Next Article in Journal
Pharmaceutical Potential of High-Altitude Plants for Fatigue-Related Disorders: A Review
Previous Article in Journal
Discovery and Diagnosis of a New Sobemovirus Infecting Cyperus esculentus Showing Leaf Yellow Mosaic and Dwarfism Using Small-RNA High Throughput Sequencing
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Plants
Volume 11
Issue 15
10.3390/plants11152003
plants-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Antimicrobial Efficiency of Some Essential Oils in Antibiotic-Resistant Pseudomonas aeruginosa Isolates

by
Luc Tran Van
1,2,
Ilinca Hagiu
3,
Adelina Popovici
1,
Florica Marinescu
1,2,
Irina Gheorghe
1,2,
Carmen Curutiu
1,2,*,
Lia Mara Ditu
1,2,
Alina-Maria Holban
1,2,
Tatiana Eugenia Sesan
1 and
Veronica Lazar
1,2
1
Microbiology & Immunology Department, Faculty of Biology, University of Bucharest, 77206 Bucharest, Romania
2
Research Institute of the University of Bucharest-ICUB, University of Bucharest, 050657 Bucharest, Romania
3
The Overlake Private School, 108th St., Redmond, WA 98053, USA
*
Author to whom correspondence should be addressed.
Plants 2022, 11(15), 2003; https://doi.org/10.3390/plants11152003
Submission received: 14 July 2022 / Revised: 27 July 2022 / Accepted: 27 July 2022 / Published: 31 July 2022
(This article belongs to the Section Phytochemistry)
Browse Figures
Versions Notes

Abstract

:
Pseudomonas aeruginosa is a non-fermentative Gram-negative opportunistic pathogen, frequently encountered in difficult-to-treat hospital-acquired infections and also wastewaters. The natural resistance of this pathogen, together with the frequent occurrence of multidrug-resistant strains, make current antibiotic therapy inefficient in treating P. aeruginosa infections. Antibiotic therapy creates a huge pressure to select resistant strains in clinical settings but also in the environment, since high amounts of antibiotics are released in waters and soil. Essential oils (EOs) and plant-derived compounds are efficient, ecologic, and sustainable alternatives in the management of various diseases, including infections. In this study, we evaluated the antibacterial effects of four commercial essential oils, namely, tea tree, thyme, sage, and eucalyptus, on 36 P. aeruginosa strains isolated from hospital infections and wastewaters. Bacterial strains were characterized in terms of virulence and antimicrobial resistance. The results show that most strains expressed soluble pore toxin virulence factors such as lecithinase (89–100%) and lipase (72–86%). All P. aeruginosa strains were positive for alginate encoding gene and 94.44% for protease IV; most of the strains were exotoxin producers (i.e., 80.56% for the ExoS gene, 77.78% for the ExoT gene, while the ExoU gene was present in 38.98% of the strains). Phospholipase-encoding genes (plc) were identified in 91.67/86.11% of the cases (plcH/plcN genes). A high antibiotic resistance level was identified, most of the strains being resistant to cabapenems and cephalosporins. Cabapenem resistance was higher in hospital and hospital wastewater strains (55.56–100%) as compared to those in urban wastewater. The most frequently encountered encoding genes were for extended spectrum β-lactamases (ESBLs), namely, blaCTX-M (83.33% of the strains), blaSHV (80.56%), blaGES (52.78%), and blaVEB (13.89%), followed by carbapenemase-encoding genes (blaVIM, 8.33%). Statistical comparison of the EOs’ antimicrobial results showed that thyme gave the lowest minimum inhibitory concentrations (MIC) and minimum biofilm eradication concentrations (MBEC) in P. aeruginosa-resistant isolates, making this EO a competitive candidate for the development of efficient and ecologic antimicrobial alternatives.

1. Introduction

Pseudomonas aeruginosa is a versatile Gram-negative, aerobic, rod-shaped bacterium that belongs to the bacterial family Pseudomonadaceae, a member of γ-proteobacteria [1]. A recognized opportunistic pathogen, P. aeruginosa is responsible for severe hospital-acquired (nosocomial) and community infections, being present in chronic lungs of cystic fibrosis patients. The bacterium’s virulence depends on many cell-associated and extracellular factors, which enable host colonization, survival, invasion, and persistent infections [2]. Attached virulence factors, such as lipopolysaccharide, pili, and flagella are involved in tissue colonization, while soluble virulence factors such as alkaline protease, elastase, exotoxin A, pyoverdine, pyocyanin, and rhamnolipids are responsible for host immune evasion and persistence [3]. The ability to develop biofilms in various environments, including chronic lung infections and on implantable devices, makes P. aeruginosa the preferred Gram-negative model to investigate biofilm regulation. The study of bacterial biofilms is a medical priority since these multicellular-attached microbial communities are highly tolerant to increased antibiotic concentrations and host immune defense [2,3].
It has been estimated that at least 70% of the persistent P. aeruginosa infections are biofilm associated. Microbial tolerance associated with biofilm growth, its natural resistance mechanisms, and the acquisition of multiple resistance genes have led this opportunistic pathogen to be categorized as a superbug [2].
Multiple antibiotics administered in high amounts are often required to treat P. aeruginosa infections in critical patients. The administration of high amounts of antibiotics in clinical settings, together with their use in some agriculture and animal facilities, puts huge pressure on the selection of antibiotic-resistant strains [4]. Large amounts of antibiotics are released into hospital and municipal wastewaters due to incomplete metabolism in humans and animals or due to inappropriate disposal of unused drugs [5]. These substances finally find their ways into different natural environmental compartments.
Antibiotic-resistant genes, antibiotic-resistant bacteria, and antibiotic residues have been detected in wastewaters from community, hospitals, and animal slaughterhouses [6,7,8,9]. These factors create a high selective pressure for the development of multi-resistant bacteria strains and support the increased resistance rates [10].
In the context of high antibiotic resistance rates, alternatives to handling difficult-to-treat P. aeruginosa infections, caused by resistant strains, are urgently required. In recent years, researchers have investigated alternative physical, chemical, or biological alternatives to fight resistant bacteria, such as: (i) virulence modulators (including biofilm formation inhibitors), (ii) antimicrobial vaccines, (iii) genetically engineered bacteria, (iv) bacteriophages, (v) plant-derived compounds and essential oils (EOs), (vi) nanoparticles [11,12], (vii) plasmas, and (viii) other physic microorganism removal methods based on light or ultrasound [13].
Virulence modulation represents an essential approach in dealing with antimicrobial-resistant strains. Targeting specific virulence features would enable the control of the infectious process, and this approach could be efficient in both susceptible and antibiotic-resistant strains. Furthermore, natural plant-derived compounds and mixtures, i.e., EOs, were intensively investigated for the development of efficient antimicrobials to overcome high antibiotic resistance rates in P. aeruginosa [14,15].
Several natural and synthetic compounds have been introduced in previous studies to modulate P. aeruginosa growth and virulence. Extracts of various natural products and EOs of certain plants (i.e., lavender, eucalyptus, and citrus) have shown inhibitory effects against the growth and virulence of P. aeruginosa [16,17]. EOs are volatile compounds produced by the secondary metabolism of plants, mainly composed of terpenoids, terpenes, and aliphatic and aromatic constituents, and they have been proven to be efficient antimicrobial agents in multi-resistant bacteria and fungi [18,19] due to a broad spectrum of biocidal activity [20,21,22]. EO composition, such as phenolic content and antioxidants, significantly influence their antimicrobial properties [23].
Sage (Salvia officinalis), thyme (Thymus vulgaris), and Eucalyptus sp. are among the most important medicinal plants, used from ancient time as aromatic plants, but also in traditional medicine. Recent studies have shown that the essential oils produced by these plants, along with tea tree oil extracted from Melaleuca alternifolia, have antibacterial and antifungal effects against various pathogenic microorganisms. The total content of EOs as well as their chemical composition can vary depending on environmental factors, growth region, phenological cycle, and cultivation practices. Alpha- and beta-thujone; 1,8-cineole; camphor; and humulene are among most identified compounds in Salvia officinalis EO. Thyme essential oils (TEOs) include thymol, carvacrol, and paracymene. The antibacterial activity of Eucalyptus extracts is due to components such as 1,8-cineole, citronellal, citronellol, citronellyl acetate, p-cymene, eucamalol, limonene, linalool, β-pinene, γ-terpinene, α-terpinol, alloocimene, and aromadendrene. Tea tree oil consists of about 100 terpinenes and their alcohols, of which terpinene-4-ol, α-pinene, linalool, and α-terpineol are considered the most important for antimicrobial activity [24,25,26,27,28].
In this study, we aimed to evaluate the antimicrobial activity of four EOs with high content of antibacterial compounds on multidrug-resistant P. aeruginosa strains. Commercial tea tree, thyme, sage, and eucalyptus EOs were evaluated on planktonic and biofilm P. aeruginosa cultures obtained from hospital-acquired infections and wastewaters. Correlations between microbial isolation sources, virulence and resistance profiles, and antimicrobial efficiency of the tested EOs were also established.

