Antioxidant and antibacterial potentials of four indigenous propolis extracts against oral tooth decay pathogens, Streptococcus mutans and Candida albicans

INTRODUCTION

Tooth decay is a multifactorial process influenced by various factors, such as poor oral hygiene, dental plaque build-up, presence of cariogenic bacteria, and inadequate fluoride exposure as well as individual tooth ana­tomy and enamel composition [1]. The composition of oral microbiota plays a crucial role in maintaining oral health, and any disruptions in microbial balance can lead to proliferation of opportunistic pathogens, contributing to oral diseases [2]. Certain acid-producing bacteria in the oral cavity, including Streptococcus and Lactobacilli, are known to be major contributors to tooth decay. Streptococcus mutans, a Gram-positive, facultative anaerobic cocci-shaped bacterium commonly found in the human oral cavity, is strongly associated with the development of dental caries [1, 3]. While, among fungal biological agents, Candida albicans is responsible for oral candidiasis, an opportunistic infection that arises due to disruptions in the oral microbial flora [3]. Imbalances in the oral microbiota can create a favorable environment for the overgrowth of undesirable micro-organisms, such as C. albicans, which can exacerbate oral infections and contribute to severity of oral diseases [2, 3]. Increased microbial resistance to conventional antimicrobial agents has prompted the use of various natural bioactive compounds, which could prevent dental carries. Among these natural bioactive ingredients, the therapeutic efficacy of propolis in periodontal diseases has been widely studied, showing their ability to prevent or reduce plaque formation on the tooth surface, leading to reduced risk of dental caries and other oral infections [4, 5]. Propolis is a natural, non-toxic phytocompound produced by Apis mellifera bee by feeding on a number of botanical compounds from various plant sources [3]. These phytochemicals with wide spectrum antimicrobial actions are produced by bees to protect their hives and colonies from a number of bacterial, fungal, and viral diseases [2, 6]. The antimicrobial potency of propolis has been attributed to their hydrophilic and hydrophobic polyphenolic compounds, such as caffeic acid and ferulic acid [7].
In personalized medicine, propolis has been used for centuries, combating pathogens and treating human diseases [8]. This complex phytocompound having numerous therapeutic and health benefits, are considered a natural remedy for treating tooth decay pathogens and improving oral health [9, 10]. Propolis exert their actions on the pathogens in several ways, including forming physical barriers, inhibiting essential enzymes and proteins required for invasion of the pathogen into the host, and preventing the replication of pathogen’s processes [11]. Propolis ability to inhibit metabolic processes of pathogens, disrupts cellular organelles of the target micro-organism and components responsible for energy production, leading to the growth inhibition of the pathogen [10, 12]. In dentistry, it has been observed that fatty acids in propolis could inhibit the formation of dental carries by creating a cariostatic effect, i.e., decreasing the tolerance of micro-organisms to low pH and decelerate acid production [13]. Additionally, the waxy nature of propolis can protect sensitive teeth by sealing the enamel and dentin, covering tiny tubules to reduce sensitivities [14]. Major advantages of using propolis is its biocompatibility, meaning it is safe for use in the oral cavity without harming surrounding tissues [15].
The chemical composition of distinct propolis can vary depending on geographical location, variability in the botanical source, climatic conditions, flora at the site of collection, and bee species [8]. Changes in the composition of different propolis can be responsible for striking variability in their functionality and bio-logical activity [11, 15]. Therefore, understanding geographical variations in the propolis’ composition, and their impact on antibacterial and antifungal activities may have significant implications for the development of natural oral health products.

OBJECTIVES

The aim of the present study was to analyze and compare the differences in several important functional properties of four propolis samples collected from different regions of Iran. Ethanolic extracts of respective propolis (EEP) were analyzed for their antibacterial and antifungal activities, antioxidant potentials, total phenols, and flavonoid contents. Furthermore, in this study for the very first time, the antibacterial and antifungal properties of a mixture of all four EEP (MEEP) against oral pathogens (S. mutans and C. albicans) were evaluated. Findings of the study provided valuable insights into the variations in the key functional components and antimicrobial effectiveness of the studied propolis against common oral pathogens. MEEP with enhanced biolo­gical activities can have potential applications in food or pharmaceutical industry in future.

