Phytochemical characterization, and antioxidant and antimicrobial activities of essential oil from leaves of the common sage Salvia officinalis L. from Abha, Saudi Arabia

Leaves of S. officinalis L. were sampled from natural populations situated in Abha (southern region of Saudi Arabia; latitude: 18° 13′ 0.4692″ N, longitude: 42° 30′ 19.01″ E, altitude: 2,400 m) in winter–spring 2018. The plant is not considered an endangered or protected species, and no specific permit was required for its collection. Plants were identified by Professor Wissem Mnif (Faculty of Sciences and Arts in Balgarn, University of Bisha, Saudi Arabia), and voucher specimens (SO1) have been deposited in the Higher Institute of Applied Biology of Medenine (Tunisia). Samples were dried in the shade while ventilated at room temperature for 15 days. A disc mill machine (model FFC-45; Zhengzhou Kaishan Mechanical & Electrical Equipment Co., China) was used for decortication of the stem-free leaf samples.

The essential oil was extracted from the air-dried leaves (200 g) by hydrodistillation using a Clevenger-type apparatus (4 h). Petroleum ether was used to extract the aqueous phase (3 × 50 mL), then dried with anhydrous sodium sulfate. After filtration, the petroleum ether was removed using a rotary evaporator (model 94200; Heidolph Instruments, Germany) under reduced pressure to ensure its complete removal. Samples were stored in amber glass bottles (at 4 °C) until use [18]. The extracted oil was weighed to determine the yield as estimated based on the dry weight of the plant material.

The composition of the essential oil from leaves of Salvia officinalis L. was determined using a Hewlett Packard HP 5890 gas chromatography system coupled with an HP 5972 Mass Selective Detector (Agilent Technologies), set to scan from 20 m/z to 550 m/z. The gas chromatography-mass spectrometry (GC-MS) analysis was conducted using a DB-5 capillary column (length 25 m, internal diameter 0.25 mm, and 0.25 μm film thickness; Agilent Technologies). The carrier gas was helium at a linear velocity (u) of 30.1 cm/s with an inlet pressure of 99.8 kPa. The detector temperature was adjusted at 250 °C at a carrier gas flow rate of 1.2 mL/min. Feed oil with 1% hexane was injected in a split mode in a volume of 0.2 μL at 240 °C. The ionization energy was 70 eV. The GC-flame ionization detection (FID) peaks are indicative of various chemical constituents of the oil, which were recognized based on comparing their retention time and mass spectra with entries in the Wiley version 7.0 and National Institute of Standards and Technology 05 MS (NIST) spectral data libraries. The peak area normalization was applied to determine the component percentages [19]. The GC-MS method was validated under the supervision of the Tunisian Ministry of Research and Higher education. The precision and interassay variation were evaluated as >97%.

The essential oil phenol content was determined using a Folin–Ciocalteau reagent method as previously described [20]. In brief, the extracted essential oil was mixed with 10% Folin–Ciocalteau reagent in a ratio (10 μL:750 μL) and further mixed with 20% Na₂CO₃ solution. After 1 h of reaction at room temperature, the absorbance was determined at 765 nm with a spectrophotometer (Shimadzu). Gallic acid was used as a standard for the calibration curve, which is used to determine the total phenol content. Results are presented as μg/mL of the gallic acid equivalent (GAE).

To determine the flavonoid content, the essential oil was diluted in ethanol in a ratio of 1:10. Then, 75 μL of 5% NaOH solution was added to 250 μL of the diluted oil solution. After a few minutes of incubation, 150 μL of 7% AlCl₃ solution was added to the mixture. After 1 h of incubation at room temperature, the absorbance was determined at 510 nm using a spectrophotometer (Shimadzu). The flavonoid content was determined using a calibration curve of quercetin as a standard. Results were presented as μg/mL of the quercetin equivalent (QE) [21].

The essential oil antioxidant activity was determined using the 2,2′-diphenyl-1-picrylhydrazyl (DPPH) free radical scavenging method [22]. Essential oil was diluted to 1 mL with methanol (range between 0 μg/mL and 1,000 μg/mL), and 0.25 mL of DPPH radical solution (0.2 mM) was added [23]. The absorbance was determined at 517 nm using a spectrophotometer (Shimadzu). The antioxidant activity percentage was calculated as follows [18]: Inhibition (%)=(ODControl−ODSampleODControl)×100,{\rm{Inhibition}}\,\left( \% \right) = \left( {{{{\rm{O}}{{\rm{D}}_{{\rm{Control}}}} - {\rm{O}}{{\rm{D}}_{{\rm{Sample}}}}} \over {{\rm{O}}{{\rm{D}}_{{\rm{Control}}}}}}} \right) \times 100, where OD_Control is the absorbance of the control (DPPH solution plus methanol) and OD_Sample is the absorbance of the sample including DPPH plus the oil. The essential oil concentration that could scavenge 50% of DPPH (IC₅₀) was determined as specified by Shimada et al. [24].

