leaves were dried in a convective drying oven (Novatech, HS60-AID, Tlaquepaque, Mexico) for 24 h at 60 °C. The dried material was ground by using a high-speed blender (NutriBullet^® SERIE 900, Los Angeles, CA, USA) and sieved (150 μm diameter). The powder (flour) was vacuum-packed and stored until use [2].

2.2. Chemical Substances

The N-tert-butyldimethylsilyl-N-methyltrifluoroacetamide (MTBSTFA, >97%); L-norleucine; amino acid standards; acetonitrile (ACN); potassium persulfate; 2,2′-azinobis-(3-ethylbenzothiazoline-6-sulfonic acid (ABTS⁺); 6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid (Trolox); Bradford reagent; and sodium dodecyl sulfate (SDS) were obtained from Sigma-Aldrich (St. Louis, MO, USA). The analytical grade chemicals, such as NaOH and HCl, were obtained from Thermo Fisher Scientific Inc.

2.3. Protein Ultrasound-Assisted Extraction (UAE) and the Modification of Protein by High-Intensity Ultrasound (HIU)

A depigmentation procedure and the protein extraction were performed according to Vera-Salgado et al. [6] using a water bath (22–25 °C). The temperature of the samples was recorded before and after the HIU treatments. The extracted proteins were stored at 4 °C until the HIU treatment. For the HIU treatments, 6 g of the obtained leaf protein concentrate (LPC) was dispersed in 100 mL of deionized water by stirring. The dispersion was processed by using a sonicator FSI-1200 (Shanghai, China), which was equipped with a 16 mm diameter probe, at 24 KHz at 600, 840, and 1080 W power for 10, 15, and 20 min. The mixtures were stored at 4 °C until use.

2.4. Amino Acid Profile by Gas Chromatography–Mass Spectrometry (GC-MS)

The amino acid determination was carried out following the protocol proposed by Brion-Espinosa et al. [5]. Samples of LPC that were treated or untreated by HIU underwent acid hydrolysis with 6 M HCl for 24 h at 110 °C. Subsequently, samples were reacted with MTBSTFA (N-tert-Butyldimethylsilyl-N-methyltrifluoroacetamide), a GC derivatization reactant. In a nutshell, 100 µL of the sample and 10 µL of L-norleucine (internal standard, 0.2 mg/mL in 0.1 M HCl) were dried using nitrogen gas. The residue was dissolved in 200 µL of acetonitrile, and 200 µL of MTBSTFA was added. The solution was incubated in a glycerol bath at 100 °C for 2.5 h. The derivatization procedure was carried out in a 2 mL screw-capped vial with a PTFE lining. A GC-MS analysis was performed using an Agilent 7890A connected to an MS 240 Ion Trap (Agilent Technologies; Palo Alto, CA, USA). An Agilent J&W VF-5ms capillary column (30 m × 0.25 mm, i.d., 0.25 μm film thickness) was utilized for the separation. Helium (99.99%) served as the carrier gas, at a flow rate of 2 mL/min. The oven temperature program was initiated at 150 °C and was ramped up to 280 °C in 2 min at a rate of 3 °C per min. At 260 °C, 2 μL of the solution was injected into the GC injector port using an autosampler in split mode (20:1). The MS parameters included an ionization energy of 70 eV, a setting of full scan mode (35–650 m/z), an ion trap at 150 °C, a manifold at 80 °C, and a transfer line at 130 °C. For a combination of straight-chain alkanes (C7–C30) that were injected under the same analytical circumstances, linear retention indices were computed. In grams of amino acid per 100 g of material, the amino acid profile was given. The same procedure was followed for the L-amino acids standards.

2.5. Techno-Functional Properties Characterization

2.5.1. Loss of Hydrophobicity (Solubility)

The samples (33 μL) of LPC that were untreated or treated by HIU were placed in a 3 mL conical tube and 1 mL of Bradford reagent was added. Then, the tubes were vortexed (Hettich MIKRO 220R, Hettich, Germany) for 30 s to dissolve the sample. The protein content was determined by the Bradford method [17]. After the solubilization of the sample, 1 mL of 0.5 N NaOH solution was added. The total protein content was determined and the loss of hydrophobicity (%) was calculated (Equation (1)).

$$\mathrm{Loss\; of\; hydrophobicity}\left(\%\right)={\displaystyle \frac{P_{IP}}{P_{\mathit{Total}}}}·100$$

(1)

where P_IP is the weight of insolubilized protein and P_Total is the initial total weight of the protein sample.

