Evaluation of the antimicrobial activity and cytotoxic effect of hydroxyapatite containing Brazilian propolis

The great need to utilize biomaterials in the orthopedic and odontology fields is justified by the increase in life expectancy and factors such as trauma, injury and disease. Hence, the main use of bioceramic materials, as spare parts, is related to the degeneration of bone tissue, embracing a vast field of medicine [1, 2]. Thus, because of its similarity with the mineral component of the bones, HA is the most used calcium phosphate biomaterial. It is able to induce biological reactions that result in the formation of bone tissue over the implant surface by means of chemical bonds, enabling the implant to be covered by bone cells. Its main applications are the filling of small bone defects and the coating of ceramic and metallic materials [3–5].

However, one of the most harmful effects associated with implant surgery is infections. Infections in arthoplasties can cause the loosening of the bone/implant interface, bad positioning and wear or fracture of the material, disabling the person, thereby increasing the time of permanence at the hospital as well as health system costs [6, 7]. Among the microorganisms that cause infections in arthoplasties, the main ones are S. aureus and Staphylococcus epidermidis (50%–60% of cases) and Escherichia coli (up to 10% of cases). S. aureus is an important pathogenic agent because of its high virulence and high frequency [7]. Even though some studies indicate a reduction in infection cases with the use of prophylaxis and antibiotics, it is known that the bacteria become resistant to this therapy [8]. Within this context, natural products have several pharmacological properties, and are widely used in popular medicine as an alternative to the conventional remedies [9–11].

Propolis is a resinous natural product, composed of a complex mixture of compounds collected from plants and mixed with beeswax and saliva. The resinous material is used as protection for the hives against attacks from predators and microorganisms [12–14]. The interest in this natural product is increasing, because of its chemical, biological and pharmaceutical characteristics. Several studies have demonstrated some of the properties of propolis, such as antifungal, antiviral, antioxidant, anti-inflammatory, antitumor and antibacterial activity [15–18]. The antibacterial activity of propolis is one of the most investigated pharmacological characteristics of this material, in the development of alternative treatment to fight infections [19]. Its action has been attributed to the presence of phenolic compounds, which are able to cause functional and structural alterations, such as damage to the cytoplasmic membrane, inhibition of the synthesis of nucleic acids, energetic metabolism and the formation of biofilms. It also increases the permeability of the cell membrane, causing lysis and the death of bacteria [20–22].

Since previous studies have showed that hydroxyapatite pellets containing green and red propolis exhibit only inhibitory activity against S. aureus after 24 h [23], the question may be raised of whether or not this material may also exhibit bactericidal activity against the same microorganism. In the present work, the bactericidal effect of hydroxyapatite, in the form of powder, containing dry green and red propolis extracts, was explored. In contrast to the cited work, dry green and red propolis extracts were incorporated into the hydroxyapatite samples by means of a spray drying technique. Furthermore, the HA/propolis complex constitutes a fast system, which can be used to perform a release study. This has not been reported before in the literature for such a material, showing bactericidal characteristics over a shorter interval of time.

2.1. Preparation of ethanolic extracts of propolis (EEP)

2.2. Synthesis of hydroxyapatite

2.3. Preparation of the HA powder with propolis

2.4. Characterization

2.5. Release test

2.6. Antibactericidal activity

2.6.1. Bacterial strains

The antibactericidal analyses of the EEP and the materials were made with the use of the bacterial strain Staphylococcus aureus ATCC 25923. All the culture media and materials used in the assays were moist heat sterilized with the use of an autoclave at 120 °C and a pressure of 1 kgf cm⁻² for 20 min. Bacteria were reactivated in a TSB culture at 37 °C for 24 h. The concentration of the bacterial suspension was adjusted to 1 × 10⁸ UFC ml⁻¹, which corresponds to 0.5 on the MacFarland scale.

2.6.2. Counting of bacterial colonies

The counting of viable bacterial colonies after contact with the materials as a function of time was made by the methodology previously described [28]. Briefly, the assay was made at a concentration of 2% (m/v) in TSB medium culture containing bacterial suspension (1 × 10⁵ UFC ml⁻¹). The material was homogenized for 20 s, an aliquot was immediately collected and appropriate dilutions were made. Then, aliquots of 100 μl of all dilutions were seeded in Petri dishes containing Mueller–Hinton agar media (MH) (time zero). The test tubes with the samples were incubated at 37 °C under stirring (150 rpm) and specimens were collected at times of 0.5, 1, 2 and 4 h. For each collecting time, the solutions in the tubes were homogenized and the same procedures for time zero were repeated. Two references were used: (1) with HA; (2) with TSB and bacteria (bacterial growth).