2. Results

2.1. The Characterization of P. aeruginosa Strains

This study included 36 strains; two laboratory strains, namely, Pseudomonas aeruginosa ATCC 27853 and PA01, were purchased from American Type Culture Collection (ATCC, US), while 34 strains were isolated from clinical infections (n = 7), hospital wastewater (n = 9), and municipal wastewater (n = 18) from Romania.
Nine strains were isolated from untreated or treated (chlorine treatment) hospital wastewater from two large hospital units from Bucharest, while 18 strains were isolated from the urban wastewaters of large Romanian cities from the northeast of the country.

2.1.1. Antibiotic Susceptibility

Antibiotic susceptibility was evaluated by the disc-diffusion method. The results showed high levels of antibiotic resistance in the analyzed P. aeruginosa strains. Antibiotic resistance to piperacillin/tazobactam (TZP) was encountered in 48% of the analyzed strains, ceftazidime (CAZ) in 48%, cefepime (FEP) in 42%, aztreonam (ATM) resistance was observed for 48% of the P. aeruginosa strains, meropenem (MEM) in 38%, imipenem (IPM) in 71%, tobramycin (TOB) in 21%, amikacin (AK) in 32%, ciprofloxacin (CIP) in 33%, doripenem (DOR) in 33%, and Gentamicin (CN) in 48%, as shown in Figure 1.
The results demonstrated that the analyzed P. aeruginosa strains possess increased resistance to imipenem, but also gentamicin, aztreonam, ceftazidime, and the piperacillin/tazobactam combination.
Analysis of antibiotic resistance results of strains isolated from various sources (Figure 2) showed that P. aeruginosa strains isolated from nosocomial infections and hospital wastewaters account for higher resistance levels (55.56–100%) as compared with urban wastewaters (15.79%).
Indeed, the molecular investigation of the genes encoding for antibiotic resistance revealed that the most frequently encountered encoding genes were for extended spectrum β-lactamases (ESBLs), namely, blaCTX-M (n = 30 strains), blaSHV (n = 29 strains), blaGES (n = 19 strains), and blaVEB (n = 5 strains), followed by carbapenemase-encoding genes (blaVIM, n = 3 strains) (Figure 3).

2.1.2. Soluble Enzymatic Virulence Factors

Regarding their virulence profile, the evaluation of soluble virulence factors showed that most of the P. aeruginosa strains expressed pore forming enzymes, such as hemolysins, lipase, and lecithinase. The distribution of virulence factors seems to be unrelated to the isolation source of analyzed P. aeruginosa strains (Figure 3). In addition, about 50% of the analyzed strains showed the ability to degrade esculin in vitro (Figure 3), and this phenotype could be related to the ability of these strains to acquire iron and produce siderophores [29]. Although not significant, some modifications in the distribution of soluble virulence factors were observed among P. aeruginosa strains, depending on their isolation source. Generally, clinical and hospital wastewater isolates showed a higher percentage of positive strains as compared to urban wastewater isolates (Figure 4). Differences in virulence characteristics of microbial strains isolated from various sources were also reported in earlier studies, and they may be explained by strain diversity during infection in particular hosts [30,31].
Regarding the distribution of virulence markers in clinical and wastewater P. aeruginosa strains, the molecular analysis showed that 100% of the strains were positive for alginate-encoding gene and 94.44% for protease IV (TCF/TCR); the greatest part of the investigated strains were exotoxin producers (i.e., 80.56% for exoS gene, 77.78% for exoT gene, while the exoU gene was present only in 38.98% of the strains). Phospholipase-encoding genes (plc) were revealed as follows: plcH in 91.67% and plcN in 86.11% of the analyzed strains (Figure 5).

2.2. Antimicrobial Potential of Essential Oils

The next step of this research was to analyze the antimicrobial potential of four essential oils (tea tree, thyme, sage, and eucalyptus) by quantitative protocols to evaluate their antimicrobial potential to be considered as alternative and ecologic antimicrobials [32].
The qualitative analysis revealed that the large diameter of the inhibition zone was observed in cases of tree tea EO, followed by thyme EO regardless of isolation sources (Figure 6).
Quantitative assays revealed two key parameters, namely, minimum inhibitory concentration (MIC), and minimum biofilm eradication concentration (MBEC). These were utilized to show the efficiency of the analyzed EOs in planktonic (free floating) and attached (biofilms) bacteria cultures.
The results gave various MIC values, depending on the analyzed EO, being influenced also by the isolation source of the P. aeruginosa strains. Generally, the lowest MIC values were obtained for thyme and tea tree EOs in the tested P. aeruginosa strains, regardless of their isolation source. The average value of the minimum concentration of tea tree was 0.697%; for thyme it was 0.349%, for sage it was 4.688%, while for eucalyptus, it was 2.726%, as shown in Figure 7.
Next, we compared the minimum inhibitory concentrations of P. aeruginosa strains depending on the isolation source of the strains. The results in Figure 8 show that the mean value of the MICs for tea tree EO ranged from 0.538 to 0.759%; for thyme it was 0.246–0.470%, for sage it was 4.583–5.000%, and for eucalyptus it was 2.361–3.214%. Generally, the lowest MICs were found for thyme, followed by tea tree EOs. This trend was constant among all P. aeruginosa isolates, regardless of their isolation source.
Moreover, we used the Minitab software to statistically reveal the differences among the obtained results. Figure 9 reveals charts showing Minitab comparison of the p-values and t-values of the MICs of essential oils in the analyzed P. aeruginosa groups, in which a 95% confidence interval (CI) was assumed.
Our data demonstrated the enhanced antibacterial activity of the thyme EO in P. aeruginosa multidrug-resistant strains, as previously demonstrated [33]. It seems that MIC values of thyme EO were very low in all the analyzed P. aeruginosa groups, regardless of the isolation source. The average MICs obtained for thyme and tea tree EOs were significantly lower as compared to the average MICs of sage and eucalyptus in the analyzed P. aeruginosa strains (Figure 4).
Since the highest antibacterial efficiency in planktonic cultures, expressed as MIC value, was given by tea tree and thyme EOs, we utilized these two EOs for highlighting their impact in P. aeruginosa biofilms. It is well known that biofilm formation is a virulence and persistence trait of this opportunistic pathogen [34,35]. The minimum biofilm inhibition concentration (MBIC) assay revealed that thyme is more efficient than tea tree EO, the first being able to inhibit monospecific biofilms at lower concentrations. Our results demonstrated that the average MBIC value obtained for thyme EO in the analyzed P. aeruginosa strains was 0.028%, while tea tree EO gave an average MBEC of 0.051%, as shown in Figure 10.
When comparing the MBIC values of the two tested essential oils with the origin of P. aeruginosa strains, the results showed that the average value of tea tree was in the range of 0.028–0.052%, and that of thyme was in the range of 0.022–0.039%, as Figure 11 reveals.
We found no statistically significant correlation regarding the MBICs of the tested EOs (thyme and tea tree) among P. aeruginosa strains isolated from different sources (Figure 12).
However, overall biofilm inhibition efficiency was higher for thyme EOs, since the average MBIC for this oil was significantly lower than the average MBIC obtained for tea tree in the evaluated P. aeruginosa strains (Figure 13).
These data suggest that the analyzed EOs could be highly efficient in both planktonic and biofilm cultures, and also in antibiotic-resistant strains, which are frequent in clinical settings, but also in the environment and wastewaters.

