1. Introduction
Growing interest has been observed in exploring natural sources’ phytochemical composition and functional properties in recent years [
1]. The by-products of agro-industries, such as plant leaves, roots, and stems, are emerging as promising renewable resources rich in cellulose, hemicellulose, and lignin [
2]. Their complex matrix traps and binds bioactive substances such as phenolic compounds, amino acids, proteins, minerals, and lipids, which offer numerous health benefits and have potential applications in various industries, in particular in the food and food supplement industries [
3]. Moreover, consumers’ awareness of diet and health correlations is rising steadily [
4]. If prepared appropriately, plant by-products can deliver prebiotic, anti-diabetic, and antioxidant functions. For example, hydrolytic treatment has enabled the extraction of previously non-extractable polyphenols, resulting in higher phenolic recovery in plant-by product extracts [
5]. However, different plant materials and their morphological parts possess miscellaneous compositions, which can fluctuate positively and negatively, accentuating the need for a fundamental analysis. Furthermore, the rising interest in by-product development is fully aligned with the United Nations Sustainable Development Goals (SDGs)—in particular, SDG No. 12, Responsible Consumption and Production, which aims to cut food loss and waste in food production by up to 50%, and SDG, No. 2, End Hunger, achieving food security and improved nutrition, and promoting sustainable agriculture [
6]. Following these rationales, investigating the possible functionality of plants is crucial, and sea buckthorn (
Hippophae rhamnoides L.),
Aralia cordata Thunb. (
A. cordata), and industrial hemp (
Cannabis sativa L.) are plant species that have gained attention due to their rich phytochemical composition and promising functional properties [
7,
8,
9,
10,
11].
The sea buckthorn (
H. rhamnoides) is a valuable plant known for its rich phytochemical composition in both its berries and leaves. The berries contain essential compounds such as carotenoids, flavonoids, phenolic acids, fatty acids, ascorbic acid, and tocopherols, contributing to their powerful antioxidant, anti-inflammatory, and anti-cancer properties. Additionally, the leaves are rich in flavonols (rutin, Q-3-
O-galactoside, I-3-
O-glucoside, K-3-
O-glucoside, K-3-
O-rutinoside, quercetin, and kaempferol), phenolic acids (gallic acid, caffeic acid,
p-coumaric acid, ferulic acid) vitamins, proteins, and minerals [
12,
13], which have been found to have anti-obesity, hypoglycaemic, antioxidant, antibacterial, anti-inflammatory, antidiabetic and anti-cardiovascular disease properties [
14,
15]. While the berries are commonly used for their nutritional and medicinal benefits, the leaves, utilised in herbal preparations, also possess significant therapeutic potential.
Meanwhile,
A. cordata is a plant that is highly valuable due to its rich content of bioactive compounds such as polyphenols, anthocyanins, carbohydrates, and vitamins [
7,
16]. These compounds have been found to possess various pharmacological properties, including anti-inflammatory, cardiovascular, and nervous system protection, compound metabolism regulation, and antibacterial, antiviral, and antioxidant properties. Moreover, the plant has been traditionally used to treat ailments such as hepatitis and rheumatoid arthritis. In a previous study,
A. cordata roots and leaves were found to possess significant antioxidant activity and total phenolic content [
7].
Hemp (
C. sativa L.) is a widespread plant species of the
Cannabaceae family found in different habitats. The interest in hemp has remarkably increased due to specific phytochemicals in its leafy anatomical parts. More than 70 biologically active and unique to Cannabis terpenophenolic compounds, phytocannabinoids, have been found [
17]. Several research studies showed the health-promoting and medicinal properties of phytocannabinoids. Among them, Δ
9-tetrahydrocannabinol (Δ
9-THC) is a well-known natural psychotropic compound; for this reason, only the approved cultivars of
C. sativa accumulating less than 0.2–0.3% of Δ
9-THC are officially allowed in Canada, the USA, and many European countries. Hemp leaves are rich in phytochemicals, like cannabinoids, terpenes, phenolic compounds, and their biosynthetic routes. Cannabinoids represent the most studied group of compounds, mainly due to their wide range of pharmaceutical effects in humans, including psychotropic activities [
18].
Human health, from a holistic viewpoint, requires a comprehensive approach. A highly balanced and functional nutrition has to deliver a whole spectrum of particular properties such as antioxidant, antidiabetic, and antimicrobial ones, among others, together with the nutritional value of the food. Hence, products with a high nutritional value can be enriched with plant by-products [
19]. Recently, the antidiabetic activity of extracts has been garnering attention. This activity can also be associated with obesity prevention. More than 650 million adults and 340 million children suffer from obesity, and food is one of the key factors in preventing it [
20]. For example, the study of berry pomace extracts showed a high α-glucosidase and pancreatic lipase inhibition capacity, which are referred to as antidiabetic properties [
21].