The percentage of bacterial reduction (%R) for each collecting time was determined according to equation (1).

$$\begin{eqnarray} &&\% R=\left(\tfrac{{C}_{0}-C}{{C}_{0}}\right)\times 100\end{eqnarray} \tag{ 1 }$$

where C₀ is the average number of colony-forming units (CFU) of the HA reference, and C is the average number of CFUs of the following materials: HA-GP20, HA-GP8, HA-RP20 and HA-RP8.

2.6.3. Determination of minimal inhibitory concentration (MIC) and minimal bactericidal concentration (MBC)

The MIC for the EEP was determined according to the methodology previously described [29]. Briefly, the EEP (serial dilutions with concentrations varying from 12.5−1600 μg ml⁻¹) were added to the TSB medium containing the bacterial suspension (1 × 10⁵ UFC ml⁻¹) and then incubated at 37 °C for 24 h under stirring (150 rpm). Four control solutions were used: (1) with ethanol 80% (negative control); (2) with gentamicin at 25 μg ml⁻¹ (positive control); (3) with phosphate-buffered saline (PBS) (bacterial growth); (4) with TSB medium (without inoculum) and ethanol 80% (white).

In order to determine the CIM regarding the materials, release was made in a TSB medium of 2% (m/v) at 37 °C for 24 h under stirring (150 rpm). After release, the material was centrifuged (3500 rpm for 5 min) and the supernatant from each material was serially diluted using the TSB medium. After dilution, the bacterial suspension (final concentration of 1 × 10⁵ UFC ml⁻¹) was added, followed by incubation at 37 °C for 24 h under stirring (150 rpm). Four control solutions were used: 1) with HA supernatant (negative control); 2) with gentamicin at 25 μg ml⁻¹ (positive control); 3) with TSB medium and bacterial suspension (bacterial growth); 4) with TSB medium (without inoculum) and PBS (white). After incubation (EEP and materials), the absorbance was read in a microplate reader at 620 nm (Multiskan FC, Thermo Scientific, USA). MIC was considered to be the lowest concentration range where there was no visible bacterial growth (absorbance smaller than 0.05). In order to improve visualization at the plate, 5 μl of resazurin dye was added (3 mg ml⁻¹). In this case, the blue color indicates no bacterial growth, while the pink color indicates the presence of viable bacteria.

The CBM was determined according to the methodology previously described [29]. Aliquots (100 μl) from the suspensions that showed absorbance lower than 0.05 were seeded in dishes containing Mueller–Hinton agar media (MH) and incubated at 37 °C for 24 h. CBM was considered to be the lowest concentration where no visible bacterial growth was observed on the dish.

2.7. Cell culture

The assay was made according to the methodology previously described [30]. The CHO-k1 cell line was cultivated in an RPMI 1640 culture medium (Gibco) supplemented with glutamine (2 mM) and fetal bovine serum (10%) in the presence of penicillin (100 U ml⁻¹), streptomycin (100 μg ml⁻¹) and amphotericin B (0.025 μg ml⁻¹) (SCM—supplemented culture medium). Cultivation was made in a CO₂ (5%) atmosphere at 37 °C for 72 h (Incubator CCL-170B, ESCO, USA). After this period, the supplemented medium was taken away and the cells were removed using a trypsin solution (Gibco TrypLE Express, phenol red 12605).

2.7.1. Cell viability

The materials HA, HA-GP8 and HA-RP8 were mixed with the SCM at a proportion of 2% (m/v), according to the ISO 10993-12 standard [31], and incubated at 37 °C in a CO₂ atmosphere (5%) for 24 h. After this period, the specimens were homogenized and centrifuged at 13 000 rpm for 5 min, and the supernatant was reserved for the assay.

Three controls were used: (1) alumina (negative control); (2) phenol solution 0.2% (positive control); and (3) SCM (white). After 24 h, the specimens and the controls (negative and positive) were serially diluted to the following concentrations: 100%, 50%, 25%, 12.5% and 6.25%.

