This study was approved by the Ethics Committee of Gonabad University of Medical Sciences, Gonabad, Iran, under approval number IR.GMU.REC.14004.030. Collection of Propolis samples was performed with the permission and verbal informed consent of the participating beekeepers. A. simplex L3 larvae were recovered from naturally infected fish obtained from local fish markets, and no live vertebrate animals were experimentally infected, handled, or euthanized specifically for this study. All procedures involving infected fish tissues and parasite isolation were conducted in accordance with the institutional biosafety and laboratory safety regulations of Gonabad University of Medical Sciences. The study materials were handled using standard microbiological and parasitological safety precautions to minimize biological risk and environmental contamination.

This study was conducted during June 2023 to December 2024 at the Department of Pathobiology, Faculty of Veterinary Medicine, University of Tabriz, Tabriz, Iran.

Propolis samples were collected during the peak resin-gathering season (June–August 2023) from ten beehives located in four Iranian provinces: Tehran (3 hives), Kermanshah (2 hives), Neyshabour (3 hives), and South Khorasan (2 hives). The botanical origin was verified through consultation with local beekeepers. Raw Propolis was immediately transported on ice and stored at −20°C ± 1°C until processing (maximum 2 weeks). The collected Propolis was immediately ground into a fine powder using an electric blender (MKM6003, Bosch, Germany). To facilitate extraction, 100 g of powdered Propolis was mixed with 400 mL of 70% ethanol and sonicated for 2 h. The mixtures were then filtered through Whatman cellulose filters (Whatman Ltd., Buckinghamshire, UK), and the resulting filtrates were dried using a rotary evaporator (40°C, 40 mbar) until complete solvent evaporation. Extraction yields were: Tehran 18.2% ± 0.6%, Kermanshah 15.3% ± 0.5%, Neyshabour 17.1% ± 0.4%, and South Khorasan 19.4% ± 0.7% (mean 17.5% ± 0.9%). The dried extracts were stored at 4°C ± 1°C in dark amber glass bottles, protected from light, and kept in a desiccated environment. Under these conditions, the extracts remained stable for 6 months, with <5% loss of activity. Dried extracts were re-dissolved in 70% ethanol before each experiment to prepare a 10 mg/mL stock solution. Working concentrations (0.25, 0.5, 1.0, and 2.0 mg/mL) were prepared by diluting the stock with distilled water, resulting in final ethanol concentrations of ≤0.7% (v/v), which is below the toxicity threshold for nematodes. A control experiment confirmed that 0.7% ethanol alone showed no significant larval mortality (p > 0.05), indicating that the observed effects were due to the bioactives of Propolis, not the solvent. Before each experimental series, extracts were assessed for clarity, pH (4.5–6.5), and optical density at 280 nm to ensure quality consistency. Following extraction and initial quality control in Iran, the dried hydroalcoholic Propolis extracts from the four geographic regions were prepared for international shipment to a collaborating laboratory in Spain, where the larvicidal assays would be performed.

For the in vitro assays, third-stage larvae (L3) of Anisakis simplex were collected from the visceral organs (including the digestive tract, liver, gonads, and mesenteries) of naturally infected intermediate fish hosts (S. japonicus, Atlantic mackerel) obtained from local fish markets in Spain (n = 30 fish examined; 93% naturally infected). Larvae were isolated under sterile conditions and rinsed three times with sterile 0.9% NaCl solution. Taxonomic identification was confirmed using a stereomicroscope (Olympus, Okayama-shi, Okayama, Japan) based on morphological criteria: third-stage larva with characteristic features including a body length of 250–400 μm, a prominent ventricle in the esophagus, visible cecum, and genital primordium. Only active, motile larvae demonstrating coordinated muscular contractions and responding positively to gentle mechanical stimulation were selected and immediately transferred to sterile physiological saline (0.9% NaCl) maintained at room temperature (up to 4 h post-isolation). Three independent biological replicates were prepared to ensure robustness and reproducibility, each from larvae collected from different fish individuals from separate market sources to minimize potential genotypic or phenotypic bias associated with single-source parasite populations. Upon arrival of the Iranian Propolis extracts in Spain (shipped as described above in the Propolis section), the dried extracts were re-dissolved in 70% ethanol to prepare working solutions. The purity and chemical composition of the transported samples were verified prior to use by gas chromatography–mass spectrometry (GC-MS) analysis conducted in Iran, confirming that no degradation occurred during transit. Subsequently, in vitro larvicidal assays against A. simplex L3 larvae were performed in the Spanish collaborating laboratory using the collected larvae and the re-dissolved Propolis extracts.