3. Discussion

P. aeruginosa is one of the most versatile and naturally resistant opportunistic pathogens. This Gram-negative bacteria model is also recognized for its ability to develop biofilms and a high tolerance to antimicrobials and host immune systems [36]. Our study showed that the strains isolated from Romanian wastewaters and also hospitals presented high resistance rates to most of the tested antibiotics. These findings are in accordance with previous studies reported by our group [37,38,39]. Our results showed a slight reduction in the susceptibility levels of P. aeruginosa to cabapenem antibiotics (imipenem, meropenem, doripenem), of which the imipenem resistance level (IPM) is worth mentioning (71%), as compared to previously reports [18,39].
Very recently, Buzila et al. demonstrated in a retrospective 11-year study from North-Eastern Romania a very similar imipenem resistance level in P. aeruginosa strains recovered from various intra-hospital infections [40].
Similar to our obtained results, Tatu et al., in 2017, showed comparable resistance levels to third- and fourth-generation cephalosporins (ceftazidime and cefepime) in P. aeruginosa strains recovered from ICU patients admitted in one large hospital unit in Bucharest [41]. Saviuc et al., in 2016, highlighted a comparable resistance level in MDR P. aeruginosa nosocomial strains isolated from the same hospital unit (75%) and demonstrated the presence of the blaTEM ESBL-encoding gene in analyzed strains [38]. Merezeanu et al. and Porumbel et al., in 2016, demonstrated a high repertoire of virulence markers (i.e., plcH, exoU, exoT, exoS, algD) and a lower resistance level to carbapenems encoded by the blaIMP gene and a significant collection of P. aeruginosa recovered from hospitalized patients in Bucharest [42,43]. In 2015, Dortret et al. revealed also a high resistance level to carbapenem in P. aeruginosa strains isolated from intra-hospital infections from the central part of our country (Cluj Napoca Infectious Disease hospital), demonstrating the spread of carbapenem-resistant clones in Romania and their corresponding carbapenemase encoding genes: ST 2026-VIM-2-producing P. aeruginosa, a very related clone to the epidemic ST233 clone; and the ST 1982-IMP-13-producing strain, associated with class 1 integrons [44]. Other authors also reported during that period a comparable carbapenem resistance level encoding by different carbapenemase-encoding genes (i.e., VIM-4; VIM-2) in P. aeruginosa strains isolated from hospitalized patients in the south or northeast of Romania [45,46]. This result is somewhat different than the conclusion from the study of Khilasa Pokharel et al. (2019) showing that antibiotics such as imipenem, meropenem, piperacillin/tazobactam, ciprofloxacin, gentamicin, amikacin, and tobramycin are considered to be good choices for treating infections caused by this organism [47].
Production of virulence traits is a survival approach for pathogens to evade the host immune mechanism, resulting in pathogenesis mostly at the initial stage of colonization and acute infection. A large amount of virulence factors as well as cell-associated or secreted compounds of minimal and high molecular weight have been documented as vital in establishing infections by P. aeruginosa [48]. Along with high acquired resistance rates, P. aeruginosa also presents virulence factors that are responsible for their ability to colonize, persist, and invade the host. Biofilm formation represents the most important virulence trait [49], enabling chronic and persistent infections that are highly tolerant to antimicrobials and host defenses [50]. Additionally, enzymatic virulence factors are essential for the progress of infectious processes. Our results showed that almost all P. aeruginosa tested strains isolated from all sources produced lipases and lecithinases, which are pore-forming toxins [51]. Correlating virulence profiles and infection clinical outcomes could be very useful for setting up efficient preventive and therapeutic procedures for patients with positive P. aeruginosa cultures [51].
The pathogenicity of P. aeruginosa is multifactorial; the detection of different virulence genes in P. aeruginosa isolates suggests that they can be linked with different levels of inherent virulence and their propensity to cause infection. However, it is worth noting that virulence and other phenotypic traits such as resistance genes can contribute to the survival of organisms, as well as in disease spread and severity.
In the context of increasing antibiotic resistance rates, alternative antimicrobial strategies are urgently needed [52]. One efficient approach is to utilize agents to modulate bacterial virulence (i.e., production of virulence enzymes, toxins, and biofilm formation ability) without interfering with microbial fitness [53]. This approach would contribute to lower the rates of selecting resistant mutants, because most of the virulence factors needed for bacteria in the progression of the infectious process are not vital molecules [54]. Therefore, their modulation would not impact the population fitness, which is one of the major triggers for resistant mutant selection [55,56,57]. In our study we demonstrate that the tested EOs could impact P. aeruginosa biofilm formation when utilized in subinhibitory concentrations. This aspect could be further exploited in investigating natural factors to control this important virulence factor and control the infectious process and transition to persistent diseases. The fact that the MBICs of thyme and tea tree EOs are very reduced (ranging from 0.051% to 0.028%) in both susceptible and antibiotic-resistant strains, present in wastewaters and clinical samples, serves to suggest these natural products as efficient alternatives in the management of resistant P. aeruginosa infections. Our results concluded that EOs are effective, ecological, and sustainable alternatives, which could be used for the control of various infections, including those involving resistant pathogens.

4. Materials and Methods

4.1. Microbial Strains and Growth Conditions

Pseudomonas aeruginosa ATCC 27853 and PAO1 strains were purchased from American Type Culture Collection (ATCC, Manassas, Virginia, US), and 34 strains of P. aeruginosa were isolated from intrahospital infections (HAIs) (n = 7), hospital sewage tanks from two big hospital units from Bucharest encoded 1 and 2 (n = 9), and municipal (urban) wastewaters (n = 18) from Bucharest, Romania, during 2018–2020 (Table 1). All strains used were identified by the Maldi-Tof Mass Spectrometry method, both those from hospital infections and those isolated from wastewater. Strains from hospital sewage tanks and urban wastewater were isolated by the membrane filtration method using chromogenic culture media supplemented with antibiotic (ChromID ESBL agar and ChromID CARBA agar; BioMérieux SA, Marcy-l’Etoile, France) for the selection of antibiotic-resistant P. aeruginosa strains.
Bacteria strains are maintained as glycerol stocks in the Laboratory of Microbiology, Faculty of Biology, and University of Bucharest, Romania. Glycerol stocks were streaked on nutritive agar to obtain fresh cultures to be used for all further studies.

4.2. Essential Oils (EOs)

The study was performed employing the following four EOs: thyme (Thymus vulgaris), eucalyptus (Eucalyptus globulus Labillardiere), sage (Salvia officinalis), and tea tree (Tea Melaleuca alternifolia). The EOs were diluted as 1% stock solutions in dimethylsulfoxide (DMSO). Additionally, DMSO was used as a control for the experiments. The EOs were obtain from a Romanian distributor (Hofigal, https://hofigal.eu/, accessed on 24 July 2022), and all the other chemical used were purchased from Sigma–Aldrich Co., Baden-Württemberg, Germany, through MERCK, Romania.

4.3. Characterization of Microbial Strains

4.3.1. Antibiotic Susceptibility (Disc Diffusion Kirby Bauer Method)

The phenotypic susceptibility assay was performed by the disc-diffusion method, as recommended by CLSI 2020 [58] for P. aeruginosa.

4.3.2. Evaluation of Soluble Enzymatic Factors Implicated in Bacterial Virulence

The expression of the soluble virulence factors was highlighted by biochemical tests, using specific culture media, with built-in substrate, following a protocol previously reported by Georgescu et al. (2016) [50] and Preda et al. (2021) [31].
A number of eight soluble virulence factors was analyzed: amylase, lecithinase, lipase, caseinase, gelatinase, DNase, hemolysin, and esculin hydrolysis.
Briefly, bacterial strains were seeded on agar containing the substrate for each of the mentioned enzyme, and the as-prepared Petri dishes were incubated for 24–48 h at 37 °C. After incubation, results were read as a visible manifestation of the substrate (i.e., hemolysis, precipitate, halo, color change) appearing around the inoculated bacteria colonies.