In order to isolate bioactive compounds and guarantee product quality, efficacy, and safety, the extraction process is essential in the food and pharmaceutical industries. Conventional extraction methods, such as soxhlet extraction, solid–liquid extraction, and maceration, involve subjecting the raw material to high temperatures with chemical solvents. While this practice enhances the yield of compounds, it concurrently diminishes the quality and degrades valuable compounds, due to prolonged exposure and considerable energy consumption [
22,
23]. Lately, innovative technologies, also known as “green technologies”, have been developed for extracting bioactive compounds without using toxic chemicals [
24]. These eco-friendly techniques are directly related to the environment and consumer health [
25]. Notable innovative methods for extracting bioactive compounds include supercritical fluid, microwave-assisted, enzyme-assisted, ultrasound-assisted, pressurised liquid, and pulsed electric field techniques [
26]. Enzyme-assisted extraction has been observed to offer multiple benefits compared to conventional extraction techniques, due to its eco-friendly nature and improved ability to extract specific compounds under gentle processing conditions, shorter extraction times, and reduced solvent, as well as energy, usage [
27]. Additionally, various factors, including the type and concentration of enzymes, extraction duration, substrate-to-solvent ratio, pH, and temperature, should be considered to obtain extracts with higher yields and improved properties [
28].
The aim of this research is to evaluate and compare sea buckthorn leaf, industrial hemp leaf, and
A. cordata leaf and root extracts obtained by different methods, namely, enzyme-assisted extraction (EAE) and solid–liquid extraction (SLE). This study covers the selection of the raw material and the preparation, extraction, and evaluation of phytochemical composition to achieve a high yield of biologically active substances and effective antioxidant, antidiabetic, and antimicrobial activities.
Figure 1 provides an overview of the overall experimental design for the production and characterization of a total of 14 extracts.
2. Materials and Methods
2.1. Plant Material
Sea buckthorn (Hippophae rhamnoides L.) and Aralia cordata plants were grown at the Institute of Horticulture, Lithuanian Research Centre of Agriculture and Forestry experimental fields (55°08′ N, 23°80′ E). Industrial hemp (Canabis sativa L.) leaves were obtained from a local food factory, “Allive Europe” (Voskoniai, Lithuania). Fresh A. cordata, sea buckthorn, and industrial hemp leaves were harvested in 2021. The collected leaves and roots were instantly frozen at −35 °C before freeze-drying. The samples were lyophilised in Zirbus lyophiliser (Zirbus Technology GmbH, Bad Grund, Germany) at 0.01 mbar pressure and −85 °C condenser temperature. The freeze-dried samples were ground to a powder (particle size 0.2 mm) using a Retsch 200 knife mill (Haan, Germany) and stored in a sealed container before the analysis.
2.2. Enzyme Products
Cellulase and Viscozyme L were purchased from Sigma-Aldrich (Steinheim, Germany). Cellulase was produced from Trichoderma reesei and declared to have ≥700 units/g. Viscozyme L is a cellulolytic enzyme complex from Aspergillus aculeatus. Furthermore, the manufacturer states that the enzyme mixture contains a wide range of arabanase, Cellulase, β-glucanase, hemicellulase, and xylanase. The product is declared to have ≥100 FBGU/g.
2.3. Enzyme-Assisted Extraction (EAE) and Solid-Liquid Extraction (SLE)
Enzyme-assisted extraction (EAE) was carried out as outlined by Puzeryte et al. under optimal conditions [
8].
Aralia cordata leaves and roots, sea buckthorn leaves, and industrial hemp leaves were extracted using EAE at a 1:20 (
w/
v) ratio and mixed to obtain a homogeneous suspension. The suspension pH, 4.9, was adjusted using 6 M HCl and 0.5 M NaOH, and the enzymes Viscozyme L and Cellulase 1% (
v/
w of dry matter) were added. EAE was carried out at 45 °C in an incubator for 3:15 h. After extraction, the enzyme was deactivated by heating the hydrolysed material at 95 °C for 10 min. Then, the separation of the suspension into liquid and solid fractions was accomplished using a filter (200 mesh). Sample extracts (liquid fraction) were frozen and stored at −35 °C before analysis. Furthermore, the extract to be used as a control was prepared under optimal conditions without enzyme addition. SLE of the tested plant sources was performed in a ratio of 1:20 (
w/
v) with H
2O for 24 h in the dark. All the experiments were performed in triplicate.
2.4. Protein Content by Lowry Assay
The protein content of the sample extracts was determined using the colorimetric method described by Lowry et al. [
29].
2.5. Determination of Sugars and Organic Acids
Sugars and organic acid concentrations were determined through high-performance liquid chromatography (HPLC). Briefly, the samples were centrifuged and the supernatants were collected, filtered through 0.22 µm filters, and injected into the HPLC system for analysis. The HPLC system consisted of an Aminex HPX-87H cation exchange column (300 × 7.8 mm) (Bio Rad Laboratories Pty Ltd., CA, USA), coupled to refractive index and ultra-violet detectors. Throughout the analysis, the column was maintained at 40 °C, and the mobile phase utilised was 5 mM H2SO4 at a flow rate of 0.7 mL min−1. The compounds were identified by the retention time and quantified through the area of the corresponding peak by interpolation of calibration curves determined from the respective standards.
2.6. Determination of Total Phenolic Content
The total polyphenol content (TPC) in the extracts was determined according to the Folin–Ciocalteu method [
30], using gallic acid (GA) as the standard, according to the method of Bobinaite et al. [
31]. The reagent was prepared by diluting a stock solution with ultra-pure distilled water (1/10,
v/
v). The samples (1.0 mL, three replicates) were introduced into test cuvettes, followed by 5.0 mL of Folin–Ciocalteu’s phenol reagent and 4.0 mL of Na
2CO
3 (7.5%). The system was then placed at ambient temperature for 1 h. The absorbance of all the samples was measured at 765 nm using a Cintra 202 (“GBC Scientific Equipment”, Knox, Australia) spectrophotometer. The total concentration of phenolic compounds was determined from the calibration curve and expressed in mg of gallic acid equivalents in 100 mL of extract.
2.7. Evaluation of Phenolic Compounds in Extracts Using the UHPLC-ESI-MS/MS Technique
The qualitative and quantitative content of phenolic compounds in the tested
A. cordata leaf and root, sea buckthorn leaf, and industrial hemp leaf samples extract were evaluated using ultra-high-performance liquid chromatography (UHPLC) coupled to a mass spectrometer. The method employed was described and validated in an article by Gonzalez-Burgos et al. [
32]. The phenolic content of the sample extract was analysed using a liquid chromatography system, “Waters ACQUITY UPLC
® H–Class”, with a tandem quadrupole mass detector, “Xevo TQD” (Waters, Milford, MA, USA). The compounds were separated using a “YMC Triart C18” (100 Å, 100 × 2.0 mm; particle size 1.9 μm) with a pre-column. Both qualitative and quantitative analyses were performed. The mass spectrometry parameters for the phenolic compound analysis are presented in
Table 1.
2.8. Determination of Antioxidant Activity
An ABTS
•+ radical cation decolourization assay was applied according to the methodology described by Re et al. [
33]. An amount of 2 mL of ABTS
•+ (2,2′-azino-bis(3-ethylbenzthiazoline-6-sulphonic acid)) solution, with an absorbance of 0.800 ± 0.02, was mixed with 20 μL of the samples. After 30 min, the absorbance of each sample was measured at 734 nm using a Cintra 202 spectrophotometer (GBC Scientific Equipment, Knox, Braeside, VIC, Australia) to determine the decrease in absorbance.
A ferric ion reducing antioxidant power (FRAP) assay was conducted following the method described by Benzie and Strain [
34], with some modifications. The FRAP solution was made by combining TPTZ (0.01 M dissolved in 0.04 M HCl), FeCl
3 × 6H
2O (0.02 M in water), and acetate buffer (0.3 M, pH 3.6) at a ratio of 1:1:10. A quantity of 2 mL of the freshly prepared FRAP reagent was mixed with 20 μL of the samples. After 30 min, the absorbance increase was determined at 593 nm using a Cintra 202 (GBC Scientific Equipment, Knox, Australia) spectrophotometer.
All antioxidant activity assays were calculated using Trolox calibration curves and expressed as the μmol of the Trolox equivalent (TE) per one millilitre of extract (µmol TE/mL of extract).
2.9. Oxygen Radical Absorbance Capacity Assay (ORAC)
The ORAC assay was conducted following the procedure established by Dávalos, Gómez-Cordovés, and Bartolomé [
35] with some modifications. Firstly, a phosphate buffer (75 mM, pH 7.4) was used, and 20 μL of the sample (after dilution), along with 120 μL of fluorescein (1166.1 µM), was added to a black microplate (Nunc, Denmark). The mixture was pre-incubated at 37 °C for 15 min. After that, 60 μL of 2,2′-azobis-(2-methylpropionamidine)-dihydrochloride (AAPH) (46.6 mM) was added rapidly and then incubated at 37 °C for 140 min. The microplate was read at 1 min intervals using a Multidetection plate reader (Synergy H1, Vermont, USA) at 458 nm and 528 nm. The software used was the Fluostar Control 1.32 R2 version. The calibration curve was made using Trolox (0.0002 to 0.0016 μmol TE/mL) as an antioxidant standard, and the results were expressed in mmol TE/mL of extract. A blank was prepared by using the phosphate buffer instead of a sample. All analyses were carried out in triplicate.
Normalised antioxidant curves were obtained by multiplying the original data by fluorescenceblank,t=0/fluorescencesample,t=0, and dividing by the blank curve corresponding to the same assay.
2.10. α-Glucosidase Inhibition Assay
The α-glucosidase inhibitory activity was measured to assess antidiabetic potential according to the procedure described by Kwon et al. [
36], with slight modifications. To begin, 50 μL of the sample was mixed with 100 μL of 0.1 M phosphate buffer (pH = 6.9) containing α-glucosidase solution (1.0 U/mL) in each well. The mixture was then incubated at 25 °C for 10 min. Afterwards, 50 μL of 5 mM
p-nitrophenyl-a-D-glucopyranoside solution in 0.1 M phosphate buffer (pH = 6.9) was added to each well. Subsequently, the absorbance measurements of the reaction mixtures were taken at 405 nm using a multi-detection plate reader (Synergy H1, VT, USA), after a further 5 min of incubation at 25 °C.
For the experiment, 50 μL of buffer solution was used as the negative control, and 50 μL of acarbose at a concentration of 10 mg/mL was used as the positive control. All assays were performed in triplicate. The inhibition of α-glucosidase was calculated as follows:
2.11. Antibacterial Analysis
In order to determine the antimicrobial activity of
A. cordata leaves and roots, as well as the leaves of sea buckthorn and industrial hemp extracts (SLE and EAE_Visc), a well diffusion test method was used based on Kaewchomphunuch et al. [
37] with minor modifications. A wide range of pathogenic bacteria were tested, which are listed in
Table 2. Each microorganism was grown on Tryptic Soy Agar plates (TSA) at 37 °C for 24 h prior to analysis. The extracts were freeze-dried and then resuspended in 2 mL of water before testing. Agar well diffusion assays were performed in 90 mm diameter Petri dishes containing Mueller Hinton Agar to a depth of 4 ± 0.5 mm. Bacterial suspensions were obtained with a cell concentration of 1 McFarland (around 3 × 10
8 CFU/mL) in saline solution (0.9% NaCl, Honeywell, Fluka, Cambridge, UK). This suspension was spread uniformly using a sterile cotton swab. Six wells with a 5 mm diameter were used in the agar plates. Then, 50 μL of the test extracts and 10 μL of the positive control, ampicillin, at a concentration of 100 µg/mL was placed into each well using a sterile pipette. The plates were then incubated for 24 h at 37 °C. After incubation, the diameters of the growth inhibition zones were measured in millimetres to the nearest 0.1 mm. Each experiment was repeated three times, and the means and standard deviations were calculated.
2.12. Scanning Electron Microscopy (SEM)
This study examined the structure and changes in the samples that were hydrolysed by the Viscozyme L enzyme, using scanning electron microscopy (SEM). The samples, both before and after EAE_Visc, were freeze-dried and mounted on SEM pins using double-sided adhesive carbon tape (NEM tape; Nisshin, Tokyo, Japan). They were then coated with gold/palladium and visualised in a PhenomTM XL G2 (Thermo Fisher Scientific, Eindhoven, The Netherlands) SEM. These analyses were conducted at an accelerating voltage of 5 kV using the secondary electron detector (SED).
2.13. Statistical Analysis
All analyses were performed in triplicate. MS Excel 2020 (Redmond, WA, USA) calculated the mean values and standard deviations. Afterwards, a one-way ANOVA was performed, followed by Tukey’s HSD test to compare the means that showed significant variation (p < 0.05). These calculations were performed using GraphPad Prism 8.0 software (GraphPad, San Diego, CA, USA).