Antibacterial, antibiofilm, and cytotoxic activities and chemical compositions of Peruvian propolis in an in vitro oral biofilm

Background: Natural products with antibacterial potential have begun to be tested on biofilm models, bringing us closer to understanding the response generated by the complex microbial ecosystems of the oral cavity. The objective of this study was to evaluate the antibacterial, antibiofilm, and cytotoxic activities and chemical compositions of Peruvian propolis in an in vitro biofilm of Streptococcus gordonii and Fusobacterium nucleatum. Methods: The experimental work involved a consecutive, in vitro, longitudinal, and double-blinded study design. Propolis samples were collected from 13 different regions of the Peruvian Andes. The disk diffusion method was used for the antimicrobial susceptibility test. The cytotoxic effect of propolis on human gingival fibroblasts was determined by cell viability method using the MTT (3-[4,5-dimethylthiazol-2-yl]-2,5 diphenyl tetrazolium bromide) assay, and the effect of propolis on the biofilm was evaluated by confocal microscopy and polymerase chain reaction (PCR). Results: The 0.78 mg/mL and 1.563 mg/mL concentrations of the methanolic fraction of the chloroform residue of Oxapampa propolis showed effects on biofilm thickness and the copy numbers of the srtA gene of S. gordonii and the radD gene of F. nucleatum at 48 and 120 hours, and chromatography (UV, λ 280 nm) identified rhamnocitrin, isorhamnetin, apigenin, kaempferol, diosmetin, acacetin, glycerol, and chrysoeriol. Conclusions: Of the 13 propolis evaluated, it was found that only the methanolic fraction of Oxapampa propolis showed antibacterial and antibiofilm effects without causing damage to human gingival fibroblasts. Likewise, when evaluating the chemical composition of this fraction, eight flavonoids were identified.


Introduction
Fusobacterium nucleatum (Choi et al., 2020;Geng et al., 2018) and Streptococcus gordonii (Geng et al., 2018) are some of the most important bacteria in periodontal and peri-implant disease, especially in patients who are smokers (Lazar et al., 2015). The scientific community has made considerable efforts to discover new antimicrobial agents that can disrupt the complex structures of subgingival biofilms. One of the main sources of new molecules with antimicrobial activity is products of natural origin (Arbia et al., 2017;Lee et al., 2017;Slobodníková et al., 2016). The search for new antimicrobial agents has gained momentum, particularly due to increased rates of antibiotic resistance to periodontal pathogens (Kouidhi et al., 2015;Rams et al., 2014;Van Winkelhoff et al., 2005). Socransky et al. (1988) found that the subgingival microbiota can be classified into different complexes based on the association of different microbial species with healthy or diseased sites. If left intact, gingival biofilms create conditions that can favour colonization by orange-complex species, namely, Fusobacterium, Campylobacter, Prevotella, Parvimonas, and some Streptococcus species (for example, S. constellatus). The orange complex is associated with the transition from health to disease due to a change in environmental conditions that favours colonization by Porphyromonas gingivalis, Tannerella forsythia, and Treponema denticola, which are red-complex species that are considered the main pathogens in periodontal disease (Socransky et al., 1988).
Propolis is a resinous mixture obtained by bees from tree buds, sap exudates or other plant sources and then processed in the hive as a sealant for small holes (Bueno- Silva et al., 2017;Santos et al., 2019). Propolis has been considered a promising source of bioactive molecules with pharmacological properties, including antimicrobial (Alencar et al., 2007;Bueno-Silva et al., 2013Dantas Silva et al., 2017), anti-inflammatory (Markiewicz-Żukowska et al., 2012;Trusheva et al., 2004), antioxidant, and antiparasitic activities (Bueno- Silva et al., 2015). Propolis can have different geographical origins (Freires et al., 2016;Lotti et al., 2010;Piccinelli et al., 2011), and although they may share similar physical characteristics, the chemical compositions of propolis isolates from different sites cannot be the same because the composition depends on the place, time, method of collection, and plants of origin (Becerra et al., 2019). The chemical composition of propolis is closely related to the resins and balsams of the plant sources used to produce it. More than 300 chemical components of propolis have been identified (Barbosa Bezerra et al., 2017;Bryan et al., 2016), and the main groups are waxes, polyphenols (phenolic acids, flavonoids), and terpenoids (Bonamigo et al., 2017;De Freitas et al., 2018;El-Guendouz et al., 2018;Hochheim et al., 2020;Popova et al., 2015;Sve cnjak et al., 2020;Touzani et al., 2019;Zhang et al., 2017). Some research studies have demonstrated an effect of propolis on oral bacteria biofilms. Veloz et al. (2019a) compared some biological properties of individual compounds and mixtures of these compounds from Chilean propolis, such as their antimicrobial and antibiofilm activities against S. mutans cultures. Their results suggest that the polyphenols pinocembrin and apigenin exhibit antimicrobial activity against S. mutans at low concentrations. Martins et al. (2019) evaluated the antibacterial activity, antibiotic effect, and cytotoxic potential of mouthwashes containing Brazilian red propolis in vitro. The results showed that red propolis mouthwashes decreased viable cells to 44% after 1 minute of contact with fibroblasts. Thus, red propolis extract was concluded to exhibit antibacterial activity against Streptococcus spp. and Lactobacillus casei and had cytotoxic and antibiofilm effects similar to those of chlorhexidine.