4.3.3. Molecular Characterization of The Strains

DNA Extraction

The DNA from P. aeruginosa strains was extracted using an adapted alkaline extraction method. More specifically, 1–5 colonies of bacterial cultures were suspended in 1.5 mL tubes containing 20 µL solution of NaOH 0.05 M (sodium hydroxide) and SDS 0.25% (sodium dodecyl sulphate) and heated on a thermoblock at 95 °C for 15 min for the permeabilization of the bacterial cell wall. The following step was the addition of 180 μL of TE buffer (TRIS + EDTA) 1X and centrifugation at 13,000 rpm for 3 min. After centrifugation, the supernatant was taken in a new tube and store at −20 °C until use [31,50].

PCR for Virulence and Antibiotic Resistance Genes Identification

Genomic DNA was used as a template for the PCR screening of 7 virulence genes encoding for protease IV (TCF/TCR), three exoenzymes (exoS, exoT, exoU), two phospholipases (plcH (hemolytic phospholipase C), plcN (non-hemolytic phospholipase C), and for alginate (algD), and also for β-lactam (ESBL and carpapenemase encoding genes), aminoglycosides, and quinolone resistance. Primers and amplification programs are presented in Table 2, Table 3, Table 4 and Table 5 [39,42,43]. The amplification products were visualized by electrophoresis on a 1% agarose gel stained with the SYBRSafe DNA and specific weight marker (100pb, Ladder, ThermoScientific, Bucharest, Romania).

4.4. Antimicrobial Evaluation of the EOs

4.4.1. Antimicrobial Qualitative Assessment

The in vitro qualitative screening of the antimicrobial activity of the tested EOs was performed by an adapted agar diffusion technique (as recommended by CLSI, 2020) as previously described [59]. Briefly, we obtained a bacterial suspension of 0.5 McFarland density (1.5 × 108 CFU (colony forming units)/mL) in PBS (phosphate buffered saline) from each P. aeruginosa fresh culture developed on nutritive agar. Microbial suspensions were utilized to swab inoculate Mueller Hinton agar distributed in 90 mm diameter Petri dishes. Five microliters of each EO, prepared as a 1% solution in DMSO (dimethyl sulfoxide), was drop-added on the P. aeruginosa-inoculated Petri dishes and then incubated for 16–18 h at 37 °C.
The results were explained by measuring the diameters of the inhibitory regions that appear around the EOs and denoting them as "mm".

4.4.2. Minimum Inhibitory Concentration (MIC) Evaluation (Microdilution Method)

For establishing the MIC (minimum inhibitory concentration) values of the evaluated EOs, we utilized a microdilution method performed in nutritive broth. The sterile broth was added in sterile 96 well plates, and binary dilutions of each tested EO were performed in a final volume of 150 μL. After realizing the binary dilutions, 15 μL of microbial suspension adjusted to an optical density of 0.5 McFarland (1.5 × 108 CFU/mL) was added in each well. The MIC values were established by naked eye analysis and spectrophotometric measurement (Absorbance at 600 nm).

4.4.3. Biofilm Formation and MBEC (Minimum Biofilm Eradication Concentrations) of the EOs

Monospecific biofilm formation was evaluated by the crystal violet microdilution method. Briefly, 96-multi well plastic plates containing binary dilutions of the tested EOs in a final volume of 150 µL in nutritive broth were inoculated with 15 µL microbial suspensions of 108 CFU/mL prepared in sterile saline. The as-prepared plates were incubated for 24 h at 37 °C. After incubation, the wells were discarded, washed with PBS to remove unattached cells, and the bacterial cells adhered to the plastic walls were stained by 1% violet crystal solution for 15 min. The colored biofilm was thereafter fixed by cold methanol for 5 min and resuspended by 33% acetic acid solution. The Abs at 490 nm of the blue suspension was measured using an ELISA reader. The obtained absorbance values were proportional with the intensity of biofilm formation on the wells of the plates. MBICs (minimum biofilm inhibitory concentrations) were established by dividing the average Abs 490 value for untreated control by two. The Abs 490 nm/2 of the untreated control represents the reference value for the calculation of MBEC for each EO. Therefore, MBECs for each of the tested EOs was established as the lowest concentration that gives an Abs 490 nm value corresponding to (less than) half of the Abs 490 nm value of the untreated control.
Each experiment was performed in triplicate and repeated on at least three separate occasions.

4.5. Statistical Analysis of the Results

Statistical analysis was performed in Minitab® Statistical Software, with a significance level of p < 0.05 (95% confidence level). In order to be able to calculate the significance in two sample-paired t-tests, all MIC and MBIC values that were not quantifiable (>50%) were considered as the next two-fold dilution factor (100%).

5. Conclusions

This study reports on virulence and resistance characterization of recently isolated P. aeruginosa strains obtained from areas of concern in Romania, such as hospital infections, hospital sewage, and wastewater treatment plants. Although profiling of antibiotic resistance and virulence factors has been done in the past, constant updating is needed for learning about virulence changes and the occurrence of resistant strains in this highly versatile opportunistic pathogen.
Since antibiotic-resistant P. aeruginosa represents one of the most common etiologies in Romanian intrahospital infections, finding efficient alternatives in fighting such infections is a national and international priority.
Our study has shown that two essential oils, namely, tea tree and thyme, are particularly effective against P. aeruginosa strains, regardless of their isolation source and antibiotic resistance. These EOs showed much-reduced MIC values and were able to inhibit biofilm formation when utilized in sub-inhibitory concentrations.
Although preliminary, our results reveal that plant-derived EOs could be efficiently utilized for the therapy of P. aeruginosa infections, covering susceptible and resistant strains, as well as planktonic- or biofilm-embedded bacteria. This study supports the idea that natural plant compounds and EOs are a resourceful choice for the development of efficient antimicrobials, which could be intensively exploited by future studies to develop efficient and ecologic antimicrobials, based on natural products, such as plant extracts and EOs.

Author Contributions

Conceptualization, V.L., T.E.S., and A.-M.H.; methodology, F.M., C.C., L.M.D., I.G., A.P., and L.T.V.; software, I.H.; validation, L.T.V. and A.-M.H.; formal analysis, L.T.V.; investigation, A.-M.H. and F.M.; data curation, L.T.V., I.G., C.C., A.P., L.M.D., and F.M.; writing—original draft preparation, L.T.V., C.C., and I.G.; writing—review and editing, T.E.S. and A.-M.H.; supervision, V.L. and T.E.S. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Acknowledgments