The cytotoxicity was determined by the indirect method of neuter red incorporation, according to the ISO 10993-5 [32]. Briefly, cells (1 × 10⁴ cells/well) were cultivated in SCM and incubated at 37 °C in a CO₂ atmosphere (5%) for 24 h. Afterwards, the previous treatments (specimens and controls) and their respective dilutions were added, and the dish was incubated at 37 °C in a CO₂ atmosphere (5%) for 24 h. Following this, the neuter red solution (50 μg ml⁻¹) was added, and the dish was again incubated at 37 °C in a CO₂ atmosphere (5%) for 3 h, in order to incorporate the neutral red. After the neuter red was removed, an extraction solution (50% ethanol:1% acetic acid:49% distilled water) was added. Absorbance readings were made at 540 nm and the results were expressed in terms of cell viability (%CV), which was calculated according to equation (2).

$$\begin{eqnarray} &&\% {\rm{C}}{\rm{V}}=\left(\tfrac{{\rm{A}}{\rm{b}}{\rm{s}}\,{\rm{s}}{\rm{p}}{\rm{e}}{\rm{c}}{\rm{i}}{\rm{m}}{\rm{e}}{\rm{n}}\,}{{\rm{A}}{\rm{b}}{\rm{s}}\,{\rm{w}}{\rm{h}}{\rm{i}}{\rm{t}}{\rm{e}}}\right)\times 100\end{eqnarray} \tag{ 2 }$$

where 'Abs specimen' is the absorbance of the specimen at 540 nm, and 'Abs white' is the absorbance of the white at 540 nm.

2.8. Statistical analysis

Figure 1 shows the morphology of materials HA (A), HA-GP20 (B), HA-GP8 (C), HA-RP20 (D) and HA-RP8 (E). In the HA powder (figure 1(A)), agglomerates and apparently small particles with a spherical shape can be observed. On the other hand, in the HA specimens with different amounts of dry green and red EEP, a lower degree of agglomeration and apparently spherical particles is observed. So, the addition of propolis seems to lower the degree of particle agglomeration.

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FTIR analyses for dry EEP; the materials HA and HA with propolis are shown in figure 2. The FTIR spectra were measured for pure dry EEP (figure 2(A)), in order to identify characteristic functional groups of flavonoids and phenolic compounds, which are mostly responsible for the antimicrobial activity [22]. Table 1 shows signals from FTIR and their attributions. For HA, it is possible to observe characteristic peaks of groups ${{{\rm{PO}}}_{4}}^{-3}$ (1090, 1041, 964, 602 and 574 cm⁻¹) and O−H (3571 cm⁻¹). Furthermore, the presence of H₂O, characterized by the peak at 3431 cm⁻¹, and ${{{\rm{CO}}}_{3}}^{-2},$ with peaks at 1459, 1410 and 874 cm⁻¹ are also observed. The spectra obtained for specimens HA-GP20 and HA-GP8 (figure 3(B)), and HA-RP20 and HA-RP8 (figure 3(C)), show that there is a peak at 2929 cm⁻¹, which is characteristic of aromatic rings. The peaks observed at 1606 and 1257 cm⁻¹ for the specimens with green propolis (figure 3(B)) and 1619 and 1292 cm⁻¹ for the specimen with red propolis (figure 3(C)), which are attributed to the existence of C=O and C−O−C bonds, respectively, suggest the presence of phenolic acids and flavonoids in the analyzed materials (table 1). From these results, it is possible to say that the dry extracts of green and red propolis were incorporated into the hydroxyapatite, with the presence of total phenolic and flavonoid compounds in the obtained biomaterial.

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Table 1.  The FTIR signal attribution for materials HA and HA with green and red propolis.

HA	HA-GP	HA-RP	Attribution	Reference
3571			Axial deformation for O-H	[33–35]
3431			Axial deformation for H2O	[33–35]
	2929	2929	Axial deformation for C-H of aromatic ring	[36]
1640			Out-of-plane angular deformation for H2O	[33–35]
	1606	1619	Axial deformation for C=O	[36, 37]
1459 e 1410			Axial symmetrical deformation for ${{{\rm{CO}}}_{3}}^{-2}$	[34, 35, 38]
	1257	1292	Asymmetric axial deformation for C-O-C	[36]
874			Asymmetric axial deformation for ${{{\rm{CO}}}_{3}}^{-2}$	[39]
1090 e 1041			Asymmetric axial deformation for PO4	[33, 35, 37, 38, 40]
964			Symmetric axial deformation for PO4	[33–35, 39]
602 e 574			Out-of-plane angular deformation for PO4	[33, 35, 37, 39, 40]

