Evaluation of in vitro bioactivity profile of bee propolis extracts delivered by yeast glucan particles

Evaluation of in vitro bioactivity profile of bee propolis extracts delivered by yeast glucan particles Ad´ ela Brejchov´ a a,1 , Eva Kr´ alov´ a a,1 , Ondˇ rej Strnad b , Petra Tˇ reˇ sˇ n´ akov´ a b , Giyaullah Habibullah b , Jana Rýparov´ a Kvirencov´ a c , Vojtˇ ech Hrbek c , Jitka Viktorov´ a b , Denisa Lizoˇ nov´ a a,* , Frantiˇ sek ˇ Stˇ ep´ anek a,** a University of Chemistry and Technology Prague, Department of Chemical Engineering, Technick´ a 3, Prague 6, 166 28, Czech Republic b University of Chemistry and Technology Prague, Department of Biochemistry and Microbiology, Technick´ a 3, Prague 6, 166 28, Czech Republic c University of Chemistry and Technology Prague, Department of Food Analysis and Nutrition, Technick´ a 3, Prague 6, 166 28, Czech Republic ARTICLE INFO Keywords: Propolis Glucan particles Encapsulation Spray drying Dissolution tests Composition In vitro bioactivity ABSTRACT Due to its antibacterial, anticancer and immunomodulatory properties, propolis is widely used in the treatment of various diseases. However, its poor water solubility and bioavailability reduce its effectiveness in vivo. A solution may be found by combining enhanced propolis extraction with loading the extract into a suitable carrier. Here, we investigated glucan particles, obtained from the cell walls of baker’s yeast (Saccharomyces cerevisiae), as promising porous carriers capable of increasing the dispersion of propolis and, thereby, its solubility. First, propolis extracts were prepared using a modified sonication method (70 % ethanol at a propolis/ethanol ratio of 1:10); three types of raw propolis differing in source location and year of harvest were used for comparison. Next, lyophilized propolis extracts were loaded into the glucan particles by spray drying and slurry evaporation to ensure the resulting propolis extract content of 10 wt %. ATR-FTIR spectroscopy and SEM confirmed the successful loading into the glucan particles, and UV–VIS spectrophotometry was used to determine encapsulation efficiency. Dissolution tests showed that the loading into glucan particles led to the formation of a supersaturated solution of propolis extract and the enhancement of its dissolution rate. Subsequently, the biological activity of the encapsulated propolis extract was investigated. Overall, the encapsulated propolis extract showed much higher anti-inflammatory, antioxidant and antimicrobial activity than pure propolis. Thus, our results indicate that the combination of propolis extract with glucan particles increases propolis solubility and bioactivity. 1. Introduction Propolis is a resinous substance produced by honeybees (Apis mellifera L.) to protect their hives from diseases caused by fungi, yeast, and bacteria [1–3]. Propolis composition varies depending on its geographic location and the plants from which it is collected but typically consists of more than 850 chemical components [4,5], mostly flavonoids, phenols, and essential oils [6]. These include, for example: chrysin, apigenin, kaemferol, quercetin, pinocembrin, galangin, 10-hydroxyl-2-decenoic acid, and cinnamic acid. Chrysin is a potent flavone known for its strong anti-inflammatory and antioxidant properties [7,8]. It reduces the inflammation of the immune system, thereby decreasing damage caused by macrophages, neutrophils, and other immune-inflammatory reactions. Chrysin also lowers levels of TNF α , IL-12, and IL-6 and downregulates pro inflammatory enzymes such as COX-2 and NO synthase [7]. Apigenin is a flavone notable for its potential anti-cancer, anti-inflammatory, and neuroprotective effects. It induces apoptosis, inhibits cell proliferation, and blocks angiogenesis in various cancer lines. Apigenin also reduces inflammation by inhibiting the COX-2 and modulating NF-κB signaling, which decreases IL-8 and TNF α production [9,10]. Kaempferol, a flavonol, exhibits significant antioxidant activity and has been associated with reduced risk of chronic diseases such as cardiovascular disease and cancer [11,12]. Quercetin, one of the most abundant flavonoids in the human diet, is known for its anti-inflammatory, antihistamine, and antiviral properties [12,13]. Pinocembrin, a flavanone found * Corresponding author. ** Corresponding author. E-mail addresses: [email protected] (D. Lizoˇ nov´ a), [email protected] (F. ˇ Stˇ ep´ anek). 1 Equal contribution. Contents lists available at ScienceDirect Journal of Drug Delivery Science and Technology journal homepage: www.elsevier.com/locate/jddst https://doi.org/10.1016/j.jddst.2025.107490 Received 17 July 2025; Received in revised form 3 September 2025; Accepted 4 September 2025 Journal of Drug Delivery Science and Technology 114 (2025) 107490 Available online 5 September 2025 1773-2247/© 2025 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY license ( http://creativecommons.org/licenses/by/4.0/ ). prominently in propolis, is well-recognized for its antimicrobial and anti-inflammatory activities. By inhibiting the production of pro-inflammatory cytokines, pinocembrin effectively promotes wound healing and combats infections, making it a valuable therapeutic compound [12]. Galangin, is a flavonol distinguished by its antioxidant, anticancer, anti-inflammatory and antimicrobial effects, protecting cells from oxidative damage and inhibiting the growth of various pathogens [14,15]. 10-hydroxyl-2-decenoic acid is a fatty acid derivative known for its immunomodulatory effects, promoting the proliferation of immune cells and enhancing host defense mechanisms [16]. Cinnamic acid is a phenolic compound with antioxidant and anti-inflammatory properties. It has been investigated for its potential in preventing oxidative stress-related diseases [17]. Thus, propolis has very promising therapeutic potential, including anti-inflammatory, antioxidant, and antimicrobial activities [2,3]. However, it also has unfavorable physicochemical properties which, together with its inherent chemical variability, limit its direct application due to poor solubility and bioavailability [18]. Propolis cannot be used as a raw material, that’s why purification by extraction is usually employed [19]. Extraction of propolis is the most popular technique with the use of various types of solvents such as ethanol, glycerin, propylene glycol, and polyethylene glycol [20]. The most popular one is ethanolic extraction, which is simple and effective while rich in biologically active compounds and