MATERIAL AND METHODS

COLLECTION OF PROPOLIS AND PREPARATION OF ETHANOL EXTRACTS

Propolis samples were collected during spring 2021 from four geographically different regions of Iran, such as Karaj, Khorramabad, Hamedan, and Damavand. Frozen propolis samples were crushed manually, and their EEP were prepared by slight modifications in the method described by Al-Ani et al. [3]. Briefly, 10 grams of crushed propolis was mixed with 100 ml of 70% ethanol, and kept in dark for 24 hrs. at room temperature on a magnetic stirrer. Samples were filtered using Whatman filter paper (No. 4.0), and later centrifuged for 30 min at 4,000 × g. The collected supernatant was dried in a rotary evaporator at 50°C with low pressure. Mixed EEP (MEEP) samples were prepared by mixing all four EEP in equal proportions. All samples were kept in dark, at refrigerated temperature until further use.

ANTIBACTERIAL AND ANTIFUNGAL ACTIVITIES

Antibacterial and antifungal activities of EEP and MEEP were determined by agar-well diffusion meth-od [6]. S. mutans strain ATCC 25175 was grown in BHI (Brain Heart Infusion, Merck, Germany) broth under micro-aerobic conditions for 24 hrs. at 37ºC. C. albicans strain ATCC 10231 was grown in SDA (Sabouraud dextrose agar; HiMedia, Mumbai, India) for 48 hrs. at 37°C. Fresh colonies were suspended in 2 ml of sterile phosphate buffer, standardized to 0.5 McFarland turbidity standard (1.5 × 10⁸ CFU/ml), and spread evenly on the entire surface of agar plates with a sterile cotton swab. Wells (6 mm) were punched into the overlaid agar medium using crock borer, and filled with 50 µl of EEP suspensions diluted in dimethyl sulfoxide (DMSO). After incubation at 37°C, halo zone diameters around the wells were measured in millimeters. Normal saline solution (0.89%) was used as negative control, while gentamycin sulphate (10 µg/ml) and amphotericin B (16 µg/ml) were employed as positive controls for the respective bacterial and fungal pathogens.

MINIMUM INHIBITORY, BACTERICIDAL, AND FUNGICIDAL CONCENTRATIONS

A broth micro-dilution method was used to determine minimum inhibitory (MIC) of EEP and MEEP against S. mutans and C. albicans, according to the Clinical and Laboratory Standards Institute (M07-A9) [16]. In brief, 20 µl of microbial suspension was added to each well of a 96-well micro-titer plates, resulting in a final bacte­rial concentration of 1 × 10⁵ CFU/ml, with 0.2% chlorhexidine gluconate solution used as control. Different concentrations of EEP (i.e., 10, 25, 50, 75, 100, 125, 150, 175, 200, and 225 μg/ml) were prepared by mixing 50 mg of EEP with 500 μl of DMSO, and further diluting 1 ml of the suspension in 9 ml of 1 × PBS. The micro-plates were incubated in ambient air at 37°C for 24 hrs., and later observed visually for the presence of turbidity in respective wells. MIC values were defined as the lowest concentration, at which no microbial precipitate or turbidity was observed. Minimum bactericidal (MBC) and fungicidal concentration (MFC) were defined by culturing 50 μl aliquots of respective dilutions from the micro-titer plates, showing no growth (equal to or greater than the recorded MIC) on respective agar plates [17]. After incubation at 37°C for 24 hrs., the plates were observed for the appearance of visible colonies. MBC and MFC were the lowest concentrations, which pre-vented visible growth.

ANTIBIOFILM INHIBITION ASSAY

Antibiofilm effect of EEP samples against the two pathogens was tested with crystal violet assay using flat-bottomed 96-well polystyrene micro-titer plates [9]. Freshly grown cultures (0.5 McFarland turbidity standard) were dispensed into the wells of 96-well micro- titer plates containing BHI broth and different concentrations of EEP samples. After incubation at 37°C for 48 hrs., the plates were washed with distilled water, the wells air-dried and stained with 1% (w/v) crystal violet. After room temperature incubation for 15 min, unabsorbed stain was removed by washing the plates thoroughly with sterile distilled water, and absorbance was read at 540 nm. Staphylococcus aureus ATCC 25923 (biofilm producer) was used as positive control, S. epidermidis ATCC 122228 (non-biofilm producer) was employed as negative control, and inoculated broth medium was used as sterility control. Percentage of biofilm inhibition was calculated with the following formula:
(OD growth control – OD sample)/OD growth control) × 100,
where OD is optical density. Results were recorded as adherent when an OD ≥ 0.05 was obtained.

