Functional and Antimicrobial Properties of Propolis from Different Areas of Romania

Abstract

Propolis is a complex resinous substance produced by Apis mellifera L. through a process of mixing tree resins with saliva and beeswax. This substance plays a crucial role in the hive’s defence against a range of pathogenic agents, maintaining a consistent internal temperature and aseptic environment for the bee colony. The objective of the present study was to assess the chemical composition and antibacterial characteristics of five hydroalcoholic propolis extracts sourced from diverse geographic regions within Romania. This study shows that the biological and functional properties of propolis extracts are related to the plant resources in the vicinity of the hives, and this relates to greater or lesser bioactivity of the extracts; therefore, to standardise the extracts, it is essential to catalogue the plant essences in the proximity of the hives. The antimicrobial activity of propolis extract from each apiary was evaluated against five Gram-positive, five Gram-negative bacteria, and one fungal strain, using the difuzimetric method and minimum inhibitory concentration (MIC). The results showed some variability, supporting the hypothesis that not only may the botanical origin influence the properties of propolis but also that a higher number of flavonoids influences the higher antimicrobial activity in the extracts.

1. Introduction

Propolis is a complex resinous substance that honeybees (Apis mellifera L.) produce by mixing tree resins with saliva and beeswax. The resins produced by plants are intended to protect buds and shoots and, depending on the season and the plant, have a variable composition that gives propolis different chemical and antimicrobial characteristics [1,2].

The geographical origin has a significant impact on the chemical composition of propolis, which is influenced by a number of factors, including the local flora, climatic zones, and time of year [3,4]. The main challenge has been, and continues to be, the identification of the plant-derived compounds collected by bees to produce propolis; these botanical products are, in fact, secreted or exuded from different parts of the plant, either as a result of wound formation or as a secretion [4]. In areas with high plant diversity, honeybees have access to a variety of resinous or secretory materials throughout the active season. However, in areas that are intensively cultivated, the landscape may undergo a transition to a monoculture, which can result in a reduction in the availability of these compounds during the active period [5,6]; consequently, the variability of these compounds may also be significantly diminished. In the case of hives situated in urban areas, the composition of propolis is likely to be influenced by the botanical species present in that location [7,8]. Given the distance that the honeybee is required to traverse, it may be reasonably assumed that the principal materials for propolis are also gathered from the area surrounding the hive. Honeybees are capable of identifying these compounds and can transport them to the hive, thereby maintaining an aseptic environment and contributing to the hive’s overall health [5,9].

Propolis defends the hive, as bees use it to seal small openings and smooth out the interior spaces of hives, thus achieving a constant internal temperature and an aseptic environment [10]. The most representative compounds present in propolis include phenolics, terpenoids, amino acids, sugars, minerals, and vitamins. Given their beneficial properties, bee products have a long history of being employed for the disinfection and healing of wounds, as well as for their antimicrobial effects. In the context of the emergence of new pathologies associated with multidrug-resistant pathogens (MDR) and the irrational use of antibiotics, there is a growing need for extensive research and development of new antibiotics or complementary drugs on a global scale [11]. According to a report by the World Health Organisation, the use of active ingredients not belonging to traditional medicine is rapidly growing [12]. In this regard, laboratory tests have demonstrated that propolis and its constituents (flavonoids, galangin, and pinocembrin) are effective in the treatment of Gram-positive and Gram-negative pathogenic bacteria [11,13].

The phytochemical profile of propolis is inextricably linked to its geographical and botanical origins [14,15], thereby influencing its therapeutic efficacy. The specific plants used by bees for resin collection have been shown to influence the flavonoids, phenolic acids, and other bioactive compounds present in the resulting propolis [16,17]. The flavonoids and phenolic compounds present in propolis from temperate regions have been linked to strong antibacterial and antifungal activities. In contrast, tropical propolis often contains terpenes that target different pathogens [18,19]. Furthermore, the physical properties of propolis, including colour, are indicative of floral diversity, with specimens ranging from light yellow to dark brown [20]. Geographical and botanical factors exert a profound influence on the bioactivity of propolis, even within the same region [16,17,21]. The resins present in Romanian propolis are derived from deciduous plants belonging to the genera Quercus, Ulmus, Picea, and Salix; they appear to contain elevated levels of phenolic compounds, although there may be variations depending on the geographical origin, harvesting period, beekeeping practices, and extraction method [22,23]. This study aimed to evaluate the functional properties of five hydroalcoholic propolis extracts from different areas of Transylvania; characteristics such as the wax content of the crude propolis, the residue of the ethanolic extract (balsam), chemical composition (flavonoids and phenolic compounds), antimicrobial activity, antibiofilm, and minimum inhibitory activity (MIC), as well as antioxidant activity, were investigated. The chemical composition of the five propolis samples is responsible for their antioxidant and antimicrobial effects.

2. Materials and Methods

2.1. Study Area and Land Use

The area covered by this work includes five apiaries located in the Transylvanian area, specifically in the counties of Cluj, Alba, Sălaj, and Harghita (

Table 1. List of the apiaries included in the study.

Sample ID	County	Apiary	Coordinates
C1	Cluj	USAMV apiary	46°45′37.16″ N; 23°34′13.47″ E
C2	Cluj	Palocsay apiary	46°45′35.83″ N; 23°37′22.57″ E
A	Alba	Alba apiary	46°20′31.68″ N; 24°1′23.13″ E
S	Sălaj	Sălaj apiary	46°58′0.3″ N; 23°1′0.4″ E
H	Harghita	Corund apiary	46°27′27.3″N; 25°16′11.8″ E

System of coordinates WGS84 EPSG 4236.

). For each apiary, maps were generated for the image-based study of land use and land cover (Figures S1–S5), paying particular attention to the vegetation cover of the areas within a radius of 1200 and 3000 m from the apiaries. The maps were constructed using the program QGIS (Quantum GIS ver. 3.30.0, Hertogenbosch; http://www.qgis.org (accessed on 9 September 2024) and the Corine Land Cover 2018 with a level III resolution (https://land.copernicus.eu/en, accessed 9 September 2024). This study was conducted in collaboration with beekeepers affiliated with the APHIS-DIA (University of Agricultural Sciences and Veterinary Medicine Cluj Napoca, Romania) laboratory collaborators network. The 3000 m radius area, which we defined as the maximum flight area, was found to encompass 2.82743 hectares, while the 1200 m radius area, defined as the economically viable flight area, was determined to be approximately 452.38 hectares. The radius areas were selected based on the available literature regarding our local honeybee populations [24,25]. For each apiary, the surface area for each radius was analysed, and the percentage of each land class was calculated. To facilitate the visualisation of these data, pie charts were generated for each location radius for analysis.

2.2. Propolis Sampling and Storage

Raw propolis samples, approximately 70 g for each location, were collected by scraping the inner surfaces of 30–40 hives with a stainless-steel spatula. The sample collection period was between late June 2024 and early July 2024, from four counties in Romania (Cluj, Alba, Sălaj, and Harghita). Immediately after collection, the samples were stored at −20 °C in the freezer until further analysis. After sample grinding, a preliminary colour assessment was made for the raw propolis; the colour of the samples varied from light orange to dark brown.

2.3. Methodologies and Reagents

The analyses were carried out in compliance with ISO/DIS 24381 (Bee Propolis—Specifications) [26], focusing on propolis ethanolic extraction, percentage of balsam (ethanol extractables), wax content, total phenolic, and flavone/flavonoids content. All reagents used were of analytical grade.

2.4. Propolis Ethanolic Extracts (PEE) and Balsam Percentage (Evaluation of Dry Extract)

The frozen propolis was reduced to a powder using an electric grinder to allow the maximum homogenisation of the samples. Briefly, 1 g and 10 g of propolis from each sample were weighed using an analytical balance (accurate to 0.001 g) and suspended in 40 mL of 80% (v/v) ethanol in a 100 mL conical flask. The samples were left under mechanical agitation until the next day, when they were filtered through a qualitative paper filter and the extraction was repeated. The extraction process lasted 3 days with a final volume of 100 mL. All samples for both concentrations were extracted in duplicate.

The balsamic fraction was obtained as a percentage of soluble ethanol compounds. Dry glass dishes were weighed using an analytical balance (accuracy of 0.0001 g) and conditioned to constant weight in an oven at 105 °C for 90 min. Subsequently, 2 mL of each propolis extract was placed in the drying dishes and subjected to the same heat treatment until constant weight (maximum difference of 2 mg between two consecutive weighings). The percentage of dry matter content was calculated using the following equation:

$$X_1= {\displaystyle \frac{m_1}{m_2}}\times Dil. \times 100$$

where X₁ is the ethanol/water content extract in the sample (count by dry matter) in g/100 g, m₁ is the average mass of the propolis sample in grammes, m₂ is the mass of the dry extract in grammes, and Dil. represents the dilution factor. For both concentrations, the assessment was performed in duplicate. The tests were conducted in triplicate.

