Propolis Nanoemulsion Enhance the Antimicrobial Effect on Multidrug-Resistant Strains of Pseudomonas aeruginosa in Combination with Ciprofloxacin

Abstract

Background: Antimicrobial alternatives, including nanoparticles, are needed to combat bacteria as antibiotic resistance threatens global health. The aim of this research was to determine the antibacterial efficacy of a synthesized propolis nanoamulsion (PN) as a natural reducing agent and its synergistic effect in combination with ciprofloxacin (CP) against multidrug-resistant (MDR) P. aeruginosa biofilm. Methods: Scanning electron microscopy (SEM), dynamic light scattering (DLS) and zeta potential (ZP) were used to measure the mean particle size and nanoparticle size distribution. Iranian propolis (IP) was analyzed using gas chromatography-mass spectrometry (GC-MS). Antibacterial activity against MDR strains of P. aeruginosa was also evaluated. Furthermore, the synergistic in vitro effect of IP and PN combined with CP and the antiadhesion activity were evaluated using the checkerboard method and biofilm metabolic activity, respectively. Results: SEM, DLS and ZP revealed a size of 161.4 nm and a negative surface charge value of −37.4 mV. GC-MS analysis revealed the existence of four flavonoid compounds. This study showed that PN was more effective against MDR P. aeruginosa than IP with a minimum inhibitory concentration (MIC) value of 3.2-15 mg/mL. The combination of PN and CP had a synergistic effect, reducing the dosage of each compound and the time required to destroy the bacteria compared to using them separately. Furthermore, IP and PN suppressed the adhesion of MDR P. aeruginosa biofilm. Conclusions: According to the results, propolis nanoamulsion could enhance the antimicrobial effect, reduce the viability of bacteria and affect the biofilm formation of MDR P. aeruginosa. In combination with CP, it led to a stronger effect of the drug on the organism.

Keywords

antibacterial biofilms nanoamulsion multi-drug resistant propolis

Introduction

Pseudomonas aeruginosa (P. aeruginosa) is a gram-negative opportunistic pathogen and the most common bacterium causing hospital-acquired (nosocomial) infections, particularly ventilator-associated pneumonia (VAP), cystic fibrosis, and immunodeficiency disorders.¹ It is intrinsically resistant to various antimicrobial agents, including aminoglycosides, quinolones and β-lactams.² P. aeruginosa utilized multiple mechanisms to acquire antibiotic drug resistance, primarily horizontal gene transfer and biofilm formation.³ A wide variety of intrinsic and acquired resistance mechanisms exist within P. aeruginosa populations, leading to the emergence of new strains with multidrug resistance patterns (MDR). Currently, in addition to MDR strains, the development and spread of extensive and pandrug-resistant phenotypes (XDR and PDR) represent a major challenge for patient treatment, as it leads to increased morbidity, mortality and longer hospital stays.⁴ In 2018, a new term for difficult-to-treat resistance (DTR) was proposed. P. aeruginosa with DTR is characterized by resistance to all β-lactamases, carbapenems and quinolones.⁴ Combination therapy may represent a therapeutic option for the treatment of these highly resistant bacteria, thereby reducing the mortality rate of patients with severe infections.⁵ On the other hand, excessive use of antibiotics can lead to antimicrobial resistance, which often renders the antibiotic ineffective against microorganisms. Studies have shown that natural products serve as effective therapeutic agents against pathogenic bacteria.⁶

Propolis is a natural substance collected by honeybees from various plants and mixed with their salivary secretions to seal cracks in their hives. It also serves as a disinfectant to prevent infectious diseases from developing and spreading within the beehive.⁷ Propolis exhibits antibacterial activity either by directly affecting microorganisms or by stimulating the immune system.⁸ To date, over 300 chemical components, including phenols, flavonoids, and aromatic compounds, have been identified in propolis.⁹ Flavonoids such as galangin, pinocembrin, apigenin, kaempferol, pinostrobin, chrysin, and quercetin, as well as phenolic acids like caffeic, ferulic, cinnamic, and coumaric acids, have been identified in propolis samples from various geographical regions.¹⁰ Many flavonoids inhibit pathogenic bacteria through two main mechanisms: acting on the cell membrane and inhibiting DNA gyrase in gram-negative bacteria.¹¹ Research has shown that the geographical origin of bees and the time of collection influence the composition of propolis.¹² Recently, propolis has gained significant attention for its wide range of biological functions, particularly its antimicrobial activity, which can help lower the required dosages of antimicrobial drugs.¹³

