Formulation and characterization of nanoemulsion from Alhagi maurorum essential oil and study of its antimicrobial, antibiofilm, and plasmid curing activity against antibiotic-resistant pathogenic bacteria

Abstract

Nanoemulsion technology is an alternative candidate to overcome antibiotic resistance in pathogenic bacteria. The aim of this research was nanoemulsion production from the essential oil of Alhagi maurorum and the characterization of this nanostructure. Nanoemulsion of essential oil from A. maurorum was prepared using the ionotropic gelation method and chitosan as a nano-carrier. Scanning Electron Microscopy (SEM) was used to characterize the synthesized nanoparticles. The effect of nanoemulsion on the antibacterial, antibiofilm, and plasmid curing of six antibiotic-resistant pathogenic bacteria (P. aeruginosa, E. coli, S. aureus, K. pneumonia, A. baumannii, B. cereus) was evaluated. The results of this study showed that nanoparticles had a spherical shape and smooth topology. The mean size were 172 ± 4 nm and Zeta potentials was +28.6 mv. The results of antibacterial activity confirmed that nanoemulsion of essential oil had higher inhibition against bacteria compared to free essential oil. Also, this nanoemulsion had antibiofilm activity. Minimum Inhibitory Concentration (MIC) and Minimum Bactericidal Concentration for Biofilm (MBCB) were determined for nanoemulsion against the biofilm of pathogenic bacteria. The results have shown that the MIC value for A. baumannii is 12.5 mg ml ⁻¹ and for E. coli this value is 1.75 mg ml ⁻¹. This finding means that MIC values were highest for A. baumannii and lowest for E. coli. Statistical analysis demonstrated that the inhibitory effect of nanoemulsion against bacterial biofilm was significant (P < 0.05). This nanoemulsion also had a remarkable effect the curing of R-plasmid of three antibiotic-resistant bacteria. According to GC-MS analysis of A. maurorum essential oil, the main compounds were oxygenated sesquiterpenes and hydrocarbons. Nanoemulsion of A. maurorum had the potential to use as suitable antimicrobial agents against antibiotic-resistant bacteria.

Introduction

Despite the dramatic advances in medical knowledge and technology, infectious diseases account for significant morbidity and mortality worldwide [1]. Extensive administration of antibiotics has contributed to emerge and spread antibiotic-resistant bacteria. Also, when bacteria enter biofilm forms the resistance to antimicrobial agents increase.

A biofilm, as the predominant bacterial life-style, is a system structured by surface-attached bacteria that are exposed by their extracellular polymer matrix [2]. It is formed usually in the aqueous solutions in the presence of different microorganisms and solid surfaces. One of the key benefits of such biofilms for bacteria is an increase in resistance to antimicrobial agents compared to sporadic status [3].

There are some mechanisms involved in the antibiotic resistance within the biofilms, such as the formation of a physical barrier by exopolymeric materials, the presence of inactive bacteria inert to antibiotics, and the expression of unique resistance genes [4]. The ability of bacterial biofilm to resistance with antimicrobial agents caused the researcher to screen new compounds with the natural sources to combat biofilm of pathogenic bacteria. [5]. Among these, natural substances derived from plants containing bioactive compounds have shown potential antimicrobial properties [6].

Abundant secondary metabolites are present in the plants, which play a protective role against microbial pathogens. The human beings have been using globally over 35,000 plant species for medicinal purposes. Typically, such plant materials have shown safe and have not any side effects compared to chemical compounds [7].

The essential oils have been extracted as antimicrobial agents from plants have been recognized since old time. Recently, the medicinal plants have attracted significant attention owing to antioxidant and antibacterial properties and because of having high levels of secondary metabolites, including phenolic and flavonoid compounds, glycosides and alkaloids [8].

Natural essential oils contain a mixture of numerous volatile molecules [7]. They have many applications while being highly sensitive to environmental factors. The stability of essential oils could increase by encapsulation technology [9].

In the present era, one of the newly emerged promising approaches is a nanotechnology that has been able to formulate biologically active medicines and to enhance the bioavailability of phytomedicines. Herbal medicine nanonization has opened a new window for dealing with novel herbal medications. The active ingredients present in the extracts often has low absorption and poor bioavailability and cannot pass through the cell membrane due to their very high molecular size or low solubility in water [10].

A few medicinal plant extracts have been developed for clinical applications. Integration of herbal drugs with nanotechnology have some benefits such as decreased administration dosage and enhanced bioactivity. Among these, nanoemulsion technology has been introduced as a strong candidate to deliver effectively poorly soluble drugs [11].

Genes responsible for antibiotic resistance are usually located on extrachromosomal genetic determinants known as plasmids with DNA sequences [12], which can be transferred to surrounding bacteria. These elements are often responsible for antibiotic resistance. The therapeutic strategy using antibiotics-related plasmids can be effective in managing the onset and spread of antibiotic resistance (R-plasmids) [12].