Study design
The experimental work involved a consecutive, in vitro, longitudinal, and double-blinded study design. The decision criteria method was used for the present investigation (Andersen et al., 2019;Bentz et al., 2013). Based on the results

REVISED Amendments from Version 1
The new version includes the participation of a co-author: Jaeson Calla, who was in charge of designing the protocols for the measurement of oral biofilms by confocal microscopy, as well as image capture and interpretation.
Any further responses from the reviewers can be found at the end of the article obtained in each experiment, it was decided to continue the trials only with those propolis that showed antibacterial activity, antibiofilm and were not cytotoxic to gingival fibroblasts.

Propolis sampling
The propolis samples were collected from fixed apiaries and wild flora around beehives. The collection process was performed by scraping the boxes with a bevelled plastic spatula and placing them in airtight glass containers covered with dark plastic bags until they were delivered to the Biopolymers and Metallopharmaceuticals Research Laboratory (LIBIPMET) of the National Engineering University, Lima. All propolis were masked by a person not involved in the study and labelled with letters until after data processing was completed. They were then stored refrigerated at 4°C until processing.
Samples were collected in 13 Andean regions of Peru, considered as static apiaries (they are not mobilized by seasonal changes) ( Table 1).

Preparation of the crude extracts of propolis
The 13 samples were removed from refrigeration, and after waiting two hours for them to reach room temperature (RT), they were thinly cut and then macerated in absolute ethanol using a volume of 100 mL per 10 grams of propolis. The samples were then incubated at RT for 24 hours. The macerate was filtered using a glass funnel with a 20-cm diameter and sterile cotton, and the sample was collected in a glass dish that was transferred to an extraction hood for evaporation of the ethanol (until a paste was formed) (Markiewicz-Żukowska et al., 2012).