Recent advances in nanotechnology have enabled the production of various materials at the nanoscale. Nanoparticles (NPs) are a broad class of materials with at least one dimension under 100 nm, utilized for diverse applications.¹⁴ Currently, therapeutic NPs dominate the NP formulations used in clinical settings. They enhance the accumulation of pharmacologically active agents in diseased tissues, thereby increasing therapeutic efficacy and reducing side effects by minimizing exposure to healthy tissues. NPs can readily penetrate biofilms and disrupt bacterial cell walls and membranes, significantly impacting resistant bacteria.¹⁵ Due to the toxicity of many engineered nanoparticles, there is a need for safer alternatives, such as plant-based nanoparticles. The synthesis of nanoparticles from plant extracts is a fast, safe, cost-effective, and environmentally friendly method that avoids producing toxic derivatives.¹⁶ Multiple studies have demonstrated varying degrees of reduction in P. aeruginosa biofilm formation by propolis and its nanoparticles.^17–21 This study aimed to prepare and characterize propolis nanoemulsion (PN) using ethanol extract of Iranian propolis (IP), a natural compound rich in phenolic acids and flavonoids, against MDR P. aeruginosa. Additionally, we sought to determine the antibacterial efficacy of PN and its synergistic effect in combination with ciprofloxacin (CP) against MDR strains of P. aeruginosa biofilm.

Methods

Propolis Sampling and Extraction

Crude propolis, produced by Apis mellifera bees was collected on May 2020, in north of Iran (polour) latitude 35°50′ 49′′ N and longitude 52°2′ 55′′ E, altitude of 2300 m. There are 139 plant species in this area. The vegetation of this area mainly consists of Compositae, Lamiaceae, Rosaceae, Leguminosae, Cruciferae, Umbelliferae, Liliaceae, Scrophulariaceae, Caryophyllaceae, and Plumbaginaceae. The sample was kept in polypropylene jars at −4 °C. According to our previous study, ethanolic extract of IP was provided.⁹

Preparation of Propolis Nanoemulsion (PN)

Preparation of nanoemulsion of propolis was carried out using high energy emulsifying method. IP (0.6 g) was mixed in Tween 80 (10%), distilled water, and 80% ethanol (8:10:2) using an ultrasonic homogenizer and heated at 100 rpm for 2 h until emulsion was formed. For the preparation of nanoemulsions, the sample was treated with an ultrasonic homogenizer by using 75 W ultrasonic waveform generator and frequency 30 kHz for 20 min and the final solution was stored at room temperature. Field emission scanning electron microscopy (FE-SEM) (MIRA3, TESCAN-XMU, Czech Republic) analysis and dynamic light scattering using a Zetasizer Nano (DLS, HORIBA) were used to determine the size, distribution of NPs and dispersity index. Moreover, the zeta potential of PN was measured at 25 °C using a SZ-100 Nanoparticle Analyzer.

GC–MS Analysis

IP sample was analyzed by GC–MS equipment (7890B-5977B MSD, Agilent) with gas chromatography-mass spectrometry (GC-MS) column DB −5Ms (30 m × 0.25 mm × 0.25 µm), and the carrier gas was helium at a flow rate of 1 mL/min. One microliter of the sample was injected, and the injector temperature was 250 °C in split-less mode. The oven temperature was initially held at 50 °C for 1 min and augmented at a rate of 8 °C/min to 120 °C, and then ramped at a rate of 6 °C/min to 250 °C, ultimately to 250 °C at 15 min. The solvent delay time was 0 to 3 min, and the total GC–MS running time was 47 min.⁹ Peaks were recognized with NIST Mass Spectral Database (11 Variant).

Antibacterial Assay

P. aeruginosa were collected from Yahyanejad Hospital in Babol. Strains of P. aeruginosa were identified using routine differential diagnoses and biochemical test kits API 20E (BioMerieux, Marcy, France). Antibiotic susceptibility was performed according to the Kirby-Bauer disk diffusion method. Based on the CLSI description, MDR P. aeruginosa was defined when it was found non-susceptible to at least one agent in three or more antimicrobial categories.¹² Finally, the antimicrobial action of IP and PN against 10 strains of MDR P. aeruginosa and P. aeruginosa (PTCC 1430) (as a strain sensitive to antibiotics) was measured by disk diffusion and the broth dilution. The antibiotic CP (according to CLSI) was considered as control. The tests were carried out in triplicate.²²