2.5. Wax Content in Raw Propolis (Petroleum Ether Extractables)

For each propolis sample, 2 g of crude propolis (w₁) was weighed into a cellulose cartridge. The extraction process took place in a Soxhlet apparatus for 6 h, mixing the propolis with 90 mL petroleum ether; in the end, the extract was evaporated to dryness under reduced pressure (maximum 50 °C). Finally, the residue was allowed to cool in a desiccator to constant weight (w₂). The wax content, expressed as dry matter, was calculated as follows:

$$Z_1= {\displaystyle \frac{w_2}{w_1}}\times 100$$

where Z₁ is the wax content in raw propolis in g/100 g, w₁ is the mass of the raw propolis sample in grammes, and w₂ is the mass of dry residue extract in grammes. The tests were performed in triplicate.

2.6. Total Phenolics (TPC) and Flavone/Flavonol (TFC) Content

The Folin–Ciocalteu colourimetric method (adapted for propolis analysis) was employed to determine the TPC. An aliquot of the PEE (200 µL) was combined with 1.5 mL of ultrapure water, 400 µL of Folin–Ciocalteu reagent, 600 µL of a 20% sodium carbonate solution, and the requisite volume of ultrapure water to reach a total volume of 5 mL. Following 30 min of incubation at room temperature and in the dark, the absorbance at 760 nm was determined. A gallic acid calibration curve was constructed for the quantification of total phenols, with the following final concentrations: 1.0, 2.0, 3.0, 4.0, 5.0, 6.0, and 8.0 µg/mL. The results were expressed as mg of gallic acid equivalents (GAE)/g, and all samples were analysed in triplicate.

To determine the flavone/flavonol content, a 25 mL volumetric flask was prepared with 1 mL of PE, 15 mL of methanol, and 500 µL of a 5% aluminium chloride solution. The volume was then adjusted to 25 mL with methanol. The solution was then maintained at room temperature and in the dark for 30 min, after which the absorbance was measured at 425 nm. A quercetin calibration curve was constructed to quantify the total phenols with the following final concentrations: 1.0, 2.0, 3.0, 4.0, 7.0, and 10.0 µg/mL. The results were expressed as mg of quercetin equivalent (QE)/g, and all samples were analysed in triplicate.

2.7. Antioxidant Activity Assays

2.7.1. ABTS^·+ Assay

The antioxidant activity of PEEs using the ABTS^·+ method was carried out according to the method described by Mărgăoan et al. [27]. The test measured the antioxidant activity of the extracts as a function of the reaction between the radical cation ABTS^·+ and antioxidant compounds in the extracts. ABTS^·+ was dissolved in pure methanol at a concentration of 7 mM. The ABTS radical cation was obtained by reacting the methanolic ABTS solution with a 2.45 mM potassium persulfate solution. The mixture was kept in the dark for 24 h before use. The solution of the radical cation was diluted to an optical density of 0.700 (OD₇₃₄) before the assay. ABTS was determined by adding 30 µL of the sample to 170 µL of ABTS· + solution, and the OD was measured at 734 nm after 6 min in the dark using a spectrophotometer (BioTek Instruments, Winooski, VT, USA). Trolox was used as the standard for the calibration curve, and the antioxidant activity was expressed as Trolox Eq./g. Additionally, ascorbic acid was used as a positive control.

2.7.2. DPPH·Assay

The activity of PEEs was also evaluated using the DPPH assay according to the method of Urcan et al. [28]. A total of 40 µL of ethanolic extract of propolis was added to 200 µL of DPPH solution (0.02 mg/mL). Ascorbic acid was used for positive control. The absorbance of the samples was measured at 517 nm after 15 min using a spectrophotometer (BioTek Instruments, Winooski, VT, USA). Trolox was used as a standard for the calibration curve, and the antioxidant activity was expressed as Trolox Eq./g.

2.7.3. FRAP Assay

The method described by Urcan et al. [29] was used to evaluate the antioxidant activity of PEEs via their ferric-reducing power. Briefly, 300 µL of FRAP reagent was mixed with 10 µL of the sample and the same amount of ultrapure water, and the same procedure was performed for the positive control. After five minutes of incubation at 37 °C, the antioxidant power was measured at 593 nm using a microplate reader (BioTek Instruments, Winooski, VT, USA). Trolox was used as the standard for the calibration curve, and the antioxidant activity was expressed as Trolox Eq./g.

2.8. Chromatographical Conditions

The phenolic profile of propolis samples was carried out according to Cucu et al. [30]. The samples were analysed using high-performance liquid chromatography (HPLC) combined with a photodiode array detector (PDA). The separation of the phenolic compounds was performed using a Teknokroma Mediterranean Sea 18 column (150 × 46 mm) with an intern diameter of 5 µm and a flow rate of 1 mL/min. The binary gradient elution system consisted of (A) pH 2.5 water (adjusted with orthophosphoric acid) and (B) acetonitrile. The temperature of the column was kept at 28 °C, and the separation was monitored in a wavelength range of 220–400 nm [31]. The identity of each peak was corroborated by superimposing the spectrum of the peak in question, its retention time, and the spectrum of the corresponding standard (protocatechuic acid, p-OH benzoic acid, catechin, vanillic acid, chlorogenic acid, caffeic acid, vanillin, p-cumaric acid, ferulic acid, naringin, rosmarinic acid, quercitrin, quercetin, naringenin, apigenin, kaempferol, caffeic acid phenethyl ester (CAPE), chrisyn, pinocembrin, and galangin).

Furthermore, the peaks were identified by comparing their retention times (Rt) with the retention times of the pure standards. The standard of individual compounds (1000 mg/L) were dissolved in HPLC-grade methanol. Each phenolic compound was quantified from the samples’ peak areas and the corresponding calibration curve of each phenolic. The results were expressed as μg/g matrix (ppm).

2.9. Antimicrobial Activity

The antimicrobial efficacy of 10% propolis extracts was assessed against the following indicator bacteria: Enterococcus faecalis ATCC 29212, Staphylococcus aureus ATCC 25923, Staphylococcus aureus MRSA ATCC 43300, Staphylococcus epidermidis ATCC 12228, Listeria monocytogenes ATCC 35152, Klebsiella pneumoniae ATCC 13883, Escherichia coli ATCC 25922, Pseudomonas aeruginosa ATCC 27853, Salmonella enterica ATCC 25928, Acinetobacter baumanii ATCC 19606, and Candida albicans ATCC 10231 (ATCC, American Type Culture Collection, Manassas, VA, USA). The above pathogenic strains were aerobically grown in 10 mL BHI broth at 37 °C for 16 h; bacterial suspensions were adjusted to a concentration of 0.5 McFarland to give a cell density of ~10⁸ CFU/mL. The antimicrobial activity of propolis samples was performed according to Iorizzo et al. [32] with some modifications. Briefly, 15 mL of BHI agar was inoculated with a 0.5 McFarland of pathogens into Petri dishes (Ø 90 mm). Wells of 0.6 mm diameter were cut in the plates, and 50 µL of propolis extract was added to the wells. As a negative control, 50 µL of 80% ethanol was used, while as a positive control, vancomycin (30 µg) was used for Gram-positive bacteria, gentamicin (10 µg) for Gram-negative bacteria, and fluconazole (25 µg) for yeasts. The plates were observed, and antimicrobial activity was recorded as the diameter (mm) of the zone of clear inhibition (ZOI) around the inoculated wells after incubation at 37 °C for 24–48 h. All the tests were performed in triplicate.

2.10. Minimum Inhibitory Concentration (MIC)

The MIC of propolis extracts was evaluated using the microdilution technique, based on the Clinical and Laboratory Standards Institute (CLSI) guidelines M07-A9 [33] with slight modifications [34]. For this assay, bacterial and fungal strains were cultured in Brain Heart Infusion (BHI) broth (for bacteria) and Sabouraud Dextrose Broth (SDB) (for fungi) containing various concentrations of propolis extracts ranging from 50 mg/mL to 0.01 mg/mL. The tests were conducted in 96-well microplates, and the cultures were incubated at 37 °C for 24 h. The bacterial inoculum was standardised to a density of 10⁸ CFU/mL using the 0.5 McFarland scale and subsequently diluted to achieve a final concentration of 5 × 10⁵ CFU/mL in the wells, while the fungal inoculum was prepared to a density of 10⁶ CFU/mL using the same standardisation technique. After the incubation period, 30 µL of 0.01% (w/v) resazurin solution was added to each well. Following a 30 min incubation [34], a visual inspection was carried out to detect any colour changes. A shift from purple to pink indicated microbial growth. Positive controls (bacterial or fungal growth without extract), negative controls (sterile culture medium), and ethanol controls were included in each experiment to ensure reliability. The MIC was defined as the lowest concentration of propolis extracts that completely inhibited visible microorganisms’ growth. Each bacterial and fungal strain was tested in three independent experiments.