Alhagi maurorum is a perennial shrub plant belonging to the Fabaceae family, with various names of camelthorn, camelthorn-bush, Caspian manna, and Persian manna plant. This medicinal herb is found natively to regions from the Mediterranean to Russia, as well as Australia, Southern Africa, and the western United States [13]. The heavily branched plant has gray-green thicket with long spines along the branches. Diaphoretic, laxative, gastroprotective, diuretic, expectorant, anti-diarrheal and antiseptic properties have been reported for this plant, as well as it is effective to heal rheumatism and hemorrhoids [14].

In this research antimicrobial activity of A. maurorum essential oil in free and nanoemulsion was evaluated. The antimicrobial activity of A. maurorum had been described by some researchers previously. For instance, Lagharia et al. [15] evaluated the antimicrobial properties of methanol extract from flowers of A. maurorum against E. coli and S. aureus. The extract had remarkable antibacterial properties against these bacteria [15, 16].

Sulaiman [17] investigated the antimicrobial properties of methanol extract form A. maurorum leaves and flowers. They concluded that the antibacterial effect of leaves extract was more than flower extract.

In the present study, we extracted the essential oil from the tropical A. maurorum plant, and this essential oil was formulated as nanoemulsion with chitosan as a carrier. This research aimed to optimize nanoemulsion production from A. maurorum and characterization of this nanostructure. Evaluation of the activity of this nanoemulsion against some antibiotic-resistant planktonic and biofilm bacteria and plasmid curing efficiency was another purpose of this research.

Materials and methods

Plant collection, identifications and essential oil extraction

The collection of plant samples was performed in Kerman province, Iran. There are not any permissions necessary to collect plant materials. Plant taxonomists identified collected plants and the plant voucher sample (SBUK0981) was prepared as a deposit from the herbarium at Shahid Bahonar University of Kerman, Iran. The samples of leaves and stems (70 g of each part) were dried in dark, powdered by an electric blender (Bosch MSM66150), and subjected to dry steam distillation for three hours. The essential oil was concentrated by n-hexane solvent (HPLC grade, Merck; Germany) and dried by Na₂SO₄ until obtaining analytical reagent grade anhydrous and then stored at 4 °C in sealed vials before use [18].

Bacterial strains used

The bacteria used in this study included six species, two Gram-positive bacteria: Bacillus cereus MCCKM569, Staphylococcus aureus MCCKM890 and four Gram-negative bacteria: Pseudomonas aeruginosa MCCKM971, Escherichia coli MCCKM854, Acinetobacter baumannii MCCKM345, and Klebsiella pneumonia MCCKM467. The bacterial strains were prepared from the Microbial Culture Collection of Kerman Medical University (MCCKM).

Antibiotic susceptibility test

The sensitivity of these bacteria was determined to various antibiotics by the disc diffusion method. The bacteria were cultured on the Muller Hinton Broth (MHB) medium (Merck, Germany). Each bacterial suspension was grown to OD (Optical Density) of 0.08 at 600 nm (10⁶ CFU ml⁻¹) and cultured onto Muller Hinton Agar (MHA) by swabs. Antimicrobial susceptibility test was performed by 10 antibiotics: Gentamicin, 10 mg (CN10), Tetracycline, 30 mg (TE 30), Penicillin G, 1 IU (P 1), Erythromycin, 15 mg (E 15), Oxacillin, 1 mg (OX 1), Bacitracin,10 IU (BA 10), Rifampicin, 5 mg (RD 5), Ampicillin, 10 mg (AMP 10), Ciprofloxacin, 5 mg (CIP 5), Streptomycin, 10 mg (S 10). After 24-h incubation at 37 °C, the Zone of Inhibition (ZOI) was calculated [19]. All antibiotics were purchased from Sigma-Aldrich.

Preparation of unloaded chitosan nanoparticles

Unloaded chitosan (deacetylation degree of 0.8 and medium molecular weight, Sigma- Aldrich) nanoparticles were prepared by ionotropic gelation method [20]. A solution of chitosan biopolymer (3 mg ml⁻¹, 10 ml) was obtained by the desolation of polymer powder in the aqueous solution of acetic acid (0.5% v/v) at room temperature for 30 min and 8100×g. The polymer solution filtered by a 0.45-μm syringe filter was added to 200 μl of tween 80 (4% v/v) and stirred at 40 °C for 60 min. Subsequently, Pentasodium TriPolyPhosphate (TPP, Sigma GmbH) solution (1 mg ml⁻¹, 10 ml) was poured dropwise into the polymer solution and stirred for 2 h. Finally, the nanoparticles collected by centrifugation with 6000×g for 15 min were washed with PBS buffer three times [21].