Disk Diffusion Method

Bacterial isolates were inoculated into Brain Heart Infusion broth (BHI broth, Merck, Darmstadt, Germany) and incubated at 37 °C for 24 h. Suspension of bacteria was compared with 0.5 McFarland, then using sterile swab, the bacteria were spread on MHA medium (Mueller Hinton agar). The filter paper discs (6 mm diameter) were impregnated with 20 of different concentrations of IP (25-100 mg/mL) and PN (1.8-7.5 mg/mL), after drying, the discs were located on the seed plates. The pure solvent (DMSO) and CP disk (5 μg, HiMedia) were also used for control. After incubation, the plates were observed for zone of inhibition.

Minimum Inhibitory Concentration (MIC)

First, 100 μL of MHB was added to all wells of 96-well microplate (200 μL), followed by 100 μL of different concentrations of IP (25-200 mg/mL), PN (0.8-15 mg/mL) and CP (0.07-15 mg/mL). Then, the bacterial suspension (10⁵ CFU mL⁻¹) was added to all wells. The numbers of wells in each plate were allocated to negative control (MHB medium), positive control (MHB medium + bacteria), control for the solvents (DMSO and Tween), and control for extract (extract + media). The minimum inhibitory concentration (MIC) value was described using visual observation, the lowest concentration of the samples that suppressed bacterial growth. For determine the minimum bactericidal concentration (MBC), 5 μL of three previous wells was transferred on MHA, and after overnight incubation, the lowest concentration that revealed no visible bacterial growth (99% inhibition) was regarded as MBC.²³

Time–Kill Assay of IP and PN

Time-kill assay of IP and PN against MDR P. aeruginosa strains was carried to determine the ideal incubation time.²⁴ Briefly, tests were performed in BHI broth using a 96-well microplate (200 μL) containing a bacterial suspension of 5 × 10⁵ CFU/mL. The first and second wells contained samples at sub-MIC, and the third well was regarded as growth control. Next, 10 μL of each dilution was transferred to a BHI agar at 0, 4, 6, 8, 10, 12 and 24 h to measure bacterial growth. The procedure was done in triplicate. A graph of log CFU/mL was plotted versus time.

Checkerboard Method

The Checkerboard testing method was used to evaluate synergism among IP and PN with CP against MDR P. aeruginosa strains. Briefly, twofold serial dilutions of each sample were made in a microplate by diluting IP or PN across the x-axis while individual CP was titrated across the y- axis, at a concentration range of zero MIC to 2 × MIC. Then, added to 100 μL of bacterial suspension (0.5 McFarland), and the MIC was read as the lowest dilution without any turbidity. The fractional inhibitory concentration index (FICI) was determined as FICI = FICA (MIC A combination/MIC A single) + FICB (MIC B combination/MIC B single).²⁵

Assessment of Anti-Biofilm Activity

The anti-biofilm action of IP and PN against MDR P. aeruginosa strains was evaluated by biofilm metabolic activity (the MTT assay) according to our previous study with some modifications.¹² First, 100 μL of BHI with 2% sucrose (w/v) containing 0.5× MIC of IP or PN and 100 μL of the suspension of isolates (5 × 10⁵ CFU/mL) were added to all wells (96-well microplate, 200 μL). The control wells were contained BHI/sucrose 2% w/v (untreated) or solvent (control). After 24 h of incubation at 37 °C, the wells were washed twice with PBS and dried. Then, 100 μL of MTT solution was added to the wells, and the samples were incubated for 3 h at 37 °C. The formazan formed was dissolved in 150 μL of DMSO. Finally, the absorbance of each well was determined at 570 nm using a microtiter plate spectrophotometer (Rayto, RT- 2100C, Chinese). The inhibition percentage was determined as [(OD control- OD sample)/ OD control] × 100. Three independent assays were performed.

Morphological Observation by SEM

MDR P. aeruginosa strains were cultured in the presence of IP and PN 0.5× MIC). After 24 h, the wells were rinsed in PBS three times and fixed into glutaraldehyde (4%) 24 h at room temperature, and then washed with PBS. The biofilms were dehydrated with serial concentrations of 50%, 70%, 90% and 100% of ethanol solutions for 15 min, and dried overnight. The samples were then coated with gold prior to SEM analysis (SNE- 4500M, SEC CO., LTD, Suwon, Korea).¹²

Statistical Analysis

All experiments were performed at least three times. Data were analyzed using GraphPad software v 9 (CA, USA) and shown as mean ± SD. The results were evaluated by one-way analysis of ANOVA variance, followed by Tukey's multiple comparison tests. Values of P < .05 were regarded significant.