2.11. Statistical Analysis

The statistical analysis was conducted using RStudio (R software version 4.3.0; R Core Team 2023). All the data obtained from the three independent experiments are expressed as the mean ± standard deviation (SD). Statistical analysis was performed using an analysis of variance (ANOVA). The obtained data, normally distributed, were analysed using Tukey’s post hoc tests with ANOVA. Statistical significance was attributed to p-values < 0.05.

3. Results

3.1. Landscape Complexity

A land use analysis was conducted in the localities from which the samples were sourced with the objective of establishing potential links between land use, the quality of the raw propolis, and its properties. For each apiary, the percentage of land classes within a radius of 1200 and 3000 m surrounding the apiary was calculated (

Table 2. Land cover expressed as a percentage for each land class identified in the 1200 and 3000 m radius areas.

			Locations
	CLC ID	Land Class Description	C1	C2	A	S	H
Economically viable flight area (1200 m radius)	112	Discontinuous urban fabric	76.73	27.82	6.13	18.53	0.00
	121	Industrial or commercial units	0.00	48.86	0.00	0.00	0.00
	133	Construction sites	0.00	2.09	0.00	0.00	0.00
	211	Non-irrigated arable land	20.71	7.83	0.00	5.32	0.00
	222	Fruit trees and berry plantations	0.00	3.50	8.65	0.00	0.00
	231	Pastures	2.56	0.00	27.68	24.46	27.40
	242	Complex cultivation patterns	0.00	9.90	19.49	29.96	0.00
	243	Land principally occupied by agriculture, with significant areas of natural vegetation	0.00	0.00	16.30	15.84	0.00
	311	Broad-leaved forest	0.00	0.00	6.39	5.89	29.05
	312	Coniferous forest	0.00	0.00	3.43	0.00	0.00
	313	Mixed forest	0.00	0.00	0.00	0.00	28.39
	321	Natural grasslands	0.00	0.00	0.00	0.00	13.98
	324	Transitional woodland-shrub	0.00	0.00	0.00	0.00	1.17
	411	Inland marshes	0.00	0.00	11.93	0.00	0.00
Maximum flight area (3000 m radius)	112	Discontinuous urban fabric	61.82	46.20	4.56	5.15	0.00
	121	Industrial or commercial units	4.47	13.36	0.00	1.11	0.00
	133	Construction sites	0.01	3.42	0.00	0.00	0.00
	141	Green urban areas	7.17	0.90	0.00	0.00	0.00
	211	Non-irrigated arable land	0.99	12.12	27.27	9.89	0.00
	222	Fruit trees and berry plantations	0.02	16.19	1.40	0.34	0.00
	231	Pastures	5.51	1.38	47.65	31.32	20.51
	242	Complex cultivation patterns	4.14	5.57	4.15	17.04	6.38
	243	Land principally occupied by agriculture, with significant areas of natural vegetation	0.00	0.19	6.05	6.42	0.00
	311	Broad-leaved forest	14.80	0.27	2.53	23.92	26.90
	312	Coniferous forest	0.80	0.00	1.62	0.00	1.86
	313	Mixed forest	0.00	0.38	0.00	0.00	14.29
	321	Natural grasslands	0.00	0.00	0.00	0.00	25.94
	324	Transitional woodland-shrub	0.00	0.00	0.00	4.80	4.12
	411	Inland marshes	0.00	0.00	4.78	0.00	0.00
	511	Water courses	0.27	0.00	0.00	0.00	0.00

). The number and percentage of land classes identified in each area varies depended on the radius and complexity level.

With regard to the C1 apiary, it can be observed that within the 1200 m radius, there are only three land classes. Of these, 77.73% represents discontinuous urban fabric, 20.71% non-irrigated arable land, and 2.56% pastures. However, for the 3000 m radius, a considerable number of land classes are found. The dominant land class remains discontinuous urban fabric, which accounts for 61.82% of the total area, followed by broad-leaved forest (14.8%), green urban areas (7.17%), pastures (5.51%), industrial or commercial units (4.47%), and complex cultivation patterns (4.14%). Among the remaining surfaces, coniferous forests account for 0.80%.

A different situation is identified in apiary C2 as follows: for the 1200 m radius, a total of six land classes are identified, with 48.86% representing industrial or commercial units. The remaining land cover classes are discontinuous urban fabric (9.9%), complex cultivation patterns (7.83%), non-irrigated arable land (3.5%), fruit trees and berry plantations (2.09%), and construction sites (1.77%). At the 3000 m radius, the dominant land class is discontinuous urban fabric, which accounts for 46.2% of the total area, while 13.36% represents industrial or commercial units.

Other land classes comprise 16.19% fruit trees and berry plantations, 12.12% non-irrigated arable land, 5.57% complex cultivation patterns and 3.42% construction sites. The remaining land classes are as follows: 1.38% pastures, 0.90% green urban areas, 0.38% mixed forest, 0.28% broad-leaved forest, and 0.19% land principally occupied by agriculture, with significant areas of natural vegetation.

In the case of the apiary situated in Alba County (A), the 1200 m radius reveals the presence of eight land classes, among which, 27.68% are identified as pastures, 19.49% as complex cultivation patterns, and 16.3% as land principally occupied by agriculture. Additionally, there are significant areas of natural vegetation (11.93%), inland marshes (8.65%), fruit trees and berry plantations (6.39%), broad-leaved forest (6.13%), and discontinuous urban fabric (3.43%), as well as coniferous forest (1.38%). In the 3000 m radius, nine land classes are identified. The most prevalent land class is pasture, representing 47.65% of the total area, followed by non-irrigated arable land (27.27%), land principally occupied by agriculture with significant areas of natural vegetation (6.05%), and inland marshes (4.78%). Moreover, the land cover composition includes 4.56% discontinuous urban fabric, 4.15% complex cultivation patterns, 2.53% broad-leaved forest, 1.62% coniferous forest, and 1.4% fruit trees and berry plantations.

The Sălaj County apiary (S) encompasses seven land classes within a 1200 m radius. The proportion of land exhibiting complex cultivation patterns is 29.96%, while 24.46% comprises pastures and 18.53% of discontinuous urban fabric. The land is predominantly agricultural, with natural vegetation comprising 5.89% forest, 5.32% arable land, and 0.003% woodland. In the 3000 m radius, nine land classes are observed as follows: pasture (31.32%), broad-leaved forest (23.92%), complex cultivation patterns (17.04%), and non-irrigated arable land (9.89%). The land is primarily utilised for agricultural purposes, with the presence of natural vegetation, woodlands, and fruit trees.

Finally, apiary H, situated in Harghita County, is characterised by a high level of spontaneous vegetation. In the radius of 1200 m, five land classes are revealed, with 29.05% comprising broad-leaved forest, 28.39% mixed forest, 27.40% pastures, 13.98% natural grasslands, and 1.17% transitional woodland shrub. For the 3000 m radius, a total of seven land classes were discerned. The predominant land class remains that of broad-leaved forest, representing 26.9% of the total area, followed by natural grasslands at 25.94%, pastures at 20.51%, mixed forest at 14.29%, complex cultivation patterns at 6.38%, transitional woodland-shrub at 4.12%, and 1.86% coniferous forest.

Maps and pie charts illustrating land use were made for all apiaries (Supplementary Figures S1–S10 and Supplementary Maps S1–S6). Additional information regarding the possible vegetation present for each land class can be found in Supplementary Table S1.)

3.2. Wax and Balsam

The quality of samples, and therefore the percentage of wax present in the sample, was evaluated before the preparation of the hydroalcoholic propolis extracts (

Table 3. Percentage of wax content in raw propolis samples and percentage of balsam in PEEs. The data are expressed as mean ± SD (n = 3). Different lowercase letters (a–e) in each row indicate significant differences p < 0.05.

Quality Parameter (%)	Propolis Extracts
	C1	C2	A	S	H
Wax	41.2 ± 0.2 b	47.2 ± 0.6 a	20.0 ± 0.3 e	31.9 ± 0.3 d	39.5 ± 0.7 c
Balsam 1%	49.4 ± 0.5 c	44.7 ± 0.9 d	61.6 ± 0.6 a	51.9 ± 0.8 b	51.6 ± 0.9 b
Balsam 10%	50.0 ± 0.9 c	41.0 ± 0.9 d	64.8 ± 0.2 a	54.8 ± 0.5 b	54.1 ± 0.5 b

). The samples showed a variable wax content with statistically significant differences. Sample A had a wax content of 20%, while for the other samples, wax content varied between 39.5% (H) and 47.2% (C2). Significant differences among samples were observed.