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In order to verify the maximal release of propolis along time, besides comparing the absorption peaks of releases with the peaks of green and red EEP, UV–visible spectrophotometry analyses were also made (figures 3–5). Green and red EEP showed peaks of maximum absorption at 289 and 280 nm, respectively (figure 3). Materials containing green propolis showed a peak of maximum absorption at 289 nm, which is the same value found for the green EEP, indicating the presence of the same substances (not necessarily the same amounts) in the extract as in the release in the TSB buffer solution. Moreover, both specimens containing green propolis showed maximum propolis release starting at 30 min (figures 4(A) and (B)). Materials with red propolis showed a peak at 280 nm (in agreement with the peak for the red EEP); however, another absorption peak at 333 nm was also observed, indicating a difference in chemical composition between red EEP and the release in the TSB buffer solution. This suggests that some substances might have been retained in the HA (figure 5). The extract release of the specimen with the lowest amount of red propolis (HA-RP8) started at 30 min (figure 5). On the other hand, for the specimen with the highest amount of red propolis (HA- RP20), the maximum release was verified starting at 4 h (figure 5(B)).

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The total content of phenolics and flavonoids in the EEP, as well as the releases, were determined in order to verify the maximum release of these compounds and to compare them with the values found for green and red EEP. Considering the results obtained from spectrophotometry, a time of 4 h was adopted as the period for maximum propolis release for all specimens; the results are shown in table 2. Both the EEP and the releases showed a high content of total phenolics, and the highest values were observed for the materials with higher amounts of propolis (HA-GP20 and HA-RP20). On the other hand, the total amount of flavonoids released by the materials was lower when compared to their respective extracts. The specimens with red propolis showed a higher release of total flavonoids.

Figure 6 shows the percentage bacterial reduction along time for all specimens in relation to the reference (HA). Materials containing green propolis showed a bacterial reduction of 24.2% (HA-GP20) and 19.8% (HA-GP8) starting from the initial time (zero hour) (figure 6(A)), while the values for materials containing red propolis were 10.4% (HA-RP20) and 39.6% (HA-RP8), as shown in figure 6(B). Starting from 0.5 h of contact, materials with a higher amount of propolis (HA-GP20 and HA-RP20) showed 100% of bacterial reduction, while materials with a lesser content showed 92.6% (HA-GP8) and 95.8% (HA-RP8). Finally, starting from 1 h of contact with the bacterial suspension, it is possible to observe 100% bacterial reduction for all materials with both green and red propolis (figures 6(A) and (B)). The results found in this assay demonstrate that after maximum extract release, i.e. 100% release, all specimens showed a bactericidal effect in the time intervals ranging from 0.5 to 1 h.

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In order to investigate the antibacterial activity more deeply, and to compare the results obtained for the EEP, both the MIC and MBC were determined against S. aureus for the materials after 4 h of release. The red EEP showed greater inhibitory activity (12.5 μg ml⁻¹) when compared to the green EEP (100 μg ml⁻¹), and both EEPs showed the same bactericidal activity (800 μg ml⁻¹). As expected, the materials containing red propolis showed greater inhibitory and bactericidal activity when compared to the materials containing green propolis (table 3). However, all materials with green and red propolis showed less inhibitory activity and greater bactericidal activity when compared to the EEP. Materials with red propolis showed more significant differences in terms of MIC and MBC when compared to the EEP.

According to the results obtained regarding antibacterial activity, the materials with less green and red propolis, i.e. HA-GP8 and HA-RP8, were selected to determine cell viability. The cell viability curves for the specimens and controls are shown in figure 7. The materials (HA-GP8 and HA-RP8) and the positive control (phenol) showed similar profiles; increased concentrations resulted in decreased cell viability. On the other hand, the material HA and the negative control (alumina) did not show cytotoxic effect, with cell viability above 50%. The IC₅₀ values were 387.1 μg ml⁻¹ and 84.8 μg ml⁻¹ for the specimens HA-GP8 and HA RP8, respectively, suggesting a higher cytotoxic activity for the material with red propolis.