Preparation of the liquid-liquid partitions of propolis
The crude extract (CE) was solubilized with a mixture of chloroform (CHCl 3 ):methanol (4:1 v:v). Approximately 50 mL of the mixture was needed to completely solubilize each sample. Next, 30 mL of ultrapure water was added to the assembled separation column, followed by solubilized propolis and approximately 250 mL of CHCl 3 . The separation column was left for six hours until both phases were translucent. We added 250 mL of CHCl 3 and repeated the previous procedure. The chloroform phases were combined in the same beaker. The aqueous phase was recovered in a 100-mL beaker. Both collected phases were taken to the extraction hood for chloroform evaporation (Eldeniz et al., 2007;Santos et al., 2002;Verma et al., 2018).
In the same assembled separation column as above, 300 mL of 1-butanol (BuOH) was added, and then the aqueous phase obtained above was poured into the column. The formation of two phases (lower aqueous and upper butanolic) was observed. They were allowed to stand for approximately three hours until both phases were completely translucent. Then, both phases were separated and collected in beakers of 500 mL (butanolic phase) and 100 mL (aqueous phase). They were immediately transferred to the extraction hood for the evaporation of the BuOH. Once a paste had formed, 50 mL of water was added. Each sample was frozen at À70°C for 24 hours and lyophilized twice. The aqueous phase was frozen at À70°C for 24 hours (in an ultralow-temperature freezer), and then the sample was lyophilized in a 2.5-L Obtaining the propolis fractions from the chloroform residue (RCHCl 3 ) After the propolis partitions were obtained, RCHCl 3 was fractionated using Sephadex LH-20 as the stationary phase and the solvents petroleum ether (PE), dichloromethane (DCM), and methanol (MeOH) as the mobile phase. RCHCl 3 propolis was first solubilized with PE. Open-column chromatography was performed with Sephadex LH-20 as the stationary phase (100 g) in PE. The solubilized sample was added to the column and eluted with PE (450 mL), followed by DCM (350 mL), and finally with MeOH (500 mL), thus yielding the PE fraction, dichloromethane fraction, and methanolic fraction. The fractions obtained were dried in an extraction hood for 24 hours (Mancarz et al., 2019;Yang et al., 2011).
Finally, the three dry fractions were transferred to the Bacteriology Laboratory of the Universidad Peruana Cayetano Heredia for antimicrobial sensitivity tests.
The challenge was performed by embedding a swab with the previously prepared inoculum, spreading the inoculum on the surface of the agar four times, allowing it to stand for five minutes, and then placing Whatman No. 3 filter paper discs (6-mm diameter) impregnated with 10 μL of each of the propolis extracts 25 mg/mL on top but away from one another. This procedure was repeated four times to yield plates for each of the extracts (CE, partitions, and fractions). A 0.12% chlorhexidine solution was used as a positive control, and dimethyl sulfoxide (DMSO) and Milli-Q water (1:1) were used as negative controls. The evaluation of antibacterial activity was carried out by means of inhibition zones in millimeters using a Truper calibrator (Millones- Gómez et al., 2020).

First decision criteria
After evaluating the CEs of the 13 propolis samples, we decided that partitions would only be performed on the extracts that exhibited zones of inhibition on both strains studied, and then we could perform cytotoxic evaluations against human gingival fibroblasts.
Evaluation of the cytotoxic effect of propolis on cell line HGF-1 (ATCC ® CRL-2014) Cytotoxicity was evaluated in the dentistry laboratory of the Center for Research and Development in Health Sciences of the Universidad Autónoma de Nuevo León, Mexico. The cytotoxic effect of propolis on human gingival fibroblast cell line (HGF-1 ATCC ® CRL-2014, American Type Culture Collection; USA) was determined by adapting the cell viability method for microplates using the MTT assay as described below (Devequi-Nunes et al., 2018;Poggio et al., 2017). Cytotoxicity responses were classified as severe (30%), moderate (30-60%), mild (60-90%), or noncytotoxic (> 90%) (Poggio et al., 2017).

Second decision criteria
After evaluating cytotoxicity, we decided that only residues that showed cell viability greater than 90% in the HGF-1 cell line (ATCC ® CRL-2014) would be fractionated and evaluated for antimicrobial activity.

Minimum inhibitory concentration
The method used was microdilution in broth using 96-well microtiter plates. Chlorhexidine at 0.12% was the positive control, and diluted DMSO solution with Milli-Q water (1:1) was the negative control. The entire procedure was repeated four times for each propolis sample with the largest inhibition zones (Millones- Gómez et al., 2020). The minimum inhibitory concentration of propolis was deemed to be the concentration in the well where no development was observed (turbidity) using a spectrophotometer.