Results

Analysis of IP

The results of the GC-MS analysis of the ethanol extract of IP showed the presence of four flavonoid compounds, including tectochrysin, galangin, pinostrobin, and naringenin from three groups: flavones, flavonols, and flavanones (Table 1). Among them, galangin (55.5%) was the main component of IP.

Table 1.

Content of Flavonoids Found in Ethanolic Extract of Iranian Propolis Using GC/MS Analysis.

Peak	Compounds	*R.T. min	Molecular Formula	% of total
1	Pinostrobin chalcone	32.066	C₁₆H₁₄O₄	15.099
2	Galangin	33.256	C₁₅H₁₀O₅	55.509
3	Tectochrysin	34.973	C₁₆H₁₂O₄	13.216
4	Naringenin	36.552	C₁₅H₁₂O₅	16.177

*RT: Retention time (minutes).

Nanostructure Preparation

Figure 1 illustrates the process of PN preparation. The size of NPs was determined using SEM and dynamic light scattering (DLS). FE-SEM images demonstrate that the PN has a nearly spherical morphology (Figure 2A). Figure 2B presents the DLS data of the PN at room temperature with an average size of 161.4 nm and only one peak (100% intensity). Zeta potential (ZP) was calculated from electrophoretic mobility at 25 °C, demonstrating a negative value (−37.4 mV), which was adequately high to prevent PN aggregation (Figure 2C).

Figure 1.

Schematic representation of PN preparation.

Figure 2.

(A) FE-SEM image of the PN; (B) size distributions of the PN; (C) the zeta potential of PNP.

Antibacterial Assay

In this study, a total of 10 isolates of MDR P. aeruginosa were considered, all of which were resistant to ciprofloxacin, levofloxacin, gentamicin, amikacin, meropenem, and imipenem, and two isolates were resistant to piperacillin/tazobactam (TZP). P. aeruginosa (PTCC 1430) was sensitive to all antibiotics. Table 2 depicts the disk diffusion assay based on inhibition zone size (mm). IP and PN inhibited P. aeruginosa (PTCC 1430) at lower concentrations than the MDR strains, and inhibition zone increased with increasing concentrations to 17 ± 2 mm and 15 ± 2 mm respectively.

Table 2.

Zone of Inhibition (mm) of Antibiotics, IP and PN Against MDR P. aeroginosa by Disc Diffusion Method.

Strains	Sample^a	Antibiotics^b							IP(mg/mL)		PN(mg/mL)
		CP	LEV	GM	AMK	IMI	MEM	TZP	100	200	3.2	7.5
P3	SPC	R	R	R	R	R	-	- ^c	- ^d	11 ± 0.5	-	11 ± 0.5
P5	UC	R	R	R	R	R	-	-	-	10 ± 0.5	-	11 ± 1
P7	UC	R	R	R	R	R	-	-	-	11 ± 1	-	11 ± 0.5
P9	CT	R	R	R	R	R	-	-	-	11 ± 0.5	-	11 ± 1
P10	UC	R	R	R	R	R	-	-	-	11 ± 1	-	11 ± 1
P11	SPC	R	R	R	R	R	-	-	-	12 ± 1	-	11 ± 0
P12	UC	R	R	R	R	R	-	R	-	11 ± 1	-	11 ± 0.5
P13	WC	R	R	R	R	R	-	-	-	10 ± 0.5	-	11 ± 1
P14	WC	R	R	R	R	R	-	R	-	11 ± 1	-	11 ± 0.5
P15	UC	R	R	R	R	R	-	-	-	11 ± 0.5	-	11 ± 0
P 1430	-	S	S	S	S	S	I	S	15 ± 2	17 ± 2	12 ± 1	15 ± 1

^a UC (urine culture), SPC (sputum culture), CT (Chest tube), WC (wound culture) “R” resistant; “S” Susceptible. ^b CP (Ciprofloxacin, 5 µg(, LEV (Levofloxacin, 5 µg), GM (Gentamicin, 10 µg), AMK (Amikacin, 30 µg), IMI (Imipenem, 10 µg), MEM (Meropenem, 10 µg) and TZP (Piperacillin/tazobactam, 110 µg), IP (Iranian propolis), PN (propolis nanoparticles). - ^c Not checked., ^d Zone of inhibition was not seen. Data are presented as mean ± SD (n = 3).