2.5.2. Foaming Properties

The foaming capacity (FC) and foaming stability (FS) of the LPC samples that were treated or untreated were determined according to the methodology used by Calderón-Chiu et al. [2]. A sample aliquot (6 mL) of a dispersion protein (1%, w/v in deionized water) was placed in a conical tube (15 mL) and homogenized (16,000 rpm, 2 min, 25 °C) by using Ultra-Turrax (IKA T10, Staufen, Germany) to incorporate air bubbles. Then, the total solution was immediately transferred into a 15 mL glass graduated cylinder; after 30 s the total volume was enregistered and the FC (%) was calculated (Equation (2)).

$$FC\left(\%\right)={\displaystyle \frac{A_0-B}{B}}·100$$

(2)

where A₀ is the volume after homogenization (mL) and B is the volume before homogenization (mL).

For the the FS (%) determination, the previously homogenized sample was used. The samples were left at rest for 10 min, the volume was recorded, and the FS (%) was calculated (Equation (3)).

$$FS\left(\%\right)={\displaystyle \frac{At-B}{B}}·100$$

(3)

where At is the volume after rest (mL) and B is the volume before homogenization (mL).

2.5.3. Emulsifying Properties and Droplet Size Distribution

The analyses of the emulsifying properties were carried out by using the turbidimetric method. Briefly, in a 15 mL test tube, 2 mL of olive oil was mixed with 6 mL of the treated or untreated LPC samples in deionized water. Initially, the samples were homogenized using an Ultra-Turrax disperser at 10,000 rpm for 1 min. Then, 50 μL from the bottom of the tube was extracted after 0 and 10 min and diluted 100 times in a 0.1% SDS solution. The sample was stirred for 10 s on a magnetic stirring plate at 150 rpm. The absorbance was measured at 500 nm from the emulsion at the time (t) of 0 (A₀) and 10 min (A₁₀) by using a spectrophotometer (Cary 50 Bio UV–Visible, Varian, Mulgrave, Australia). The emulsifying activity index (EAI) and emulsion stability index (ESI) were determined by Equations (4) and (5), respectively [18].

$$EAI={\displaystyle \frac{2·2.303·DF·A_0}{c·f·10000}}$$

(4)

$$\mathit{ESI}={\displaystyle \frac{A_0}{A_0-A_{10}}}·t$$

(5)

where DF is the dilution factor (100), c is the mass of the sample (g), and f is the mass fraction of olive oil in the emulsion (0.25 g).

2.5.4. Size Distribution

The emulsion particle size distribution was measured using a Mastersizer 3000 (Hydro EV, Malvern, Worcestershire, UK). Briefly, the emulsion was added to the Hydro EV unit dropwise until it reached an 8–12% laser obscuration. For the dispersed phase (olive oil), the refractive index was adjusted to 1.46, and for the dispersant (water), it was set to 1.33. The dispersions were measured five consecutive times at 25 °C in the diffractometer. Mastersizer software (version 3.60, Worcestershire, UK), was used to characterize the particle size of each sample in terms of the volume distribution (%) vs. droplet diameter; the surface-weighted mean diameter (d_[3,2]; Equation (6)); the volume-weighted mean particle diameter (d_[4,3]; Equation (7)); and the polydispersity index (PDI; Equation (8)).

$$d_{\left[\mathrm{3,2}\right]}={\displaystyle \frac{\sum n_i{d}_i^3}{\sum n_i{d}_i^2}}$$

(6)

$$d_{\left[\mathrm{4,3}\right]}={\displaystyle \frac{\sum n_i{d}_i^4}{\sum n_i{d}_i^3}}$$

(7)

where $d_i$ is the droplet diameter and $n_i$ is the number of oil droplets having a diameter $d_i$.

$$PDI={\displaystyle \frac{d_{90}-d_{10}}{d_{50}}}$$

(8)

d₁₀, d₅₀, and d₉₀ are the diameters at 10, 50, and 90% of the cumulative droplet distribution, respectively [19].