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It is also possible to compare the IC₅₀ values found in this assay with the MIC and MCB values for the specimens HA-GP8 and HA-RP8 (table 4). Both specimens (HA-GP8 and HA-RP8) showed MIC values bellow IC₅₀, indicating that the concentration needed to inhibit bacterial growth regarding S. aureus does not have a cytotoxic effect on the cell line used in this work. However, the extract concentration needed to cause bacterial death (MBC) is higher than IC₅₀, indicating the possible cytotoxic effect of materials at such a concentration.

In this study, extracts of green and red propolis were incorporated into HA powder by spray drying in order to obtain a biomaterial (HA) containing a bactericidal agent with a high contact surface. This material can be used as a fast release system for the antibacterial agent. All materials containing propolis showed significant antimicrobial activity against S. aureus. Comparing the results shown in the present work with those shown in the work by Grenho et al [23], several differences can be pointed out. Regarding the antibacterial effect, whereas in the reference, using pellets obtained via immersion, only inhibitory activity against S. aureus was observed after 24 h, in the present work with HA/propolis powder obtained by spray drying, a fast bactericidal action was observed in a time-lapse of 0.5 to 1 h (figure 6), with a faster bactericidal effect for the higher propolis concentrations. Furthermore, in the work by Grenho et al [23], reductions of 45% and 61% in the amount of viable bacteria (S. aureus) for HA with green propolis (12 and 25 μg ml⁻¹, respectively) and 99% for HA with red propolis for the same concentrations, were observed. Moreover, whereas the remaining bacteria populations reported in the literature were 10⁸ and 10⁶ UFC for the materials containing green and red propolis, respectively [23], in the present work a 100% bacterial population reduction was observed.

The antibacterial properties of different kinds of propolis are attributed to the presence of total phenolics and flavonoids as demonstrated in other studies [29, 41, 42]. Hence, the faster bactericidal action regarding S. aureus showed by HA-GP20 and HA-RP20 indicates that this effect should be related to the higher amount of phenolic compounds released by these materials (table 2). Likewise, a fast bactericidal action showed by all materials is in agreement with the results found in the spectrophotometric analysis, which indicates the fast release of green and red propolis (figures 4 and 5). In fact, the presence of such compounds can be confirmed by the FTIR peaks (figure 2), which are characteristic of aromatic rings, phenolic acids and flavonoids, as shown in table 1. Gatea et al [43] produced gold nanoparticles with propolis and observed a peak at 1632 cm⁻¹, attributed to the presence of acid groups. Siripatrawan and Vitchayakitti [44] produced chitosan with propolis and observed peaks at 2930, 1647 and 1257 cm⁻¹, indicating the presence of phenolic compounds incorporated into the material.

Furthermore, characteristic peaks of water and carbonate were observed for the produced HA (table 1). Other authors have already observed the presence of adsorbed water in HA obtained by chemical precipitation because of the air humidity. They also reported the presence of carbonate adsorbed from the atmosphere during the synthesis process, since it is carried out in an open system [33–35, 38, 39, 45]. Additionally, the morphology of the materials with propolis points to an alteration of the physical properties of the materials (figure 1), indicating that the incorporation of propolis into the materials causes a decrease in the degree of particle agglomeration.

The antibacterial activity of the HA with green and red propolis can be confirmed by the values of MIC and MBC against S. aureus (table 3). The materials with red propolis showed greater bactericidal and inhibitory activity when compared to the materials with green propolis. The antibacterial activity of the former is attributed to its chemical composition, especially the presence of flavonoids—mainly isoflavones such as daidzein, formononetin and biochanin A [10, 24, 46]. Thus, the higher inhibitory and bactericidal activities showed by HA with red propolis could be related to the higher amount of total flavonoids present in this release (table 2). Moreover, all materials with green and red propolis showed less inhibitory activity (higher MIC) and higher bactericidal activity (smaller MBC) when compared to the green and red EEP. This characteristic of the compounds present in the release from the specimens may be related to their interaction with the HA, suggesting that the released phenolic and flavonoid compounds have a greater bactericidal effect. Furthermore, materials with red propolis showed more significant differences regarding MIC and MBC when compared to the red EEP (table 3). Additionally, the absorption profiles for red EEP and the releases for HA with red propolis (figures 3 and 5) could help explain those differences. The peak at 333 nm could be related to the chemical nature of the extract, since red propolis has a high amount of polar substances in its composition [24], suggesting that some of these substances could be retained in the HA, which is hydrophilic [47], while other substances would be released in the buffer solution because of their affinity with water, which is a consequence of the formation of hydrogen bonds with phenolics and flavonoids present in the propolis. In fact, with the use of ethanol/water it is possible to 'dissolve' the flavonoids present in water through intermolecular interactions, mainly hydrogen bonds [48].