The financial support of the Research Projects PN-III-P4-ID-PCCF-2016-0114 and PN-III-P1.1-TE-2021-1515 (TE 112/2022) awarded by UEFISCDI is gratefully acknowledged. The funding had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Todar, K. Todar’s online textbook of bacteriology. Univ. Wis.-Madison Dep. Bacteriol. 2008. Available online: https://www.worldcat.org/title/todars-online-textbook-of-bacteriology/oclc/846984317?referer=di&ht=edition (accessed on 24 July 2022).
  2. Khalifa, A.B.H.; Moissenet, D.; Thien, H.V.; Khedher, M. Virulence factors in Pseudomonas aeruginosa: Mechanisms and modes of regulation. Ann. Biol. Clin. 2011, 69, 393–403. [Google Scholar]
  3. Chifiriuc, M.C.; Savin, I.; Lazar, V. Virulence markers of Escherichia coli strains isolated from hospital and poultry wastewater. Biointerface Res. Appl. Chem. 2013, 3, 579–587. [Google Scholar]
  4. Pruden, A.; Pei, R.; Storteboom, H.; Carlson, K.H. Carlson, Antibiotic resistance genes as emerging contaminants: Studies in Northern Colorado. Environ. Sci. Technol. 2006, 40, 7445–7450. [Google Scholar] [CrossRef]
  5. Nagulapally, S.R.; Ahmad, A.; Henry, A.; Marchin, G.L.; Zurek, L.; Bhandari, A. Occurrence of ciprofloxacin, trimethoprim sulfamethoxazole, and vancomycin-resistant bacteria in a municipal wastewater treatment plant. Water Environ. Res. 2009, 81, 82–90. [Google Scholar] [CrossRef]
  6. Zhang, T.; Zhang, M.; Zhang, X.; Fang, H.H. Tetracycline resistance genes and tetracycline resistant lactose fermenting Enterobacteriaceae in activated sludge of sewage treatment plants. Environ. Sci. Technol. 2009, 43, 3455–3460. [Google Scholar] [CrossRef] [PubMed]
  7. Savin, M.; Alexander, J.; Bierbaum, G.; Hammerl, J.A.; Hembach, N.; Schwartz, T.; Schmithausen, R.M.; Sib, E.; Voigt, A.; Kreyenschmidt, J. Antibiotic-resistant bacteria, antibiotic resistance genes, and antibiotic residues in wastewater from a poultry slaughterhouse after conventional and advanced treatments. Sci. Rep. 2021, 11, 16622. [Google Scholar] [CrossRef] [PubMed]
  8. Karkman, A.; Do, T.T.; Walsh, F.; Virta, M.P. Antibiotic-Resistance Genes in Waste Water. Trends Microbiol. 2018, 26, 220–228. [Google Scholar] [CrossRef] [Green Version]
  9. Li, J.; Cheng, W.; Xu, L.; Strong, P.J.; Chen, H. Antibiotic-resistant genes and antibiotic-resistant bacteria in the effluent of urban residential areas, hospitals, and a municipal wastewater treatment plant system. Environ. Sci. Pollut. Res. Int. 2015, 22, 4587–4596. [Google Scholar] [CrossRef]
  10. Kunhikannan, S.; Thomas, C.J.; Franks, A.E.; Mahadevaiah, S.; Kumar, S.; Petrovski, S. Environmental hotspots for antibiotic resistance genes. Microbiol. Open 2021, 10, e1197. [Google Scholar] [CrossRef]
  11. Rajendrachari, S.; Ceylan, K.B. The activation energy and antibacterial investigation of spherical Fe3O4 nanoparticles prepared by Crocus sativus (Saffron) flowers. Biointerface Res. Appl. Chem. 2020, 10, 5951–5959. [Google Scholar]
  12. Tyagi, P.K.; Mishra, R.; Khan, F.; Gupta, D.; Gola, D. Antifungal Effects of Silver Nanoparticles Against Various Plant Pathogenic Fungi and its Safety Evaluation on Drosophila melanogaster. Biointerface Res. Appl. Chem. 2020, 10, 6587–6596. [Google Scholar]
  13. Zarif, M.E.; Yehia, S.A.; Biță, B.; Sătulu, V.; Vizireanu, S.; Dinescu, G.; Holban, A.M.; Marinescu, F.; Andronescu, E.; Grumezescu, A.M.; et al. Atmospheric pressure plasma activation of hydroxyapatite to improve fluoride incorporation and Modulate Bacterial Biofilm. Int. J. Mol. Sci. 2021, 22, 13103. [Google Scholar] [CrossRef] [PubMed]
  14. Pereira, V.; Dias, C.; Vasconcelos, M.; Rosa, E.; Saavedra, M. Antibacterial activity and synergistic effects between Eucalyptus globulus leaf residues (essential oils and extracts) and antibiotics against several isolates of respiratory tract infections (Pseudomonas aeruginosa). Ind. Crops Prod. 2014, 52, 1–7. [Google Scholar] [CrossRef]
  15. Ragno, R.; Papa, R.; Patsilinakos, A.; Vrenna, G.; Garzoli, S.; Tuccio, V.; Fiscarelli, E.; Selan, L.; Artini, M. Essential oils against bacterial isolates from cystic fibrosis patients by means of antimicrobial and unsupervised machine learning approaches. Sci. Rep. 2020, 10, 2653. [Google Scholar] [CrossRef] [PubMed]
  16. Winska, K.; Maczka, W.; Lyczko, J.; Grabarczyk, M.; Czubaszek, A.; Szumny, A. Essential Oils as Antimicrobial Agents-Myth or Real Alternative? Molecules 2019, 24, 2130. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  17. Davis, S.C.; Perez, R. Cosmeceuticals and natural products: Wound healing. Clin. Dermatol. 2009, 27, 502–506. [Google Scholar] [CrossRef]
  18. Ait-Ouazzou, A.; Loran, S.; Bakkali, M.; Laglaoui, A.; Rota, C.; Herrera, A.; Pagan, R.; Conchello, P. Chemical composition and antimicrobial activity of essential oils of Thymus algeriensis, Eucalyptus globulus and Rosmarinus officinalis from Morocco. Sci. Food Agric. 2011, 91, 2643–2651. [Google Scholar] [CrossRef]
  19. Nazarparvar, M.; Shako, A.; Ranjbariyan, A. Chemical Composition and Antimicrobial Activity Against Food Poisoning of Alcoholic Extract of Nigella Sativa L. Biointerface Res. Appl. Chem. 2020, 10, 6991–7001. [Google Scholar]
  20. Zaïri, A.; Nouir, S.; Khalifa, M.A.; Ouni, B.; Haddad, H.; Khelifa, A.; Trabelsi, M. Phytochemical Analysis and Assessment of Biological Properties of Essential Oils Obtained from Thyme and Rosmarinus Species. Current pharmaceutical biotechnology. 2020, 21, 414–424. [Google Scholar] [CrossRef]
  21. Mayaud, L.; Carricajo, A.; Zhiri, A.; Aubert, G. Comparison of bacteriostatic and bactericidal activity of 13 essential oils against strains with varying sensitivity to antibiotics. Lett. Appl. Microbiol. 2008, 47, 167–173. [Google Scholar] [CrossRef] [PubMed]
  22. Bucková, M.; Puskarova, A.; Kalaszova, V.; Kisova, Z.; Pangallo, D. Essential oils against multidrug resistant gram-negative bacteria. Biologia 2018, 73, 803–808. [Google Scholar] [CrossRef]
  23. Ghavam, M.; Manca, M.L.; Manconi, M.; Bacchetta, G. Chemical composition and antimicrobial activity of essential oils obtained from leaves and flowers of Salvia hydrangea DC. ex Benth. Sci. Rep. 2020, 10, 15647. [Google Scholar] [CrossRef] [PubMed]
  24. Aljabeili, H.S.; Barakat, H.; Abdel-Rahman, H.A. Chemical Composition, Antibacterial and Antioxidant Activities of Thyme Essential Oil (Thymus vulgaris). Food Nutr. Sci. 2018, 9, 433–446. [Google Scholar] [CrossRef] [Green Version]
  25. Cai, C.C.; Ma, R.J.; Duan, M.W.; Lu, D.J. Preparation and antimicrobial activity of thyme essential oil microcapsules prepared with gum Arabic. RSC Adv. 2019, 9, 19740–19747. [Google Scholar] [CrossRef] [Green Version]