Third decision criteria
After determining the minimum inhibitory concentration of fractions with zones of inhibition on both strains studied, we evaluated the biofilm considering those concentrations with antibacterial and nontoxic effects on the HGF-1 cell line (ATCC ® CRL-2014).
Formation of a biofilm of S. gordonii and F. nucleatum Each strain was inoculated from a colony into 15 mL of Trypticase soy broth and incubated at 37°C in anaerobiosis until reaching the exponential growth phase of S. gordonii (four hours and 30 minutes) and F. nucleatum (eight hours), which was shown by an optical density of 550 nm and 0.125 nm with 150 Â 10 6 cells/mL (Sánchez et al., 2011).
Subsequently, the surfaces of Thermo Scientific Nunc Lab-Tek Chamber Slides were prepared with 30 μL of poly-Llysine incubated at RT for 30 minutes and then allowed to dry at 37°C for 24 hours under sterile conditions. Then, 300 μL of artificial saliva was added for incubation at 4°C for 16 hours. The sheets were washed with 300 μl of phosphatebuffered saline (PBS 1Â) twice. A total of 280 μL of brain heart infusion (BHI) + 2.5% sucrose was added. Finally, 10 μL of S. gordonii + 10 μL of F. nucleatum was inoculated, and then each slide was incubated for 24 hours at 37°C under anaerobic conditions (Sánchez et al., 2011). S. gordonii (Ayala et al., 2016;Cayo et al., 2016) and F. nucleatum (Jacinto, 2018; Sánchez, 2018) were identified using specific primers:

Evaluation of the effect of propolis on the biofilm
To evaluate the effect of propolis on the biofilm, 10 μL of S. gordonii + 10 μL of F. nucleatum was inoculated, and then each plate was incubated for 24 hours at 37°C under anaerobic conditions. The propolis fraction with the greatest antibacterial effect and lowest toxicity against HGF-1 cells was used.
The first dose of the propolis extract was administered 24 hours after its formation, where the supernatant was carefully removed from each well. Two washes were performed without exerting much force with 300 μL of PBS (1Â). Then, 300 μL of propolis was added, followed by incubation for 1 minute at RT. The plate was removed and washed up to the surface of each well with 300 μL of PBS (1Â) twice. Next, 300 μL of sterile BHI + SAC 2.5% culture medium was added, and the plate was incubated at 37°C for 24 hours under anaerobic conditions (Sánchez et al., 2011).
The following groups were formed: -Group 1 (Sg + Fn): 300 μL of propolis was pipetted in each well every 24 hours until evaluation at two days and five days.
-Group 2 (negative control): 300 μL DMSO + Milli-Q water (1:1) was pipetted in each well every 24 hours until evaluation at two days and five days.
-Group 3 (positive control): 300 μL of 0.12% chlorhexidine was pipetted in each well every 24 hours until evaluation at two days and five days.
-Three repetitions of each group were performed.
The final exposure of the slide occurred when the biofilms reached 120 hours of maturation, where the Nunc Lab-Tek Chamber Slide™ system fixation and staining procedure was performed with fluorochromes for confocal microscopy.

Fixation and staining of slides
The removable chamber was taken off the surface of the Lab-Tek slide, placed horizontally to fix each surface of each well with 30 μL of 4% paraformaldehyde (4%) that was added over the entire surface of the biofilm, and incubated RT for 20 minutes. Then, three gentle washes were performed with 1Â PBS. Five microlitres of the L13152 LIVE/DEAD ® BacLight bacterial viability kit was added at a concentration ratio of 1:1 with fluorochromes to the surface of the biofilm. The cells were stained with 5 μL of VECTASHIELD ® HardSet™ Antifade Mounting Medium with DAPI (4 0 ,6diamidino-2-phenylindole).
H-1500-10.This last step was performed to differentiate the nucleic acids (DNA) in the bacteria and conserve the samples (Sánchez et al., 2011).