TOTAL PHENOL CONTENT

Total phenolic compounds in the prepared ethanolic propolis extracts were determined using Fo-lin-Ciocalteu’s method [9]. To 1.5 ml of distilled water and 0.4 ml of Folin-Ciocalteu’s reagent (2N), 0.2 ml of EEP extract was added. After 5 minutes, 0.6 ml of 20% sodium carbonate solution (w/v) was applied, and the tubes were incubated for 2 hrs. in dark at room temperature. Absorbance was measured at 760 nm by spectrophotometer (Agilent Carr UV-Vis 4000). 0.2 N Folin-Ciocalteu’s reagent (100 μl), 600 μl of distilled water, and 300 μl of 7.5% sodium carbonate were used as a blank sample. Standardization curve’s reference was gallic acid, and findings were expressed in mg gallic acid (mg GA/g EEP).

TOTAL FLAVONOID CONTENT

Aluminum chloride was utilized to measure total flavonoid content (TFC) of EEP samples, which were then converted to quercetin equivalents, according to previously described method [18]. In sterile falcon tubes, EEP samples were added to equal amounts of 2% AlCl3, and after homogenizing, the samples were placed in dark for 15 minutes at room temperature. Calibration curve was generated using quercetin (standard solutions of 5, 10, 15, 20, 25, 75, and 100 g/ml, 90% (v/v) ethanol). Absorbance was measured by spectrophotometer (Agilent Carr UV-Vis 4000) set at 435 nm. Blank consisted of equal amounts of 2% AlCl3 and distilled water. Based on calibration curve, TFC of the respective EEP samples was estimated in milligrams of quercetin equivalents (mg QE/g EEP).

ANTIOXIDANT POTENTIALS

Antioxidant capacity of EEP was evaluated using DPPH radical (2, 2-diphenyl-1-picrylhydrazyl) scavenging method [19]. DPPH (3 ml) was added to 1 ml of different EEP concentrations (i.e., 10, 25, 50, 75, 100, 125, 150, 175, 200, and 225 μg/ml). After 30 minutes of incubation in dark, the absorbance at 517 nm was measured by spectrophotometer (Carry UV-Vis 4000, Agilent). Antioxidant activity was assessed as a percentage of radi­cal oxidation inhibition.
% Inhibition = Abs_Control – Abs_Extract /Abs_Control × 100,
where Abs_Control is the absorbance of the control sample, and Abs_Extract is the absorbance of the extract. Results were expressed as the percentage inhibition.

HEMOLYTIC ACTIVITY

Percentage hemolysis of human erythrocyte cells by EEP was determined using previously described method [20]. Peripheral blood drawn from healthy human volunteers was poured into sodium cit-rate-containing tubes, and subsequently centrifuged at 1,500 × g for 10 min; the collected erythrocytes were washed thrice with a saline solution. For the test, 10% erythrocyte suspension was prepared in saline to obtain a final 2.5% after treatments. The erythrocytes were pre-incubated at 37°C for 30 min in the presence of different concentrations of EEP. Control included erythrocyte suspensions with 0.9% NaCl, while total hemolysis was induced by incubation of the erythrocyte suspensions with distilled water, and again incubated at 37°C with periodical stirring. After 0, 8, 12, 24, and 48 hours, the aliquots were centrifuged (1,500 × g for 10 min), and the absorbance of supernatant was read at 540 nm. The percentage hemo­lysis was measured using the formula below: A/B × 100, where A is the sample absorbance, and B is the total hemolysis.

STATISTICAL ANALYSIS

Data were analyzed using SPSS software v. 20.0, and one-way ANOVA test was employed to compare in vitro antimicrobial activity of the different propolis extracts against the respective pathogens. All data were obtained from three independent experiments.