Moreover, the typical UV–visible absorption spectra of flavonoids must have two absorption bands with maximum peaks in the ranges from 240–285 nm and 300–550 nm [41], suggesting the presence of significant amounts of flavonoids released in the TSB buffer for HA with red propolis. On the other hand, the absorption profile observed for HA with green propolis suggests the presence of the same substances (not necessarily with the same amounts) for the extract as well as for the release in TSB buffer (figures 3 and 4). Previous works have already demonstrated that this absorption band is attributed to the presence of phenolic and flavonoid compounds [49–52].

Phenolic compounds are considered to be acid groups, and they can be deprotonated in a slightly alkaline medium, generating phenoxide ions and increasing their solubility in an aqueous medium [53], suggesting that the high release of total phenolics from HA specimens (table 2) could also be related to the pH of the TSB buffer. On the other hand, the lower amount of total flavonoids released in the TSB buffer observed in this work when compared to the EEP, could be related to the structure of these compounds, such as the nature and size of the substitutes bonded to the aromatic ring, which would determine its affinity with HA or water.

The production of biological devices with antimicrobial characteristics, and the association of propolis with biomaterials, has been the subject of several pieces of work. In general, the most used materials are biopolymers [54–57]. Hydroxyapatite has also been associated with other antimicrobial substances, such as silver, copper, nickel, cobalt and zinc nanoparticles [58–60], or synthetic drugs such as chlorhexidine [61]. However, there is only one report in the literature regarding hydroxyapatite and propolis [23], and in that study, despite the total amount of phenolics and flavonoids not being measured, the presence of specific phenolics and flavonoids extracted in the methanol was observed. Regarding the release of propolis in the TSB buffer, since no quantification of phenolics and flavonoids is made in [23], a comparison with the other data found in the literature can be made. Machado et al [62] evaluated red, green and brown EEP from different regions of Brazil. They found values ranging from 110.92–300.36 mg GAE g^–1 DE for total phenolics and from 24.4–57.6 mg QE g^–1 DE for total flavonoids, with the best results observed for red propolis. Brazilian origin green and brown EEP, produced by Bittencourt et al [63], had total amounts of phenolics of 185.52 and 48.24 mg GAE g^–1 DE, respectively. The EEP from the Brazilian state of Paraná produced by Schmidt et al [64], with different extraction conditions, had values of total phenolics ranging from 147.5–231 mg GAE g^–1 DE, and total flavonoids ranging from 1.9–11.7 mg QE g^–1 DE. There is significant variation in the chemical composition of propolis produced in different regions of Brazil; thirteen types have already been characterized as a function of their locations and the plant species present in the area [13, 22]. Therefore, this context must be taken into account when comparing the results obtained in different works. Variations in the amounts of total phenolics and flavonoids found by the authors, when compared to the values reported in the present work, must be related to the phytogeographic conditions of the specimens.

Furthermore, the greater antibacterial activity of red EEP against S. aureus, when compared to green EEP, has already been reported in previous works. Machado et al [62] investigated the antibacterial activity regarding the S. aureus of EEP from different regions of Brazil. They verified that the red EEP had greater inhibitory and bactericidal activities, and this result was attributed to the greater amount of total phenolics and flavonoids present in the specimen. Grenho et al [23] evaluated the antibacterial activity against the S. aureus of nanohydroxyapatite containing green and red propolis. The authors found that for the same concentrations the material with red propolis was more effective concerning bacterial growth. The greater inhibitory activity of the red compared to the green propolis was attributed to the difference in the chemical composition of the extracts—especially the presence of cinnamic acid and chrysin in the red propolis.