In the MIC method, the average MIC for 9 MDR strains was 32.5 ± 11 and 7.3 ± 3 for IP and PN, respectively. PN with MIC values of 3.2-15 mg/mL was more effective against MDR P. aeruginosa than IP with a MIC value of 25-50 mg/mL (P < .001). Also, CP with MIC values of 0.31-1.25 mg/mL showed antibacterial activity against MDR P. aeruginosa (Table 3). In addition, no significant difference was found between PN and CP in terms of antibacterial activity against MDR P. aeruginosa (P = .1).

Table 3.

The Minimum Inhibitory Concentration (MIC), Minimum Bactericidal Concentration (MBC) Values of Samples Against MDR P. aeruginosa.

Strains	IP(mg/mL)		PN(mg/mL)		CP(mg/mL)
Strains	MIC	MBC	MIC	MBC	MIC	MBC
P3	50	>100	3.2	7.5	0.31	7.5
P5	50	>100	7.4	15	0.62	15
P7	25	100	7.4	15	1.25	15
P9	25	100	7.4	15	0.62	15
P10	25	100	3.2	15	0.31	15
P11	25	100	7.4	15	1.25	15
P12	50	>100	7.4	15	0.62	15
P13	25	100	15	>15	1.25	>15
P14	25	100	7.4	>15	0.62	>15
P15	25	100	7.4	15	0.62	15
P 1430	25	50	3.2	7.5	0.00078	0.0015

Abbreviations: IP, Iranian propolis; PN, propolis nanoparticles; CP, Ciprofloxacin.

Time-Kill Assay

The average bacterial density of the initial inoculum in all experiments was 5-6 log₁₀ CFU/mL. For the control group, the isolates grew to 9-10 log₁₀ CFU/mL by 24 h. Figure 3 shows the time-kill curve for P. aeruginosa isolates at different sub-MICs of IP, PN, and CP. At 24 h, the colony forming unit (CFU) of P. aeruginosa (PTCC 1430) and MDR P. aeruginosa decreased, respectively, in the cultures treated with PN (1.6 and 3.2 mg/mL), IP (25 and 12.5 mg/mL) and CP (0.0001-0.0003 mg/mL) relative to the control group. Other sub-MICs of IP and PN did not result in fewer bacteria than the control group.

Figure 3.

A; Growth curve of P. aeruginosa (PTCC 1430) and B; MDR P. aeruginosa in presence of IP and PN. P. aeruginosa cells were grown in presence or absence of samples at subMIC for 24 h at 37 °C.

The Synergistic Effect of IP and PN Combined with CP

The ΣFIC results of P. aeruginosa (PTCC 1430) indicated the synergistic antibacterial effect of PN combined with CP. As shown in Figure 4A, PN (0.4 mg/mL) combined with CP (0.00019 mg/mL) displayed a synergistic effect as the FICI value dropped below 0.5, whereas CP combined with IP (ΣFIC = 0.75) showed an additive effect. The checkerboard results demonstrated the additive effects of MDR P. aeruginosa isolates (FICI: >0.5 to ≤1) of PN and IP combined with CP. Only MDR P. aeruginosa strain (P3) showed synergistic antibacterial effect, when PN (0.8 mg/mL) combined with CP (0.07 mg/mL) (FICI <0.5) (Figure 4B).

Figure 4.

Synergistic effect of IP or PN with CP. Schematic checkerboard of P. aeroginosa (PTCC 1430) (A) and MDR P. aeruginosa (P3) (B) growth inhibition with varying concentrations of IP or PN with CP. Bacterial load (log₁₀ CFU/mL) over 24 h, (C) P. aeruginosa (PTCC 1430) and (D) MDR P. aeruginosa (P3) for CP regimen.