obtaining none or low amounts of wax [3,19]. To overcome limitations regarding its strong flavor, poor solubility, and low bioavailability, encapsulation techniques are frequently employed [21–23]. Encapsulation is a process of enclosing the active ingredient into a suitable carrier to protect or improve the properties of the active ingredient. Of the available encapsulation techniques, spray drying appears to be one of the best options because of its wide availability of equipment, low processing cost, and final product stability and quality [24,25]. Moreover, spray drying has previously been used with success for producing amorphous solid dispersions, thereby enhancing solubility and absorption of poorly soluble drugs [21,26]. Several papers have described the spray-dried encapsulation of propolis extract in various carriers. The most widely used carriers are maltodextrins, vinal gum/Arabic gum, and inulin [22,25,27]. Maltodextrins are commonly used for their low viscosity, high water solubility, high bioactive compound retention and good protection of the active ingredient against oxidation [21,22,25,28,29]. Nevertheless, their low emulsifying capacity is a big limitation for poorly water-soluble molecules [22,25]. The use of gums has been shown to change the release kinetics in encapsulation systems. They have good emulsifying ability in aqueous solutions and are odorless and tasteless [22,30,31]. However, the use of such a substance on its own does not provide sufficient protection of the active ingredient [22]. Inulin is known for its prebiotic and dietary fiber effects, and low cost, but its use is hampered by its poor water solubility and low encapsulation efficiency [22,25,32]. More recently, interest has turned to biocompatible drug carriers from natural sources, particularly glucan particles, whose own biological activity could be beneficial [33]. As carriers, glucan particles can promote immunomodulatory effects by interacting with the immune system through Peyer’s patches located in the gut [34–36]. These polysaccharides can be inexpensively and easily obtained from the cell walls of baker’s yeast (Saccharomyces cerevisiae) which consist mainly of (1,3)-β-D-glucan and (1,6)-β-D-glucan in side chains [33,37,38]. As they are living organisms, consistently produced under controlled conditions [37], the size of the yeast cells is highly conserved (5.1 ±1.9 μ m [39]). Glucan particles are porous, inherently insoluble in ethanol and aqueous media but are partially hydrophilic and easily dispersed in water [40]. Furthermore, the encapsulation of poorly soluble compounds in glucan particles has been shown to improve their dissolution properties [40], and protect against evaporation, oxidation and light [27]. Yet, no study has investigated the encapsulation of propolis extract in yeast glucan particles. Here, we encapsulate propolis extracts in glucan particles by spray drying in order to improve their bioactivity and dissolution properties. In in vitro tests, we study the anti-inflammatory, antioxidant, and antibacterial potential of three different propolis samples and compare it to the composites obtained by loading the propolis extracts into the glucan particles. We show that the method of propolis extract delivery employing glucan particles has great potential for its direct application to treat infections and promote wound healing. 2. Materials and methods 2.1. Materials Three different samples of propolis were collected. The first sample (PP 1) was collected in the Czech Republic, ˇ Cesk´ a Lípa, 2018. Other samples were collected in Prague (Czech Republic) in the years 2020 (PP 2) and 2021 (PP 3). Instant baker’s yeast, S.I. Lesaffre (France), was purchased from a local grocery shop. Hydrochloric acid, phosphoric acid, acetone, and isopropanol were from Lach-Ner (Czech Republic). Sodium hydroxide, sodium dihydrogen phosphate dihydrate, methanol and ethanol were from Penta (Czech Republic). Sodium dodecyl sulfate was from SigmaAldrich (USA). Standards of 8 substances of interest, in terms of biological activity, were purchased from Sigma-Aldrich (USA): 10-hydroxyl-2-decenoic acid, cinnamic acid, chrysin, apigenin, kaempferol, quercetin, pinocembrin, galangin. Deionized water was obtained from a Milli-Q purification system supplied by Millipore (USA), HPLC grade methanol from Honeywell (Germany), formic acid from VWR Chemicals (Great Britain), ammonium formate from Merck (Germany) and APCI Positive and Negative Calibration Solutions for the SCIEX X500 System from SCIEX (Canada). TNF alpha Monoclonal Antibody (14-7423-85), TNF alpha Polyclonal Antibody, Biotin (13-7341-85), IL-6 Monoclonal Antibody (147061-85), IL-6 Monoclonal Antibody, Biotin (13-7062-85), Avidin and HRP conjugate (434423) were purchased from Thermo Fisher Scientific (USA) 3,3 ′ ,5,5 ′ -Tetramethylbenzidine Liquid Substrate (T4319), Griess reagent (modified; G4410) and Resazurin sodium salt were from Merck (Germany). 2.2. Collection of propolis Propolis was collected from one of the authors’ (F.ˇ S.) own bees (Apis melliphera carnica, descendants of mother no. CCZ KR1 180201, with morphometric analyses K7821-K7824), stationed at Prague - Hodkoviˇ cky (235 m.a.s.l., beekeeping station reg. no. CZ 90217199) characterised mainly by decorative park and forrest trees, garden fruit trees and graden flowers. Samples from the 2020 and 2021 seasons were used. Both seasons were characterized by typical Central European summer weather, with no anomalies in temperature or precipitation. 2.3. Preparation of dry ethanolic extract of propolis For the propolis extraction, a modified combination of methods published by Khacha-ananda et al. [41] and Pobiega et al. [42] was used. First, crude propolis was frozen with liquid nitrogen and crushed with a pestle to obtain fine particles and facilitate material processing. Then, a solution of propolis in 70 % ethanol (ratio 1:10) was prepared and stirred (300 rpm) overnight, in the dark, at 30 ◦C. Then, the solution was sonicated for 30 min and cleared of particulate impurities by centrifugation (5000 rpm, 10 min). The solution was cooled in the fridge for a couple of hours and filtrated using Whatman® qualitative filter paper, Grade 1 (cellulose filters, 11 μ m). To remove residual wax, the solution was left in the freezer overnight and filtered one more time through a Millipore Express® PLUS membrane filter (polyethersulfone membrane, 0.22 μ m). By this method, all the visually detectable wax A. Brejchov´ a et al. Journal of Drug Delivery Science and Technology 114 (2025) 107490 2 was removed. The solvent was evaporated in a vacuum evaporator (200 rpm) at 30 ◦C. The pure propolis sample was lyophilized to remove residual water. The dried extract was weighted to get yield and finally stored in amber glass vials at 4 ◦C. From this point on, the term “propolis” or “PP” refers to the ethanolic extract of propolis used in the experiments, even if not explicitly stated as “extract” in every instance. 