The larvicidal activity of ethanolic Propolis extracts against A. simplex L3 larvae was tested in vitro using a standardized protocol. For each concentration tested (0.25, 0.5, 1.0, and 2.0 mg/mL), ten larvae were placed in each well of a 6-well microplate (Falcon™, Corning, USA) containing 2 mL of sterile physiological saline (0.9% NaCl). Each concentration was examined in five technical replicates across three independent biological replicates, resulting in 15 replicate wells per concentration. Microplates were incubated at room temperature (22°C ± 2°C) in darkness to maintain consistent environmental conditions and prevent light-induced changes in larval behavior. Propolis extract solutions were checked for quality using high-performance liquid chromatography (HPLC; Agilent 1200 series, USA) before each experimental run to ensure extract consistency and concentration accuracy. Larval viability was monitored at regular intervals (2, 4, 6, 8, 12, 24, and 48 h) with an inverted stereomicroscope (Olympus, Okayama-shi Okayama, Japan) at ×40 magnification. Larvae were assessed based on standardized criteria: alive (showing coordinated muscular contractions and responding to mechanical stimulation), immobile (no spontaneous movement but responding to gentle stimulation), or dead (no motility and no response to mechanical stimulation). Larval mortality percentage was calculated at 24 and 48 h post-exposure as (number of dead larvae/total larvae per well) × 100. Lethal time values (LT₅₀ and LT₁₀₀), indicating the time needed to reach 50% and 100% larval mortality, were calculated using probit analysis with 95% confidence intervals in GraphPad Prism version 6 (GraphPad Software, San Diego, CA, USA). Using three biological replicates and five technical replicates per condition ensured reliable reproducibility and accounted for biological variability (differences between larvae from different fish sources) and technical variability (procedural differences).

The antioxidant potential of Propolis extracts from different geographic regions was assessed using the 2,2-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging assay, a standard method for measuring free radical scavenging ability in natural products. Various concentrations of Propolis extract (0.25, 0.5, 1.0, and 2.0 mg/mL) were prepared by serial dilution in 70% ethanol and adjusted to a final volume of 40 μL with dimethyl sulfoxide (DMSO; Sigma-Aldrich, USA). A freshly prepared DPPH solution (0.1 mM in absolute ethanol; Sigma-Aldrich, USA) (2.96 mL) was added to each Propolis sample, resulting in a final DPPH concentration of 0.099 mM. The reaction mixtures were incubated in the dark at room temperature (22°C ± 2°C) for exactly 20 min to ensure stable radical-antioxidant balance, which is essential for reproducibility. The absorbance was measured at 517 nm using a spectrophotometer (UV-1900i, Shimadzu, Japan), with a blank cuvet containing 70% ethanol serving as a reference. Three independent replicates were analyzed for each Propolis concentration, each with three technical replicates (n = 9 measurements per concentration). The radical scavenging activity (% RSA) was calculated using the following formula:

% RSA = [(Absorbances (Abs) control − Abs sample) / Abs control] × 100,

where “Abs control” is the absorbance of the DPPH solution without extract (negative control), and “Abs sample” is the absorbance measured in the presence of Propolis extract.

IC₅₀ values (concentration of extract needed to achieve 50% radical scavenging activity) were calculated using non-linear regression analysis (sigmoidal curve fitting) with 95% confidence intervals in GraphPad Prism version 6 (GraphPad Software). Ascorbic acid (vitamin C; Sigma-Aldrich, USA) served as a positive control standard at concentrations of 0.01–1.0 mg/mL to validate assay performance and facilitate comparison of Propolis antioxidant potency with published reference values. To reduce environmental variation, all measurements were performed under consistent ambient conditions and within the same experimental session.