The Time Kills the Effect of PN in Combination with CP

The mean bacterial loads (log₁₀ CFU/mL) over 24 h for P. aeruginosa isolates treated with PN combined with CP regimen are shown in Figure 4. PN (0.4 mg/mL) and CP (0.00019 mg/mL) produced greater killing effect (≥2 log₁₀ CFU/mL) against P. aeruginosa (PTCC 1430) at 24 h post-treatment relative to either treatment alone (Figure 4C). PN (low doses of 0.2 and 0.8 mg/mL) combined with CP (0.00019 and 0.00009 mg/mL) exhibited no synergistic effects relative to either treatment alone. In investigating MDR P. aeruginosa (P3), as an isolate with a synergistic effect, PN (0.8 mg/mL) and CP (0.07 mg/mL) reduced the colony count at 24 h (Figure 4D). The combination of low-dose PN with CP was not bactericidal against any of the isolates at 24 h. However, PN (0.2 and 0.4 mg/mL) combined with CP (0.00019, and 0.07 mg/mL) reduced the number of P. aeruginosa (PTCC 1430) and MDR P. aeruginosa (P3), respectively, compared to the control.

IP and PN Reduced the Metabolic Activity of MDR P. aeruginosa Isolates

The ability of IP and PN to inhibit biofilm formation in MDR P. aeruginosa isolates (P3, P9, and P12) after 24 h of growth stages was evaluated using the MTT method (Figure 5A). The metabolic activity of MDR P. aeruginosa isolates after IP treatment decreased to 19.24 ± 9.43% at sub-MICs, whereas PN exhibited better antibiofilm action against the isolates with a 44.56 ± 5.75% reduction in metabolic activity relative to the control (P < .001). Only in the P12 strain, no significant difference was observed in the IP compared to the control (P = .07).

Figure 5.

A; Effect of IP (25, 12.5 mg/mL) and PN (1.6 and 3.2 mg/mL) on the biofilm formation at 24 h of growth phase of P. aeruginosa (PTCC 1430) and MDR P. aeruginosa at sub-MIC level. The data represent an average of triplicate experiments ± SD (n = 3) and ***P < .001 and **P < .01 indicated in compare between samples. # The all treatments of PN had significant difference in compare with the untreated control (P < .001) except P12 strain. B, C and D; The scanning electron micrograph of MDR P. aeruginosa (9P) biofilm formed after 24 h of incubation. Control (B) and in the presence of sub- MIC levels of IP (C) and PN (D).

SEM analysis indicated the impact of IP and PN on the activity of MDR P. aeruginosa (Figure 5). Figure 5B displayed biofilm formation for the control sample after 24 h, while a reduction in the number of MDR P. aeruginosa (isolates) was detected following treatment with sub-MICs of IP and PN (Figure 5C and D).

Discussion

Propolis is a non-toxic natural substance with diverse biological properties due to the phenolic and flavonoid compounds it contains.¹² Four flavonoid compounds were found in IP: galangin, pinostrobin chalcone, tectochrysin and naringenin. The flavonoid galangin is consistently present in propolis, the most abundant compound (55%) in IP, which exhibits antibacterial activity by inhibiting bacterial RNA polymerase (RNAP).²⁶ Pinostrobin was found to inhibit Escherichia coli (E. coli), Bacillus subtilis (B. subtilis), Shigella dysenteriae (S. dysenteriae) and methicillin-resistant Staphylococcus aureus (MRSA) with MIC values in the range of 250 to 500 µg/mL.²⁷ Chrysin and tectochrysin were active against Acinetobacter baumannii, by causing membrane disruption and a change in the bacterium's membrane potential.²⁸ In addition, chrysin resulted in growth inhibition of E. coli, Staphylococcus aureus (S. aureus), and Enterococcus faecalis (E. faecalis), but failed to inhibit P. aeruginosa up to 250 µg/mL. Instead, chrysin derivatives increased antibacterial activity and inhibited P. aeruginosa with MIC values ranging from 25 to 200 µg/mL.²⁹ Tectochrysin was able to inhibit Gram-positive strains, including S. aureus and all Streptococcus species.³⁰ Naringenin and its derivatives significantly attenuated biofilm formation in MRSA and Δagr mutants and showed a synergistic effect in combination with oxacillin.^31,32 These data suggest that the synergistic effect of the main components of IP may be associated with the inhibition of MDR P. aeruginosa. Herein, IP with MIC values of 25-50 mg/mL could inhibit MDR P. aeruginosa. However, the MIC values of various geographical origins for ethanolic extracts of propolis against P. aeruginosa were found to be 1252 µg/mL (32-7910 µg/mL).³³ In this study, the effective dose of IP was increased due to the resistance of MDR P. aeruginosa strains to more than five antibiotics. In all experiments, higher concentrations of IP and PN inhibited MDR strains more effectively than P. aeruginosa (PTCC 1430). Additionally, converting IP to PN reduced the inhibitory dose required against P. aeruginosa isolates, indicating that nanoparticles exhibit greater inhibitory activity against Pseudomonas strains.