2.4. Preparation of glucan particles Glucan particles (GPs) were obtained from baker s yeast by a multistep procedure, based on the work of Saloˇ n et al. [37]. This procedure involves the removal of internal organelles by several steps: alkali extraction, acidification, and washing with deionized water, isopropanol, and acetone. Firstly, 150 g of dry baker s yeast was gradually poured into the 1M sodium hydroxide solution, mixed with T10 basic Ultra-Turrax (IKA), and stirred for 1 h at 90 ◦C. Then, the suspension was centrifuged (10,000 rpm, 3 min) and the supernatant was discarded. This alkali extraction was repeated two more times with a fresh 1M sodium hydroxide solution. The remaining sediment was resuspended in distilled water and by using 35% hydrochloride acid, the pH was adjusted to reach values between 4 and 5. The suspension was continuously stirred for 2 h at 75 ◦C. After 2 h, the supernatant was discarded and the sediment was resuspended, mixed, and centrifuged (7000 rpm, 10 min). These partial steps were repeated three times with deionized water, four times with isopropanol, and two times with acetone. The final product was left in a fume hood for acetone evaporation to obtain a dry powder, which was then lyophilized for 48 h, and stored in a refrigerator, protected from air humidity. 2.5. Encapsulation by spray drying Propolis (at a ratio of 1:9 with respect to GPs) was dissolved in ethanol. GPs were then added to the solution, and the resulting suspension was homogenized using T10 basic Ultra-Turrax (IKA). The volume of used ethanol was calculated to achieve the final concentration 20 mg/mL of GPs in suspension. The initial loading of propolis extract was set to 10 wt % relative to the total mass of the formulation. This value was selected as a starting point based on literature data [26], suggesting that such a loading is likely to ensure complete encapsulation of the active compound. The spray drying was performed on a Mini Spray Dryer B-290 (Büchi AG, Switzerland) with an ultrasonic nozzle under a nitrogen atmosphere. The process parameters, set according to the work of Ruphuy et al. [26], were: inlet gas temperature 120 ◦C, flow rate of the suspension 5 mL/min, ultrasonic nozzle set to 2.4 W, outlet gas temperature 70 ◦C. The final product, comprising dry propolis-loaded glucan particles, was stored in a refrigerator. Further studies with higher propolis loading will be considered if sufficient encapsulation efficiency and promising biological activity are observed. 2.6. Encapsulation by rotary evaporator Similarly to the spray-drying method, the concentration of propolis (PP) was initially set to achieve 10 wt % loading in the final GPs/PP composites, to evaluate the feasibility of the encapsulation process, with the intention to increase the loading in case of favorable results. GPs (10 mg/mL) were added into the ethanolic solution of propolis, and the suspension was homogenized in a round bottom flask using T10 basic Ultra-Turrax (IKA). After the homogenization, the suspension was placed in a rotary evaporator. Ethanol was removed by slowly decreasing the pressure, starting from the atmospheric pressure up to 80 mbar, by setting the rotary evaporator to 175 rpm and the water bath heated to 60 ◦C. The resulting powder was lyophilized for 48 h to remove residual moisture. 2.7. Characterization of GPs/PP composites GPs/PP composites, a material formed by combining glucan particles as a carrier and propolis as the active ingredient, were characterized based on their physicochemical properties. 2.7.1. Scanning electron microscopy The surface morphology and shape of propolis extracts, empty GPs, and GPs/PP composites were studied by Scanning Electron Microscope (SEM) Joel JCM-5700. Prior to the analysis, the samples were coated with a thin layer (~5 nm) of gold with an Emitech K550X. 2.7.2. Size distribution The size distribution of GPs was evaluated by the laser scattering particle size distribution analyzer Horiba Partica LA-950V2. Before the measurement, GPs were dispersed in a 1:1 (v/v) mixture of distilled water and ethanol, and subjected to sonication. 2.7.3. X-ray powder diffraction The crystallinity of propolis extracts, empty GPs, and GPs/PP composites was evaluated by powder X-ray diffraction using a PANalytical X’Pert PRO diffractometer with High Score Plus software. Scans were taken from 5◦to 50◦2θ angle. 2.7.4. Attenuated total reflectance Fourier transform infrared spectroscopy Characteristic bands of propolis extracts, empty GPs, and GPs/PP composites were identified by a Nicolet iZ10 FTIR (Fourier Transform Infrared) module with Attenuated Total Reflectance (ATR-FTIR) on a ZnSe crystal. The spectra were collected as 64 co-added scans in the range of 4000-700 cm −1 with a resolution of 4 cm −1 . 2.7.5. Nitrogen sorption measurements The nitrogen sorption, at liquid nitrogen temperature (77 K), was measured to determine the specific area and pore distribution of empty GPs and GPs/PP composites. The specific area was evaluated by the BET method, and the pore diameters were calculated using BJH method from the desorption part of the isotherm. The measurements were carried out on a 3Flex 5.01 instrument from Micromeritics, USA. 2.8. Composition of propolis extracts and composites 2.8.1. Sample preparation In the case of both PP and GPs samples, propolis extracts were prepared in methanol with a concentration of 10 mg/mL. To evaluate the concentration, extracted ion chromatograms of 10x or 100x diluted extracts of these samples were used for the majority of monitored analytes, as needed, to ensure that the intensity of the evaluated analyte was within the range of the calibration curve for the given standard. GPs/PP composites were prepared independently in three separate batches and each sample was analyzed twice, to evaluate the homogeneity of the preparation process. 