2.5.5. Antioxidant Properties

To evaluate the antioxidant properties of jackfruit protein hydrolyzates, the 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS) assay was used [6,20]. The antioxidant properties were expressed as a percentage of the radical scavenging activity (%RSA, Equation (9)) and in the mg of Trolox equivalents per gram of the dry sample (mg TE/LPC). For the second stage of the analysis, Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid) was utilized for the calibration curve $y=0.0713x+0.8561; R^2=1.00$. Each sample was analyzed three times.

$$\%\mathrm{RSA}{\displaystyle \frac{A_{\mathit{Control}}-A_{\mathit{Sample}}}{A_{\mathit{Control}}}}\times 100$$

(9)

where A_Control is the absorbance of the ABTS⁺ solution and A_Sample represents the absorbance of the reaction (ABTS⁺ with sample).

Subsequently, curves representing the radical-scavenging activity (RSA, %) y-axis versus the sample concentration (mg/mL) x-axis were plotted for each sample. The corresponding point at 50% of the antioxidant activity with the x-intercept was defined as the IC₅₀ value. A linear regression equation of curves was used for this purpose [2].

2.6. Statistical Analysis

An experimental design (2³) was used to analyze the yield of the modified proteins. A one-way analysis of variance (ANOVA) and the post-hoc Tukey test for the mean comparison (p ≤ 0.05) were used for the data analyses of the techno-functional properties of the samples. The STATISTICA software performed all the tests (version 12.0, StatSoft, Inc., Tulsa, OK, USA, 2011).

3. Results and Discussion

3.1. The Yield of Protein and the Amino Acid Composition

The moisture (8.64%), protein (24.06%), lipid (8.01%), ash (19.39%), and carbohydrate (29.92%) content of the feedstock material was previously reported [2]. The protein yield from jackfruit leaves increased as the intensity did (

Table 1. The yield of modified proteins from jackfruit leaves and their solubility.

Treatment	Time	Protein Content (%, w/w)	Solubility (%)
Control	0	22.0 ± 3.1 e	6.88 ± 0.23 e
600 W	10	25.4 ± 2.9 de	7.94 ± 1.09 de
	15	29.3 ± 3.1 d	9.16 ± 0.89 d
	20	30.1 ± 4.3 d	9.41 ± 0.56 d
840 W	10	27.9 ± 1.1 d	8.72 ± 0.43 d
	15	38.6 ± 2.6 c	12.06 ± 1.25 c
	20	49.4 ± 5.1 b	15.44 ± 0.98 b
1080 W	10	53.0 ± 5.2 ab	16.56 ± 1.56 ab
	15	58.0 ± 2.4 a	18.13 ± 1.33 a
	20	50.2 ± 4.1 b	15.69 ± 0.76 b

Different lower-case letters in the same column are significantly different, according to Fisher’s LSD test at p ≤ 0.05.

). The higher yield (58.0 ± 2.4%) was obtained at 1080 W and 15 min. The yields obtained using HIU were between three and eight times higher than those obtained using the same source by the maceration process (1 N HCl) and by UAE (42 kHz for 20 min) [6]. Previously, it was reported that by increasing the power from 100 to 400 W, the extraction yields of pecan proteins were enhanced from 20.5 to 45.5%. However, increasing the power to 500 W decreased the yield of pecan proteins to under 25% [21]. In other studies, as the power increased, it was possible to obtain a decrement in protein extraction linked to protein denaturation, owing to hydrolysis or aggregation related to the formation of disulfide bridges through the oxidation of sulfur-containing amino acids in the protein, induced by hydroxyl-free radicals generated at a high ultrasound (US) power [10,21]. The high yields linked to HIU are related to the energy input during the extraction process, which can lead to a higher yield of proteins. The increased energy input and disruption of the solid matrix can accelerate the release of compounds, owing to the enhancement of mass transport, resulting in a higher extraction rate [10].

Regarding the temperature during the extraction process, the initial temperature of the samples was 22 ± 2 °C; after the treatments, an increase of 5 ± 2 °C was observed, without exceeding 60 °C. The total temperature increment was less than 7 °C in all treatments. This behavior indicates that the processing was carried out under adequate temperature conditions. Temperature increments up to 60 °C that are caused by HIU could negatively affect the protein structure [22].

The composition of the proteins obtained from jackfruit with an intensity of 1080 W was assessed by GC-MS. Using this technique, 15 of the 20 amino acids (AA) were identified in the proteins from jackfruit leaves, of which seven (Val, Leu, Ile, Met, Thr, Phe, and Lys) corresponded to essential amino acids (EAAs,

Table 2. The amino acid profile of the modified proteins from jackfruit leaves obtained by HIU at 1080 W.