Preparation of A. maurorum essential oil loaded in chitosan nanoparticles (Nanoemulsion)

Essential oil of A. maurorum loaded in chitosan nanoparticles was prepared the same way as the preparation method for unloaded chitosan nanoparticles. A solution of chitosan biopolymer (3 mg ml⁻¹, 10 ml) was obtained by dissolution of polymer powder in an aqueous solution of acetic acid (0.5% v/v) at room temperature for 30 min and 8100×g. The filtration of the polymer solution was done by 0.45 μm syringe filter and then added to 200 μl of tween 80 (4% v/v) and stirred at 40 °C for 60 min. Next, 10 mg of essential oil was poured in 500 μl of ethanol to be dissolved and appended dropwise into the chitosan-tween 80 (Merck KGaA) solution and stirred for 60 min. Subsequently, the TPP solution (1 mg ml⁻¹, 10 ml) was poured dropwise into the polymer solution, and stirred 2 h. At last, the centrifugation with 6000×g for 15 min was performed to collect the nanoparticles and washing with PBS buffer (pH = 7) was performed three times for further analysis [22].

Determination of entrapment efficiency (EE %) and yield of nanoparticles

Nanoparticles (Nanoemulsion) were centrifuged with 6000×g for 15 min. Then, the supernatant was removed and filtered by a 0.2 μm filter. The unloaded essential oil was extracted by solvent and the amount of unloaded essential oil in the supernatant was determined gravimetrically. Entrapment efficacy was determined and calculated by this equation.

$$\left(\left(\mathit{\text{Total}}\mathit{EX}-\mathit{\text{Unloaded}}\mathit{EX}\right)÷\mathit{\text{Total}}\mathit{EX}\right)\times 100$$

In where, total EX is the total amount of essential oil, and unloaded EX is the amount of remained essential oil in the supernatant. The nanoparticle suspension was centrifuged and then the resulting pellet was freeze-dried to calculate the produced nanoparticle yield. The weight of powder was measured and nanoparticle^’s yield was calculated [23].

Analysis of nanoparticles size, zeta potential and scanning Electron microscopy (SEM)

Size and Zeta analyzer (Beckman Coulter) was used to calculate the particle size and Zeta potential values. The nanoparticle suspension was added to distilled water to introduce to the size and Zeta analyzer. The nanoparticles morphology was investigated using an EM3200 scanning electron microscope. The protocol for preparation of samples for SEM analysis was as follows: First, the dispersion of the nanoemulsion in different the solvents like water, 2-propanol, ethanol, and DMSO was analyzed. The DMSO was selected as solvent because the best solubility take place in this solvent. Then, the different dispersions are ultra-sonicated to separate the particles and the most stable dispersion is chosen for SEM characterization. Finally, a drop of the stable solution is taken on the SEM grid and dried in a vacuum oven and the grid is inserted in the SEM machine [22].

Antibacterial assay of nanoemulsion and free essential oil by agar well plate method

The agar well plate was used instead of disc diffusion to assay antibacterial activity because the nanoemulsion and free essential oil had hydrophobic nature. This substrate was first dissolved in DMSO and then nanoemulsion and free essential oils were disseminated into agar well to assess the antibacterial activity [24].

The overnight bacterial culture (0.5 Mac-Farland) spread on MHA (Mueller Hinton Agar) plates. A sterile blade was used to build several wells with 8 mm in diameter and about 1.5 cm in width in each MHA plate. The nanoemulsion and free essential oils were prepared at a concentration of 100 mg ml⁻¹ as follows: the 100 mg of nanoemulsion or essential oil (powder, solid) was dissolved into 1 ml of DMSO. Then, a sterile syringe was used (when it completely dissolved) to inject approximately 40 μl of produced concentrations (100 mg ml⁻¹) into the wells and left at 30 °C for 2 h for diffusion. Also, the extract-free inoculum was considered as a negative control. The Zone of Inhibition (ZOI) values were calculated by a ruler after the incubation of MHA plates at 37 °C for 24 h. All tests were performed in triplicate and the mean ZOI values were used for analysis [19].

Determination of minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC)

The MIC values were determined by the macro-broth dilution method proposed by the Clinical and Laboratory Standards Institution (CLSI) using the MHB medium by overnight bacterial cultures at a concentration of 5 × 10⁵ CFU ml⁻¹ [25]. The nanoemulsion washed before MIC testing to eliminate residual in the medium then dried to be able to weight them for MIC testing. Different concentrations of nanoemulsion and free essential oil (dissolved in DMSO) were prepared from serial two-fold dilutions by dissolving free essential oil and nanoemulsion stock concentration (100 mg ml⁻¹) in sterilized MHB medium. One milliliter of each standard bacterial suspension, various nanoemulsion concentrations and free essential oil (0.05–100 mg ml⁻¹) were mixed in 1 ml of MHB-containing tubes, followed by incubation at 37 °C for 18 h. The MIC value was the concentration of the first tube lacking obvious growth sign. Controls included MHB plus nanoemulsion or free essential oil, cells plus MHB with sterile distilled water, and MHB plus solvents.

The MHB tubes that had no visible growth of bacteria were culture onto MHA plates and subsequently incubating at 37 °C for 24 h. The MBC value was calculated as follows: the concentration of nanoemulsion that inhibits bacterial growth and no colony found in the MHA plate [24, 26].