Chemical composition of propolis
The major chemical components of the propolis fraction with effects on the biofilm were analysed through proton-and C-13 nuclear magnetic resonance ( 1 H NMR and 13 C NMR) at the Analytical Center of the Institute of Chemistry of the University of São Paulo, Brazil. A 25-mg sample was solubilized in MERCK brand deuterated methanol, and NMR spectra were recorded in a 500-MHz Bruker spectrometer operating at 500 MHz to obtain the hydrogen spectra and operating at 125 MHz for the C-13 spectra (Figures 4 and 5).
Then, a 2-mg sample of FMeOH from RCHCl 3 of Oxapampa propolis was solubilized in methanol (CH3OH), filtered through a filter with a pore size of 0.45 μm, and transferred to a 2-mL vial. The sample was analysed by liquid

Results
The objective of this study was to evaluate the antibacterial effect of the CEs of 13 propolis isolates from the Peruvian Andes (Table 2). The microscopy images and raw data are available in Underlying data (Millones-Gómez, 2021). The  (1), isorhamnetin (2), apigenin (3), kaempferol (4), diosmetin (5), acacetin (6), glycitein (7), and chrysoeriol (8). . When performing post hoc comparisons of propolis with chlorhexidine, the difference in their average ranges was not statistically significant; thus, they are listed in Table 2 with the same letter.
When evaluating the CE partitions with the greatest antibacterial effect (Table 3), the Kruskal-Wallis H test indicated a difference between some of the experimental propolis partitions and/or controls in the growth inhibition of both S. gordonii (p < 0.001) and F. nucleatum (p < 0.001). Likewise, the Z test revealed that, together with chlorhexidine (S. gordonii: 14.45 AE 0.30 and F. nucleatum: 14.40 AE 0.18 mm), the partition that exceeded the average inhibition in both types of bacteria was the chloroform residue from Oxapampa (9.40 AE 0.28 and 9.95 AE 0.19 mm). The chloroform residues from Saylla (10.2 AE 0.14 mm) and Yura (9.27 AE 0.05 mm) showed above-average S. gordonii inhibition. The butanolic residues from Saylla (9.98 AE 0.21 mm) and Oxapampa (9.45 AE 0.19 mm) showed above-average inhibition of F. nucleatum. When performing post hoc comparisons of the best propolis with chlorhexidine, the difference in their average ranges was not statistically significant; thus, they are written in Table 3 with the same letter.   Table 5. Antibacterial effects of selected propolis fractions against S. gordonii and F. nucleatum.

Zones of inhibition (mm)
S When evaluating the cytotoxicity of the propolis partitions on human gingival fibroblast cell line (HGF-1 ATCC ® CRL-2014, American Type Culture Collection; USA) considering 10% Triton X-100 (negative control) and the medium (positive control) (Table 4), we generally observed that increasing the dilution decreased the cell viability of the partitions of the propolis evaluated. The chloroform partition of Oxapampa propolis showed its highest percentage of cell viability (92.64%) at the 0.78 mg/mL dilution.
When evaluating the antibacterial effects of the fractions of the chloroform partition of Oxapampa propolis (Table 5) by the Kruskal-Wallis H test, a difference was identified in the inhibition of S. gordonii (p < 0.01) and F. nucleatum (p < 0.01) prepared in the different dilutions. The methanolic fraction of the chloroform residue of Oxapampa propolis (12.55 AE 0.19 and 12.15 AE 0.19 mm) had greater inhibition power against each bacterium than when combined with chlorhexidine (14.55 AE 0.19 and 14.33 AE 0.19 mm), exceeding the average. In addition, post hoc analysis revealed that both fractions did not differ from each other but had greater inhibitory power towards both bacteria.
When evaluating the concentrations of 0.781 and 1.563 mg/mL of the methanolic fraction of the chloroform residue of Oxapampa propolis on biofilm thickness by confocal microscopy, the Kruskal-Wallis H test indicated that there was a difference in the thickness of the biofilm formed with the different treatments, both at 48 hours (p < 0.01) and at 120 hours (p < 0.001). The biofilm formed with 0.12% chlorhexidine (8,100 AE 1,149 and 45,200 AE 5,718 μm) was not thinner than the average, as shown in Table 6, at either 48 hours or 120 hours (p > 0.05). The biofilms with lower-than-average thickness were those formed with 0.78 mg/μL propolis (8.026 AE 1.609 and 6.840 AE 1.675 μm) and 1.56 mg/μL propolis (7.374 AE 1.620 and 9.248 AE 0.679 μm). Post hoc tests indicated that these did not differ from chlorhexidine in the biofilm formed at 48 hours; but at 120 hours, although they did not differ from each other, propolis made with a concentration of 0.78 mg/μL was thinner than that made with chlorhexidine.
When evaluating the effect of the methanolic fraction of the chloroform residue of Oxapampa propolis on the number of bacteria by quantifying the number of copies of the srtA gene of S. gordonii and the radD gene of F. nucleatum in the biofilm by PCR, the Kruskal-Wallis H test indicated that there was a difference in the number of S. gordonii (p < 0.01, in both periods) and in the number of F. nucleatum (p < 0.05 in both periods) under the different treatments. The concentration of propolis of 1.56 mg/mL decreased the amount of S. gordonii both at 48 (6.347 AE 0.410) and at 120 hours (5.157 AE 0.202) and the amount of F. nucleatum at 120 hours (5.747 AE 0.618) compared to the average, as did 0.12% chlorhexidine. However, the post hoc analysis indicated that this concentration showed no difference from chlorhexidine in the control of the development of both bacteria in the times evaluated (Table 7).