Following the hydroalcoholic extraction, another quality parameter of propolis was evaluated—the amount of balsam (content of matter soluble in ethanol) for 1% and 10% PEEs (Table 3). The proportion of balsam present in the propolis extracts was found to differ significantly between the analysed samples, with a range from 49.4% (C1) to 61.6% (A) for the 1% extracts and 50% (C1) to 64.8% (A) for the 10% extracts.

3.3. TPC and TFC

TPC showed values ranging from 150.4 to 125.4 mg (GAE)/g, with the highest values observed for samples A, C1, and H. Extract C2, on the other hand, showed the lowest value of total phenolic compounds (125.4 mg (GAE)/g). For TFC, the highest values were 16.9 mg (QE)/g (C1) and 13 mg (QE)/g (A); the lowest value for total flavonols was observed for extract H (1.5 mg (QE)/g). Data for TPC and TFC are shown in

Table 4. Total phenolic content and total flavonols of the 1% PEEs. The data were expressed as the mean ± SD (n = 3). Different lowercase letters (a–e) in each row indicate significant differences p < 0.05.

	C1	C2	A	S	H
TPC	149.8 ± 0.8 a	125.4 ± 1.0 c	150.0 ± 0.4 a	146.3 ± 0.6 b	149.7 ± 0.6 a
TFC	16.9 ± 0.5 a	9.8 ± 0.8 c	13.0 ± 0.8 b	5.9 ± 0.5 d	1.5 ± 0.01 e

TPC: mg GAE/g fresh weight; TFC: mg QE/g fresh weight.

.

3.4. Phenolic Profiles of PEEs

Twenty phenolic compounds (phenolic acids and flavonoids) were quantified in the propolis samples (

Table 5. Phenolic profiles of the five propolis ethanolic extracts. The data were expressed as the mean ± SD (n = 3).

	Propolis Extracts
Phenolic Compounds (µg/g)	C1	C2	A	S	H
Protocatechuic acid	0.40 ± 0.02	0.37 ± 0.04	0.41 ± 0.2	0.62 ± 0.04	0.20 ± 0.03
p-OH-benzoic	3.76 ± 0.4	3.20 ± 0.2	9.15 ± 1.1	5.76 ± 1.1	7.16 ± 1.2
Catechin	n.d.	n.d.	0.24 ± 0.1	n.d.	1.60 ± 0.08
Vanillic acid	0.50 ± 0.02	0.61 ± 0.03	0.85 ± 0.2	1.3 ± 0.05	n.d.
Clorogenic acid	n.d.	n.d.	n.d.	0.54 ± 0.02	n.d.
Caffeic acid	42.41 ± 2.1	22.93 ± 1.6	77.95 ± 0.7	20.76 ± 0.6	28.15 ± 2.1
Vanillin	6.58 ± 1.7	8.84 ± 0.7	18.69 ± 1.8	27.26 ± 1.9	22.80 ± 2.9
p-cumaric acid	64.12 ± 2.6	57.76 ± 1.5	148.41 ± 6.1	188.33 ± 3.0	295.08 ± 3.9
Ferulic acid	31.12 ± 1.1	66.51 ± 3.6	96.90 ± 2.9	214.13 ± 2.71	330.40 ± 4.8
Naringin	1.62 ± 0.1	n.d.	n.d.	n.d.	n.d.
Rosmarinic acid	0.85 ± 0.03	n.d.	0.98 ± 0.2	6.85 ± 1.3	2.76 ± 0.3
Quercitrin	0.45 ± 0.02	n.d.	1.19 ± 0.1	n.d.	2.27 ± 0.1
Quercetin	11.13 ± 1.0	4.25 ± 0.2	n.d.	n.d.	n.d.
Naringenin	55.41 ± 1.9	17.20 ± 2.2	21.19 ± 1.10	8.84 ± 0.7	n.d.
Apigenin	22.46 ± 2.6	9.77 ± 0.6	15.89 ± 1.67	13.48 ± 1.2	4.41 ± 0.8
Kaempferol	7.97 ± 1.4	n.d.	n.d.	n.d.	n.d.
CAPE	116.77 ± 2.2	n.d.	180.61 ± 5.1	44.15 ± 0.9	n.d.
Chrysin	188.48 ± 3.9	83.22 ± 3.0	158.61 ± 3.7	38.2 ± 2.3	9.27 ± 1.1
Pinocembrin	357.86 ± 3.7	208.13 ± 2.8	180.31 ± 4.1	n.d.	n.d.
Galangin	43.07 ± 2.1	19.80 ± 1.7	29.43 ± 2.83	3.65 ± 0.2	n.d.

). The results demonstrated that samples A, S, and H exhibited the highest concentration of phenolic acids, while C1 and C2 displayed a higher abundance of flavonoids. The highest value of protocatechuic acid was observed in extract S, 0.62 μg/g, followed by A (0.41 μg/g), C1 (0.40 μg/g), C2 (0.37 μg/g), and H (0.20 μg/g). The highest concentration of p-OH-benzoic acid was observed in the PE from Alba County (A), with a value of 9.15 μg/g, followed by H (7.16 μg/g) and then S (5.76 μg/g). The lowest values were observed for C1 and C2, at 3.76 and 3.2 μg/g, respectively. Catechin was identified in only two of the extracts, namely H (1.6 μg/g) and A (0.24 μg/g). The highest concentration of vanillic acid was observed in extract S (1.3 μg/g), followed by extract A (0.85 μg/g). Extract C2 exhibited a concentration of 0.61 μg/g, while C1 demonstrated a concentration of 0.50 μg/g. With regard to chlorogenic acid, this was only identified in extract S (0.54 μg/g) among the five samples analysed. Caffeic acid was identified in all the analysed samples. The highest concentration was observed in extract A (77.95 μg/g), followed by C1 (42.41 μg/g). The concentrations of this phenolic acid in samples H, C2, and S were found to be 28.15 μg/g, 22.93 μg/g, and 20.76 μg/g, respectively. Vanillin was identified in all samples, with the highest concentration observed in extract S (27.26 μg/g), followed by H (22.8 μg/g) and A (18.69 μg/g). Lower concentrations were recorded for extracts C2 (8.84 μg/g) and C1 (6.58 μg/g). About p-cumaric acid, notable quantities are evident across the five extracts. Extract H exhibits the highest concentration of this compound, at 295.08 μg/g, followed by S (188.33 μg/g) and A (158.41 μg/g). The extracts with the lowest concentration of the specified substance were C1 (64.12 μg/g) and C2 (57.76 μg/g). Similarly, ferulic acid was present in all propolis extracts, with the highest concentration observed in extract H, at 330.4 μg/g, followed by S (214.13 μg/g) and A (96.90 μg/g). Extracts C1 and C2 exhibited a lower concentration of this compound compared to the other three. Naringin was identified in only one extract, C1, at a concentration of 1.61 μg/g. Rosmarinic acid was detected in four of the extracts, excluding extract C2. The highest concentration was observed in sample S, at 6.85 μg/g, followed by extract H, at 2.76 μg/g. For sample A, 0.98 μg/g was detected, and for C1, 0.85 μg/g. The presence of quercitrin was confirmed in three of the five samples subjected to analysis. The highest concentration was observed in sample H, with a value of 2.27 μg/g, followed by sample A, with a concentration of 1.19 μg/g, and extract C1, with a concentration of 0.45 μg/g. The presence of quercetin was confirmed in extract C1 at a concentration of 11.13 μg/g, while extract C2 exhibited a concentration of 4.25 μg/g. Four of the five extracts were identified as containing naringenin. The highest quantity was identified in extract C1, with a concentration of 55.41 μg/g, followed by extract A (21.19 μg/g), C2 (12.20 μg/g), and S (8.84 μg/g).

Apigenin was detected in the extracts at different concentrations; the highest quantity was observed in extract C1, with a value of 22.46 μg/g, followed by extract A (15.89 μg/g).

The propolis extract from Sălaj (S) exhibited a concentration of 13.48 μg/g of apigenin, while the C2 demonstrated a concentration of 9.77 μg/g. The propolis from Harghita County (H) exhibited the lowest concentration among the tested extracts, with a concentration of 4.41 μg/g. Only one of the five analysed samples contained kaempferol, with a value of 7.96 μg/g for the sample C1. The analysis revealed the presence of CAPE in three of the examined extracts. The extract from A exhibited the highest concentration, with a value of 180.61 μg/g, followed by C1 with 116.77 μg/g and S with 44.15 μg/g. Chrysin was identified in all five extracts, albeit in varying quantities. The extract C1 exhibited the highest concentration, with a value of 188.48 μg/g, followed by extract A, which had a concentration of 158.61 μg/g. Extract C2 had a concentration of 83.22 μg/g, while the extract from S had a concentration of 38.2 μg/g. The lowest concentration of chrysin was found for sample H (9.27 μg/g). The highest concentration of pinocembrin was observed in extract C1, with a value of 357.86 μg/g, followed by C2 (208.13 μg/g) and A (180.31 μg/g). Additionally, galangin was identified in C1, with a concentration of 43.07 μg/g, followed by the extract from A with 29.43 μg/g, the extract from C2 with 19.8 μg/g, and the lowest amount detected was in S, with 3.65 μg/g.