This study used Tween 80 (10%) to stabilize NPs by preventing their condensation by orienting them around the PN prepared using ultrasonic waves. Particle size and ZP are crucial variables that affect antibacterial activity, providing antimicrobial properties against microorganisms.³⁴ DLS analysis of PN revealed the distribution of particles within the nanometer range. The PN (average size = 161.4 nm, MIC value: 3.2-15 mg/mL) demonstrated greater efficacy against MDR P. aeruginosa compared to IP. The unique electrical and physical properties, increased strength and stability, and larger surface area of nanomaterials drive the interest in their processing and application.³⁵ In our previous study, PN (250 μg/mL) proved more effective against E. faecalis than IP using the MIC method.²³ In another study, these nanoparticles inhibited the growth of S. mutans with a MIC value of 25 mg/mL.³⁶ Various mechanisms, such as cell membrane damage leading to protein and DNA misfolding and the production of reactive oxygen species (ROS), have been proposed to explain the antibacterial properties of NPs.³⁷ Further experiments are needed to elucidate the exact mechanisms involved in the antibacterial action of NPs. Rozak et al reported that Au@AgNPs of propolis (555 nm) exhibited antibacterial activity against bacterial growth, with MIC/MBC values against P. aeruginosa OL375153 being 31.25 µg/mL. Additionally, the time-kill curve indicated antibacterial efficacy against all tested bacteria with treatment at 50 µg/mL for 5 h.³⁸ Abdelsattar et al (2023) investigated the antimicrobial action of three types of PNs, including Ag-CS NPs, Pro-CS NPs, and Ph-CS NPs, using phages. Among the NPs, Ag-CS NPs (380-420 nm), when combined with phages, exhibited reduced MIC values of 62.2, 31.2, and 15.6 μg/mL against Staphylococcus sciuri (S. sciuri), Salmonella enterica serotype Typhimurium (S. Typhimurium), and P. aeruginosa, respectively. These findings suggest that PNs, in combination with phages, could effectively inhibit bacterial growth and biofilm formation.³⁹ Therefore, propolis nanoparticles (AgNPs, ZnO NPs, and SeNPs) demonstrated rapid release of phenolic compounds and exhibited superior antimicrobial activity against various bacteria.⁴⁰ The propolis nanoemulsion synthesized in this study also showed effective antibacterial properties against MDR P. aeruginosa. Further research is needed to optimize the synthesis of propolis-based nanoparticles, enhance their efficiency, control particle size, and explore their applications in medicine and healthcare.⁴¹

Ciprofloxacin resistance in most bacterial isolates is caused by chromosomal mutations or reduced drug accumulation due to the overexpression of efflux pump systems. Mutations in the target genes GyrAB and parCE in P. aeruginosa decrease the affinity of DNA gyrase or topoisomerase for this drug. Ciprofloxacin may become more effective in photodynamic therapy against biofilm communities of microorganisms and multidrug-resistant isolates when combined with antibiotics from different classes, nanoparticles, natural products, and bacteriophages. Functionalization of antibiotic nanomaterials improves efficacy by enhancing drug delivery characteristics and creating interesting synergistic effects.⁴² Nanoparticles have emerged as a novel tool to help overcome antibiotic resistance in bacterial infections.⁴³ Although studies have investigated the combination of propolis and its NPs with antibiotics against bacteria, there are no reports on their effect against drug-resistant strains, particularly MDR P. aeruginosa. This study employed a checkerboard assay in a 96-well microplate (200 μL) to evaluate the effect of two antimicrobial combinations, providing a good estimate of CP and PN under synergistic conditions. In the time-kill assay, the colony-forming units (CFU) of MDR P. aeruginosa isolates decreased at 3.2 mg/mL for PN and at 50 mg/mL for IP compared to the control at 24 h. However, only one MDR P. aeruginosa isolate (P3) exhibited a synergistic antibacterial effect for PN (0.8 mg/mL) combined with CP (0.07 mg/mL) with a ΣFIC < 0.5. Consequently, the effective compounds in IP and their transformation into NPs could reduce the MIC and dose of CP against MDR P. aeruginosa. In Archi et al's study, propolis and PN (156.8 nm) combined with CP exhibited a synergistic antimicrobial effect against P. aeruginosa (PAO1), with a lower concentration of PN required compared to IP when combined with CP.¹⁷ Similarly, another report demonstrated that PN combined with the chlorophyllin-phycocyanin mixture or toluidine blue (TBO) produced a synergistic effect against S. mutans.³⁶ These findings suggest that PNs enhance the potency of antimicrobial activity synergistically.