Assay of biofilm inhibition by nanoemulsion and free essential oil

The slightly modified method of O’Toole and Kolter [27] was used to assess the biofilm formation. Three concentrations of nanoemulsion and free essential oil were prepared from serial two-fold dilutions by dissolving nanoemulsion stock concentration (100 mg ml⁻¹) in Tryptic Soy Broth medium (TSB; Merck, Germany). The same amount (100 μl) of each concentration (12.5–50 mg ml⁻¹) was poured into the wells of a 96-well polystyrene microtiter plates. Bacteria were cultured into TSB medium overnight and re-suspended them to reach 0.2 optical density (OD). Then, each well was added by 100 μl of these suspensions, and incubated at 37 °C for 24 h. Controls included TSB plus nanoemulsion (essential oil control), TSB plus bacteria (positive control), TSB plus sterile distilled water (negative control) and TSB only (media control).

The formation of biofilm was quantitated by reading the OD of biofilm stained by Crystal Violet (CV, Merck, Germany) at a wavelength of 630 nm. After 24 h of exposure, media were discarded and each well was washed with PBS three times to clean non-adherent cells. The adhered cell was fixed by adding 150 μl of methanol 96% for 15 min. The contents of the wells were then removed by the sampler, and 200 μl of CV (1%) was appended to the wells for staining for 20 min. Additional dye was removed by washing with tap water and the plates were dried in air at room temperature. To measure the OD of adhered cells, the CV was re-solubilized with 160 μl of acetic acid glacial (33%, Merck, Germany) and the OD was measured with a microplate reader (ELX-800, Biotec, India) at 630 nm. The percentages of biofilm inhibition by various concentrations of nanoemulsion and free essential oil were calculated employing as stated below the ratio of the OD630nm values with and without the nanoemulsion [28].

$$\%\text{inhibition}=\frac{\left(\mathit{OD}\text{negative control}-\mathit{OD}\text{media control}\right)-\left(\mathit{OD}\mathit{\text{test}}-\mathit{OD}\text{essential oil control}\right)}{\left(\mathit{OD}\text{negative control}-\mathit{OD}\text{media control}\right)}\times 100$$

Destruction of biofilm by nanoemulsion and free essential oil

The biofilm samples were prepared by adding 100 μl of stationary-phase bacterial cultures (0.5 Mac-Farland) to TSB medium contained in the wells of a 96-well polystyrene microplate under sterile conditions and then incubating at 37 °C for 24 h. Following the formation of biofilm, the medium was discarded slowly and non-adherent cells were evacuated by rinsing with PBS three times. The impacts of nanoemulsion and free essential oil on formed biofilm were assessed by the same procedure as described above except that each nanoemulsion and free essential oil was added to the well-selected concentrations (12.5–50 mg ml⁻¹) and it were incubated for 24 h at 37 °C. The CV staining method was used to analyze further destruction of formed biofilm. The destruction levels of biofilm structures in the presence of different concentrations of nanoemulsion and free essential oil was calculated employing the formula as described in the previous section [29].

Assessment of biofilm the metabolic activity

The method proposed by Ramage et al. [30] was applied to evaluate the effect of nanoemulsion and free essential oil on metabolic activity of preformed biofilm. First, the PBS was used to rinse twice the preformed biofilms, followed by adding nanoemulsion and free essential oil (12.5–50 mg ml⁻¹) incubating for 24 h at 37 °C. Then, each well was appended by 50 μl of Triphenyl Tetrazolium Chloride (TTC, Merck, Germany) solution to allow inducing a reaction in the dark at 37 °C for 3 h. The microplate reader was used to read the OD at a wavelength of 490 nm. The biofilm metabolic activity level was measured in the presence of different concentrations of nanoemulsion and free essential oil by comparing the OD of control and treated inoculation with nanoemulsion and free essential oil [30].

Plasmid curing activity of nanoemulsion

The approach previously described by Deshpande et al. [31] was used for the plasmid treatment. In brief, three antibiotic-resistant bacteria (P. aeruginosa, E. coli, A. baumannii) grown in the exposure of nanoemulsion at different concentrations (10–200 μg ml⁻¹) for 24 h at 37 °C were cultured onto MHA medium to obtain pure colony’s; the cultivation was repeated onto antibiotic-containing MHA and MHA media. Putative cured derivatives were the colonies non-grown in the presence of antibiotics. The agarose gel electrophoresis of the DNA plasmid preparations was applied to determine the plasmid loss. The large-scale alkaline lysis method was utilized to isolate the plasmid, followed by 0.8% agarose (type 1; Sigma) electrophoresis of 25 μl of plasmid samples at 150 V, 60 mA for 5.5 h in TBE buffer [31].