Amino Acid (g/100 g Protein)	TBDMS-Derivatized Amino Acid	LRI	Extraction Time (min)			Suggested Intake (mg/kg of Weight)
			10	15	20
Alanine	L-Alanine, N-(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	1521	3.54 ± 0.009 a	11.42 ± 0.091 d	19.83 ± 0.134 f	NA
Glycine	Glycine, N-(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	1546	0.79 ± 0.004 j	6.32 ± 0.023 i	13.22 ± 0.032 k	NA
Valine	L-Valine, N-(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	1635	1.73 ± 0.008 d	8.51 ± 0.011 e	20.47 ± 0.122 e	24
Leucine	L-Leucine, N-(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	1674	1.71 ± 0.005 e	10.26 ± 0.026 d	26.75 ± 0.321 c	42
Isoleucine	L-Isoleucine, N-(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	1706	1.06 ± 0.002 i	7.18 ± 0.009 h	16.94 ± 0.105 g	19
Proline	L-Proline, 1-(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	1742	3.22 ± 0.001 b	28.91 ± 0.102 a	25.94 ± 0.438 d	NA
Methionine	L-Methionine, N-(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	1938	ND	1.26 ± 0.002 l	3.79 ± 0.058 l	19
Serine	L-Serine, N,O-bis(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	1956	1.17 ± 0.003 h	7.52 ± 0.023 g	13.94 ± 0.098 j	NA
Threonine	L-Threonine, N,O-bis(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	1982	1.46 ± 0.002 g	8.56 ± 0.045 e	15.37 ± 0.110 i	20
Phenylalanine	L-Phenylalanine, N-(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	2053	0.69 ± 0.001 k	6.15 ± 0.018 j	16.50 ± 0.201 h	33
Aspartic acid	L-Aspartic acid, N-(tert-butyldimethylsilyl)-, bis(tert-butyldimethylsilyl) ester	2120	1.88 ± 0.005 c	14.89 ± 0.029 c	29.00 ± 0.348 b	NA
Hydroxyproline	L-Proline, 4-[(tert-butyldimethylsilyl)oxy]-1-(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	2147	ND	1.08 ± 0.008 m	2.08 ± 0.065 m	NA
Glutamic acid	L-Glutamic acid, N-(tert-butyldimethylsilyl)-, bis(tert-butyldimethylsilyl) ester	2237	1.53 ± 0.007 f	21.09 ± 0.128 b	40.16 ± 0.984 a	NA
Lysine	L-Lysine, N2,N6-bis(tert-butyldimethylsilyl)-, tert-butyldimethylsilyl ester	2339	ND	2.74 ± 0.002 k	25.34 ± 0.872 d	38
HAA	-	-	12.05	73.85	126.94	-
AAA	-	-	0.69	6.15	16.50	-
EAA	-	-	6.64	44.64	125.16	-
NCAA	-	-	3.41	35.98	69.15	-
TAAC	-	-	18.77	135.87	269.32	-

NA: not apply (i.e., they are non-essential amino acids); HAAs: hydrophobic amino acids; AAAs: aromatic amino acids; EAAs: essential amino acids; NCAAs: negatively charged amino acids; TAAC: total amino acid content; ND: not detected. Different lower-case letters in the same column are significantly different, according to Fisher’s LSD test at p ≤ 0.05.

). These EAAs were reported in the proteins and modified proteins of jackfruit leaves that were obtained by macerations and UAE [6]. However, the concentration of the EAAs obtained by HIU was significantly higher (approximately six times higher). Using emerging technologies such as HIU to valorize agro-waste is an important alternative method for providing proteins to malnourished populations. The protein concentrations in this study ranged from 22 to 58% for dry weight. Thus, the protein content in jackfruit leaves is 1.2–3.4 times higher than that of beef (17.4%), fish (19.2–20.6%), and chicken (19–24%) and is 1.23 times higher than that of entire eggs (47%) [23]. This makes jackfruit leaves a concentrated and efficient source of high-quality protein with a similar content of EAAs to Arthrospira sp., a microalga that has been called a “superfood” by the World Health Organization. The proteins of this microalga can have eight of the nine EAAs in concentrations of 10 to 55 mg of each amino acid per gram of powder [24].