  26. Ilie, C.; Golet, I.; Craciunescu, M.; Hogea, E.; Popescu, R.; Popescu, R. Florin George Horhat. Chemical Composition and Antimicrobial Activity of Essential Oil of Western Romanian Salvia officinalis. Rev. Chim. 2016, 67, 1. [Google Scholar]
  27. Yazgan, H. Investigation of antimicrobial properties of sage essential oil and its nanoemulsion as antimicrobial agent. LWT-Food Sci. Technol. 2020, 130, 109669. [Google Scholar] [CrossRef]
  28. Bachir, R.G.; Benali, M. Antibacterial activity of the essential oils from the leaves of Eucalyptus globulus against Escherichia coli and Staphylococcus aureus. Asian Pac. J. Trop. Biomed. 2012, 2, 739–742. [Google Scholar] [CrossRef] [Green Version]
  29. Gasser, V.; Kuhn, L.; Hubert, T.; Aussel, L.; Hammann, P.; Schalk, I.J. The Esterase PfeE, the Achilles Heel in the Battle for Iron between Pseudomonas aeruginosa and Escherichia coli. Int. J. Mol. Sci. 2021, 22, 2814. [Google Scholar] [CrossRef]
  30. Cheng, R.A.; Eade, C.R.; Wiedmann, M. Differences in virulence mechanisms, disease severity and host adaptations contribute to the success of nontyphoidal salmonella as a foodborne pathogen. Front. Microbiol. 2019, 10, 1368. [Google Scholar] [CrossRef] [Green Version]
  31. Preda, M.; Mihai, M.M.; Popa, L.I.; Dițu, L.-M.; Holban, A.M.; Manolescu, L.S.C.; Popa, G.-L.; Muntean, A.-A.; Gheorghe, I.; Chifiriuc, C.M.; et al. Phenotypic and genotypic virulence features of staphylococcal strains isolated from difficult-to-treat skin and soft tissue infections. PLoS ONE 2021, 16, e0246478. [Google Scholar] [CrossRef] [PubMed]
  32. Mohammed, H.B.; Rayyif, S.M.I.; Curutiu, C.; Birca, A.C.; Oprea, O.C.; Grumezescu, A.M.; Ditu, L.-M.; Gheorghe, I.; Chifiriuc, M.C.; Mihaescu, G.; et al. Eugenol functionalized magnetite nanoparticles modulate virulence and persistence in Pseudomonas aeruginosa clinical strains. Molecules 2021, 26, 2189. [Google Scholar] [CrossRef] [PubMed]
  33. Trojanowska, D.; Mrowiec, P.; Soja, Ł.; Budak, A. Activity of Thyme oil (Oleum thumi) against multidrug resistant Acinetobacter baumannii and Pseudomonas aeruginosa. Acta Pol. Pharmaceutica. Drug Res. 2016, 73, 975–981. [Google Scholar]
  34. Moradali, M.F.; Ghods, S.; Rehm, B.H.A. Pseudomonas aeruginosa Lifestyle: A Paradigm for adaptation, survival, and persistence. Front. Cell. Infect. Microbiol. 2017, 7, 39. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  35. Cotar, A.-I.; Chifiriuc, M.-C.; Dinu, S.; Bucur, M.; Iordache, C.; Banu, O.; Dracea, O.; Larion, C.; Lazar, V. Screening of Molecular Virulence Markers in Staphylococcus aureus and Pseudomonas aeruginosa Strains Isolated from Clinical Infections. Int. J. Mol. Sci. 2010, 11, 5273–5291. [Google Scholar] [CrossRef] [Green Version]
  36. Hamad, M.N.F.; Marrez, D.A.; El-Sherbieny, S.M.R. Toxicity Evaluation and Antimicrobial Activity of Purified Pyocyanin from Pseudomonas aeruginosa. Biointerface Res. Appl. Chem. 2020, 10, 6974–6990. [Google Scholar]
  37. Gheorghe, I.; Novais, A.; Grosso, F.; Rodrigues, C.; Chifiriuc, M.C.; Lazar, V.; Peixe, L. Snapshot on carbapenemase-producing Pseudomonas aeruginosa and Acinetobacter baumannii in Bucharest hospitals reveals unusual clones and novel genetic surroundings for blaOXA-23. J. Antimicrob. Chemother. 2015, 70, 1016–1020. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  38. Saviuc, C.; Gheorghe, I.; Coban, S.; Drumea, V.; Chifiriuc, M.C.; Banu, O.; Bezirtzoglou, E.; Lazăr, V. Rosmarinus officinalis essential oil and eucalyptol act as efflux pumps inhibitors and increase ciprofloxacin efficiency against Pseudomonas aeruginosa and Acinetobacter baumannii MDR Strains. Rom. Biotechnol. Lett. 2016, 21, 11782–11790. [Google Scholar]
  39. Gheorghe, I.; Czobor, I.; Chifiriuc, M.C.; Borcan, E.; Ghiță, C.; Banu, O.; Lazăr, V.; Mihăescu, G.; Mihăilescu, D.F.; Zhiyong, Z. Molecular screening of carbapenemase-producing Gram negative strains in Roumanian intensive care units during one year survey. J. Med. Microbiol. 2014, 63, 1303–1310. [Google Scholar] [CrossRef]
  40. Buzilă, E.R.; Năstase, E.V.; Luncă, C.; Bădescu, A.; Miftode, E.; Iancu, L.S. Antibiotic resistance of non-fermenting Gram-negative bacilli isolated at a large infectious diseases hospital in north-eastern Romania, during an 11-year period. Germs 2021, 11, 354–362. [Google Scholar] [CrossRef]
  41. Tatu, L.A.; Merezeanu, N.; Pantea, O.; Gheorghe, I.; Popa, M.; Banu, O.; Cristea, V.C.; Carmen, M.C.; Chifiriuc, M.C.; Lazar, V.; et al. Resistance features of Pseudomonas aeruginosa strains isolated from patients with infectious complications of cardiovascular surgery. Biointerface Res. Appl. Chem. 2017, 7, 2004–2008. [Google Scholar]
  42. Porumbel, I.; Gheorghe, I.; Popa, M.; Banu, O.; Lazăr, V.; Chifiriuc, M.C. Virulence and resistance profiles among Pseudomonas aeruginosa isolated from patients with cardiovascular diseases in Bucharest. Biointerface Res. Appl. Chem. 2016, 6, 1176–1179. [Google Scholar]
  43. Merezeanu, N.; Gheorghe, I.; Popa, M.; Chifiriuc, M.C.; Pântea, O.; Lazăr, V.; Banu, O.; Bolocan, A.; Grigore, R.; Berteșteanu, S.V. Virulence and resistance features of Pseudomonas aeruginosa strains isolated from patients with cardiovascular diseases. Biointerface Res. Appl. Chem. 2016, 6, 1117–1121. [Google Scholar]
  44. Dortet, L.; Flonta, M.; Boudehen, Y.-M.; Creton, E.; Bernabeu, S.; Vogel, A.; Naas, T. Dissemination of carbapenemase-producing Enterobacteriaceae and Pseudomonas aeruginosa in Romania. Antimicrob. Agents Chemother. 2015, 59, 7100–7103. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  45. Cardoso, A.Á.; de Lima e Borges, E.E.; de Souza, G.A.; da Silva, C.J.; Pires, R.M.O.; Dias, D.C.F.S. Seed Imbibition and Germination of Plothymenia Reticulata Benth. (Fabaceae) Affected by Mercury: Possible Role of Aquaporins. Acta Bot. Bras. 2015, 29, 285–291. [Google Scholar] [CrossRef] [Green Version]
  46. Mereuță, A.I.; Bădescu, A.C.; Dorneanu, O.S.; Iancu, L.S.; Tuchiluș, C.G. Spread of VIM-2 metallo-beta-lactamase in Pseudomonas aeruginosa and Acinetobacter baumannii clinical isolates from Iași, România. Rev. Română Med. Lab. 2013, 21, 389–396. [Google Scholar] [CrossRef] [Green Version]
  47. Pokharel, K.; Dawadi, B.R.; Bhatt, C.P.; Gupte, S. Prevalence of Pseudomonas aeruginosa and its Antibiotic Sensitivity Pattern. J. Nepal Health 2019, 17, 109–113. [Google Scholar] [CrossRef]
  48. Abdelghany, A.M.; Ayaad, D.M.; Mahmoud, S.M. Antibacterial and Energy Gap Correlation of PVA/SA Biofilms Doped With Selenium Nanoparticles. Biointerface Res. Appl. Chem. 2020, 10, 6280–6288. [Google Scholar]
  49. Ciofu, O.; Tolker-Nielsen, T. Tolerance and Resistance of Pseudomonas aeruginosa Biofilms to Antimicrobial Agents-How Paeruginosa Can Escape Antibiotics. Front. Microbiol. 2019, 10, 913. [Google Scholar] [CrossRef] [Green Version]