Gas chromatography-mass spectrometry (GC-MS)

The GC-MS was used to analyze the chemical composition of essential oil. The chromatography device (Agilent 6890 UK) equipped with an HP-5MS capillary column (30 × 0.25 mm ID × 0.25 mm film thickness) was adjusted for the initial temperature of 50 °C, temperature ramp of 5 °C min⁻¹, 240 °C min⁻¹ to 300 °C (holding for 3 min), and injector temperature of 290 °C. The helium as carrier gas and the split ratio was 0.8 ml min⁻¹. The components of plant essential oil were identified by mass spectra assembled by a Wiley 5 mass spectra computer library or with authentic compounds [15].

Statistically analysis

Data from differences in parameters between control, and treated groups were analyzed by SPSS Version 18.0 software analysis of variance (ANOVA) followed by the Duncan multiple range test. Statistically difference levels were the P values less than 0.05, 0.01 and 0.001 for all tests that were performed in triplicate and P value 0.05 was regarded as statistically significant.

Results

Characterization of chitosan nanoparticles

The results of morphological analysis by SEM showed nanoparticles with monomodal distribution and spherical (Fig. 1A). The mean size of the unloaded chitosan nanoparticles was 163 ± 9 nm (Fig. 1B). The suspension of chitosan nanoparticles was stable and there was no aggregate. In addition, zeta potential was +30 mV for the unloaded nanoparticles (Fig. 1C).

Fig. 1.

a Scanning electron micrograph of unloaded chitosan nanoparticles b Size distribution of unloaded chitosan nanoparticles and c Zeta potential of unloaded chitosan nanoparticles (+30 mV)

Characterization of essential oil of A. maurorum loaded in chitosan nanoparticles (nanoemulsion)

Results of morphological analysis by SEM showed loaded chitosan nanoparticles with monomodal distribution and spherical (Fig. 2A). The mean size of essential oil loaded in chitosan nanoparticles was 172 ± 11 nm (Fig. 2B). The suspension of loaded chitosan nanoparticles was stable and there was no aggregate. Also, the zeta potential was +28.6 mV for essential oil loaded nanoparticles (Fig. 2C).

Fig. 2.

a Scanning electron micrograph of A. maurorum essential oil that loaded in chitosan nanoparticles, nanoparticles with spherical shape were observed b Size distribution of essential oil loaded in chitosan nanoparticles (Nanoemulsion) and c Zeta potential of essential oil loaded in chitosan nanoparticles (+28.6 mV)

Determination of yield and encapsulation efficacy (EE %)

The yield of unloaded nanoparticles and essential oil loaded nanoparticles was 8 mg (26.6%) and 12 mg (30.1%), respectively. About 40% of essential oil (4 mg) was entrapped into the chitosan nanoparticles.

Antibiotic susceptibility profile of bacteria

The sensitivity and resistance of six pathogenic bacteria were determined against 10 traditional antibiotics. The results were presented in (Table 1). As shown, multi-drug resistance confirmed and all the bacterial strains were at least resistant to one antibiotic. However, among these six pathogenic bacteria, A. baumannii was multi-drug resistant to 4 antibiotics namely: Tetracycline, Penicillin, Bacitracin, and Ampicillin.

Table 1.

The selected bacterial species were tested to resistant or susceptibility to known antibiotics. The Zone of Inhibition measured as mm (ZOI)

Antibiotics	Gentamicin	Tetracycline	Penicillin	Erythromycin	Oxacillin	Bacitracin	Rifampicin	Ampicillin	Ciprofloxacin	Streptomycin (ZOI mm)
Bacteria
S.aureus	18	19	0	22	17	10	16	0	19	R
B.cereus	0	22	18	16	28	30	14	0	17	19
P.aeruginosa	18	24	0	14	25	23	12	16	R	R
A.baumannii	22	0	0	20	0	17	21	12	10	R
E.coli	13	26	21	0	14	0	18	0	9	17
K.pneumoniae	10	23	17	13	0	28	0	8	12	16

Antibacterial activity

The difference in antibacterial activity levels of essential oil before and after encapsulated in chitosan nanoparticles was studied by the agar well plate method. As shown (Table 2), the negative control (DMSO) and chitosan had no antibacterial activity. The antibacterial activity was smaller significantly in free essential oil than in loaded essential oil in nano-chitosan (nanoemulsion). The best antibacterial activity of loaded essential oil was seen against A. baumannii (29 mm) and the lowest antibacterial activity was shown by P. aeruginosa (15 mm).

Table 2.

The antimicrobial effect of nanoemulsion and free essential oil from A. maurorum against six planktonic bacteria and MIC/MBC values

Bacteria	Antimicrobial effect of Nanoemulsion (mm)	Antimicrobial effect of Free essential oil (mm)	DMSO (Negative Control mm)	Erythromycin (Positive control mm)	Bacteria	Nanoemulsion (mg ml −1)		Free essential oil (mg ml −1)
S.aureus	26 ± 0.1	13 ± 0.5	0	22		MIC	MBC	MIC	MBC
B.cereus	24 ± 0.4	11 ± 0.2	2	16	B.cereus	1.75	6.25	3.125	12.5
P.aeruginosa	15 ± 0.2	6 ± 0.1	0	14	P.aeruginosa	3.25	12.5	6.5	25
A.baumannii	29 ± 0.6	15 ± 0.3	3	20	A.baumannii	12.5	25	25	50
E.coli	18 ± 0.3	7 ± 0.8	1	0	E.coli	1.75	3.25	3.125	6.5
K.pneumoniae	22 ± 0.7	8 ± 0.3	0	13	K.Pneumoniae	6.25	25	12.5	50
					S.aureus	6.25	25	12.5	50