Propolis effectively inhibits P. aeruginosa biofilm formation, has anti-inflammatory benefits, and serves as an adjuvant in antibiotic therapy against infectious diseases.⁴⁴ It significantly reduces the number of metabolically active viable cells in P. aeruginosa biofilms.^19–21 In our previous study, biofilm formation was reduced by 27.2% with IP (500 μg/mL) and by 54.2% with PN (50 μg/mL) compared to the control in the MTT assay.²³ The current study corroborates these results, showing that the metabolic activity of MDR P. aeruginosa isolates in biofilm decreased after treatment with sub-MIC levels of IP, while PN exhibited superior antibiofilm effects. Although no significant differences were observed among strains treated with PN, IP's biofilm inhibition was notably more effective against MDR P. aeruginosa isolate P3 compared to isolates P9 and P12. This variability likely stems from differences in drug resistance, as bacteria develop diverse defense mechanisms through mutation and selection. Ong et al found that chitosan-PNs significantly suppressed biofilm formation by Staphylococcus epidermidis (S. epidermidis), reducing viability to 25% compared to ethanol and ethyl acetate extracts of propolis.⁴⁵ This enhanced biofilm inhibition can be attributed to the smaller size and larger surface-area-to-volume ratio of the NPs, which facilitate better biofilm penetration and more efficient bacterial killing.⁴⁴

As previously mentioned, antibiotic resistance in P. aeruginosa represents a significant challenge in antibiotic treatment. Based on the results of this study, PN synergistically increased antimicrobial activity. Therefore, combining natural compounds such as propolis and propolis nanoemulsion with antibiotics that have low side effects could be an effective strategy to reduce the incidence of drug resistance. Reportedly, certain compounds in propolis, particularly phenolic compounds, are responsible for their bioactivity against various diseases. The current study has a number of limitations as the effects of the four flavonoid compounds identified in IP as well as the antibacterial mechanisms of PN combined with CP against MDR P. aeruginosa were not investigated in this study. Further research is needed to elucidate the mechanisms underlying the beneficial properties of propolis. It is also true that although the present study investigated the antibacterial efficacy of PN and its synergistic effect in combination with ciprofloxacin (CP) against MDR strains of P. aeruginosa biofilm in vitro, in vivo studies are the next step to confirm the therapeutic benefit of PN.

Conclusions

This study demonstrated that a nanoemulsion can be prepared from the ethanol extract of propolis, and this nanoemulsion exhibits effective antibacterial properties against MDR P. aeruginosa in vitro. The PN showed more potent antibacterial activity and was more effective in suppressing biofilm formation against MDR P. aeruginosa. Furthermore, the combination of PN and CP enhanced the drug's effect on the organism. This synergistic combination reduced the required dosage of each compound and shortened the time needed to eradicate the bacteria compared to using the compounds separately. Combining PN with antibiotics can help lower the antibiotic dosage while disrupting biofilms, thus increasing treatment efficacy. Given the rise of antibiotic-resistant strains and the side effects of antibiotics, it is crucial to conduct more rigorous and practical research on using natural compounds with low side effects to treat such infections.

Footnotes

Acknowledgments

The authors are grateful to Babol University of Medical Sciences for its support in this research.

Author Contributions

The study was designed by ZS and FA. Experimental work, data collection and analysis were performed by F A, S K, BN S, ZE and Z M. Manuscript and figures were prepared by FA, S K and M JA. All authors have read and agreed to the final version of the manuscript.

Declaration of Conflicting Interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Ethical Approval

This study was approved by the Research Ethics Committee of Babol University of Medical Sciences, Babol, Iran (IR.MUBABOL. HRI. REC. 1396.151).

Funding

This work was supported by grants from Babol University of Medical Sciences under research project No 140013321.

Data Availability

The datasets used and/or analysed during the current study available from the corresponding author on reasonable request.

ORCID iD

Fariba Asgharpour

Statement of Human and Animal Rights

This article does not contain any studies with human or animal subjects.

Statement of Informed Consent

There are no human subjects in this article and informed consent is not applicable.

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