RESULTS

The findings from the agar-well diffusion assay (Table 1) provided valuable insights into the antibacterial and antifungal activities of the tested propolis samples, collected from different regions of Iran. Based on the measured inhibitory zone diameters, significant differences were observed in the antibacterial effects of EEP samples (F_{5, 27} = 10.835, p < 0.0001). Additionally, the pattern and the extent of sensitivity of the two pathogens against the respective EEP varied significantly (F_{1, 27} = 7.355, p = 0.011). EEP from the region of Karaj (EEP-K) demonstrated the strongest inhibitory effects (Figure 1) against the tested bacterial pathogens (21.0 ± 1.15), but were weaker compared with the effects of gentamycin sulphate (23.0 ± 1.33). While EEP from the Damavand region (EEP-D) was the least effective against S. mutans (15.0 ± 1.72), but had the highest inhibitory effects against the fungal agent, i.e., C. albicans (18.0 ± 1.34). No significant differences in antifungal activity of EEP-K and EEP-D were observed, and their effects were comparable to amphotericin B (p > 0.05). The differences in the antagonistic effects of EEP from Khorramabad (EEP-KH) and EEP from Hamedan (EEP-H) regions were insignificant (p > 0.01). Similarly, insignificant differences were recorded in the antifungal effects of EEP-K and EEP-D against C. albicans. However, the mixture of all four propolis extracts (MEEP) showed significantly wider inhibitory zone of inhibition compared with individual EEP, and these differences were significant. Overall, the tested propolis extracts were more effective against the bacterial pathogen compared with the fungal strain (F_{1, 56} = 6. 6735, p = 0.024). MIC and MBC values of the tested EEP against the pathogens are depicted in Table 2. EEP-K was the most effective against the tested pathogens, and showed the lowest MIC and MBC values (125 µg/ml and 150 µg/ml, respectively) against S. mutans compared with other tested extracts. Even though EEP-D appeared the least effective against S. mutans, it showed the highest MIC and MBC values against the pathogen (225 µg/ml and 250 µg/ml, respectively). However, EEP-D demonstrated the least MIC and MBC values against the fungal agent (150 µg/ml and 175 µg/ml, respectively). Moreover, the inhibitory effects of EEP-D against C. albicans as seen by their MIC (150 µg/ml), appeared to be stronger then amphotericin B (175 µg/ml). Among the tested propolis ex-tracts, EEP-KH was the least effective in inhibiting the pathogen, showing the highest MIC and MBC values (200 µg/ml and 250 µg/ml, respectively). All the tested EEP had higher MBC values compared with MIC values against both the bacterial and fungal pathogens, while C. albicans appeared more resistant compared with S. mutans to the actions of EEP (p < 0.05). As observed during agar-well diffusion assay, MEEP provided significantly enhanced antimicrobial effects against the tested pathogens, compared with individual EEP (p < 0.05). The least MIC and MBC values were recorded for MEEP against S. mutans (100 µg/ml and 150 µg/ml, respectively) and C. albicans (150 µg/ml and 200 µg/ml, respectively). MIC values of MEEP against S. mutans and C. albicans were lower than the effects of gentamycin (125 µg/ml) and amphotericin B (150 µg/ml) against these pathogens.
The ability of different concentrations of EEP to eliminate the biofilms of S. mutans and C. albicans was evaluated, and their percentage of inhibition was estimated (Figure 2). All the tested EEP were able to inhibit the biofilms of both the pathogens at all used concentrations; however, significant differences were recorded in the inhibitions of the bacterial biofilms compared with the fungal biofilms (p < 0.05). EEP-K were the most effective in eliminating S. mutans biofilms (≥ 150 µg/ml). Although EEP-D were the least effective in eliminating S. mutans biofilm, they were the most effective in the removal of C. albicans biofilms (p < 0.05). EEP-D eliminated approximately 80% of C. albicans at the concentration of 200 µg/ml. Total phenol and flavonoid contents of EEP samples were determined (Table 3). Significant differences were observed in the amounts of phenolic compounds in the tested EEP (F_{3, 6} = 12194377.2, p = 0.0001). Similarly, the total amounts of flavonoids in the four EEP samples varied (F3, 6 = 1257551.29, p < 0.0001). Overall, EEP-K samples had the highest level of total phenolic content (TPC) content, while EEP-D had the highest TFC content. The least contents of phenols and flavonoids were determined in EEP-KH and EEP-H samples (p < 0.05), respectively.
Table 4 presents the results of antioxidant capacity of the tested EEP samples. The tested EEP demon-strated significant antioxidant activity at different tested concentrations (p < 0.05). All EEP showed above 50% of antioxidant percentage at only 50 µg/ml. EEP-D (98.11 ± 1.23) and EEP-K (97.76 ± 1.48) indicated the highest radical scavenging activity at the highest used concentration (225 µg/ml), but the differences between the two were insignificant (p > 0.05). None of the EEP extracts showed hemolysis of red blood cells at concentration of 250 µg/ml at the tested time intervals (Table 4).
EEP-H and EEP-D at the concentration of 275 µg/ml indicated a slight hemolytic percentage (0.78%) after 48 hours of incubation.