*Unloaded chitosan was 3 mm ZOI

The MIC and MBC of nanoemulsion were determined in comparison with free essential oil using the macro-broth dilution method against six bacteria (Table 2). According to the results, the MIC and MBC values of essential oil loaded in chitosan nanoparticles were smaller than free essential oil. In the other hand, the nanoemulsion can inhibit the growth of all pathogenic bacteria at a lower concentration than free essential oil.

Antibiofilm activity

Figures 3, 4, and 5 show the inhibitory effects of the nanoemulsion and free essential oil on preventing biofilm formation, destruction, and inhibition of its metabolic activity.

Fig. 3.

The effect of different concentrations of free essential oil (a) and nanoemulsion (b) from A. maurorum against biofilm formation of pathogenic bacteria

Fig. 4.

The effect of different concentrations of free essential oil (a) and nanoemulsion (b) from A. maurorum against the destruction of biofilm structures

Fig. 5.

The effect of nanoemulsion and free essential oil on dehydrogenase enzyme activity of six pathogenic bacteria

The maximum and minimum inhibitory effects of nanoemulsion were observed on A. baumannii (98%) and E. coli (35%) (Fig. 3). The destruction of bacterial biofilm treated with different concentrations of free essential oil and nanoemulsion had diverse results. The A. baumannii biofilm was susceptible (90%) but P. aeruginosa biofilm (22%) had resistant (Fig. 4) biofilm structure among tested bacteria. In the overall antibiofilm activity of the nanoemulsion was bigger than free essential oil.

The effect of nanoemulsion and free essential oil on inhibition of dehydrogenase enzyme of pathogenic bacteria was illustrated in Fig. (5). As shown in this figure, the metabolic activity of bacteria in biofilm was significantly reduced, the maximum decrease was obtained for S. aureus (82%) and the minimum decrease for K. pneumoniae (55%).

The effect of different concentrations of nanoemulsion and free essential oil on biofilm shown that the best concentration that inhibits biofilm formation is 12.5 mg ml⁻¹. Also, when the concentration of nanoemulsion increased the antibiofilm activity had increment patterns (Figs. 3,4 and 5).

The statistical analyses are shown in (Table 3). As shown in this table the response of a different types of bacteria to the inhibitory activity of essential oil was significant in three levels (P˂ 0.05, P˂ 0.01, and P˂ 0.001). Also, the inhibitory effect of free essential oil against the biofilm of bacteria was not significant. However, the inhibitory effect of nanoemulsion against bacterial biofilm was significant in three levels (P˂ 0.05, P˂ 0.01, and P˂ 0.001).

Table 3.

The data obtained from different experiment were statistical analysis by Duncan’s test

Sources Change	Df			MS			Sig
	Biofilm formation	Destruction biofilm	Inhibition enzyme	Biofilm formation	Destruction biofilm	Inhibition enzyme	Biofilm formation	Destruction biofilm	Inhibition enzyme
1. Bacteria type	5	5	5	0.123	0.047	0.048	*	**	**
2. Free essential oil	1	1	1	0.091	0.247	0.002	–	–	–
3. Nanoemulsion	5	5	5	0.0121	0.090	0.037	*	***	*
4. Concentration of free essential oil	10	10	15	0.042	0.007	0.009	–	–	–
5. Concentration of nanoemulsion	2	2	3	0.014	0.065	0.003	*	**	–
Total	25	25	32

Df: degree of freedom

MS: min square

Sig: signification

Plasmid curing activity of nanoemulsion

The effect of nanoemulsion on curing and deletion of R-plasmid of three antibiotic-resistant bacteria were shown in Fig. (6). This Figure confirmed that all R-plasmid present in the P. aeruginosa and A. baumannii were cure completely but this nanoemulsion cannot cured R-plasmid present in E. coli. As shown in Fig. (6) P. aeruginosa has three different plasmids that cured after treatment with nanoemulsion also A.baumannii has two different plasmids that completely cured after nanoemulsion treatment however two different plasmids in E. coli present after nanoemulsion treatment. In the other hands, this nanoemulsion had not any curing effect on a resistant plasmid of E. coli.

Fig. 6.