Discussion
The present study aimed to evaluate the antibacterial effect of CEs, partitions, and fractions of Peruvian propolis and the effects of fractions on the thickness of in vitro biofilms of S. gordonii and F. nucleatum. Of the 13 CEs of propolis evaluated, only four showed antibacterial effects on both strains. Moreover, the methanolic fraction of Oxapampa propolis showed an effect on the thickness of the biofilm created in vitro. The results of the present investigation showed that of the 13 Peruvian propolis isolates evaluated, four showed antibacterial effects on both bacterial strains. These results differ from those of other investigations that have shown the strong antibacterial power of propolis against all evaluated microorganisms (Becerra et al., 2019;Bueno-Silva et al., 2013;Dantas Silva et al., 2017;Trusheva et al., 2004). Although several studies of Peruvian propolis have shown effects on oral bacteria, all of them focused on other types of oral bacteria. Examples include the La Libertad propolis against Streptococcus mutans (Becerra et al., 2019;Sánchez, 2018); Huacho and Cajabamba propolis against Streptococcus mutans (Ayala et al., 2016;Cayo et al., 2016); and propolis from Arequipa, Huánuco, and Huaraz against Streptococcus mutans and Streptococcus oralis (Jacinto, 2018;Vargas, 2019). This study is the first to evaluate the effects of Peruvian propolis on bacteria linked to the growth of subgingival biofilms.
In the present study, after identifying the CEs of the four propolis with the greatest zones of inhibition on each bacterium evaluated, partitions were performed, and the strongest antibacterial effects were found in the butanolic and chloroform residues of all the propolis studied. Although there are no studies with partitions of Peruvian propolis, the few studies carried out on propolis from other sources indicate that some solvents have greater affinity for certain components with biological effects (Flemming et al., 2016). For example, the amounts of flavonoids and terpenes in different extracts were found to decrease in the following order: chloroform extract > ethanolic extract > butanolic extract (Osés et al., 2020;Palomino et al., 2009).
In Peru, as in many countries, beekeepers prepare by hand the products derived from the hive. Many therapeutic effects have been attributed to propolis (Narimane et al., 2020). However, many of these products could be used without any control, with several of them being toxic for human consumption; therefore, the toxicity must be correctly recognized and the product used reasonably (Sun et al., 2015). In the present study, toxicity to gingival fibroblasts was evaluated by the percentage of cell viability (Poggio et al., 2017). Only the chloroform residue of Oxapampa propolis was found to be nontoxic; the results of the MTT assay, with more than 92% cell viability, suggest good biocompatibility. Although few studies have evaluated the cytotoxicity of propolis on this cell line, they show that these are not very toxic (Pobiega et al., 2019;Santos et al., 2019).
When evaluating the fractions of the chloroform partition, only the methanolic fraction had an inhibitory effect, with a minimum inhibitory concentration of 0.78 mg/mL on both strains evaluated. Although no studies of Peruvian propolis have been reported, studies of foreign propolis have found similar results (Nani et al., 2018;Zeng & Jiang, 2010), demonstrating that methanolic extraction can produce a higher concentration of compounds with biological activity (Flemming et al., 2016).