Table 6. Total quantity of phenolics and flavonoids obtained by summing their individual quantities. Data are expressed in μg/g as the mean ± SD (n = 3).

	Propolis Extracts
	C1	C2	A	S	H
Phenolic acids	259.9 ± 41.5	151.38 ± 29.9	515.2 ± 72.7	482.45 ± 88.4	663.7 ± 157.2
Flavonoids	688.5 ± 120.6	342.37 ± 79.3	406.6 ± 79.6	61.18 ± 15.3	15.95 ± 3.5
Flavonoids + Phenolic acids	948.4 ± 92.4	493.75 ± 59.5	921.91 ± 72.7	546.63 ± 72.1	679.7 ± 135.0

presents the total content of phenolic acids and flavonoids, calculated by summing the individual compound quantities. Extract H exhibits the highest overall value for total phenolic acids, with a concentration of 663.75 μg/g, followed by extract A (515.29 μg/g), S (482.45 μg/g), and C1 (259.95 μg/g). With regard to flavone/flavonol content, extract C1 exhibits the highest value among the analysed extracts, with a value of 688.5 μg/g, followed by extract A with 406.62 μg/g and extract C2 with 342.37 μg/g. The extract from S has a flavonoid content of 61.18 μg/g, while the extract from H has the lowest flavonoid content, with 15.95 μg/g.

It is evident that the flavonoid and phenolic acid content present in the five extracts varies considerably. The highest value was recorded for C1 (948.45 μg/g), followed by A (921.91 μg/g). Notwithstanding the closeness of the values observed, C1 demonstrated a higher overall antimicrobial activity than A.

3.5. Antioxidant Activity

The antioxidant activity was evaluated using three methods, and the values are presented in

Table 7. Antioxidant activity of the propolis ethanolic extracts. The data (mean ± SD; n = 3) are expressed as Trolox Eq./g. Different lowercase letters (a–e) in each row indicate significant differences (p < 0.05).

Antioxidant Assay	Propolis Extracts
	C1	C2	A	S	H	PC
FRAP	5.4 ± 0.1 d	4.8 ± 0.09 d	7.4 ± 0.2 b	5.4 ± 0.3 d	6.2 ± 0.2 c	58.3 ± 1.0 a
DPPH	29.8 ± 0.8 c	24.9 ± 0.5 e	31.6 ± 0.3 b	26.4 ± 0.6 d	28.8 ± 0.5 c	79.6 ± 0.4 a
ABTS	34.0 ± 0.3 b	25.2 ± 1.90 e	33.4 ± 0.2 b	29.2 ± 0.3 d	31.0 ± 0.2 c	49.5 ± 0.7 a

PC: positive control.

. The values are expressed as mg Trolox Eq./g. The FRAP values of the ethanolic extracts of crude propolis ranged from 4.8 (C2) to 7.4 (A) mg Trolox Eq./g. The method based on the reduction in the stable free radical DPPH was used to evaluate the ability to scavenge free radicals, and its values ranged from 24.9 (C2) to 31.6 (A) mg Trolox Eq./g. In addition to FRAP and DPPH, the 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS) assay was also used to evaluate the antioxidant capacity of the same propolis extracts with values ranging from 25.2 (S) to 34.0 mg (C1) Trolox Eq./g. It can be observed that the positive control exhibits a markedly higher result for all three tests in comparison to the 1% propolis extract.

3.6. Antimicrobial Activity and MIC

The PEEs were evaluated for their antimicrobial activity against five Gram-positive pathogens (

Table 8. Antimicrobial activity of propolis ethanolic extracts against the tested Gram-positive pathogenic bacteria. The data (mean ± SD; n = 3) are expressed as ZOI (mm). Different lowercase letters (a-f) in each column indicate significant differences p < 0.05.

Propolis Extracts	Gram-positive Indicator Strains
	E. faecalis ATCC 29212	S. aureus ATCC 25923	S. aureus MRSA ATCC 43300	S. epidermidis ATCC 12228	L. monocytogenes ATCC 35152
C1	10.4 ± 0.3 b	12.8 ± 0.2 b	9.4 ± 0.4 b	11.1 ± 0.2 b	10.4 ± 0.4 b
C2	5.4 ± 0.3 d	12.2 ± 0.2 c	9.0 ± 0.3 bc	8.1 ± 0.2 d	8.1 ± 0.1 d
A	5.3 ± 0.1 c	11.0 ± 0.2 d	8.5 ± 0.4 c	8.8 ± 0.2 c	9.2 ± 0.2 c
S	8.4 ± 0.3 d	7.3 ± 0.3 f	6.7 ± 0.2 d	6.9 ± 0.1 e	7.5 ± 0.4 d
H	5.9 ± 0.2 d	8.2 ± 0.2 e	7.2 ± 0.2 d	8.1 ± 0.3 d	6.5 ± 0.3 e
Vancomycin (30 µg)	15.3 ± 0.9 a	19.0 ± 0.08 a	18.0 ± 0.08 a	20.0 ± 0.1 a	17.0 ± 0.1 a

), five Gram-negative pathogens, and one yeast pathogen (

Table 9. Antimicrobial activity of propolis ethanolic extracts against the tested Gram-negative pathogenic bacteria and yeast strain. The data (mean ± SD; n = 3) are expressed as ZOI (mm). Different lowercase letters (a–e) in each column indicate significant differences p < 0.05.

Propolis Extracts	Gram-negative and Yeast Indicator Strains
	S. enterica ATCC 25928	K. pneumoniae ATCC 13883	P. aeruginosa ATCC 27853	E. coli ATCC 25922	A. baumanii ATCC 19606	C. albicans ATCC 10231
C1	7.4 ± 0.2 d	3.1 ± 0.1 d	10.5 ± 0.3 a	4.9 ± 0.1 d	7.5 ± 0.4 b	12.5 ± 0.6 b
C2	11.1 ± 0.2 b	5.7 ± 0.2 c	9.1 ± 0.3 c	6.1 ± 0.1 c	7.5 ± 0.4 b	8.1 ± 0.2 d
A	11.5 ± 0.4 b	5.5 ± 0.1 c	9.4 ± 0.4 bc	4.9 ± 0.05 d	6.5 ± 0.2 c	8.9 ± 0.2 c
S	8.3 ± 0.2 c	7.2 ± 0.3 b	8.3 ± 0.2 d	7.1 ± 0.2 b	7.9 ± 0.1 b	7.1 ± 0.3 e
H	3.8 ± 0.1 e	0 ± 0 e	7.1 ± 0.1 e	4.2 ± 0.2 e	3.2 ± 0.1 d	7.3 ± 0.3 e
Gentamycin (10 µg)	19.0 ± 0.2 a	18.0 ± 0.08 a	10.0 ± 0.08 ab	20.0 ± 0.1 a	16.0 ± 0.08 a	-
Fluconazole (25 µg)	-	-	-	-	-	28.0 ± 0.2 a

). Among Gram-positive bacteria, for E. faecalis ATCC 29212, the inhibition values ranged from 5.3 mm (A) to 10.4 mm (C1). The inhibition values for S. aureus ATCC 25923 ranged from 7.3 mm (S) to 12.8 mm (C1), while for methicillin-resistant S. aureus ATCC 43300, the highest inhibition value, 9.4 mm, was obtained from extract C1. Regarding S. epidermidis ATCC, the lowest inhibition value was observed with extract S (6.9 mm), while the highest inhibition halos were produced by extracts A and C1 (8.8 mm and 11 mm, respectively). The lowest inhibition values for L. monocytogenes ATCC 35152 were observed with sample H (6.5 mm). In the case of Gram-negative bacteria, specifically S. enterica ATCC 25928, the lowest recorded inhibition was 3.8 mm (H), while inhibition values of 11.1 and 11.5 mm were observed for samples C2 and A, respectively. For Gram-positives, the inhibition levels observed with the positive control ranged from 15.3 mm (E. faecalis) to 20 mm (S. epidermis), with significant differences between the PEEs and the control.