The plasmid curing activity of nanoemulsion on three antibiotic-resistant bacteria. Lane (1): 10 Kb DNA ladder, Lane (2): Plasmid profile of P. aeruginosa Lane (3): P. aeruginosa treated with nanoemulsion, Lane (4): Plasmid profile of A. baumannii, Lane (5): A. baumannii treated with nanoemulsion, Lane (6): Plasmid profile of E. coli, Lane (7): E. coli treated with nanoemulsion, Lane (8): Negative control (H₂O)

Chemical composition of A. maurorum essential oil

The constituents of A. maurorum essential oil were determined by GC-MS. The results were presented in (Table 4). As shown in this table twenty-three (23) active ingredients were detected in the essential oil of this plant. The main compounds included oxygenated sesquiterpenes (24%), and hydrocarbon (19%). Major constituents of the tested extracts were as follow: 2-Nonadecanone, Octadecane, Propanamide, Trichloroacetic acid, Tetramethyl-2-hexadecen-1-ol, Squalene, and Octacosane.

Table 4.

Chemical composition of A. maurorum essential oil

Compoundsa	RTb	Area	Percent
2-Nonadecanone	4.973	9.599e+5	5.09
9-Octylheptadecane	5.842	4.038e+5	2.10
Tetrahydropyridin-2-one, 5-methyl-	6.716	4.742e+4	6.16
Octadecane	8.803	7.312e+6	0.50
3,4-Dimethyl-3-pyrrolin-2-one	10.153	4.563e+4	0.94
Hexadecanoic acid methyl ester	11.284	6.315e+4	1.14
4H-Pyran-4-one, 2,3-dihydro-3,5-dihydroxy-6-methyl	12.140	6.169e+4	1.17
Propanamide, N-(2,6-dimethylphenyl)	14.199	7.458e+4	3.98
1,2,3-Benzenetriol	18.894	1.726e+6	1.28
Pentanoic acid-trimethyl-3-carboxyisopropyl, isobutyl ester	23.785	1.234e+5	1.58
Trichloroacetic acid, decyl ester	24.665	6.843e+5	5.44
Tetramethyl-2-hexadecen-1-ol	28.976	1.946e+5	7.2
Neophytadiene	29.625	1.285e+7	5.17
Cyclododecanone, 2-methylene-	29.860	7.562e+5	5.29
Estra-1,3,5(10)-trien-17β-ol	31.455	1.287e+5	1.50
Hentriacontane	34.322	2.186e+5	0.49
9,12,15-Octadecatrienoic acid	34.859	2.121e+5	5.01
Terpene-related compounds (include: Diterpenes, Hemiterpenes, Monoterpenes, Retinoids, Meroterpenes)	35.456	4.382e+5	7.62
Methyl (Z)- eicosatetraenoate	40.889	1.136e+6	5.17
Squalene	48.897	7.349e+4	7.82
Trepenoids (include: Triterpenoids, Triterpenoids, Triterpenoids)	51.27	2.142e+7	5.98
Nonacosane	54.38	1.242e+6	5.12
Octacosane	56.14	5.842e+7	5.09
Total	–	–	86.93

a: Names according to NIST and Wiley mass spectral libraries, and by comparing their relative retention indices

b: Retention Time

c: Percentage composition of oils was computed from peak areas of GC; compounds were identified with a resemblance percentage above 90%

Discussion

The plant essential oils with wide and complex chemical compositions have been consuming for various medical purposes, such as cough, malaria, diarrhea, severe headaches, and seizures. However, their use is limited due to unstable, volatile and water-insoluble nature [32].

The resistance of bacteria to different antimicrobial compounds is one of the health and socioeconomic challenging concerns on microorganisms throughout the world. This event is faster than the speed of progress in new technology and drug developments in the pharmaceutical industries, highlighting the importance of novel strategies in this regard such as nano-encapsulation of bioactive compounds with antimicrobial activities [33, 34]. The plant essential oils are unique sources of bioactive compounds capable of inhibiting drug resistance growth of life-threatening pathogenic microorganisms. Nanoencapsulation as a part of nanotechnology has seen successful applications in some industries, such as pharmaceuticals and medicine [35].

The protective effects of nanocarriers for essential oil molecules have been reported against environmental factors, including pH, oxygen, and light. They act as a barrier between the molecule, and the environment [10], stabilizer for volatile molecules, support against oxidative degradation, evaporation and photo-degradation [36]. Moreover, the antimicrobial activity of biomaterials like essential oils can be enhanced by the nanocarriers through their elevated cellular interactions with microbes, owing to their very small size that enhances cellular uptake [37].

There is great interest among researchers to transform essential oils of plants into nano-formulation. Many plant essential oils have been modified into nanoemulsion. For instance, Sienkiewicz et al. [38] encapsulated thyme essential oil (Thymus vulgaris) into chitosan-benzoic acid nanogel that has strong antifungal, antiviral and antibacterial activities. They evaluated the antimicrobial properties against several pathogenic microorganisms, including Salmonella Typhimurium, Yersinia enterocolitica, Shigella flexneri, Listeria monocytogenes, Shigella sonnei, Salmonella cholereasuis, Aspergillus niger, Staphylococcus, Enterococcus, Escherichia and Pseudomonas [38].