Chemical profiling of Propolis extracts from each geographic region was conducted using GC–MS to identify and quantify volatile and semi-volatile bioactive compounds. Dried Propolis extracts (100 mg per region) were dissolved in HPLC-grade hexane (Merck KGaA, Darmstadt, Germany) in a 1:1 ratio (w/v) to facilitate the extraction of lipophilic compounds. The mixtures were vortexed at maximum speed for 5 min, then incubated at 22°C ± 2°C for 15 min to achieve complete phase separation between the residual Propolis components and the hexane layer. The upper hexane phase containing extracted bioactive compounds was carefully transferred into sterile glass vials using a Pasteur pipette to avoid contamination from the lower aqueous phase. The hexane extracts were filtered through 0.22 μm Polytetrafluoroethylene (PTFE) syringe filter (Millipore, USA) to remove particulates and ensure sample purity before analysis. Aliquots of 1 μL of each filtered hexane extract were injected into the GC–MS instrument (Agilent 7890A gas chromatograph coupled with Agilent 5975C mass selective detector; Agilent Technologies, Santa Clara, CA, USA) using a split ratio of 50:1 to prevent column overloading. Separation was performed using a J&W DB-5 ms capillary column (30 m × 0.25 mm i.d., 0.25 μm film thickness; Agilent Technologies, USA) with helium as the carrier gas at a flow rate of 1.0 mL/min. The temperature program was set as follows: initial temperature of 50°C for 2 min, then ramped at 8°C/min to 280°C and held at 280°C for 5 min (total run time: 34.75 min). The mass spectrometer was operated in electron ionization mode at 70 eV with a scan range of 40–600 m/z 40–600. Data acquisition and processing were performed using the Agilent ChemStation software (Agilent Technologies, USA). Compound identification was achieved by comparing the obtained mass spectra and retention times with those of reference standards in the National Institute of Standards and Technology mass spectral library and with published literature on the chemical composition of Propolis. The relative abundance (%) of each identified compound was calculated from peak area integration, enabling quantitative comparison of chemical profiles across geographic regions and correlation with larvicidal and antioxidant activities.

The cytotoxicity of Propolis extracts was evaluated using the MTT colorimetric assay on African green monkey kidney fibroblast cells (Vero cells; ATCC CCL-81; Geniranlab Co., Iran) to determine non-toxic concentration ranges suitable for therapeutic application. Vero cells (1 × 10⁴ cells/well) were seeded into 96-well tissue culture plates (Falcon™, Corning, USA) and incubated for 24 h at 37°C in a humidified atmosphere with 5% CO₂ for 24 h to allow cell monolayer attachment and stabilization. Cell confluence was verified microscopically (approximately 70%–80% confluence) before treatment to ensure consistent starting conditions. Cells were then exposed to different concentrations of Propolis extract (0.25, 0.5, 1.0, and 2.0 mg/mL) prepared in serum-free Dulbecco’s modified Eagle medium (DMEM; Sigma-Aldrich, USA) in four independent replicates per concentration for 48 h at 37°C in 5% CO₂. Untreated cells (serum-free DMEM alone) served as the negative control (100% viability reference), and cells treated with 10% dimethyl sulfoxide (DMSO; Sigma-Aldrich, USA) served as the positive control (maximum toxicity reference). After 48 h of incubation, 50 μL of serum-free medium and 50 μL of freshly prepared MTT solution (5 mg/mL in phosphate-buffered saline; Thiazolyl Blue Tetrazolium Bromide, 98% purity; Sigma-Aldrich, USA) were added to each well and incubated at 37°C in darkness for exactly 3 h to allow viable cells to metabolize MTT. The resulting formazan crystals were dissolved by removing the supernatant and adding 150 μL of high-purity DMSO (≥99.9%; Sigma-Aldrich, USA) per well, followed by gentle pipetting to ensure complete dissolution and homogenization. The absorbance was measured at 570 nm using a microplate spectrophotometer (UVILINE 9600, AquaLabo, France) with a reference wavelength of 630 nm to subtract the background absorbance. The percentage of cell viability was calculated using the following formula:

% Cell Viability = [(optical density (OD)₅₇₀ of treated sample − OD₅₇₀ of blank) / (OD₅₇₀ of negative control − OD₅₇₀ of blank)] × 100,

where OD₅₇₀ represents optical density measured at 570 nm. Cell viability data were analyzed using three independent experimental replicates, each with four technical replicates per concentration, generating n = 12 measurements per concentration. IC₅₀ values (concentration inducing 50% cell death) were calculated using nonlinear regression analysis (sigmoidal curve fitting) with 95% confidence intervals in GraphPad Prism version 6 (www.graphpad.com). Non-toxic concentration ranges were defined as those that demonstrated≥80% cell viability relative to untreated controls, the accepted safety threshold for evaluating natural products.

All experiments involved three independent biological replicates, each with five technical replicates per condition (n = 15 per group). Data are shown as mean ± standard error of the mean (SEM). Statistical analyses were conducted using GraphPad Prism version 6.0. Before analysis, normality was checked with the Shapiro-Wilk test and homogeneity of variances with Levene’s test (α = 0.05); data that were not normally distributed were analyzed using the Mann-Whitney U test or Kruskal-Wallis test as appropriate. Non-linear regression with sigmoidal curve fitting was used to determine LC₅₀ and IC₅₀ values with 95% confidence intervals. Two-way analysis of variance (ANOVA) was used to assess differences between groups, considering concentration (0.25, 0.5, 1.0, 2.0 mg/mL) and time or region as factors, followed by Tukey’s post hoc test for multiple comparisons. Interaction effects between concentration and time/region were evaluated; if significant (p < 0.05), pairwise comparisons were performed separately for each time point or region. Effect sizes (eta-squared, η²) were calculated for ANOVA results and interpreted as: negligible (<0.01), small (0.01–0.06), medium (0.06–0.14), and large (>0.14). Correlations between chemical composition (GC–MS data) and biological activities were assessed using Pearson’s correlation (parametric) or Spearman’s rank correlation (nonparametric) with 95% confidence intervals; significance was set at |r| or |ρ| > 0.7 and p < 0.05. A p-value < 0.05 was deemed statistically significant (two-tailed). All confidence intervals (CIs) were at 95% for means, IC₅₀, LC₅₀, and correlations. Graphs, including dose–response curves, box plots, and scatter plots, were generated using GraphPad Prism to evaluate data distribution and verify statistical assumptions. The raw data and full statistical output were preserved for transparent reporting.

Propolis extracts showed a strong larvicidal effect against A. simplex L3 larvae, which depended on dose and time, indicating powerful antiparasitic activity across all tested concentrations (Table 1). At 24 h post-exposure, the highest concentration tested (2 mg/mL) caused complete larval mortality (100.0% ± 0.5%), while lower concentrations exhibited a graded dose-dependent response: 90.0% ± 2.1% mortality at 1 mg/mL, 80.0% ± 3.7% at 0.5 mg/mL, and 0% at the lowest concentration (0.25 mg/mL). After 48 h, all tested doses, including the lowest (0.25 mg/mL), resulted in complete or nearly complete larval mortality (100 ± 0%), showing a significant increase in efficacy over time.

The regional variation in Propolis potency highlights the significance of geographic origin in natural product pharmacology. The calculated IC₅₀ values (the concentration needed to achieve 50% larval mortality) after 24 hours varied notably among Propolis samples from different Iranian regions (Figure 1). Tehran extract demonstrated the highest potency (IC₅₀: 20.65 ± 1.2 µg/mL), followed by Kermanshah (74.73 ± 3.5 µg/mL), Neyshabour (84.93 ± 4.1 µg/mL), and South Khorasan (111.23 ± 5.8 µg/mL). This 5.4-fold difference in larvicidal activity between the most potent sample from Tehran and the least potent from South Khorasan shows that biological activity can be significantly affected by regional differences in chemical makeup. Such regional variation has important implications for standardization and quality control in Propolis-based drugs.

The antioxidant activity of Propolis extracts was assessed using the DPPH radical scavenging assay, a standard spectrophotometric method for measuring free radical scavenging ability (Figure 2). A noticeable increase in antioxidant capacity with higher concentrations was observed, with larger Propolis doses leading to significantly greater DPPH radical inhibition (Figure 3). The dose–response relationship clearly shows that as the concentration of Propolis rises, the percentage of DPPH scavenging increases substantially, indicating that the extract’s antioxidant potential depends on concentration and reaches a saturation point. This antioxidant activity aligns with Propolis’s anti-inflammatory and cytoprotective properties, suggesting that both larvicidal and antioxidant effects may be interconnected through the modulation of reactive oxygen species (ROS).

GC–MS analysis identified six major volatile and semi-volatile compounds in the phytochemical makeup of the Propolis extract (Figure 4). The most abundant compound was n-hexane (64.32% of peak area), followed by cyclopentanemethyl (16.62%), hexadecanoic acid (6.03%), cyclohexane (3.27%), pyrrolidine (2.16%), and 2-methylpentane (2.16%), which together made up 94.56% of the detected components. The high presence of hydrocarbons and fatty acids (around 86.97% of the main components) indicates a lipophilic-enriched chemical profile that stands out from typical Propolis profiles, which are usually rich in flavonoids and phenolic acids (Table 2). This unique chemical makeup may directly relate to the observed strong larvicidal activity, as lipophilic compounds are known to disrupt parasite cell membranes and weaken cuticle integrity.

Cytotoxicity was assessed using the MTT cell viability assay on African green monkey kidney (Vero) cells exposed to Propolis extracts for 24, 48, and 72 h across a concentration range (0.25–2.0 mg/mL). Propolis showed minimal toxicity toward mammalian cells at all tested concentrations and time points, with cell viability remaining high throughout all exposure conditions (Table 3 and Figure 5). This favorable safety profile, combined with strong antiparasitic efficacy, supports a promising therapeutic window for developing Propolis-based anthelmintic products.

The renewed interest in natural products over recent decades reflects a growing demand for safer, eco-friendly, and effective alternatives to synthetic drugs, especially in managing infectious and parasitic diseases [20]. Propolis, a resinous substance produced from plant materials by honeybees, has become one of the most versatile bee-derived products, with recognized antimicrobial, antioxidant, anti-inflammatory, and antiparasitic properties [17]. Traditionally used to treat wounds, ulcers, and sore throats, its pharmacological potential is now being confirmed through modern scientific methods [18, 21]. The present study aimed to evaluate the larvicidal and antioxidant activities of hydroalcoholic Propolis extract against A. simplex L3 and to assess its cytotoxicity in vitro.

Propolis showed a strong larvicidal effect against A. simplex L3 depending on dose and time. Complete larval death occurred at 2 mg/mL within 24 h, and lower doses became fully effective after 48 h. This gradual increase in effectiveness highlights Propolis’s potential as a natural larvicide that works over time. The IC₅₀ values varied among Propolis samples from different Iranian regions, with the Tehran extract being most potent, which suggests that geographical differences in chemical makeup influence biological activity [22]. These results align with earlier studies showing that Propolis’s bioactivity varies by region, likely due to differences in local flora and bee diversity. Notably, in this study, Tehran Propolis was about 5.4 times more potent than South Khorasan Propolis, emphasizing the importance of geographic standardization in new product development. This has practical implications for selecting bioresources and ensuring quality control when developing Propolis-based treatments [23].

Although no previous studies have tested Propolis against Anisakis, similar antiparasitic effects have been observed against Schistosoma mansoni [24] and protozoa, including Acanthamoeba [25] and Plasmodium falciparum [26]. Silva et al. [24] showed that Brazilian red Propolis significantly reduced parasite motility, achieved 100% mortality in vitro, and lowered oviposition. Paula et al. [27] found that Brazilian green Propolis decreased hepatic granulomas and egg loads in infected mice. These findings support the current results and reinforce the idea that Propolis has broad-spectrum antiparasitic activity, possibly by disrupting the parasite cuticle and interfering with vital metabolic processes [28].

However, not all helminths respond similarly to Propolis treatment. Rana et al. [29] found no significant effect of ethanolic Propolis extract against Gastrothylax crumenifer, as it did not cause substantial mortality even at high concentrations. This contrast highlights the species-specific mode of action and underscores the importance of targeted testing before applying Propolis broadly across different parasitic groups. The ineffectiveness against G. crumenifer may be due to structural or physiological differences in the tegument, metabolism, or resistance mechanisms of the parasite [30]. This pattern unique to each species suggests that the effectiveness of Propolis may depend on parasite-related factors, including cuticle composition, antioxidant enzyme systems, and metabolic pathways, which vary among helminth species.

Interestingly, combining Propolis with other agents, such as Selenium nanoparticles (SeNPs), seems to boost its effectiveness. Sarhan et al. [31] showed that mixing SeNPs and Egyptian Propolis significantly lowered the adult and larval burdens of Trichinella spiralis, while also reducing inflammatory infiltrates and markers of angiogenesis. These findings suggest that synergistic formulations could enhance the therapeutic performance of Propolis, especially against resistant or tissue-dwelling helminths. Such approaches may be particularly relevant for zoonotic helminths like Anisakis, where systemic and localized inflammation contribute to clinical symptoms [32]. The potential to improve Propolis efficacy through formulation strategies opens new avenues for creating more powerful natural anthelmintic products for treatment purposes.