The content of each amino acid increased between four and nine times when the extraction time was increased from 10 to 15 min. Conversely, when the extraction time was increased from 15 to 20 min, in most cases the quantification of the AAs increased between 1.5 and 12 times (Table 2). Therefore, the increment of the extraction time promotes a higher content of AAs, including EAAs, hydrophobic amino acids (HAAs), negatively charged amino acids (NCAAs), and aromatic amino acids (AAAs). This means that HUI promotes the release of AAs as the extraction time increases. HUI enables the obtainment of yields that are 2.7 to 5.4 times higher after 15 and 20 min, respectively, than yields obtained from the same source using a conventional ultrasound bath (42 kHz for 20 min) [6].

3.2. The Techno-Functional Properties of Proteins

3.2.1. Loss of Hydrophobicity (Solubility)

The organoleptic properties of some foods strongly depend on the functional characteristics of proteins. In this study, the solubility increased as the HIU power did (p ≤ 0.05) but it was not affected by the extraction time (p ≥ 0.05; Table 1). The proteins treated by HIU can be more soluble, owing to the high energy caused by the sonomechanical effects breaking down protein chains and modifying the side groups of AAs, leading to changes in protein structure and functionality [25]. The enhanced mass transport effects of the ultrasound (US) can expose the internal hydrophobic residues of proteins, making these polymers more soluble [25,26]. The solubility improvement generated by HIU technology depends on various factors, including the protein source’s inherent characteristics, the US’s intensity and amplitude, and the pH, temperature, ionic strength, and extraction time [6,26]. These variables can affect the physicochemical properties of proteins and contribute to their increased solubility when subjected to HIU treatment [21,25,26].

3.2.2. Foaming Properties

The foaming capacity (FC) refers to the ability of a substance, such as a protein, to form and stabilize foam. It measures the amount of interfacial area that can be created by whipping the substance. It is determined by the ability of the protein to adsorb quickly to air–water interfaces and lower the interfacial tension, trapping more air and increasing the foam volume [9]. With the application of HIU, it was possible to obtain a modified protein with a modified FC, which was not observed in the untreated LPC (control, Figure 1). The FC average at 10 min increased according to the increase of power. However, owing to the dispersion of the data, it was not possible to identify significant differences under the different conditions. Nevertheless, it is important to consider that at 15 min, the mean of the FC under all the assessed powers decreased in comparison to the modified protein that was obtained after 10 and 20 min (Figure 1). Our results are like those obtained from modified proteins of the same source using UAE (35 ± 1.67% FC, at 42 kHz for 20 min) [6]. Studies have shown that HIU treatment can improve the foaming properties of various proteins, including soy protein isolate (SPI), whey protein microgel, pea protein isolate (PPI), and chickpea cooking water (aquafaba) [11,13]. The treatment can increase the foaming ability and stability of proteins, enhancing foam expansion, foam stability, and the emulsifying activity index [27]. In the case of PPI, the FC increased from 145.6% to 200.0% as the intensity increased from 50 to 90% (20 kHz for 30 min) [11]. The FC of a hydrolyzed protein is strongly linked to the obtention method and the conditions used in the structural modification. The method affects the exposition of HAAs, the hydrolysis degree, and the presence of carbohydrates like sugars, which are factors that affect the FC [9]. It has been reported that as the size of the modified proteins decreases the FC does also. As the protein structure decreases in size, the number of smaller molecules also decreases, which may result in the protein not being able to stabilize air bubbles as effectively, leading to a decrease in the FC [2]. US pretreatments, in terms of power and exposure time, can reduce the size of modified rapeseed protein [28] and milk protein concentrates compared to samples that were not treated with sonication [29]. The FC of protein hydrolysates treated with HIU can vary significantly, depending on the duration of the treatment. During the initial stages of treatment (for example, at 10 min), proteins undergo partial unfolding, which increases their exposed hydrophobic regions. This unfolding is crucial for enhancing the FC, as it reduces the surface tension at the air–water interface, allowing for better foam formation [11]. However, at 15 min, the extent of unfolding may reach a point where excessive cavitation and mechanical stress lead to protein aggregation or denaturation, which can negatively impact the FC. After 15 min of treatment, the particle size becomes small (Figure 2), leading to a loss of structural integrity and a reduced FC [13]. By 20 min, the system may adapt or stabilize in a way that improves foaming again. Therefore, this phenomenon could suggest that there is an optimal treatment time at which the balance between particle size and protein structure is maintained for the maximum FC [15].