The authors encapsulated the essential oil of Mentha piperia in chitosan-cinnamic acid nanogel. They evaluated the antifungal activity of this nanoemulsion against Aspergillus flavus. The conclusion was that nanogel could protect essential oils from environmental factors through increasing stability and performance as antifungal agents [18].

[39–41].

According to the findings of this work, this plant the free essential oil exhibited lower antibacterial activity against six antibiotic-resistance pathogenic bacteria compared to nanoemulsion. Our results in free form of essential oil are according to results reported by other researchers described above. But until now there is not research that explained the antibacterial activity of A. maurorum nanoemulsion as reported by this study and this research is the first report in this field.

In this study, the antibiofilm properties of free and nanoemulsion of A. maurorum essential oil were studied. Our results confirmed that the antibiofilm properties of essential oil of this plant were dramatically increased in nanoforms. The reasons for this observation can be interpreted as follow: medicinal extract nanoization can enhance the dissolution velocity, wetting, particle surface area, and saturation solubility, highlighting further bioavailability owing to enhanced in vivo release as only solubilized particles absorbed via lipophilic cell membrane [42].

These nanosystems could enhance the properties of herbal extracts, and decrease the dose of administration and complications. The bioactive substances can be dispersed at an adequate concentration by the nano-systems within the whole period of treatment, especially in the functional sites, which is impossible by the traditional medications. The plant extract efficiency can be maximized and their adverse effects can be minimized by the nanonization significantly. The SEM images prepared from nanoemulsion surface morphology exhibited smooth topology and spherical-shaped particles [42, 43].

There is no any research been carried out on antibiofilm and plasmid curing activity of essential oil of A. maurorum and this research had innovation in this regard. The chemical composition of A. maurorum extract was described by some researchers. Our findings in this research are according to compounds described previously for this plant.

Plasmids are the extrachromosomal elements that are responsible for development and spread of antibiotic resistance in bacteria. Their role assumes even more significance in the nosocomial environment as plasmid encoded resistance to multiple antibiotics can be transferred from one host to another by inter species transfer modes such as conjugation and/or transformation. Such acquired resistance can make otherwise sensitive pathogens resistant to multiple antibiotics. Plant derived compounds have been previously reported as plasmid curing agents.

For example, Patwardhan et al. [44] studied plasmid curing activity of Plumbago zeylanica root extracts against A. baumannii and E. coli. They concluded that the root of this medicinal plant as an effective plasmid curing agent capable of suppressing development and spread of antibiotic resistance.

In this research plasmid curing activity of nanoemulsion from A. maurorum were studied. Our results confirmed that this nanoemulsion had plasmid curing activity. To the best of our knowledge this is the first report of plasmid curing activity of this nanoemulsion constructed from A. maurorum.

Ability of this nanoemulsion to cure plasmid encoded antibiotic resistance in Acinetobacter and E. coli strains is particularly significant since Acinetobacter and E. coli strains are known to act as a reservoir of natural or acquired antibiotic resistance genes in the nosocomial environment facilitating in the spread of antibiotic resistance genes to more pathogenic bacteria.

The results of GC-MS analysis revealed components present in the A. maurorum mixture. Some of these components may be have antimicrobial activity. But determination of which compounds have antimicrobial activity need further research. Sesquiterpenes were the most abundant compounds in the essential oil of A. maurorum. Terpenes could be one of the components responsible for the antibacterial activity since it was reported by other studies that tested different plants [19, 45]. The major effect of this component is anti-diarrhea because of water absorption, and protein precipitation [46].

Phenolic compounds, fatty acids and esters were other compounds obtained from the essential oil of A. maurorum. These compounds are known to exhibit antibacterial activities [47, 48]. On the other hand, the hydrophobic character of phenolic compounds can potentially impair cellular function and membrane integrity. The capacity of phenolic compounds to chelate transition the metals-ligand also lowers the reactivity of metal ions by forming and insert metal-ligand complex. Chelation of transition metals such as iron and copper reduces bioavailability for bacterial growth [49, 50].

The limitation of this research was as follows: we cannot assay the toxicity effect of nanoemulsion produced in this research because we did not have the equipment of cell culture in our laboratory. it is proposed that this important assay will do in the future.

Conclusions

The nanoemulsion that constructed in this study had a sufficient inhibitory effect against antibiotic-resistant bacteria in planktonic and biofilm forms. Also, this nanoemulsion had a good ability to cure and delete R plasmid. In conclusion, this nanoemulsion could be beneficial as a suitable antimicrobial compound against antibiotic-resistant bacteria in the medicine and food industry.

Authors’ contributions

In this research Dr. Sadatfar collect the plant samples and prepared the essential oil. Also, Dr. Hassanshahian has done the experimental section of work and write the article. and Miss Masoumi done statistical analysis of the research.

Funding

This work was financially supported by the Research and Technology Institute of Plant Production (RTIPP) of the Shahid Bahonar University of Kerman. We are thankful for their collaborations.

Availability of data and material

Not applicable.

Compliance with ethical standards

Ethics approval and consent to participate

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Consent for publication

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Competing interests

Not applicable.