GC–MS analysis showed that n-hexane, cyclopentanemethyl, and hexadecanoic acid were the main compounds in the Propolis extract. This chemical profile differs from earlier reports where flavonoids and phenolic acids were the primary components [21, 33, 34]. The presence of hydrocarbons and fatty acids in the current samples may indicate regional differences in flora, harvesting techniques, and bee species [35]. These lipophilic compounds might contribute to disrupting membranes in parasites, resulting in loss of motility and eventual death [36].

The distinctive lipophilic-enriched chemical profile observed in this study differs from conventional polyphenol-rich Propolis, suggesting that this phenotype may represent a new category of Propolis with enhanced antiparasitic potential. The high abundance of lipophilic compounds, making up approximately 86.97% of the main constituents detected, likely helps penetrate parasite membranes, damaging their structure and causing osmotic imbalance. However, the exact mechanism of action still needs to be clarified and requires further research using molecular and ultrastructural methods, including transcriptomic analysis and transmission electron microscopy examination.

The extract’s strong antioxidant capacity, as shown by the DPPH assay, supports the idea that its biological effects are partly due to modulating oxidative stress [37]. Parasite killing through oxidative damage has been suggested for several natural compounds [38]. Additionally, antioxidants may help decrease host tissue damage during infection. The dual role of Propolis, combining larvicidal and antioxidant effects, provides a therapeutic benefit, especially in managing anisakiasis-related inflammation and allergic reactions [39].

This dual mechanism, involving parasite elimination through ROS-mediated stress combined with host-protective antioxidant effects, offers a potential advantage over traditional synthetic anthelmintics, which typically target parasite viability without decreasing host inflammatory responses. Such combined pharmacological activity may be especially advantageous in anisakidosis, where both parasite removal and inflammation management are clinically significant.

The MTT assay demonstrated that Propolis was non-toxic to Vero cells across all tested concentrations and exposure durations. High viability levels indicate a strong safety profile, which is crucial for further development of therapeutic or preventive applications. These results align with previous studies assessing the toxicity of Propolis in both cell cultures and animal models [40]. The combination of potent antiparasitic activity and low cytotoxicity suggests a promising therapeutic window for Propolis-based treatments. However, additional in vivo studies and toxicity tests in human tissues are needed before clinical use.

This study provides the first evidence of the larvicidal activity of Propolis against A. simplex, highlighting its potential as a natural anthelmintic agent [6]. However, several limitations should be acknowledged. First, the findings are limited to laboratory conditions, and in vivo validation in animal models of experimental anisakiasis is necessary to assess absorption, bioavailability, and systemic effectiveness. Second, although larval identification followed established morphological criteria and was conducted using naturally infected fish belonging to a known A. simplex host species [41], molecular confirmation via PCR-based ITS region sequencing would strengthen species identification [42]. Third, cytotoxicity evaluation was performed only with Vero cells, and additional safety assessments should include other mammalian cell lines, such as hepatocytes and intestinal epithelial cells, as well as primary human tissues. Fourth, GC–MS analysis primarily focused on volatile and semi-volatile compounds, and more comprehensive profiling, including phenolic compounds and terpenes, is needed for complete chemical characterization. Finally, although lipophilic-mediated membrane disruption and ROS-mediated parasite killing are suggested mechanisms, direct mechanistic evidence requires molecular docking, proteomic analysis, and ultrastructural examination.

Despite these limitations, the strong in vitro efficacy, favorable safety profile, and unique chemical makeup of Iranian Propolis suggest that this natural product is a promising candidate for further development toward therapeutic use. The exact way in which Propolis affects larvae remains unknown and may involve disruption of metabolic pathways, damage to the cuticle, or effects on neuromuscular systems. Identifying the specific active compounds responsible for this activity, including possible synergistic interactions among them, is crucial. Future research should verify these findings in in vivo models of anisakiasis, and advanced formulation techniques, such as nanoencapsulation or emulsion-based systems, should be considered to enhance the solubility, stability, and bioavailability of the active components.