On the other hand, the FS refers to the ability of a foam to maintain its structure over time. In this research, the FS of the ultrasonicated proteins was higher than that of untreated jackfruit proteins (p ≤ 0.05). In addition to this, the SF increased as the power increased (p ≤ 0.05), but it was reduced with the increment of the treatment time (p ≤ 0.05; Figure 1). This agrees with the reported PPI: in that case, the FS increased from 58.0 to 73.3% with the increasing of the intensity from 50 to 90%; after 10 min, the FS was reduced to 50.0% as the time was increased to 20 min [11]. This event can be attributed to the partial denaturalization of the modified proteins, caused by the partial unfolding of the protein structure as an effect of the HIU treatment, which promotes rapid adsorption at the freshly formed air–water interface [30]. Since the amount of HAAs increased as the treatment time did (Table 2), more hydrophobic groups were exposed. The modified proteins rich in HAAs: the increased propensity for aggregation can lead to the formation of larger aggregates, which can destabilize the foam structure by disrupting the air–water interface, promoting the coalescence of the foam [31]. These factors could explain the fact that the modified proteins’ properties were severely reduced by the treatment time, resulting in similar behavior in the FS of the untreated proteins and those treated being observed after 15 and 20 min. The mechanism behind HIU’s improvement of foaming properties is not fully understood. However, the effects of HIU on vegetable proteins can lead to structural modifications and the formation of aggregates with a determined particle size, which can enhance or decrease the foaming properties of the proteins, depending on the treatment conditions [13].

3.2.3. Emulsifying Properties

The ESI evaluates the stability of the emulsion over a designated time. The ESI decreased using HIU at 600 and 840 W. However, it increased significantly to 1080 W in comparison with the control. The higher ESI was reached at a power of 1080 W and at 15 min of treatment (Figure 2). Conversely, similar EAI values were obtained in the three assessed powers after 10 min. Few studies have reported these properties in modified proteins from jackfruit LPC. Calderón-Chiu et al. [1] investigated the emulsifying properties of jackfruit leaf protein peptides obtained through hydrolysis with pancreatin. The results demonstrated that the emulsions stabilized with the peptides had relatively good encapsulation efficiencies (40.15 ± 1.46%) and loading capacities (18.03 ± 2.78%). Conversely, the ESI in fresh emulsions was 108.15 ± 12.1 min, which decreased to 47.62 ± 4.97 and 66.12 ± 12.33 min following storage at 4 and 25 °C, respectively, for 30 days. By using the US, it was possible to increase the ESI from 46.19 ± 4.81 to 480.89 ± 10.77 min and the EAI from 33.99 ± 0.91 to 78.28 ± 0.03 m²/g [6]. The reduction in the emulsifying indexes in the modified proteins obtained by using HIU compared to those obtained with the UAE could be linked to the processing conditions. As the size of the modified protein chain is strongly affected by the power, increased hydrolysis can result in modified proteins that are too short to act as effective emulsifiers, leading to a decrease in their emulsifying power [32].

3.2.4. Particle Size Distribution

Since droplet size and dispersion influence possible coalescence processes, they rank among the most significant features of emulsions. Therefore, they can be utilized as a stand-in for stability [33]. The application of HIU promotes changes in the d_3,2, d_4,3, and the PDI; for the modified proteins, the most important reduction in the indexes’ values were acquired at 1080 W and 15 min, contributing to a monomodal distribution (Figure 2). The potential flocculation or coalescence of an emulsion can be determined by using the d_4,3 index, owing to its sensitivity to determine the presence of larger droplets; high values suggest a more unstable emulsion [34,35]. On the other hand, the d_3,2, which indicates the mean diameter at which most of the particles fall, is inversely proportional to the specific surface area of droplets [34]. The increment in the PDI could be related to the pressure in the microbubbles caused by cavitation phenomena, as this value increases as the system heterogeneity also does [36]. From the standpoint of thermodynamics, small droplets and low values of the mentioned parameters favor the system’s stability [33]. The acoustic field created by HIU waves separates the dispersed from the continuous phase. Additionally, the pressure variations caused by the cavitation compress microbubbles into micro- or nano-sized droplets, reducing the possibility of destabilizing the processes of cream production and coalescence in the system [36,37]. In addition to this, it was previously reported that the concentration of particles also plays a vital role in the emulsion stability. In dilute emulsions, smaller particles enhance stability against droplet coalescence, whereas larger particles can improve droplet stability at high-volume fractions of the dispersed phase. This suggests that the dynamics of droplet interactions differ significantly in concentrated emulsions compared to dilute systems [38]. Surfactants, such as Tween 80, are used to stabilize the droplets during emulsification and, thus, prevent the coalescence of the droplets [39].