  50. Georgescu, M.; Gheorghe, I.; Curutiu, C.; Lazar, V.; Bleotu, C.; Chifiriuc, M.C. Virulence and resistance features of Pseudomonas aeruginosa strains isolated from chronic leg ulcers. BMC Infect. Dis. 2016, 16 (Suppl. 1), 92. [Google Scholar] [CrossRef] [Green Version]
  51. Mudarisova, R.; Kukovinets, O.; Sagitova, A.; Novoselov, I. Intermolecular interactions of apple pectin modified by pharmacophores with iodine and antimicrobial activity of iodine-containing pectin materials. Biointerface Res. Appl. Chem. 2020, 10, 5724–5732. [Google Scholar]
  52. Zhao, X.; Yu, Z.; Ding, T. Quorum-Sensing Regulation of Antimicrobial Resistance in Bacteria. Microorganisms 2020, 8, 425. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  53. Foulkes, D.M.; McLean, K.; Haneef, A.S.; Fernig, D.G.; Winstanley, C.; Berry, N.; Kaye, S.B. Pseudomonas aeruginosa toxin ExoU as a therapeutic target in the treatment of bacterial infections. Microorganisms 2019, 7, 707. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Sharma, A.K.; Dhasmana, N.; Dubey, N.; Kumar, N.; Gangwal, A.; Gupta, M.; Singh, Y. Bacterial Virulence Factors: Secreted for Survival. Indian J. Microbiol. 2017, 57, 1–10. [Google Scholar] [CrossRef] [Green Version]
  55. Nolan, C.; Behrends, V. Sub-Inhibitory Antibiotic Exposure and Virulence in Pseudomonas aeruginosa. Antibiotics 2021, 10, 139. [Google Scholar] [CrossRef] [PubMed]
  56. Schroeder, M.; Brooks, B.D.; Brooks, A.E. The Complex Relationship between Virulence and Antibiotic Resistance. Genes 2017, 8, 39. [Google Scholar] [CrossRef] [Green Version]
  57. Millar, J.A.; Raghavan, R. Modulation of Bacterial Fitness and Virulence Through Antisense RNAs. Front Cell. Infect. Microbiol. 2021, 10, 596277. [Google Scholar] [CrossRef]
  58. https://clsi.org/. CLSI. Performance Standards for Antimicrobial Susceptibility Testing, 30th ed.; CLSI Supplement M100; Clinical and Laboratory Standards Institute: Wayne, PA, USA, 2020. [Google Scholar]
  59. Prodan, A.M.; Andronescu, E.; Truşcă, R.; Beuran, M.; Iconaru, S.L.; Barna, E.Ş.; Chifiriuc, M.; Marutescu, L. Anti-biofilm activity of dextran coated iron oxide nanoparticles. Univ. Politeh. Buchar. Sci. Bull. Ser. B Chem. Mater. Sci 2014, 76, 81–90. [Google Scholar]
Plants 11 02003 g001 550
Figure 1. Results of antibiotic resistance of Pseudomonas aeruginosa-evaluated strains (n = 36). (TZP: piperacillin/tazobactam; CAZ: ceftazidime; FEP: cefepime; ATM: aztreonam; MEM: meropenem; IPM: imipenem; TOB: tobramycin; AK: amikacin; CIP: ciprofloxacin; DOR: doripenem; CN: gentamicin).
Figure 1. Results of antibiotic resistance of Pseudomonas aeruginosa-evaluated strains (n = 36). (TZP: piperacillin/tazobactam; CAZ: ceftazidime; FEP: cefepime; ATM: aztreonam; MEM: meropenem; IPM: imipenem; TOB: tobramycin; AK: amikacin; CIP: ciprofloxacin; DOR: doripenem; CN: gentamicin).
Plants 11 02003 g001
Plants 11 02003 g002 550
Figure 2. The antibiotic resistance levels (%) of P. aeruginosa strains depending on their isolation sources.
Figure 2. The antibiotic resistance levels (%) of P. aeruginosa strains depending on their isolation sources.
Plants 11 02003 g002
Plants 11 02003 g003 550
Figure 3. The carbapanamase- and ESBL-encoding genes (%) expressed by P. aeruginosa strains.
Figure 3. The carbapanamase- and ESBL-encoding genes (%) expressed by P. aeruginosa strains.
Plants 11 02003 g003
Plants 11 02003 g004 550
Figure 4. Graphic representation of the distribution of soluble virulence factors expressed by the P. aeruginosa strains. The graph represents the percentage (%) of positive strains for each of the evaluated virulence factor depending on their isolation source.
Figure 4. Graphic representation of the distribution of soluble virulence factors expressed by the P. aeruginosa strains. The graph represents the percentage (%) of positive strains for each of the evaluated virulence factor depending on their isolation source.
Plants 11 02003 g004
Plants 11 02003 g005 550
Figure 5. Graphic representation of the distribution of virulence-encoding genes expressed by the P. aeruginosa strains.
Figure 5. Graphic representation of the distribution of virulence-encoding genes expressed by the P. aeruginosa strains.
Plants 11 02003 g005
Plants 11 02003 g006 550
Figure 6. Graphic representation of the average inhibition diameter zone of the EOs in P. aeruginosa strains, regardless their isolation source.
Figure 6. Graphic representation of the average inhibition diameter zone of the EOs in P. aeruginosa strains, regardless their isolation source.
Plants 11 02003 g006
Plants 11 02003 g007 550
Figure 7. Average minimum inhibitory concentration (MIC) values, expressed as percent (%) concentration of the EOs in P. aeruginosa strains, regardless of their isolation source.
Figure 7. Average minimum inhibitory concentration (MIC) values, expressed as percent (%) concentration of the EOs in P. aeruginosa strains, regardless of their isolation source.
Plants 11 02003 g007
Plants 11 02003 g008 550
Figure 8. The average MIC values of P. aeruginosa strains depending on their isolation source (MICs are expressed as percentage (%) oil in the culturing media).
Figure 8. The average MIC values of P. aeruginosa strains depending on their isolation source (MICs are expressed as percentage (%) oil in the culturing media).
Plants 11 02003 g008
Plants 11 02003 g009a 550Plants 11 02003 g009b 550
Figure 9. Charts showing comparison of the MIC values for the tested EOs among P. aeruginosa strains isolated from different sources: (a) comparison of EO MICs across hospital strains, (b) comparison of EO MICs across wastewater hospital, (c) comparison of EO MICs across wastewater urban; p < 0.05 when comparing average MICs of thyme or/and tea tree EOs to sage or/and eucalyptus.
Figure 9. Charts showing comparison of the MIC values for the tested EOs among P. aeruginosa strains isolated from different sources: (a) comparison of EO MICs across hospital strains, (b) comparison of EO MICs across wastewater hospital, (c) comparison of EO MICs across wastewater urban; p < 0.05 when comparing average MICs of thyme or/and tea tree EOs to sage or/and eucalyptus.
Plants 11 02003 g009aPlants 11 02003 g009b
Plants 11 02003 g010 550
Figure 10. Comparison of the average MBICs of tea tree and thyme EOs (%) in all the analyzed P. aeruginosa strains. p < 0.05 when comparing average MBICs of thyme to tea tree EOs.
Figure 10. Comparison of the average MBICs of tea tree and thyme EOs (%) in all the analyzed P. aeruginosa strains. p < 0.05 when comparing average MBICs of thyme to tea tree EOs.
Plants 11 02003 g010
Plants 11 02003 g011 550