2.8.2. Instrumental condition The chromatographic separation was performed using a Dionex UltiMate 3000 RS UHPLC system (Thermo Fisher Scientific, USA) with a reversed-phase BEH C18 column (2.1 ×100 mm, 1.8 μ m particle size, Waters, USA) at 60 ◦C. The mobile phase consisted of 5 mM ammonium formate and 0.1 % formic acid in (A) water/methanol (95/5, v/v) and (B) isopropanol/methanol/water (65/30/5, v/v/v) with a gradient elution: 0–2 min 90 % (A), 2–7 min 50–20 % (A), 7–13 min 20–0 % (A), 13–22 min 0 % (A), 22–22.1 min 0–90 % (A), 22.1–24 min 90 % (A). The mobile phase flow rate was 0.4 mL/min and the sample injection volume was 2 μ L. Mass spectrometric detection was performed using a TripleTOF 6600 instrument (SCIEX, Canada) equipped with a Duo Spray with a separated ESI ion source and atmospheric-pressure chemical ionization (APCI). ESI A. Brejchov´ a et al. Journal of Drug Delivery Science and Technology 114 (2025) 107490 3 was used for the measurement of extracts of the sample and APCI was used for the exact mass calibration of the instrument. The parameters of the ESI ion source were as follows: capillary voltage: +5000 V in ESI+ and - 4500 V in ESI-; collision energy 35 ±15 V in ESI+and - 35 ±15 V in ESI-; declustering potential: 80 V in ESI+and - 80 V in ESI-; desolvation temperature: 480 ◦C; curtain gas: 35 psi; drying gas pressure: 55 psi and nebulizing gas pressure: 55 psi. TOF MS method and Information Dependent Acquisition (IDA) method were employed to record full MS and MS/MS spectra at the same time. The m/z range was between 100 and 1200 Da for MS and between 50 and 1200 Da for MS/MS. An automatic m/z calibration was performed on every 6 samples using an APCI Positive or Negative Calibration Solution. Analyst TF 1.7.1 software (SCIEX, Canada) was used for instrument control and data acquisition. 2.8.3. Targeted screening of biologically active compounds SCIEX OS 1.5 software (SCIEX, Canada) was used for targeted screening of biologically active compounds. When a substance with a searched formula was detected in an extract of the sample (considering accurate mass, mass error, and isotopic pattern in MS), the measured MS/MS spectrum was compared to the literature and available databases such as PubChem (https://pubchem.ncbi.nlm.nih.gov/), ChemSpider (http://www.chemspider.com/), mzCloud (https://www. mzcloud.org/home), or METLIN (https://metlin.scripps.edu/index.ph p). The internal database of these compounds was created using LibraryView 1.0.3. software (SCIEX, Canada) and was subsequently imported into the SCIEX OS 1.5 software (SCIEX, Canada). The formula, adduct, retention time, and MS/MS spectra were saved in this library. Data evaluation was performed using SCIEX OS 1.5 software (SCIEX, Canada). In addition, standards of detected important substances were analyzed to confirm their identification and for their quantitative analysis. The identification was confirmed based on the agreement of the retention time and MS/MS fragmentation spectra. Quantification was performed based on measuring the exact m/z value, integrating the peak area, and using the external calibration of appropriate standards (0.005–10 μ g/mL in ethanol). 2.9. Validation of UV–VIS spectrophotometric method To evaluate linearity, standard stock solutions of the drug were prepared in a suitable solvent and serially diluted to obtain concentrations ranging from 1.2 to 31.7 mg/L. Each concentration was measured in triplicate using a UV–VIS spectrophotometer at the determined λ max . The Limit of Detection (LOD) and the Limit of Quantification (LOQ) were calculated based on statistical analysis of the linearity data. Blank glucan particles and solvent blanks were also analyzed to assess specificity. Precision was evaluated by intra-day repeatability across three concentration levels. For accuracy assessment, known amounts of drug were spiked into blank glucan particles and extracted using the same procedure applied to loaded samples. The extract was analyzed for recovery efficiency. 2.10. Encapsulation efficiency To investigate the amount of propolis extract incorporated into GPs, GPs/PP composites (6 mg) were mixed with 25 mL of ethanol and subjected to ultrasonic extraction for 5, 10, and 30 min. A total of nine samples were analyzed, with three replicates for each sonication time. The aim was to compare the effectiveness of different extraction times in releasing propolis from GPs into the ethanolic medium and to identify the optimal extraction time. Following this, GPs were separated by centrifugation at 10,000 rpm for 3 min. The centrifugation conditions (centrifugation speed and duration) were optimized based on preliminary experiments to ensure effective separation of the solid fraction (glucan particles) while minimizing losses of the encapsulated propolis. Finally, the supernatant was filtered through a 25 mm syringe filter with 0.45 μ m polyethersulfone membrane. The concentration of propolis in the collected filtrate was quantified using a previously validated UV–VIS spectrophotometric method above (2.9 Validation of UV–VIS spectrophotometric method). Absorbance was at 316 nm using UV–VIS spectrophotometer Analytik Jena Specord 205 and the concentration was calculated according to the calibration curve in ethanol prepared from pure propolis extracts (λ abs =316 nm; y (PP 1) =31.978 x - 0.0042, R 2 =0.9999; y (PP 2) =33.211 x - 0.0086, R 2 = 0.9996; y (PP 3) =38.859 x - 0.002, R 2 =1). The samples were measured in triplicate. The efficiency of the encapsulation process (EE) was calculated as follows: EE (%) = 100 experimental content of propolis in GPs theoretical content of propolis in GPs , where the experimental content of propolis in GPs is the amount of extracted propolis from GPs into ethanol, and the theoretical content of propolis in GPs is the theoretical amount of propolis initially weighed before the encapsulation process. 2.11. Propolis release kinetics The release kinetics of propolis was observed in two types of media. First dissolution tests were carried out in USP 2 (mini-paddle) apparatus coupled to UV–VIS spectrophotometer Specord 200 Plus (Analytik Jena, Germany), to immediately measure the rate and extent of release of propolis from GPs by measuring absorption at 316 nm. Each dissolution vessel was filled with 200 mL of phosphate buffer with pH 6.8 and preheated to 37 ◦C. Afterward, either propolis extracts or GPs/PP composites were placed into the media and continuously stirred (150 rpm). The GPs/PP composites’ weight corresponded to the dose of 8 mg of propolis. Samples were collected at pre-defined time points (0–60 min) while filtered through 0.22 μ m filters to separate the dissolved fraction from the propolis-loaded particles. Further tests were intended to show dissolution rates under sink conditions. A standard 900 mL USP 2 (paddle) round-bottom dissolution vessels consisting of phosphate buffer (pH 6.8) with the addition of 1 % sodium dodecyl sulfate (SDS) were used. The media were maintained at 37 ◦C and stirred at 75 rpm. Approximately 36 mg of dose of crude or encapsulated propolis extracts were collected at predefined points (0–48 h), filtered through 0.22 μ m filters, and measured on Lightwave II +UV–VIS spectrophotometer (Biochrom Ltd, England) at 316 nm. Final results were calculated according to the calibration curve in ethanol prepared from pure propolis extracts (λ abs =316 nm; y (PP 1) =31.633 x - 0.0006, R 2 =0.9992; y (PP 2) =35.707 x - 0.0036, R 2 =0.999; y (PP 3) =28.802 x - 0.0043, R 2 =0.9996). All measurements were carried out in triplicate. 