The particle size distribution also can be affected by the molecular weight (Mw) distribution of modified proteins from jackfruit leaf. As the duration of the HIU treatment increases, the Mw distribution of jackfruit leaf protein hydrolysates could shift to a small size. Significative reductions in the Mw by HIU have been reported in modified proteins from egg white (from 124.24 to 60.52 nm, after 15 min of treatment at 800 W, 20 kHz, and an amplitude of 30%) [40,41]; sunflower protein isolates (from 114.6 to 93.6 nm, after 30 min of treatment at 500 W, 20 kHz, and an amplitude of 25%) [42]; and in sesame protein isolates (after 6 min of treatment 20 kHz and at a 95% amplitude) [43]. These results may be linked to the acoustic cavitation effects generated by the US, which disrupt the protein structure [44]. The changes in Mw distribution are closely related to the functional properties of the modified proteins. For instance, an increased sonication time has been associated with the enhanced solubility and emulsifying properties of the modified proteins. This is because smaller modified proteins generally exhibit better solubility, hence they can stabilize emulsions more effectively than larger protein aggregates [1]. Another key factor is the concentration of the polymers in the emulsion. It has been demonstrated that the application of US (200 W for 30 s) in double emulsions based on poly (lactic acid) (60, 90, 120, and 150 mg) and pilocarpine (24, 36, 48, and 60 mg) helps to obtain a homogeneous particle size distribution. However, as the concentration of the polymers increased, the shell thickness increased from 10 to 99 nm, reducing the encapsulation efficiency from 93 to 52%. In addition to the droplet size, the shell thickness is crucial for nanoparticles designed for delivery systems. A moderate shell thickness of ~40 nm released the drug above the target level (~10 μg/mL) for over 56 days [39].

3.2.5. Antioxidant Properties

In all cases, the ABTS⁺ radical scavenging activity of the LPC treated by HIU was bigger than the non-treated LPC (

Table 3. Antioxidant properties of leaf protein concentrates treated by high-intensity ultrasound under different powers.

Time (min)	Power (W)	%RSA	mg TE/g Sample	IC50 Values of ABTS (mg/mL)
10	Control	36.99 ± 1.01 h	4.52 ± 0.21 b	ND
10	600	79.25 ± 0.56 c	4.70 ± 0.53 ab	0.032
10	840	87.60 ± 1.42 b	5.42 ± 0.02 a	0.029
10	1080	74.98 ± 1.05 e	1.99 ± 0.01 f	0.033
15	Control	28.71 ± 1.09 i	0.6 ± 0.08 h	ND
15	600	77.28 ± 1.03 d	2.59 ± 0.33 e	0.032
15	840	96.65 ± 0.56 a	1.04 ± 0.07 g	0.026
15	1080	64.72 ± 1.13 g	3.59 ± 0.28 d	0.039
20	Control	34.04 ± 1.28 h	2.34 ± 0.43 e	ND
20	600	74.66 ± 0.68 e	2.35 ± 0.12 e	0.033
20	840	71.09 ± 0.28 f	4.02 ± 0.05 c	0.035
20	1080	64.78 ± 0.71 g	4.02 ± 0.03 c	0.039

The antioxidant activity was expressed as a Percentage of Radical Scavenging Activity (%RSA) and as milligrams of Trolox equivalents per gram of sample (mg TE/LPC). Different lower letters in the same column indicate significant differences among the treatments (p < 0.05). IC₅₀ is the sample concentration (mg/mL) required to achieve 50% of antioxidant activity. ND: No determined.

). The non-treated LPC showed significantly lower antioxidant capacity than the treated proteins. However, the modified proteins extracted at 840 W showed significantly better ABTS^•+ RSA, reaching the highest antioxidant capacity (from 71 to 96.65%) in the three evaluated times.