The ethanolic extract of propolis from Sălaj (S) exhibited the greatest inhibitory effect on K. pneumoniae ATCC 13883, with an inhibition halo of 7.2 mm. With regard to P. aeruginosa ATCC 27853, the most efficacious extracts were A (11.5 mm) and C2 (11.1 mm), whereas the least efficacious was H, which exhibited an inhibition halo of 3.8 mm. In the case of E. coli ATCC 25922, the inhibition values were found to be relatively similar among the extracts, with the exception of samples S (7.1 mm) and C2 (6.1 mm). As for A. baumannii ATCC 19606, the highest inhibition value was 7.9 mm for sample S, while the lowest was recorded for extract H (3.2 mm). The yeast C. albicans ATCC 10231 was most inhibited by extracts C1 (12.5 mm) and A (8.9 mm), while the lowest value recorded was 7.1 mm (S). For Gram-negative pathogens, the inhibition values found with the positive control ranged from 10.0 to 20 mm, while for C. albicans, the antibiotic produced an inhibition halo of 28 mm.

The MIC results of the five propolis extracts showed a high variability between the strains tested, with differences between Gram-positive and Gram-negative bacteria (

Table 10. Minimum inhibitory concentration of the five propolis ethanolic extracts against the tested bacterial and yeast strains. Data are expressed as the mean value (n = 3) and as mg/mL.

		C1	C2	A	S	H	PC
Gram-positive strains	E. faecalis ATCC 29212	0.75	1.00	1.78	3.12	4.06	1.56 × 10−3
	S. aureus ATCC 25923	0.45	0.25	0.45	0.78	0.78	6.25 × 10−3
	S. aureusMRSA ATCC 43300	1.00	0.59	0.89	2.56	4.06	12.5 × 10−3
	S. epidermidis ATCC 12228	0.59	1.28	1.00	3.12	4.06	6.25 × 10−3
	L. monocytogenes ATCC 35152	0.50	1.28	1.78	4.69	5.63	3.12 × 10−3
Gram-negative strains	K. pneumoniae ATCC 13883	12.50	12.50	8.13	8.13	R	6.25 × 10−3
	E. coli ATCC 25922	1.56	0.78	3.12	5.63	11.25	1.56 × 10−3
	P. aeruginosa ATCC 27853	5.63	2.56	8.13	12.50	18.75	6.25 × 10−3
	S. enterica ATCC 25928	2.56	1.28	3.12	5.63	5.63	3.12 × 10−3
	A. baumanii ATCC 19606	6.25	8.13	11.25	11.25	12.50	6.25 × 10−3
Yeast strain	C. albicans ATCC 10231	0.39	0.39	1.03	1.28	1.50	3.12 × 10−3

R: resistant; PC: positive control.

). The most effective extract for the E. faecalis strain ATCC 29212 was identified as C1, which exhibited an average propolis extract concentration of 0.75 mg/mL. This was followed by extracts C2 (1.00 mg/mL) and A (1.78 mg/mL). The lowest response to this strain was observed for extracts S (3.12 mg/mL) and H (4.06 mg/mL). Concerning S. aureus ATCC 25923, the highest sensitivity was observed with extract C2, with an average of 0.25 mg/mL, followed by extract C1 and A, with 0.45 mg/mL. In contrast, the lowest sensitivity was observed for extracts S and H, with values of 0.78 mg/mL. For S. aureus MRSA ATCC 43300, a general increase in resistance was observed among the various test organisms to all five propolis extracts. The most effective extract remained C1, with an average of 0.59 mg/mL, followed by extract A with 0.89 mg/mL. Extract H exhibited the lowest response, with an average concentration of 4.06 mg/mL, which is five times higher than the concentration required for S. aureus ATCC 25923. The extracts with the most effective activity against S. epidermidis ATCC 12228 were C1 and A, with a mean minimum inhibitory concentration (MIC) value of 0.59 mg/mL and 1.00 mg/mL, respectively. In contrast, the extracts S and H demonstrated comparatively reduced efficacy. With regard to L. monocytogenes ATCC 35152, it is evident that there is considerable variability among the various extracts. C1 exhibited the lowest efficacy, with a mean of 0.50 mg/mL, whereas the least effective extract was H, which displayed a mean value of 5.63 mg/mL.

The results of the MIC test demonstrate that the efficacy of these extracts is diminished in comparison to the pathogens that were tested, particularly in relation to Gram-negative bacteria. With regard to the K. pneumoniae ATCC 13883, sensitivity was observed in samples A and S, with mean values of 8.13 mg/mL, followed by samples C1 and C2, which exhibited a mean value of 12.5 mg/mL. Extract H did not respond to the concentrations that were tested. In the case of the E. coli ATCC 25922, an increased sensitivity was observed for the extract derived from C2 (0.78 mg/mL), followed by C1 (1.56 mg/mL). For the extract from Alba (A), the mean MIC was 3.12 mg/mL, while for S it was 5.63 mg/mL; the extract from Harghita County (H) had the highest observed value (11.25 mg/mL), which represents the lowest level of activity against this particular strain. P. aeruginosa ATCC 27853 demonstrated a high degree of sensitivity to C2 extract, with a MIC value of 2.56 mg/mL. In contrast, the efficacy of the C1, A, and S extracts was less pronounced, with values ranging from 5.63 mg/mL to 12.5 mg/mL. The lowest efficacy was recorded for extract H, with an MIC value of 18.75 mg/mL. In the case of S. enterica ATCC 25928, the most effective extract was C2, with a minimum inhibitory concentration (MIC) of 1.28 mg/mL, followed by C1 with 2.56 mg/mL. Extract A demonstrated a relatively good response with 3.12 mg/mL. The least effective extracts for this strain were those for S and H, with 5.63 mg/mL. With regard to A. baumannii ATCC 19606, the extract exhibiting the greatest efficacy was C1, with a value of 6.25 mg/mL, followed by C2, with 8.13 mg/mL. The extracts from H and S demonstrated comparatively reduced efficacy, with values of 11.25 mg/mL and 12.5 mg/mL, respectively.

In the case of C. albicans ATCC 10231, a notable sensitivity was observed in response to all five propolis extracts. The yeast strain demonstrated a particularly high sensitivity to extracts C1 and C2 (0.39 mg/mL), followed by extract A with 1.03 mg/mL. The extracts with the highest minimum inhibitory concentration (MIC) values were S and H, with values of 1.28 mg/mL and 1.5 mg/mL, respectively. The positive control used in the MIC test varied from 1.56 µg/mL to 12.5 µg/mL depending on the strain tested.

4. Discussion

This study analysed propolis samples collected from five regions in Romania, focusing on their chemical composition, including phenolic and flavonoid content, and their bioactivity, such as antioxidant and antimicrobial properties. The results revealed significant variability influenced by geographical and botanical factors. The research area exhibits considerable diversity in landscape complexity. While the land class may not be a sufficient basis for classifying the origin of propolis, a more comprehensive analysis could provide additional insights. The availability of melliferous essences, and thus the variety of plant secretions and resins accessible to bees, may be subject to fluctuations depending on the region. Areas with a more diverse flora, where bees have access to a greater range of plant-derived materials throughout the active season, may experience less pronounced seasonal changes. Conversely, monoculture practices can lead to a reduction in plant diversity and, consequently, the variability of available materials [5,6,35]. The composition of propolis in hives located in urban settings is more likely to be influenced by the botanical species present in that area [7,8]. In the study by Pobiega et al. [8], the propolis extracts from the urban context exhibited similar compositions despite originating from disparate cities. In this particular case study, a notable discrepancy is observed between apiaries C1 and C2, indicative of significant floristic differences between the two locations. The distance between the two locations is approximately 4000 m. Consequently, the 3000 m radius encompasses a considerable portion of the radius of the other apiary, as well as a minor portion of the 1200 m radius. However, the difference between the identified phenolic compounds of C1 and C2 is notable. Additionally, there are differences between the total phenolic acids and flavonoids, with C1 having a significantly higher content. As for the propolis of the other locations, a total of 16 phenolics were identified in A, 14 in S, and 11 in H. The landscape differences between locations were significant when the area of a 1200 m radius was taken into account. Common land classes across all locations were pastures, with similar extensions in all locations, and broad-leaved forests (similar extensions in A and S). However, H had a significantly higher percentage of the aforementioned land classes compared to the other two. A more profound comprehension of the botanical provenance of propolis could prove instrumental in the endeavours to achieve standardisation. Furthermore, it is evident that climatic attributes and plant biodiversity exert a pronounced influence on the quality and quantity of propolis [35]; the climatic characteristics and plant diversity affect not only the composition of propolis but also the overall well-being of the honey bees [6,36,37].

The wax content of propolis has been observed to exhibit considerable variation across different studies, with values reported to range from 10% to 50% [38,39,40,41]. Despite the high proportion of wax present in sample C1 (41.2%), the antimicrobial activity of this propolis extract was notably high. It is likely that this pronounced activity can be attributed to the bees having access to a greater diversity of resins in that location. The area with a discontinuous urban fabric has a high plant diversity, with three different parks, the botanical garden, and the plant bioreserve of the university campus, all situated within a 1200 m radius. The quantity of balsam present was observed to vary considerably between samples; specifically, it was found that the 1% extract contained quantities of balsam between 49 and 61.6%, while the 10% extract exhibited a range of between 50 and 64.8%. The results demonstrate that the sample with the highest balsam content exhibited the lowest wax values. Moreover, the values exceed the minimum standard of 45% recommended in the literature [3]. The extract from H exhibited a higher flavonoid and phenolic acid value when compared with S and C2. However, it exhibited one of the lowest scores for antimicrobial activity, which suggests that antimicrobial activity is more influenced by flavonoids than the total phenolic acid.

Considering the phenolic composition, the variability of phenolic and flavonoid concentrations observed in this study substantiates the impact of geographical origin and the influence of the local flora. Extracts A, S, and H exhibited the highest concentrations of phenolic acids, whereas C1 and C2 exhibited a greater abundance of flavonoids. These findings are consistent with those reported by Patel et al. [42], who documented phenolic concentrations in ethanolic propolis extracts at 270 ± 9.2 mg GAE/g, and Bojić et al. [43], who observed flavonoid levels ranging from 6.83 to 55.44 mg/g, contingent on the source and extraction methodology. This reinforces the notion that phenolic and flavonoid compounds serve as crucial determining factors in propolis bioactivity. Caffeic acid was identified in all samples, with the highest concentration observed in extract A (77.95 μg/g). This finding is in accordance with the observations reported by Devequi-Nunes et al. [44], who also noted significant inter-sample variability in the content of phenolic acids. Extract S exhibited the highest concentrations of both protocatechuic acid (0.62 μg/g) and vanillic acid (1.3 μg/g).

These findings are consistent with those of Salleh et al. [45], who reported phenolic contents of 10–28.65 mg/mL in Malaysian stingless bee propolis. The present study thus provides further evidence linking phenolics to antimicrobial and antioxidant activities. Flavonoid analysis revealed that C1 is especially abundant in naringenin (55.41 μg/g), apigenin (22.46 μg/g), and pinocembrin (357.86 μg/g), which correlates with its pronounced antimicrobial activity. This is consistent with the findings of Jiang et al. [46], who highlighted the pivotal role of flavonoids in propolis bioactivity, and Elbatreek et al. [47], who reported pinocembrin’s therapeutic potential in antimicrobial applications. Extract S, which exhibits elevated levels of protocatechuic and vanillic acid, corroborates the findings of Zhao et al. [48], who reported a considerable polyphenol content in Brazilian green propolis. Similarly, the abundance of naringenin and apigenin in C1 is in accordance with the findings of Pobiega [8], emphasising the antioxidant roles of flavonoids. The presence of CAPE in extracts A, C1, and S underscores the significance of botanical diversity, as CAPE exhibits a broad-spectrum of antimicrobial and antioxidant activity [49]. The concentrations of polyphenols and flavonoids reported in this study are consistent with those reported in the literature, which highlights substantial variability across propolis samples. For example, Silva et al. [50] reported total phenolic contents ranging from 2.93 to 8.00 mg/mL in Uruguayan propolis, which is similar to the phenolic profiles observed in samples A and S. Furthermore, the findings are consistent with those of Barbarić et al. [51], who noted significant bioactivity at phenolic concentrations as low as 1 mg/mL. Furthermore, the observed variability in flavonoid content among the extracts is comparable to the findings of Bojić et al. [43], who reported a flavonoid range from 6.83 to 55.44 mg/g, depending on the extraction method and geographical origin.

The FRAP assay results ranged from 4.8 to 7.4 mg Trolox Eq./g, with extract A demonstrating the highest ferric-reducing power. These findings align with those of Duca et al. [52], who observed a FRAP value of 150 µmol FeSO₄/g for Romanian propolis, emphasising the significance of caffeic acid in antioxidant activity. Similarly, Mello and Hubinger [53] observed higher FRAP values for Brazilian green propolis, further underscoring the role of phenolic compounds in enhancing ferric-reducing capacity. The DPPH assay indicated that extract A exhibited the highest radical scavenging activity (31.6 mg Trolox Eq./g), while extract C2 exhibited the lowest value (24.9 mg Trolox Eq./g). These findings align with those reported by Devequi-Nunes et al. [44], who observed an IC50 of 12.5 µg/mL for DPPH radical scavenging in ethanolic extracts, demonstrating substantial antioxidant capacity. Silva et al. [54] observed a DPPH activity ranging from 50% to 90% for Brazilian propolis, providing evidence for the antioxidant activity of ethanolic extracts. The ABTS assay showed that extracts C1 and A exhibited the highest antioxidant activity, while extract C2 demonstrated the lowest. This trend aligns with the findings of Kamel et al. [55], who reported a TEAC value of ~1200 µmol Trolox/g for ethanolic extracts, indicating strong free radical neutralisation. Pratami [56] observed a TEAC value of 850 µmol Trolox/g for ethanolic extracts, emphasising the effectiveness of this solvent in extracting bioactive compounds with high antioxidant capacity. The findings of this study corroborate the robust antioxidant capacity of ethanolic propolis extracts, which aligns with the observations documented by Yang et al. [57]. These observations pertain to the DPPH scavenging efficacy of 65% at 1 mg/mL and the total phenolic content of 174.7 µg GAE/mg dry sample for Chinese propolis. Furthermore, the findings align with those of Mello and Hubinger [53], who observed superior antioxidant efficacy in ethanolic extracts in comparison to aqueous extracts. This superiority was attributed to the enhanced efficiency of ethanol in the extraction of polyphenols and flavonoids. The elevated antioxidant activity observed in extracts A and C1 may be attributed to their considerable phenolic and flavonoid content, as evidenced by previous analyses. This relationship is corroborated by Devequi-Nunes et al. [44], who underscored the robust correlation between total phenolic content (150.5 mg GAE/g) and antioxidant activity. The observed variability in antioxidant activity among the samples in this study is also consistent with the findings of Duca et al. [52], who identified significant regional differences in antioxidant potential due to variations in phenolic composition.

The antimicrobial activity of propolis extracts varies depending on the microorganism and origin. Among Gram-positive bacteria, E. faecalis showed the highest inhibition zones with C1, followed by S. C2, A, and H had similar but less pronounced effects. These findings align with those of Nascimento et al. [58], who recorded inhibition zones of 14 mm for E. faecalis, substantiating the efficacy of propolis against this bacterium. Kayaoğlu et al. [59] also highlighted comparable activity, validating the antimicrobial potential of the C1 extract. The C1 extract was also the most effective against S. aureus, followed by C2, A, S, and H. These findings align with those of Przybyłek and Karpiński [13], who reported inhibition zones of 10–20 mm for S. aureus. Silva et al. [20] observed 20 mm inhibition zones for S. aureus using Brazilian propolis. This shows the broad antimicrobial spectrum of propolis, particularly against Gram-positive pathogens. The C1 and C2 extracts were similarly effective against MRSA, with extract A showing a slightly lower activity. The weaker performance of S and H against this resistant strain highlights the variability in propolis chemical composition. These findings align with those of Kubilienė et al. [60], who demonstrated propolis efficacy against antibiotic-resistant strains like MRSA, reinforcing its potential as an alternative therapeutic agent. The inhibitory effects against S. epidermidis were most potent in extract C1, with the C2, A, and S extracts displaying reduced yet quantifiable activity. In contrast, the H extract demonstrated minimal inhibitory effects. This trend reflects the impact of geographical origin and extraction methods, as highlighted by Machado et al. [61], who reported significant variability in antimicrobial potency based on these factors.

The response of L. monocytogenes to propolis extracts shows that chemical diversity affects bioactivity [62]. C1 showed the strongest inhibitory effect, while A, C2, and S were less active. H was the least active. Gram-negative bacteria were less susceptible overall, as observed in previous studies [20,60]. The authors attributed this to the structural barriers of Gram-negative bacteria, particularly their outer membrane. For S. enterica ATCC 25928, the A and C2 extracts were observed to exert the greatest inhibitory effect, while extract H demonstrated the least activity. Similar findings were reported by Gülbandılar [62]. Propolis showed the greatest activity against K. pneumoniae with extract S, with an inhibition halo of 7.2 mm, which is lower than the 18 mm reported by Al-Ani et al. [63]. The moderate inhibition observed in this study highlights the need to optimise extraction methodologies to enhance efficacy against this pathogen. The A and C2 extracts showed the most notable activity against P. aeruginosa, with inhibition zones of 11.5 and 11.1 mm. Nascimento et al. [58] reported comparable outcomes with inhibition zones of ~12 mm. The effects against P. aeruginosa are consistent with the bacterium’s resistance mechanisms. E. coli inhibition zones were uniform across extracts, except for S and C2. These findings align with those of Przybyłek and Karpiński [13] and Silva et al. [20]. A. baumannii was most inhibited by extract S and least inhibited by H extract. Similarly, Al-Ani et al. [63] reported inhibition zones of 16 mm, underscoring strain-to-strain variability. The antifungal activity against C. albicans demonstrated the highest inhibition zones for C1 and A, while extract S exhibited the lowest activity. These findings are in accordance with those reported by Mattigatti et al. [64].