2.12. In vitro testing 2.12.1. Sample preparation To determine the effect of solvents on biological activities, the propolis extracts were dissolved in three different solvents - DMSO, ethanol, and cell cultivation medium. Stock solution of propolis in DMSO and ethanol was 100 g/L and maximum tested concentrations were 1 g/L, thus resulting in a final organic solvent concentration of 1 %. Stock solution of propolis in cell cultivation medium was prepared as saturated solution, the amount of propolis was determined spectrophotometrically using a propolis calibration curve. The highest tested concentration of propolis in the cultivation medium corresponded to a twice diluted saturated solution. The amount of propolis in GPs was calculated according to loading capacity. Both GPs and GPs/PP composites were dispersed in the test medium at a concentration of 20 g/L. Dissolution of was achieved by sonication for 5 min, in order to ensure A. Brejchov´ a et al. Journal of Drug Delivery Science and Technology 114 (2025) 107490 4 proper glucan particle dispersion before testing. The maximum tested concentration of GPs/PP composites was 2 g/L, which resulted in a test medium concentration of 1 %. 2.12.2. Cytotoxicity The hTERT-immortalized epithelial proximal tubular cells (RPTEC/ TERT1, CHT-003-0002Evercyte, Wien, Austria) were plated at a concentration of 1⋅10 5 cells/mL for 24 h in ProxUp2 medium (Evercyte). After that, the medium was replaced with a new one with the addition of tested substances. After 72 h, cell viability was measured by the resazurin test [43]. Briefly, the cells were incubated with resazurin solution (30 mg/L) for 30 min, then fluorescence was measured at 560/590 nm using a SpectraMax i3x Multi-Mode Microplate Reader (Molecular Devices, San Jose, CA, USA). The results were expressed as the percentage of viable cells relative to the untreated control. 2.12.3. Anti-inflammatory activity Nitric oxide (NO) production and cytokine release (TNF α and IL-6) were quantified using RAW 264.7 cells (Merck, 91062702) according to the previously described method by Kˇ ríˇ zkovsk´ a et al. [44], briefly as follows. RAW 264.7 macrophages were stimulated with LPS (100 μ g/L) for 24 h, and NO production was quantified in the supernatant using the Griess reagent (0.04 g/mL) at a 1:1 ratio, with absorbance measured at 540 nm after 15 min of incubation. Cytokine levels in the supernatant were determined by ELISA, where the supernatant was incubated with capture antibodies coated onto 96-well plates, followed by incubation with biotinylated antibodies and signal development using streptavidin–horseradish peroxidase (HRP) and tetramethylbenzidine (TMB) substrate. Concentrations of samples, that were cytotoxic, were excluded, see chapter 3.7. 2.12.4. Antioxidant activity Antioxidant activity was quantified using RAW 264.7 cells. Cells were seeded at confluency overnight. After that, the medium was replaced with a medium enriched with 12.5 mg/L concentration of fluorescein and various concentrations of the tested substances (concentrations of samples, that were cytotoxic, were excluded, see chapter 3.7). After 1 h, the medium was replaced with a solution of 2,2 ′ -Azobis (2-amidinopropane) dihydrochloride (AAPH) in PBS with a concentration of 16.3 mg/L. The fluorescence signal was read at 5min intervals for 42 h. The IC 50 values were calculated from a comparison of the guidelines according to the formula: Radical formation (%) = 100 slopesample −slopeNC slopePC −slopeNC , where slope indicates the change of fluorescence over time. The AAPH solution was chosen as the positive control (PC) and the PBS solution as the negative control (NC). 2.12.5. Antimicrobial activity Bacteria Staphylococcus aureus (DBM 7002) and Pseudomonas aeruginosa (CCM 3955) cultivated in MH (Mueller Hinton, Merck) and yeast Candida albicans (DBM 2186) cultivated in MEA (Malt Extract Agar, Merck) were incubated overnight. Microbial cultures were then prepared according to ISO 20776-1. Then the cells were added to the plate with the test substances. After 24 h, the resazurin assay was used to test cell viability [43]. The fluorescence was measured at 544/590 nm using Fluoroskan Ascent FL (Thermo Fisher Scientific). 2.12.6. In vitro wound healing assay The wound healing ability of propolis and encapsulated propolis was evaluated in vitro by scratch assay using keratinocytes (HaCaT, C0055C, Thermo Fisher Scientific) and expressed as the percentage of wound closure at different time intervals according to the previously described method [45]. In this work, HaCaT cells were seeded at the concentration of 1⋅10 5 cells/mL. The concentration of all tested samples was 5 mg/L. As a control, empty GPs and medium without samples were used. Scratch area images were captured at 24, 48, and 72 h. 3. Results and discussion 3.1. Physical appearance of propolis and glucan particles In this work, three different types of propolis, collected in ˇ Cesk´ a Lípa in 2018 (PP 1) and in Prague in the years 2020 (PP 2) and 2021 (PP 3) were used. These differ not only in physical appearance but also in composition and biological activity. Fig. 1 shows images of raw materials, so as materials after processing and lyophilization. All the raw propolis samples were dark and very sticky (Fig. 1A–C). Subsequent extraction effectively eliminated any visible wax while also improving the flowability of the final product. The final products differed in color; Fig. 1. Images of: (A) raw PP 1, (B) raw PP 2, (C) raw PP 3, (D) baker’s yeast, (E) PP 1 extract, (F) PP 2 extract, (G) PP 3 extract, (H) lyophilized GPs. A. Brejchov´ a et al. Journal of Drug Delivery Science and Technology 114 (2025) 107490 5 while PP 1 (Fig. 1E) was dark brown, PP 2 (Fig. 1F) was brighter, but still brown with orange shades, and PP 3 (Fig. 1G) had more shades of a yellow color. The yield of GPs (Fig. 1H) was 6.88 ±1.27 %, and of propolis extracts, prepared by our modified method, was: 48.64 % of PP 1, 56.84 % of PP 2, and 71.41 % of PP 3. Even though the same procedure was used for all types of propolis extracts, yield differences occurred, so it is certain that the initial content of wax and insoluble impurities has a great influence on product yield. 3.2. Characterization of GPs/PP composites The physicochemical properties of the prepared GPs/PP composites were similar for all types of propolis extracts. For the purpose of brevity, some of the following paragraphs show only the data from PP 3 samples, whereas in these cases, the data from other samples (PP 1, PP 2) can be found in the Supplementary Information. 3.2.1. Scanning electron microscopy The samples of propolis extract, empty GPs, and the encapsulated propolis in GPs by either spray drying or rotary evaporation were studied by scanning electron microscopy. Images are shown in Fig. 2. In all samples containing empty GPs (Fig. 2B), the SEM images showed the typical 2–4 μ m ellipsoidal morphology with a wrinkled surface, caused by the hydrolysis of the outer cell wall of GPs and intracellular components, which is consistent with already published articles [26,40,46]. GPs loaded with PP 3 by spray drying (Fig. 2C) or rotary evaporation (Fig. 2D) did not reveal any significant difference compared to empty GPs, which may indicate that all the propolis is comprised in the GPs/PP composites and does not form any new structures. The appearance of other propolis extracts (PP 1, PP 2) was alike after loading in GPs (see Supplementary Information 1, Figs. S1 and S2). Similar results from the successful loading of natural active ingredient into GPs were reported by ˇ Salamúnov´ a et al. [33]. 3.2.2. Size distribution Particle size distribution of pure GPs was measured using laser diffraction in a 1:1 (v/v) mixture of water and ethanol to prevent excessive swelling, which occurs when the particles are dispersed in water. The GPs were then subjected to sonication during which agglomerates were disrupted into individual particles or smaller clusters, resulting in homogenous dispersion. The resulting volume based Fig. 2. Scanning electron microscopy (SEM) images of (A) PP 3 extract, (B) empty GPs, (C) spray dried GPs/PP 3 composites, (D) rotary evaporated GPs/PP 3 composites. Scale bars represent (A) 100 μ m, and (B, C, D) 10 μ m. Fig. 3. Particle size distribution of pure GPs. A. Brejchov´ a et al. Journal of Drug Delivery Science and Technology 114 (2025) 107490 6 histogram (Fig. 3) revealed a unimodal distribution with the mojaroty of particles concentrated between 1.5 and 7 μ m. The volume mean particle size of GPs determined by laser diffraction was 3.46 ±2.20 μ m. This relatively high variation suggests residual polydispersity, likely due to natural particle variability or minor aggregation. The D 10 , D 50 , D 90 values were 1.51 μ m, 2.98 μ m, and 6.72 μ m, respectively. The observed size range falls within values commonly reported for glucan particles in the literature [26,39]. 3.2.3. X-ray powder diffraction Empty GPs were found to be fully amorphous (Fig. 4A) when analyzed by XRPD. Both propolis extracts PP 3 (Fig. 4B) and PP 2 were amorphous, while only a small percentage of crystalline form was found in PP 1 (see Supplementary Information 2, Figs. S3 and S4). All propolis extracts loaded in GPs (see GPs/PP3 composites in Fig. 4C and D) were completely amorphous, which is expressed by a characteristic band at 20◦. 3.2.4. Attenuated total reflectance fourier transform infrared spectroscopy ATR-FTIR is a frequently used method for a chemical structure characterization of GPs and their typical chemical groups per literature [47,48]. The characteristic bands for (1,3)- and/or (1,6)-linked β-glucans are 1160, 1078, 1041, and 889 cm −1 . The absorption band at 1160 cm −1 corresponds to COC and CC stretching vibrations. Absorption bands at 1070 cm −1 and 1040 cm −1 relate to the stretching vibrations of CO and CC. The band at 890 cm −1 belongs to β-glycosidic linkages (C1H). The reported region for propolis extract is between 1100 and 1800 cm −1 , which, associated with the CO group is ascribed to the flavonoids and lipids [28]. In all spectra (see Fig. 5 and Supplementary Information 3, Figs. S5 and S6), the peak in the area around 1550 cm −1 corresponded to propolis extract and confirmed successful loading. 3.2.5. Nitrogen sorption measurements Nitrogen sorption measurements can provide detailed information on, for instance, the surface area or volume of pores. For this purpose, the empty GPs and GPs/PP were analyzed by this method, and the results are shown in Table 1 and Fig. 6. The empty GPs exhibited the highest specific surface area and volume of pores, with an average pore size of approximately 9 nm. This high porosity results from the removal of internal organells and lyophilization drying process during the preparation which are crucial for their functionality in efficiently encapsulating and also releasing the particular active ingredient making them a promising candidate for drug delivery systems. After the encapsulation of propolis extracts into the porous structure of GPs, we observe a decrease in both surface area and pore volume, which is primarily caused by propolis extracts filling the pores. 3.3. Validation of UV–VIS spectrophotometric method The UV–VIS spectrophotometric method showed linearity over the Table 1 Nitrogen sorption measurements of specific surface area and desorption total pore volume of pores of empty GPs and GPs/PP composites. BET surface (m 2 /g) Pore volume (mm 2 /g) GPs 77.54 189.63 GPs/PP 1 30.97 100.00 GPs/PP 2 33.96 124.55 GPs/PP 3 33.68 110.52 Fig. 6. Barrett, Joyner & Halenda (BJH) desorption pore distribution of empty GPs and GPs/PP composites. Fig. 4. X-raypowder diffraction (XRPD) patterns of (A) pure GPs, (B) PP 3 extract, (C) spray dried GPs/PP 3 composites and (D) rotary evaporated GPs/PP 3 composites. Fig. 5. Attenuated Total Reflectance Fourier Transform Infrared (ATR-FTIR) spectra of (A) pure GPs, (B) PP 3 extract, (C) spray dried GPs/PP 3 composites and (D) rotary evaporated GPs/PP 3 composites. A. Brejchov´ a et al. Journal of Drug Delivery Science and Technology 114 (2025) 107490 7 tested concentration range (R 2 =0.999). The regression equation was y =0.0343 x +0.1895. Precision testing resulted in % RSD values ranging from 0.33 % to 0.47 %, indicating consistent performance. Accuracy values ranged from 95 % to 99 % recovery with % RSD values below 5 %, which meets the acceptance criteria defined in ICH Q2(R2) guidelines [49]. The LOD and LOQ were calculated as 1.49 mg/L and 4.50 mg/L, respectively, with LOQ corresponding to an absorbance of 0.34 AU. The evaluation of LOD and LOQ reflects standard validation practices documented in the literature [50,51]. These results confirm that the method is suitable for quantifying drug loading in GPs with acceptable sensitivity, accuracy, and reproducibility. 