In the context of ABTS^•+, the IC₅₀ value indicates the concentration of an antioxidant needed to scavenge 50% of the ABTS radicals present in each system. This value is inversely proportional to the antioxidant activity of a sample. A lower IC₅₀ value indicates higher antioxidant activity, as it means that a smaller concentration of the sample is required to scavenge the free radicals. The US breaks Van der Waals forces, hydrogen bonds, and other non-covalent bonds of the LPC [45]. The AAA composition showed that the HIU-treated LPC had higher contents of glutamic aspartic and lysine (Table 1). The acidic AAs can donate electrons and act as metal-chelating agents. Likewise, positively charged amino acids, such as lysine, can bind and neutralize negatively charged free radicals [46], which would explain the high antioxidant properties of the modified proteins compared with the controls. The same behavior was observed in lupin-modified proteins [47]. The ultrasonic waves applied during the HIU treatment to proteins break them down into smaller, modified proteins, which often exhibit enhanced antioxidant properties compared to their native proteins [41,45]. Ultrasonic waves can generate free radicals, which can interact with the protein structure, leading to the exposure of new chemical groups. The size reduction can also modify the content of the different secondary structure portions [41,45,48]. In watermelon seed proteins, HIU treatments decreased the proportion of α-helix and β-turn, increasing the β-sheet proportion [48]. These changes led to the exposure of hydrophobic amino acids, enhancing their ability to interact with and neutralize free radicals [41,45,48]. After HIU treatment, the proportion of the identified amino acids increased (Table 2). Some of them have well-demonstrated antioxidant properties, such as methionine (the sulfur atom in its structure can be oxidized, making it a scavenger of ROS), glutamic acid, and aspartic acid (both have carboxylate groups that can chelate metal ions, reducing the oxidative damage caused by metal-catalyzed radical formation) [49].

The increment of the exposition of certain AA residues contributes to the antioxidant activity, without necessarily altering the protein-binding affinity (which the IC₅₀ measures) [45,50]. This result may be beneficial in applications where improved antioxidant properties are desired, without compromising the efficiency of inhibition (IC₅₀). Even when the proteins have been modified to have a better RSA or exhibit different antioxidant behavior, their efficacy remains stable at certain concentrations. In practical applications, such as in the food or pharmaceutical sectors, this could lead to a more effective antioxidant effect without having to change the dosage, thus maintaining efficacy while improving the health benefits or shelf life of the products [51]. Therefore, these results demonstrated the potential of the HIU as a suitable method for enhancing the release of novel bioactive modified proteins with better antioxidative properties with potential applications in the industrial sector. In addition to this, an economic analysis regarding the use of US has demonstrated its profitability in the food industry [16,52].

All the assessed properties in this research are linked with the modifications provided by HIU, which generates high-frequency sound waves that create rapid pressure changes in the liquid medium. This leads to the phenomenon known as acoustic cavitation, where microscopic bubbles form and collapse violently. The energy released during the collapse of these bubbles generates localized high temperatures and pressures, which can disrupt the protein structure. This disruption primarily affects the non-covalent interactions, such as hydrogen bonds and hydrophobic interactions, that maintain the protein’s tertiary and quaternary structures [4]. This process involves the unfolding of the protein molecules, allowing hydrophobic regions to be exposed, which improves the solubility and functional properties of the modified protein. The energy imparted by HIU also increases the molecular motion of the modified protein, promoting interactions among molecules and potentially leading to new structural arrangements or aggregates that can improve the functional characteristics of the protein hydrolysates [1,2,7]. HIU alters the structural integrity of jackfruit leaf proteins through mechanisms such as acoustic cavitation, protein denaturation, fragmentation, and enhanced molecular motion.

4. Conclusions

This study demonstrates that HIU effectively enhances the techno-functional properties and antioxidant capacity of jackfruit leaf proteins. The modifications induced by HIU, particularly at 1080 W for 15 min, significantly improved the protein solubility (10%) and the foaming capacity (40%) compared to the native protein. The modified proteins obtained under the mentioned conditions contained 15 different AAs, of which, seven were EAAs, and they were able to develop emulsions with a relatively good encapsulation efficiency (40.15 ± 1.46%) and loading capacity (18.03 ± 2.78%). Furthermore, the HIU treatments improved the RSA of the modified proteins up to 45% against ABTS^•+ at stable concentrations. These findings highlight the potential of HIU-treated jackfruit leaf proteins as valuable ingredients in food formulations, offering both functional benefits and enhanced antioxidant properties for consumers. The application of HIU provides a promising approach to overcoming the limitations of plant proteins, paving the way for their broader use in the food industry. Further studies to elucidate the structural modifications obtained by HIU should be performed in regard to the obtention of modified proteins from plant proteins.