The MIC results for the five propolis extracts show variability in antimicrobial efficacy across different bacterial strains; Gram-positive and Gram-negative bacteria show significant variability, as documented in the literature. The discrepancies are likely due to the chemical composition of the extracts and the bacterial target. Among Gram-positive bacteria, S. aureus showed the highest sensitivity, with extract C2 exhibiting an MIC of 0.25 mg/mL. This aligns with the findings of Mascheroni et al. [65], who reported a similar MIC of 0.2 mg/mL. Tiveron et al. [66] noted lower values ranging from 62.5 to 125 µg/mL for red propolis. Alternatively, Al-Ani et al. [63] reported higher MICs between 1500 and 2500 µg/mL, reflecting strain variability. MRSA displayed reduced sensitivity, with extract C1 showing a MIC of 0.59 mg/mL. This value is higher than the MIC range of 80 to 100 µg/mL reported by Miorin et al. [67] but is within the upper limit reported by Stepanović et al. [68], who documented values up to 1.25%. Against E. faecalis, extract C1 exhibited the greatest efficacy with a MIC of 0.75 mg/mL. This value is higher than those reported by Vardar-Unlü et al. [69], who observed values between 0.12 and 0.25 µg/mL, and Nina et al. [70], who documented MICs ranging from 14.0 to 210.0 µg/mL, likely reflecting differences in extraction methods, chemical composition, or bacterial strain sensitivity. For L. monocytogenes, extract C1 was the most potent with an MIC of 0.50 mg/mL, consistent with Mascheroni et al. [65], who observed 0.6 mg/mL for L. innocua. Extract H, the least effective sample, displayed a MIC of 5.63 mg/mL.

In contrast to the observed efficacy of the propolis extracts against Gram-positive bacteria, the activity against Gram-negative bacteria was generally less pronounced, consistent with the established structural resistance of Gram-negative bacteria due to the presence of an outer membrane [71]. The extract C2 showed the highest efficacy against E. coli (0.78 mg/mL), while H showed the weakest (11.25 mg/mL); results that align with previous studies [72,73]. Extracts A and S were the most efficacious against K. pneumoniae, with a MIC of 8.13 mg/mL. Al-Ani et al. [63] reported lower MIC values for K. pneumoniae, indicating potential differences in sensitivity across strains and experimental conditions. The inactivity of extract H against this strain highlights the variability of propolis efficacy. P. aeruginosa ATCC 27853 showed minimal sensitivity, with the lowest MIC of 2.56 mg/mL for extract C2. The current finding is aligned with those reported by Grecka et al. [71] and Marco et al. [74]. The elevated resistance of this strain makes targeting Gram-negative pathogens with propolis extracts difficult. With regard to S. enterica, extract C2 displayed the most pronounced activity (1.28 mg/mL), followed by C1 and A; results that are in accordance with those reported by Runyoro et al. [75]. A. baumannii exhibited the greatest sensitivity to extract C1 (6.25 mg/mL). These findings are corroborated by the results of Torres et al. [34], who emphasised the general efficacy of propolis against Gram-positive bacteria rather than Gram-negative bacteria.

The MIC results show that propolis extracts vary in antimicrobial activity across bacterial strains and that Gram-positive bacteria are more sensitive than Gram-negative bacteria to the extract composition and geographical origin of raw propolis. Although the antibiotics employed as positive controls yielded greater efficacy than the propolis extracts tested in this study, it is crucial to highlight that, in contrast to the commercial formulations (20–30%), the ethanolic propolis extracts evaluated are 10%. It is also important to note that the ever-increasing emergence of multidrug-resistant pathogens requires urgent attention. The use of propolis, or a combination of propolis and antibiotics, represents a promising avenue of research, given the potential risks associated with the unrestricted use of antibiotic molecules, including the development of further antibiotic resistance [11].

The antimicrobial efficacy of propolis extracts is influenced by TPC, TFC, and individual phenolic compounds. Extracts with higher phenolic and flavonoid contents exhibited greater antimicrobial activity, supporting the hypothesis that these compounds contribute to the observed effects. Extract C1, the most antimicrobial against Gram-positive bacteria, also had the highest total phenolic and flavonoid content. C1’s activity may be due to its phenolic and flavonoid content, which disrupts bacterial cell walls and membranes. This is consistent with the findings of Kubilienė et al. [60], who demonstrated a positive correlation between TPC and antimicrobial activity in propolis. Extract H, the least antimicrobial, had the lowest TFC and moderate TPC. This shows that while TPC is important, the specific composition of phenolics and flavonoids is crucial for antimicrobial efficacy. Extract C1 displayed the highest antimicrobial activity against Gram-positive bacteria and the highest total phenolic and flavonoid content, which disrupts bacterial cell walls and membranes. This finding is consistent with that of Kubilienė et al. [60], who proved a positive correlation between TPC and antimicrobial activity in propolis. Extract H, the least potent antimicrobial, had the lowest TFC and moderate TPC. These findings suggest that the efficacy of antimicrobial activity is more dependent on the specific composition of phenolic and flavonoid compounds than on their absolute quantity. Extract H, the least antimicrobial, had the lowest flavonoid content (1.5 mg QE/g) and moderate phenolic content (125.4 mg GAE/g). TPC is a contributing factor to antimicrobial efficacy, but the specific composition of phenolics and flavonoids is of primary importance. C1’s antimicrobial activity is due to key polyphenolic compounds, including naringenin, apigenin, and pinocembrin. These compounds have antimicrobial activity. Naringenin is effective against S. aureus and S. enterica, while pinocembrin has broad-spectrum antimicrobial and antibiofilm properties. Similarly, extract A, which exhibited robust antimicrobial effects, contained elevated levels of caffeic acid, CAPE, and chrysin. These compounds are antimicrobial and synergistic. CAPE is effective against S. aureus and S. epidermidis but not E. coli and K. pneumoniae. Extract H, with low antimicrobial activity, had lower levels of bioactive compounds, including chrysin and pinocembrin. This shows that both the concentration and composition of phenolics affect propolis extract antimicrobial potential.

The antioxidant activity of the extracts, as evaluated by FRAP, DPPH, and ABTS assays, also correlates with antimicrobial activity. Extract A, with the highest antioxidant values (7.4 mg Trolox Eq./g in FRAP, 31.6 mg Trolox Eq./g in DPPH, and 34.0 mg Trolox Eq./g in ABTS), exhibited strong antimicrobial effects. Antioxidants may enhance antimicrobial activity by neutralising oxidative stress, which can weaken bacterial defences, as noted by Silva et al. [20]. The high efficacy of C1 and A against E. faecalis, S. aureus, and L. monocytogenes correlates with their high levels of naringenin, apigenin, and pinocembrin. These compounds may act synergistically to enhance antimicrobial activity. While C1 and A showed moderate efficacy against E. coli and P. aeruginosa, the overall activity against Gram-negative bacteria was lower. This could be due to the outer membrane of Gram-negative bacteria, which reduces permeability to polyphenols. Extract C2 demonstrated better activity against E. coli and S. enterica, potentially due to its specific phenolic profile, including higher quercetin levels (4.25 μg/g), which is known to disrupt Gram-negative bacterial membranes [76]. The strong antifungal activity of C1 against C. albicans may be attributed to its high chrysin content (188.48 μg/g), which has been reported to effectively inhibit fungal growth. The presence of multiple bioactive compounds in C1 and A, such as CAPE, naringenin, apigenin, and pinocembrin, likely contributes to their synergistic effects. Studies indicate that combinations of these compounds exhibit enhanced antimicrobial activity compared to individual compounds [47]. In contrast, extracts like H, which lack some of these key compounds, exhibit reduced efficacy.

5. Conclusions

A comprehensive scientific investigation was undertaken in which several parameters were taken into account for the evaluation of the five ethanolic propolis extracts. From a geographical perspective, it is essential to recognise that the foraging range of honeybees is confined to the vicinity of the hive, thereby limiting the resources available to them. In this context, the Corine land cover is a valuable tool for evaluating the soil classes present in a given area and for making a preliminary assessment. Another notable finding is the close correlation between the biological and functional properties of propolis extracts and the plant essences accessible to bees in a given area, with greater diversity resulting in enhanced properties. In order to contribute to the standardisation of propolis, it is necessary to gain a deeper understanding of the behaviour of honeybees and the decision-making processes involved in the collection of resin, as well as to carry out a more comprehensive analysis of the local botanical origin of the materials used to make propolis. The results obtained from the antimicrobial activity support the idea of using natural substances to fight antibiotic-resistant pathogens; however, further research is needed in order to better understand the possible therapeutic applications.