3.4. Encapsulation efficiency The content of propolis encapsulated in GPs was evaluated by ethanol extraction, determined by UV–VIS spectrophotometry, calculated according to the equation described in the chapter 2.10, and is presented in Fig. 7. The spectrum of raw propolis has two characteristic peaks with local maxima at 294 nm and 316 nm, whereas GPs also absorb at the wavelength of 290 nm. The spectra are provided in Supplementary Information 4, Fig. S7. Because of this spectral overlap, quantification of propolis was performed at 316 nm, where the absorbance is more specific to propolis and less affected by GPs. In the absence of GPs, propolis can also be determined at 290 nm. To optimize the ethanol extraction procedure, different sonication times (5, 10, and 30 min) were tested. The measured concentrations of propolis extracted after 5, 10, and 30 min were comparable, indicating no significant increase in yield occurred beyond 5 min. This suggests that a 10 min sonication is sufficient to achieve complete release of the encapsulated propolis from GPs. Therefore, 10 min was selected as the optimal compromise between extraction efficiency and processing time for all ethanol extraction experiments. UV–VIS spectrophotometry was selected because the system under study is a complex multi-component natural mixture. The aim of the analysis was not to separate and quantify individual components, as would be typical for UHPLC analysis, but rather to determine the total amount of propolis extract released from the particles. For this purpose, the analyte was considered as the whole extract, and UV–VIS provided sufficient sensitivity and reliability. Moreover, since biological activity is the focus of this work, it is important to highlight that active compounds act synergistically as a mixture, which is characteristic of natural products like propolis. Thus, the measured total propolis extract corresponds to the biologically active mixture responsible for the observed activity. In addition to UV–VIS quantification, UHPLC methanolic fingerprint analysis was performed (see Supplementary Information 6, Fig. S9) to confirm the completeness of propolis encapsulation. The UHPLC profiles showed characteristic peaks of propolis in the encapsulated samples, supporting the reliability of the UV–VIS method and indicating minimal loss. For all samples, the initial loading of propolis extract in the GPs was targeted to 10 wt %, based on literature data [26], which suggests that this concentration is optimal for ensuring complete encapsulation. However, the theoretical content of propolis used for calculating encapsulation efficiency was determined individually for each sample based on the exact mass of propolis extract added during the specific encapsulation process (the theoretical content of propolis in GPs). Two encapsulation methods were compared: spray drying and rotary evaporation. Although incorporation in rotary evaporator resulted in higher yields (88.49 % for PP 1, 80.89 % for PP 2, and 86.86 % for PP 3), compared to spray drying (48.91 % for PP 1, 49.73 % for PP 2, and 49.71 % for PP 3), the encapsulation efficiencies were significantly better with spray drying (Fig. 7). The lower encapsulation efficiencies, obtained using a rotary evaporator, could be attributed to insufficient encapsulation, which was visually observed during the process as sticking of the propolis to the used equipment. In addition, after encapsulation of propolis by spray drying, the powder was uniform, fine, and with good flowability, whereas rotary evaporated samples were less homogenous. For these reasons, our following work proceeded with only spray-dried composites. Additionally, the percentage of the actual loading of the propolis extract was evaluated for each encapsulation method and each propolis type. For spray drying, the loadings were: 8.61 ±0.12 % for PP 1, 9.86 ±0.09 % for PP 2, and 8.62 ±0.24 % for PP 3. In comparison, rotary evaporation resulted in lower loadings: 6.36 ±0.20 % for PP 1, 6.28 ± 0.12 % for PP 2, and 6.11 ±0.16 % for PP 3. All values were determined in triplicate and are reported as mean ±standard deviation. These results confirm the technological advantage of spray drying not only in terms of encapsulation efficiency but also in terms of the loading. The improved loading can be attributed to better process control and reduced material losses during drying, as propolis visibly adhered to equipment surfaces during rotary evaporation. 3.5. Composition of propolis extracts and composites The diverse chemical composition inherent to different propolis types makes characterizing its full chemical profile a significant challenge. To effectively manage this complexity, we selected eight standards, including chrysin, apigenin, kaemferol, quercetin, pinocembrin, galangin, 10-hydroxyl-2-decenoic acid, and cinnamic acid, which are among the most commonly represented and pharmacologically active substances found in propolis samples mainly across Europe. According to their geographical origin and the presence of these typical chemical compounds, it is assumed that the botanical origin belongs to the genus Popolus [52–54]. The use of ultra-high performance liquid chromatography (UHPLC) enabled us to quantify these compounds, which is crucial in understanding how the composition affects the therapeutic potential of these bioactive compounds. Our analyses revealed that 10-hydroxyl-2-decenoic acid and cinnamic acid were not detected in the samples (<5 and 10 μ g/g). The data presented in Fig. 8 show that galangin, chrysin, and pinocembrin were predominant in the propolis samples, consistent with other studies [1,53,55,56]. This finding also correlates with the known chemical profiles of propolis from similar geographical regions. The high content of these bioactive compounds highlights the potential health benefits, particularly antioxidant, anti-inflammatory, and antimicrobial effects. However, the overall complexity of