Figure 11. Comparison of the MBICs of thyme and tea tree EOs in P. aeruginosa strains depending on their isolation source.
Figure 11. Comparison of the MBICs of thyme and tea tree EOs in P. aeruginosa strains depending on their isolation source.
Plants 11 02003 g011
Plants 11 02003 g012 550
Figure 12. The chart of MBEC values of essential oils from different sources: (a) comparison of MBEC values of tea tree from different sources, (b) comparison of MBEC values of thyme from different sources.
Figure 12. The chart of MBEC values of essential oils from different sources: (a) comparison of MBEC values of tea tree from different sources, (b) comparison of MBEC values of thyme from different sources.
Plants 11 02003 g012
Plants 11 02003 g013 550
Figure 13. Aspect of 96 well plates after CV staining in cases of 2 P. aeruginosa strains treated with EOs (last column (12)—sterility control, the row before (11)—growth control).
Figure 13. Aspect of 96 well plates after CV staining in cases of 2 P. aeruginosa strains treated with EOs (last column (12)—sterility control, the row before (11)—growth control).
Plants 11 02003 g013
Table
Table 1. P. aeruginosa strain codes and isolation sources.
Table 1. P. aeruginosa strain codes and isolation sources.
Strain codeSourceStrain codeSource
Ps. 1ATCC 27853, USPs. 13urban wastewater
Ps. 2PAO1, ATCC, USPs. 14urban wastewater
Ps. 3intrahospital infectionPs. 15urban wastewater
Ps. 4intrahospital infectionPs. 16urban wastewater
Ps. 5intrahospital infectionPs. 17urban wastewater
Ps. 6intrahospital infectionPs. 18urban wastewater
Ps. 7intrahospital infectionPs. 19urban wastewater
Ps. 8intrahospital infectionPs. 20urban wastewater
Ps. 12intrahospital infectionPs. 25urban wastewater
Ps. 9hospital sewagePs. 26urban wastewater
Ps. 10hospital sewagePs. 27urban wastewater
Ps. 11hospital sewagePs. 28urban wastewater
Ps. 21hospital sewagePs. 29urban wastewater
Ps. 22hospital sewagePs. 30urban wastewater
Ps. 23hospital sewagePs. 31urban wastewater
Ps. 24hospital sewagePs. 32urban wastewater
Ps. 33hospital sewagePs. 34urban wastewater
Ps. 36hospital sewagePs. 35urban wastewater
Table
Table 2. Primers for virulence gene detection.
Table 2. Primers for virulence gene detection.
Virulence GenesPrimersPrimer SequencesAmplicon Size (bp)
plcHplcH-F
plcH-R		5′ GAAGCCATGGGCTACTTCAA 3′
5′ AGAGTGACGAGGGGTAG 3′		466
plcNplcN-F
plcH-R		5′ GTTATCGCAACCAGCCCTAC 3′
5′ AGGTCGAACACCTGGAACAC 3′		307
exoUexoU-F
exoU-R		5′ CCGTTGTGGTGCCGTTGAAG 3′
5′ CCAGATGTTCACCGACTCG 3′		134
exoTexoT-F
exoT-R		5′ AATCGCCGTCCAACTGCATGCG 3′
5′ TGTTCGCCGAGGTACTGCTC 3′		152
exoSexoS-F
exoS-R		5′ ATCGCTTCAGCAGAGTCCGTC 3′
5′ CAGGCCAGATCAAGGCCGCGC 3′		1352
TCF/TCRTCF/R-F
TCF/R-R		5′ TATTTCGCCGACTCCCTGTA 3′
5′ GAATAGACGCCGCTGAAATC 3′		752
algDalgD-F
algD-R		5′ ATGCGAATCAGCATCTTTGGT 3′
5′ CTACCAGCAGATGCCCTCGGC 3′		1310
Table
Table 3. Amplification program for virulence genes.
Table 3. Amplification program for virulence genes.
GenesAmplification Program
DenaturationNo. of CyclesDenaturationAnnealingExtensionFinal Extension
plcH94 °C, 3 min3094 °C, 30 s55 °C, 1 min72 °C, 1.5 min72 °C, 5 min
plcN94 °C, 3 min3094 °C, 30 s55 °C, 1 min72 °C, 1.5 min72 °C, 5 min
exoU94 °C, 2 min3094 °C, 30 s59 °C, 30 s68 °C, 1 min68 °C, 7 min
exoT94 °C, 2 min3094 °C, 30 s59 °C, 30 s68 °C, 1 min68 °C, 7 min
exoS94 °C, 5 min3594 °C, 30 s65 °C, 30 s72 °C, 2 min72 °C, 7 min
TCF/TCR94 °C, 5 min3094 °C, 30 s60 °C, 30 s72 °C, 2 min72 °C, 5 min
algD90 °C, 3 min3094 °C, 30 s51 °C, 1 min72 °C, 1.5 min72 °C, 5 min
Table
Table 4. Primers for the detection of antibiotic resistance genes.
Table 4. Primers for the detection of antibiotic resistance genes.
ARGsPrimersPrimer SequencesAmplicon Size (bp)
blaVIMIMP-F
IMP-R		5′ GATGGTGTTTGGTCGCATA 3′
5′ CGAATGCGCAGCACCAG 3′		390
blaIMPIMP-F
IMP-R		5′ GGAATAGAGTGGCTTAAYTCTC 3′
5′ GGTTTAAYAAAACAACCACC 3′		232
blaTEMTEM-F
TEM-R		5′ ATAAAATTCTTGAAGACGAAA 3′
5′ GTCAGTTACCAATGCTTAATC 3′		1080
blaCTX-MCTX-M-F
CTX-M-R		5′ CGCTGTTGTTAGGAAGTGTG 3′
5′ GGCTGGGTGAAGTAAGTGAC 3′		754
blaSHVSHV-F
SHV-R		5′ TGGTTATGCGTTATATTCGCC 3′
5′ GGTTAGCGTTGCCAGTGCT 3′		868
blaVEBVEB-F
VEB-R		5′ CGACTTCCATTTCCCGATGC 3′
5′ GGACTCTGCAACAAATACGC 3′		390
blaGESGES-C
GES-D		5′ GTT TTG CAA TGT GCT CAA CG 3′
5′ TCG CAT AGC AAT AGG CGT AG 3′		371
blaPERPER-F
PER-R		5′ AATTTGGGCTTAGGGCAGAA 3′
5′ ATGAATGTCATTATAAAAGC 3′		925
blaNDMNDM-F
NDM-R		5′ GGTTTGGCGATCTGGTTTTC 3′
5′ CGGAATGGCTCATCACGATC 3′		621
aac-3-Iaaac-3-Ia F
aac-3-IaR		5′-ATGGGCATCATTCGCACA-3′
5′-TCTCGGCTTGAACGAATTGT-3′		484
qnrAqnrA-F
qnrA-R		5′ AGAGGATTTCTCACGCCAGG 3′
5′ TGCCAGGCACAGATCTTGAC		580
qnrBqnrB-F
qnrB-R		5′ GGMATHGAAATTCGCCACTG 3′
5′ TTTGCYGYYCGCCAGTCGAA		264
qnrSqnrS-F
qnrS-R		5′ GCAAGTTCATTGAACAGGGT 3′
5′ TCTAAACCGTCGAGTTCGGCG 3′		428
Table
Table 5. Amplification program for antibiotic resistance genes.
Table 5. Amplification program for antibiotic resistance genes.
GeneAmplification Program
DenaturationNo. of CyclesDenaturationAnnealingExtensionFinal Extension
blaVIM94 °C, 10 min3695 °C, 30 s55 °C, 40 s72 °C, 5 min72 °C, 10 min
blaIMP94 °C, 10 min3695 °C, 30 s55 °C, 40 s72 °C, 5 min
blaTEM95 °C, 5 min3095 °C, 30 s52 °C, 40 s72 °C, 70 s
blaCTX-M95 °C, 5 min3095 °C, 30 s56 °C, 40 s72 °C, 60 s
blaSHV95 °C, 5 min3095 °C, 30 s56 °C, 40 s72 °C, 60 s
blaVEB95 °C, 5 min3095 °C, 30 s56 °C, 40 s72 °C, 50 s
blaGES95 °C, 5 min3095 °C, 30 s56 °C, 40 s72 °C, 50 s
blaPER95 °C, 5 min3095 °C, 30 s50 °C, 40 s72 °C, 50 s
blaNDM95 °C, 5 min3095 °C, 30 s55 °C, 40 s72 °C, 50 s
aac-3-Ia94 °C, 5 min3594 °C, 30 s55 °C, 30 s72 °C, 1 min
qnrA95 °C, 10 min3095 °C, 30 s52 °C, 40 s72 °C, 5 min
qnrB95 °C, 10 min3095 °C, 1 min54 °C, 1 min72 °C, 10 min
qnrS94 °C, 10 min3095 °C, 30 s52 °C, 40 s72 °C, 5 min
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Van, L.T.; Hagiu, I.; Popovici, A.; Marinescu, F.; Gheorghe, I.; Curutiu, C.; Ditu, L.M.; Holban, A.-M.; Sesan, T.E.; Lazar, V. Antimicrobial Efficiency of Some Essential Oils in Antibiotic-Resistant Pseudomonas aeruginosa Isolates. Plants 2022, 11, 2003. https://doi.org/10.3390/plants11152003

AMA Style

Van LT, Hagiu I, Popovici A, Marinescu F, Gheorghe I, Curutiu C, Ditu LM, Holban A-M, Sesan TE, Lazar V. Antimicrobial Efficiency of Some Essential Oils in Antibiotic-Resistant Pseudomonas aeruginosa Isolates. Plants. 2022; 11(15):2003. https://doi.org/10.3390/plants11152003

Chicago/Turabian Style

Van, Luc Tran, Ilinca Hagiu, Adelina Popovici, Florica Marinescu, Irina Gheorghe, Carmen Curutiu, Lia Mara Ditu, Alina-Maria Holban, Tatiana Eugenia Sesan, and Veronica Lazar. 2022. "Antimicrobial Efficiency of Some Essential Oils in Antibiotic-Resistant Pseudomonas aeruginosa Isolates" Plants 11, no. 15: 2003. https://doi.org/10.3390/plants11152003

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop