The chemicals used for the preparation of DESs included Betaine (99%), DL-Menthol (≥ 95%), Nile Red (≥ 98%) purchased from Sigma, Lactic acid (85%), Lauric acid (98%), Myristic acid (≥ 98%) obtained from Sigma–Aldrich, L-proline (98%) from Scharlau, D-(+)-glucose anhydrous (≥ 95%), Stearic acid (98%) from Merck/Sigma, Ethanol (96%) from Valente and Ribeiro, Methanol PA from Honeywell, Folin-Ciocalteau Reagent from Panreac, L-Ascorbic acid (99%), Quercetin (HPLC grade), which were purchased from Sigma and were used as purchased.

Cannabis sativa L. (hemp) was provided by South Hemp Tecno srl (Taranto, Puglia), providing dried threshing residues of the EU certificated variety Futura 75. The flowers and leaves were harvested in 2018 from an Italian farm located in Puglia. The material was constituted by a mixture of leaves, flowers, and seeds.

Before extraction, hemp leaves and inflorescences were grinded using a commercial blender. This method allowed a reduction of the sample size and a higher contact between the DES and the plant material, resulting in an increase of the extraction efficiency, consequently, increasing the final yield. The inflorescences, leaves, and seeds were grinded in a lab mill (IKA^® Tube-mill control) at 6,000 rpm for 45 s, with a particle size range between 1 mm and 180 μm. The samples were kept in an amber flask in a dark place to protect them from light, under room temperature.

Deep eutectic solvents were produced by the heating-stirring method. This method was selected since it is simple and allows the preparation of multiple DESs simultaneously, and it can be easily scaled up. DESs were obtained by mixing the HBAs and HBDs at the desired molar ratio as shown in Table 1. The mixtures were stirred using a magnetic stirring apparatus at 40°C until a homogenous solution was obtained. All DESs were then stored at room temperature.

Bioactive compounds from hemp samples were extracted by mixing the DES with the plant matrix in a solid-liquid ratio of 1:10 (W/W). After being briefly mixed, cannabinoids were extracted for 90 min in an ultrasonic bath [Model XUB5, Formatura (Type Solution)] (water temperature at 60°C; ultrasonic power, 100 W) in cycles of 15 min; a sample was collected for future kinetic analyses. Then centrifuged (Model Victor Nivo 3S, ILC) at 6,000 rpm for 15 min to separate the liquid from the solid phase. Each experiment was repeated three times for each DES, and the respective cannabinoids were quantified by HPLC.

The samples were prepared by diluting the obtained extracts in methanol (a 1:10 w/w ratio) and then stirred to until a homogenous solution was obtained. The solution was then filtered using a hydrophilic PTFE syringe filter with a 0.20-μm pore size (FilterLab) before analysis.

HPLC analysis of the hemp extracts was carried out using an Agilent Infinity 1100 HPLC System, and an Agilent 1100 series photodiode-array detector (DAD) for detection and recording at UV/Vis 220 nm. The cannabinoids chromatographic separations were achieved using a Kinetex C-18 column (100 mm × 4.6 mm ID and 2.6-μm particle size, 100 Å pore size). The method used for the HPLC analysis was adapted from the Cannabinoids on Raptor ARC-18 Restek LC_GN0553 methodology as described elsewhere (23).

As mobile phase A:0.1% Formic acid in water and B:0.1% Formic acid in acetonitrile were used, the solvent flow was kept constant at 1.5 ml/min with the following gradient profile:0.00–4.00 min 25% of solution A, 75% of solution B, 0.00–4.01 min 0% of solution A, 100% B of solution and 4.01–7.00 min 25% of solution A, 75% of solution B. The column oven was kept at 50°C during the run, and the injection volume was of 5 μL. The identification and the quantification of cannabinoids were based on CBD, CBDA external standards.

The total phenolic content (TPC) was determined for individual extracts using the Folin–Ciocalteu method and was adopted from Waterhouse A.L, with some modifications (24). The outcome data were expressed as mg of gallic acid equivalents per gram of hemp (mg GAE/g hemp).

The calibration curve was obtained, preparing a stock solution with concentration of 5 g/L, dissolving 0.5 g of gallic acid in 10 ml of ethanol and 90 ml of distilled water. From this solution, five standard solutions obtained, with concentrations of 50, 100, 150, 250, and 500 mg/L, The five standard solutions obtained, with concentrations of 50, 100, 150, 250, and 500 mg/L, respectively, were used to acquire the calibration standard curve. Briefly, 20 μL of extract (diluted 1:10 w/w in methanol) was mixed with 1.58 ml of distilled water and 100 μL of Folin–Ciocalteu reagent; the solution was mixed and incubated at room temperature for 7 min. Then, 300 μL of a Na₂CO₃ solution was subsequently added to the mixture and incubated at 40°C for 30 min. Afterward, the absorbance was measured, utilizing a microplate reader (Model Victor Nivo 3S, ILC) at 750 nm.

The flavonoid content of individual extracts was measured according to Navarro J.M. et al. (25). The calibration curve was prepared from a stock solution with concentration of 2 g/L, dissolving 0.2 g of quercetin in 100 ml of methanol. From this solution, five standard solutions were obtained, with concentrations of 5, 50, 100, 125, and 250 mg/L, respectively, and then used to acquire the calibration standard curve (R² = 0.998). Briefly, 125 μL of extract (diluted 1:10 in methanol) was mixed with 37.5 μL of NaNO₂ (5%) and mixed. After 6 min, 75 μL of AlCl₃ (10%) was added and incubated for 5 min after mixing. Then, 250 μL of NaOH (1 M) was added. Finally, the mixture was adjusted with 1.25 ml of distilled water. The absorbance versus a prepared blank was read at 510 nm in the microplate reader (Model Victor Nivo 3S, ILC). The total flavonoid content of the samples was expressed as mg of quercetin equivalents per gram of hemp (mg QE/g hemp). The experiments were made in triplicate.

Terpene’s identification was performed by SPME/GC-MS, using the method described by Stenerson K, with adaptations (26). In this procedure, 1 g of each hemp extract was transferred into a headspace vial. All samples were analyzed using an Equity-1 capillary GC column. Helium was used as carrier gas at 1 ml/min. The inlet temperature was set at 250°C and a split ratio of 1:30. The general extraction procedure was performed with the 100-μm PDMS fiber; the extraction time selected was 30 min during the equilibration time in a thermostatic bath at 40°C with magnetic stirring at 100 rpm. The GC oven temperature gradient started at 45°C, followed by a ramp of 2°C/min to 100°C and another increase from 5°C/min to 250°C. In this study, an analysis of the plant (hemp) was also carried out in order to verify the presence of volatile components, serving later as a means of comparison with the extractions performed with the three systems.

The removal and the quantification of waxes were carried out through a process called “winterization” of oil and were applied to the obtained extracts. The analysis was carried out in triplicate. Briefly, 1–3 g of extract was dissolved in 10–30 ml of ethanol (96%), respectively, and stirred until a homogeneous solution was obtained. The solution was then left to cool down in the freezer (-20°C) to induce wax precipitation for 24 h. The samples were then centrifugated at 6,000 rpm for 15 min to separate the waxes from the solution. The solid residue was then left to evaporate the remaining ethanol overnight and then weighted to calculate the wax concentration (% W/W). The supernatant was then transferred for a volumetric balloon to evaporate the solvent by a rotary evaporator. The recovered dewaxed extracts were kept at 4°C and in darkness until analysis.

Chlorophyll (CHL) content was determined by following the method previously described by Arnon (27). Briefly, approximately 0.1 g of extract was diluted in 10 ml of ethanol, 96% (v/v). After dilution, samples were centrifuged (6,000 rpm for 15 min) and the absorbance of the supernatants measured at 663 and 645 nm. Contents of CHL a, CHL b, and total CHL were calculated as follows:

Results of three measurements were expressed as μg total CHL/g extract.

The antioxidant activity of the extracts was measured based on their scavenging activity of the stable 1,1-diphenyl-2-picrylhydrazyl (DPPH) free radical. About 150 μL of the extracts was added to 4 ml of the DPPH working solution. After incubating for 40 min at room temperature, the absorbance of the preparations was taken at 517 nm by a microplate reader (Model Victor Nivo 3S, ILC). Sample antioxidant activity was compared to standard ascorbic acid concentrations (1–500 μg/ml). Then, the % inhibition was calculated by the following equation:

From the calibration curves, determined from different concentrations of the extracts, the IC50 was obtained. IC50 value denotes the concentration of the sample required to scavenge 50% of the DPPH free radicals.

The solubility of CBD and CBDA in PBS was determined via the shake-flask method, which consisted in dispersing an excessive amount of extract in 2 ml of phosphate-buffered saline solution (PBS Sigma Aldrich, St. Louis, MO, United States) and stirred at 300 rpm, for 24 h, at 37°C in a water bath. When the assay ended, the samples were centrifuged (Model Z 206, Hermie) at 6,000 rpm for 15 min, and the supernatant was collected. The obtained samples were filtered, using a hydrophilic PTFE syringe filter with a 0.22-μm pore size (Filter Lab, Barcelona, Spain) and analyzed by HPLC. The solubility measurements were carried out in triplicate to determine the saturation concentration of CBD + CBDA in the buffer.

The results of cannabinoids, wax, and chlorophyll extraction where all statistically treated with Graph Pad Prism 6. To indicate statistically significant differences between means, the mean value obtained from each DESs tested was compared to the control using one-way ANOVA and a confidence interval of 95% (p = 0.05). When the standard deviations of the group presented significant differences (p < 0.05), the Turkey multiple comparison test was performed to compare each mean value to the different solvents.

Seven DESs were successfully prepared as homogenous liquids, without any crystal precipitation at normal ambient temperature, excluding for Men:StA, which is solid at room temperature. All the systems appear as a transparent liquid except for Pro:Lac, which is a yellowish liquid.

The determination of the physico-chemical properties of DESs is extremely important since they have a significant influence on the solvent properties, affecting solvents suitability for specific applications. Thus, polarity, water content, density, and viscosity of the prepared systems were determined.

Experimental data on polarity obtained using Nile red as a probe and the water content measured by the Karl Fischer titrator are shown in Table 2.

Nile red is a molecule whose florescence is influenced by the polarity of where it is dissolved. The polarity evaluation of Men:StA could not be assessed since this measurement is performed at room temperature at which the solvent is in the solid state, making it impossible to read on the spectrophotometer. The higher the E_TNR value, the more non-polar the solvent is. The polarity values in Table 2 allowed us to distinguish the prepared DESs in two groups: hydrophilic (Low E_TNR) and hydrophobic (High E_TNR). Comparing the results, Lac:Gluc is the most polar solvent, followed by Pro:Lac, Bet:Lac, Men:Lac, Men:Lau, and Men:MyA, the least polar. These values are in consistence with the amount of water present in the solvents. Men:Lac presented the highest water content of all DESs; this is easily explained by the use of Lactic acid (85%) in the composition of the system, which rises the water present in the solvent, decreasing E_TNR, making it slightly more polar than the other hydrophobic DESs.

The determination of density and viscosity was carried out at atmospheric pressure from 293.15 to 338.15 K. Due to their high viscosity, it has not been possible to determine the viscosity in the whole range of temperatures for all of them. Results are shown in Tables 3, 4. The extraction efficiency is substantially influenced by the level of solvent penetration into the biomass structure; the lower the solvent density is, the more effective is its penetration into the raw material’s matrix (28). Experimental density values listed in Table 3 showed that, for all the tested systems, the increase in temperature resulted in the decrease of density. The amino acid-based DESs presented the highest density values, while menthol-based DESs presented the lowest. At 60°C, temperature at which extraction was performed Lac:Gluc presented the highest density, followed by Pro:Lac, Bet:Lac, Men:Lac, Men:Lau, and Men:StA as the lowest. From the viewpoint of the efficiency of penetration into the biomass, it can be supposed that Lac:Gluc would be the least effective.

The high viscosity of DESs can make them difficult to handle in industrial processes during processing and filtration, even though it sharply decreases when temperature increases (28). Experimental viscosity data, reported in Table 4, showed that, with an increase of temperature, a decrease on viscosity was observed. The hydrophilic DESs presented the highest viscosity, being Pro:Lac the more viscous of all systems. At 60°C, Pro:Lac presented the highest viscosity, followed by Pro:Lac, Bet:Lac, Men:Lac, and Men:Lau, as the lowest. These high values at this temperature could result in a less-effective extraction process due to problems in mass transfer and, therefore, a lower extraction yield.

The characterization of the extracts included not only the evaluation of the content in CBD and CBDA but also the quantification of other bioactive compounds, such as the TPC, total flavonoid content, and terpene content. The presence of subproducts of extraction was also evaluated, such as wax and total chlorophyll, in order to evaluate the specificity of the extraction solvents.

In this work, besides the evaluation of the potential of DESs as extractants, we also studied if their presence in the final product could potentiate the bioactivity of the extracts since many articles have claimed the use of DESs for the solubilization of poorly water-soluble drugs and transdermal drug delivery. In that way, antioxidant activity and solubility were evaluated.

Poor aqueous solubility of the terpenophenolic compound cannabidiol (CBD) is a major issue in the widespread use of this therapeutic molecule (38). As previously described, cannabinoids are highly lipophilic, resulting in very low solubility in water, which has been described in the literature to range between 2 and 10 μg/ml (39). Consequently, these compounds are not readily absorbed, and, therefore, a large dosage is required to have medical effect, presenting a major problem for product design and formulation, besides the costs associated. The development of aqueous solubility-enhanced formulations may lead to higher CBD absorption. DESs comprising or acting as solvents of active pharmaceutical ingredients (API-DESs) have been used to design polymeric drug delivery system, overcome polymorphism, enhance a dissolution rate, increase membrane permeability, and improve transdermal delivery (16). In Figure 7, the solubility values of CBD and CBDA in PBS at 37°C to simulate the physiologic conditions using the tested DESs are represented. Hydrophilic DESs present a higher CBD + CBDA solubility in PBS when compared to cannabis oil and hydrophobic DESs. Lac:Gluc and Pro:Lac presented the highest solubility of all tested DESs. These systems are capable of dissolving in PBS, acting like a carrier for cannabinoids and increasing its solubility. Hydrophobic systems, on the other hand, did not improve the solubilization of cannabinoids. Cannabinoids generally have good solubility in triglyceride lipid bases, allowing for easy solubilization in lipid formulations; since they are more stable in this phase, they do not transfer to PBS.

The statistical analysis of the solubility values showed statistically significant differences between the different solvents (one-way ANOVA, p < 0.05); Lac:Gluc and Pro:Lac showed to be statistically different from EtOH (Turkey’s multiple comparisons test, p < 0.05). These results demonstrate that these two systems can, in fact, enhance the solubilization of CBD and CBDA in PBS.