propolis makes it difficult to attribute the activity to a single molecule, which is explained Fig. 7. Encapsulation efficiency (EE) of propolis extracts (PP) after encapsulation by spray drying (solid fill) and rotary evaporation (downward diagonal). spray dried GPs/PP 1; rotary evaporated GPs/PP 1, spray dried GPs/PP 2, rotary evaporated GPs/PP 2, spray dried GPs/PP 3, rotary evaporated GPs/PP 3. The error bars represent the standard deviation (n =3). A. Brejchov´ a et al. Journal of Drug Delivery Science and Technology 114 (2025) 107490 8 by the possible synergy between the components [57]. Furthermore, to verify that all the substances contained in the propolis get inside the glucan particles during the encapsulation process, methanolic fingerprints of propolis extracts and GPs/PP composites, in positive and negative electrospray ionization modes, were measured and compared. Worth noticing is that according to the individual signals (chromatographic peaks) in the chromatographic records (see Supplementary Information 6, Fig. S9), it is evident that the transfer of substances from propolis to glucan particles, i.e. encapsulation, is qualitatively non-selective. This means that all propolis substances have been encapsulated in GPs. On the other hand, the encapsulation process is not quantitative, which is indicated by the different ratios of some compounds in GP/PP composites in comparison to propolis extracts. 3.6. Dissolution tests Dissolution profiles shown in Fig. 9 were obtained in a phosphate buffer of pH 6.8 for propolis extracts and GPs/PP composites prepared by spray drying with a propolis dose of 8 mg. We performed the study in conditions without SDS to assess the ability of propolis to create a supersaturated solution, which can improve bioavailability. Moreover, it helps to evaluate a dissolution behavior without interference. This ensures that the results reflect the propolis’s natural properties rather than being influenced by surfactants, which could artificially enhance solubility. There was a significant improvement in the dissolution rate in the case of all propolis types after their loading into GPs relative to the propolis extracts alone. Regarding propolis extracts, dissolution profiles had a similar course for all tested samples (Fig. 9). A small portion dissolved within the first 5 min, and then the dissolution slowed down, with the total dissolved fraction not exceeding 10 % over the course of 60 min. There were slight differences between the final dissolved fractions, where the PP 1 released fraction was almost 8 %, which corresponds to 3.13 ±0.23 mg/L. Propolis that was released the least, nearly 4 % (1.52 ±0.34 mg/L), was PP 2. The variety of reached concentrations could be caused by differences in propolis extract composition. A significant increase in the dissolution rate of all propolis samples was observed after the encapsulation into GPs (Fig. 9). GPs/PP 2 composites manifested the lowest release. This curve differed from others because in just 5 min, the medium became supersaturated and at this point, the concentration of active ingredient was at the highest peak and decreased from then on, which is attributed to the precipitation. After the first 15 min, the concentration of PP 2 released from GPs was 9.12 ± 0.80 mg/L and remained relatively stable throughout the rest of the dissolution test. The final concentration of PP 2 corresponds to almost 23 % of the dose. On the other hand, the remaining two composites, GPs/PP 1 and GPs/PP 3 behaved quite alike. In both cases, there was rapid release in the first 5 min, which then decelerated, but the total concentration continued to increase. After 25–30 min, the composites reached the solubility limit and remained constant. PP 1 encapsulated in GPs has achieved the highest solubility (16.27 ±1.96 mg/L) and about 41 % of the dose was dissolved. Moreover, encapsulation in GPs improves dispersion of the composites even without surfactant, which helps hydrophobic drugs with poor dispersibility to increase their solubility in polar media as was proved also in the study of Ruphuy et al. [26]. Dissolution profiles shown in Fig. 10 were performed with the same samples (propolis extracts, GPs/PP composites) as in dissolution tests before (Fig. 9) with the dose of 36 mg and with the addition of 1 % surfactant (SDS) to achieve sink conditions, allowing for dissolving the whole dose. For propolis solubility, see Supplementary Information 5, Fig. S8. PP 2 and PP3 extracts dissolved completely over the course of the experiment, reaching the maximum measured concentration at 48 h with the final measured concentrations of 40.65 ±0.76 mg/L and 41.41 ±1.22 mg/L, respectively. The concentration of propolis released from the PP 1 extract at 48 h was only 31.66 ±3.45 mg/L with the dissolution profile indicating that the concentration would likely continue to grow if measured after a longer time period. The differences in the dissolution rates of the pure extracts can be possibly attributed to the different chemical compositions of the samples indicated by the UHPLC measurements (Supplementary Information 6, Fig. S9). The encapsulation of propolis into GPs significantly enhanced the dissolution rate when compared to pure propolis extracts. In the case of all composites, most of the propolis dissolved already within the first hour of the dissolution experiment. The composites exhibited differences during the initial phase of the test, specifically in the first 30 min, where the GP/PP 1 composites released the propolis the fastest and GPs/PP 3 the slowest. After 1 h, all three profiles converged, showing similar dissolution rates for the remainder of the study period. By the 48 h mark, all composites had dissolved about 90 % of the propolis, specifically 93 % of the dose was dissolved from GPs/PP 1 (37.29 ± 0.70 mg/L) and GPs/PP 2 (37.29 ±1.40 mg/L) composites and 87 % (34.68 ±0.34 mg/L) from GPs/PP 3 composites. Fig. 8. Concentration of chrysin, apigenin, kaempferol, quercetin, pinocembrin, and galangin in propolis extracts; PP1; PP2, PP3. 10-hydroxyl-2-decenoic acid and cinnamic acid were not detected in the samples. The error bars represent the standard deviation (n =2). Fig. 9. Dissolution kinetics of propolis extracts and spray-dried GPs/PP composites in 200 mL media (pH 6.8) at 37 ◦C and mini-paddles set as 150 rpm; PP 1; PP 2; PP 3; GPs/PP 1; GPs/PP 2; GPs/PP 3. The error bars represent the standard deviation (n =3). A. Brejchov´ a et al. Journal of Drug Delivery Science and Technology 114 (2025) 107490 9