DISCUSSION

In dentistry and oral medicine, propolis has been included in many products, such as mouthwashes, gels, varnishes, cavity cleaning agents, etc. [11]. Propolis possess antimicrobial properties, and effectively targets and inhibits the growth of a wide range of bacterial and fungal pathogens [21, 22]. The bioactivities and pharmacological effects of propolis are affected by the solvent used for its extraction. Compared with water, chloroform, and ester fractions, ethanol extracts of propolis are shown to be more effective with higher antimicrobial potentials [23].
In this study, ethanol extracts of four indigenous propolis were investigated for their biological functions and antimicrobial activity. Despite variations in the plant origin and climatic condition across different Iranian regions, all the tested EEP demonstrated the ability to inhibit both the pathogens. Consistent results have been reported by other authors, who showed that various propolis samples were able to inhibit the growth of S. mutans and C. albicans; although the samples were collected from different regions and botanical sources [7, 24].
The differences observed in the antimicrobial efficacy of each EEP may suggest distinct composition of bioactive compounds present in each propolis [12, 20]. Additionally, these variations could also depend on the type of extract (ethanolic or aqueous), type of microbes, inoculum concentration, and propolis concentration in the medium [11, 20]. Notably, the propolis extract from Karaj (EEP-K) possessed the strongest antibacterial effects, while conversely, EEP from Damavand were the least effective against this pathogen. However, EEP-D had the strongest inhibitory effects against C. albicans. This differential activity against bacterial and fungal pathogens highlights the complex nature of propolis composition and its selective antimicrobial actions [10].
Similar to these findings, it has been reported that EEP are more susceptible to Gram-positive and Gram-negative bacteria, compared with fungal agent C. albicans [9, 21]. Propolis targets wide range of bacteria using several mechanisms, including its ability to cause fractional bacterial lysis (bacteriolysis), altering membrane permeability, impeding cytoplasmic membrane function, reducing the capability to form biofilms, inhibiting energy-generating pathways, and reducing bacterial resistance to certain conventional antibiotics [1, 25]. Overall, the bacterial pathogen was more sensitive to the actions of EEP compared with the fungal agent, which might be explained by differences in the outer cell membrane structure of the two. The antifungal effects of propolis has been demonstrated by Corrêa et al. [25], who showed that Brazilian propolis can exert its antifungal effects on C. albicans by damaging the integrity of cell wall and cell membrane, leading to leakage of intra- cellular organelles, and ultimately cell death. These researchers hypothesized that the antifungal efficacy of propolis is due to the capacity of polyphenols to form a complex with soluble proteins by disrupting the synthesis of chitin, which leads to cell wall disruption. Intriguingly, mixing all four propolis extracts (MEEP) together resulted in enhanced antibacterial and antifungal effects compared with EEP samples. These results indicate that combining all four propolis extracts increased the concentration of biologically active compounds in MEEP, and the synergistic interaction among key phenolic components led to enhanced antimicrobial effects [26]. The MIC values of MEEP against the pathogens appeared comparable to genta­mycin and amphotericin. Similar to these results, Kolayli et al. [27] reported that ampicillin had weaker zone of inhibition against Escherichia coli (10 mm) com-pared with the effects of propolis from Zonguldak and Konya regions of Turkey (12 mm) against this pathogen. Moreover, they demonstrated that red Brazilian propolis was more effective then ampicillin (10 mm) against Yersinia pseudotuberculosis (15 mm) and Klebsiella pneumoniae (11 mm). In another study, Vică et al. [28] reported that propolis collected from Romania had antimicrobial activity higher than the tested antibiotics. Enhanced activity of MEEP compared with antibiotics might also indicate the presence of high concentration of propolis in the medium, larger quantity of active components, and/or low concentration of the antibiotic used, higher inoculum size, and the resistance of pathogens against the respective antibiotics.
Propolis collected from different geographical location show diverse composition, with over 600 components identified to date [29]. The antibacterial activity of propolis is primarily attributed to its flavonoids and phenolic acid content [7, 28]. EEP-K samples collected from Karaj region exhibited the widest antibacterial action against S. mutans, and had the highest level of TPC. Compounds observed in Chinese popular propolis, such as himachalen, curcumene, bergamotene, and sesquicineole, were shown to influence its antibacterial effects against S. mutans [21]. However, the antifungal activity of propolis was linked to the presence of flavonoids, aromatic diterpenic compounds, and phenolic acids [22]. Among the tested etha­nolic extracts of propolis, EEP-D exhibited the highest antifungal activity, possibly due to its high TFC. However, other chemical constituents present in propolis may also play a role in its antifungal effects. For instance, com-pounds, such as 3-acetyl pinobanksin, pinobanksin-3-acetate, pinocembrin, caffeic acid, and p-coumaric acid, have been identified in propolis extracts and associated with their antimycotic properties [9, 23]. In contrast to the studies of Shokri et al. [30] and Stähli et al. [23], significant differences were observed in the MIC and MFC values of the studied EEP. The ethanolic extracts of propolis from Karaj (EEP-K) and Damavand (EEP-D) exhibited potent anti­fungal activity against C. albicans, and showed significantly lower MIC and MFC values than those reported by Shokri et al. [30]. Iranian propolis studied by these researchers had MIC and MFC values against C. albicans in the range of 2 to 20 mg/ml and 4 to 25 mg/ml, respectively. Concurrently, Stähli et al. [23] also reported MIC values below 10 mg/ml of each EEP to be effective against C. albicans strain ATCC 76615, while in another study, these values for 50 propolis collected from Northern Poland were found to be > 25 mg/ml [31]. The pre­sence of a diverse array of flavonoids and phenolic acids in propolis extracts from Karaj (EEP-K) and Damavand (EEP-D) may contribute to their strong antifungal pro­perties against C. albicans.
Antibiofilm activity of propolis has been studied by many researchers [31]. The examined propolis ex-tracts were able to inhibit the biofilms of both the tested pathogens, which is consistent with the reports of other researchers [21, 23]. S. mutans and C. albicans are key pathogens involved in biofilm formation on the teeth, contributing to dental caries. Propolis extracts are known to interfere with the biofilm formation of the pathogens, preventing them from adhering to surfaces in the oral cavity, and forming plaque that con-tributes to tooth decay and oral lesions [27]. The ability of oral bacterial and fungal pathogens to form bio-films is a significant factor in the development of dental caries, as these structures provide protection against antibiotics and host defenses.
Microbial cells growing in biofilms are 1,000 to 1,500 folds more resistant to antibiotics and biocides than planktonic cells, thus protecting the microbes from anti­biotics and host defense during infections [23]. Fun-gal biofilms, in particular, pose challenges in eradication compared with bacterial biofilms [21], and infections caused by fungal biofilms are more difficult to eradicate [32]. Therefore, targeting and disrupting microbial biofilms are crucial for preventing and treating dental caries.
All four propolis extracts in the current study demonstrated significant antioxidant activity, a common characteristic of propolis from various geographic regions [33]. The antioxidant potential of these extracts, as determined by the DPPH assay, was notably higher than in some other studies on propolis [34]. Variations in antioxidant activity of different propolis might indicate variations in the type and quantity of key phenolic compounds, such as caffeic acid, ferulic acid, and caffeic acid phenethyl ester in different propolis [32, 35]. Ensuring the safety and lack of toxicity of propolis is crucial for its potential use in medical applications [35, 36]. The evaluation of EEP on human erythrocytes revealed minimal hemolysis, with only EEP-H and EEP-D showing 1% hemolysis at the highest concentration tested (275 µg/ml). This suggests that these propolis extracts are relatively non-toxic to red blood cells. A previous study on Lebanese propolis extract also demonstrated low hemolytic activity (5%) at a concentration of 10 µg/ml, indicating its safety for red blood cells [37].