The antibacterial activity of the hydroalcoholic extracts of propolis regarding S. aureus has been reported by other authors. Campos et al [65] reported values for MIC and MBC of 550 and 1500 μg ml⁻¹ for Brazilian EEP, while Schimidt et al [66] reported average values of 650 and 1015 μg ml⁻¹ for MIC and MCB, respectively. Alencar et al [24], using red propolis extracts, obtained values of 50 and 200 μg ml⁻¹ for MIC and CBM, respectively. Grenho et al [23] produced hydroxyapatite pellets containing propolis, however the values for CIM and CBM against S. aureus were not determined. The mechanism concerning the antibacterial activity of propolis is highly complex, and it is attributed to the presence of phenolic compounds and to the synergistic effect of these compounds [42]. Thus, the antibacterial action of the chemical compounds present in propolis extracts is linked to both the functional and structural aspects of bacteria, including damage to the cytoplasmic membrane, the inhibition of nucleic acids, inhibition of the energetic metabolism and inhibition of biofilm formation. In addition, such compounds can also increase membrane permeability and inhibit bacterial motility, provoking lysis and eventual cell death [20–22].

However, the profiles of the cell viability curves for the materials HA-GP8 and HA-RP8 were similar (figure 7), indicating a possible cytotoxic effect considering the total release (100%) of green and red propolis from these materials. Moreover, the values found for IC₅₀ (table 4) suggest that HA with red propolis has a higher cytotoxic activity. Grenho et al [23] used green and red EEP solutions at lower concentrations than those used in this work, without any trace of cytotoxic activity of these materials against rat fibroblasts. As mentioned above, these authors only reported the inhibitory effect against S. aureus, after 24 h, and not bactericidal activity. So, it is possible to compare the IC₅₀ values with those related to the cytotoxic activity of the propolis extracts. Recent works utilized EEP in order to evaluate the cytotoxic activity in terms of sound cell lines [67, 68] as well as tumor cells [62, 68, 69]. The higher cytotoxic effect of red propolis extracts, compared to the green, is also reported in other works [56, 68, 70]. Lopez et al [67] evaluated the cytotoxic activity of red EEP from different states of the northeast region of Brazil in terms of sound cells, and found values of IC₅₀ between 65 and 85 μg ml⁻¹. Da Silva Frozza et al [69] reported values of IC₅₀ between 81.4 and 85.77 μg ml⁻¹ for red EEP from Sergipe, Brazil regarding HeLa tumor cells, while Cabral et al [71] found IC₅₀ values of 60 μg ml⁻¹ for EEP from the northeast of Brazil for the same tumor cell. Other studies indicate the cytotoxic activity of green propolis [62, 70]; however, the IC₅₀ value was not calculated in those works. Hence, it is possible to compare the IC₅₀ values for green propolis found in the present work to those reported for other EEPs of Brazilian origin. Machado et al [68] and Meneghelli et al [72] evaluated the cytotoxic activity of brown EEP from the southern region of Brazil in terms of sound cells and reported IC₅₀ values of 461 and 364 μg ml⁻¹, respectively, which are close to those reported in the present work.

The higher cytotoxic activity showed by HA with red propolis could well be related to the higher amount of flavonoids released by the material. Some studies indicate that isoflavones such as formononetin may be related to the cytotoxic activity showed by the red EEP [67, 69]. Similarly, the cytotoxic activity of HA with green propolis could be related to its chemical composition, with the presence of phenolic acids such as artepillin C and p-coumaric acid [51, 62].

It is also possible to compare the values for IC₅₀ to the values reported for MIC and MBC for specimens HA-GP8 and HA-RP8 (table 4). For both materials, the MIC value was below the values found for IC₅₀, indicating that the concentration needed to inhibit bacterial growth (S. aureus) does not have a cytotoxic effect regarding the cell line used in this work. However, the concentration of the extract needed to cause bacterial death (MBC) is higher than IC₅₀, which indicates the possible cytotoxic effect of the materials at this concentration.

The results obtained in this work confirm that hydroxyapatite powders with dry extracts of green and red propolis show inhibitory and bactericidal activity against S. aureus, with a rapid bactericidal effect associated with the presence of natural antimicrobial agents such as phenolic and flavonoid compounds. In addition, the results show that hydroxyapatite with red propolis has a greater antimicrobial potential due to the higher amount of flavonoids in this extract. Despite the studied biomaterials showing some cytotoxic characteristics regarding the total release of propolis from the specimens, the results indicate that the hydroxyapatite containing green and red propolis may be used in biomedical applications where S. aureus is a concern, inhibiting its growth. Nevertheless, further research is needed to establish the extent of cytotoxicity and biocompatibility allowed in these biomaterials when using them as medical devices with a high bactericidal potential.

The authors would like to thank the company Natucentro® for supplying the raw propolis used in this work.

There are no conflicts of interest.