Most bacteria reside naturally in accumulations called biofilms, which are substantially more tolerant to antimicrobials than free-floating cells (Aydin et al., 2018). A biofilm is a community of bacteria that have adhered to a surface. Once adhered, the cells begin to secrete extracellular polymeric substances composed of extracellular DNA, polysaccharides, and proteins. These extracellular substances trap nutrients and water within the biofilm, allowing cells to mature in nutrient-rich conditions while protecting them from desiccation, host immune defences, and antimicrobial agents (Cabral et al., 2009). In the present study, when evaluating the effect of the methanolic fraction of propolis on biofilm thickness, both concentrations were found to have a greater effect than chlorhexidine at 120 hours. This result may have occurred because one of the factors contributing to the resistance of biofilm-mediated infections may be a delay in the penetration of loaded antimicrobial agents due to the binding of the robust extracellular polymeric substance matrix, which prevents complete access to the entire biofilm (Das Neves et al., 2016;Li et al., 2016). However, unlike chlorhexidine, the propolis fraction is composed of more than one active component, including flavonoids, phenolic acids, terpenes, aromatic acids, and others (Sha et al., 2009). Identifying the exact chemical substances underlying a certain effect, as well as their defined biochemical mechanisms, is difficult (Hall & Mah, 2017;Khatoon et al., 2018).
The PCR results showed that the concentration of 1,563 mg/mL of the methanolic fraction of propolis decreased the copy numbers of the srtA gene of S. gordonii and the radD gene of F. nucleatum at 120 hours. Although no previous studies have evaluated the effect of propolis on biofilms of these species, some research suggests that propolis has a dosedependent effect on the inhibition of genes involved in adherence, showing a potential effect beyond bacterial growth inhibition (Veloz et al., 2019a(Veloz et al., , 2019b. The results of component identification by LC/MS of the methanolic fraction of Oxapampa propolis showed the presence of rhamnocitrin, apigenin, kaempferol, isorhamnetin, diosmetin, acacetin, glycitein, and chrysoeriol. The first three compounds have already been described to have antibiofilm effects in previous studies (Palomino et al., 2009). Previous studies have highlighted apigenin as a flavonoid with considerable antibiofilm potency (Nani et al., 2018;Palomino et al., 2009) due to its ability to increase the permeability of the bacterial membrane, resulting in cellular apoptosis (Zeng et al., 2010). Other studies have highlighted the potential of flavonoids to inhibit the formation of the extracellular polymeric matrix, exposing cells that form biofilms and increasing their susceptibility to antimicrobial therapies (Aydin et al., 2018).
The results of this study show a significant antibiofilm effect of Oxapampa propolis without causing damage to oral human cells (gingival fibroblasts), rendering it a promising option for evaluation as a new molecule for the development of new pharmacological agents that may be valuable in the dentistry field.

Luis Maita Castañeda
Faculty of Dentistry, National University of San Marcos, Lima, Peru better understand the bioactive compounds of these extracts (as indicated in the "Discussion" section), this work was logically and rigorously completed and offers valuable insight into a possible use of Oxapampa propolis as a cost-effective and readily available dental therapy presenting sufficient merit for publication.

Is the work clearly and accurately presented and does it cite the current literature? Yes
Is the study design appropriate and is the work technically sound? Yes

If applicable, is the statistical analysis and its interpretation appropriate? Yes
Are all the source data underlying the results available to ensure full reproducibility? Yes Are the conclusions drawn adequately supported by the results? Yes