CONCLUSIONS

The findings of the current study highlight the importance of regional variations, which affect the quantity and quality of propolis’ components that possibly can influence their antioxidant and antimicrobial proper-ties. The results indicates the potential antimicrobial efficacy of Iranian propolis, especially those collected from Karaj and Damavand regions. Most importantly, the synergistic actions observed after combining all four propolis extracts indicate that the functionality of propolis could be improved in combination therapy. The tested Iranian propolis extracts appeared safe and hemocompatible, supporting their potential for further studies and clini­cal use.
However, the current research is preliminary study on different Iranian propolis extracts, and further investigations with higher number of samples are highly recommended to determine the bioactive compounds present in different propolis, to better understand the correlation among the type of phenolic compounds, their amounts, and interaction between them. These findings may highlight the mechanism underlying bene­ficial effects of indigenous propolis, and provide valuable insights into their therapeutic applications in oral health, pharmaceutical industry, or food. Therefore, further studies are required.

DISCLOSURES

1. Institutional review board statement: This study was approved by the Ethics Committee of Shahid Be-heshti University of Medical Sciences (approval number: IR.SBMU.RETECH.REC.1401.321).
2. Assistance with the article: None.
3. Financial support and sponsorship: None.
4. Conflicts of interest: The authors declare no potential conflicts of interest concerning the research, authorship, and/or publication of this article.

References