We used 5 gram-positive bacteria (Enterococcus faecalis ATCC 29212, Micrococcus luteus ATCC 4698, Salmonella enterica ATCC 35664, Bacillus subtilis ATCC 6633, and Staphylococcus aureus ATCC 29213) and 5 gram-negative bacteria (Escherichia coli ATCC 8739, Klebsiella pneumoniae ATCC 10031, Shigella sonnei ATCC 29930, Agrobacterium tumefaciens ATCC 23308, and Pseudomonas aeruginosa ATCC 9027). Strains were grown on tryptone–casein–soy agar plates and cultivated at 37 °C for 24 h. All strains were obtained from the Medical Faculty of Sfax (Tunisia) and were isolated as reported in the literature [25].

A disk diffusion assay was applied to determine the antibacterial activities of the essential oil. We spread 100 μL of bacterial suspension (10⁸ colony-forming units (CFU)/mL) on trypticase soy agar contained in Petri plates. We individually impregnated 6 mm diameter paper disks (Sigma Aldrich) with 50 μL of the essential oil and placed the impregnated disks on the agar plate previously inoculated with the selected bacterial strain. Gentamicin (10 μg/disk) was used as a positive control. As a negative antibacterial control, a disk soaked in hexane was used. After incubating the plates with the disks for 1 h at 4 °C, they were then incubated at 37 °C for 24 h. For each bacterial strain, the antibacterial activity was evaluated by measuring the diameter (in millimeters) of the growth-inhibition zone [26].

The minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) for the essential oil were determined using an agar dilution method [27].

Experiments were conducted in triplicate, and results are presented as means ± standard deviation. The results were compared only for the antibacterial activity of the essential oil against various strains. Comparisons of means between groups were assessed using a one-way analysis of variance (ANOVA) with XLSTAT (version 10). Differences were considered significant at P < 0.05.

A yield of 3.24 ± 0.55% (weight/dry weight) of essential oil, with a colorless or yellowish color and a characteristic spicy odor, was obtained by extraction from S. officinalis leaves using hydrodistillation. The percentages and the retention times (RI) of the identified compounds of the essential oil are shown in Figure 1 and summarized in Table 1. A total of 26 constituents, representing 99.92% of the total oil, were identified. The essential oil was dominated by the oxygenated monoterpenoids representing a fraction of 67.36%. This fraction was composed of mainly camphor (20.3%), 1,8-cineole (15.01%), and α-thujone, with a lesser amount of carvone (6.2%), β-thujone (5.7%), and borneol (2.2%). Among the sesquiterpene fraction (22.85%), sesquiterpene hydrocarbons (11.58%) were dominated by trans-β-caryophyllene (4.0%) while viridiflorol (9.9%) was recognized as the major constituent of the sesquiterpene oxygenated portion. This essential oil also showed a low-oxygenated sesquiterpenoid content of humulene epoxide (0.86%).

Figure 1

The phenolic content was 134.3 ± 17.61 μg GAE/mL. The flavonoid content was 119.5 ± 18.75 μg QE/mL.

The essential oil had an antioxidant activity to scavenge DPPH free radicals with an IC₅₀ = 970 ± 5.5 μg/mL (Figure 2).

Figure 2

The essential oil inhibits both gram-positive (E. faecalis, M. luteus, S. enterica, B. subtilis, and S. aureus) and gram-negative bacteria (E. coli, K. pneumoniae, A. tumefaciens, and P. aeruginosa) (Table 2). For gram-positive bacteria, the highest inhibition diameter (24.8 ± 0.11 mm) was obtained for B. subtilis. This value was significantly higher (P < 0.001) than the lowest value (9.5 ± 0.47 mm) obtained for M. luteus. For gram-negative bacteria, the essential oil was highly active against E. coli ATCC 8739 and slightly active against A. tumefaciens with significantly different inhibition diameters of 25.2 ± 0.41 mm and 8.7 ± 0.28 mm, respectively. The MIC values of the essential oil ranged from 62.2 ± 3.9 μg/mL to 1398.1 ± 50.7 μg/mL for gram-positive bacteria and from 323.4 ± 69.5 μg/mL to 968.4 ± 120.6 μg/mL for gram-negative bacteria. The lowest MIC (most potent activity) was observed against B. subtilis (62.2 ± 3.9 μg/mL). The MBC values of essential oil ranged from 120.3 ± 7.6 μg/mL to 1387.4 ± 161.8 μg/mL for gram-positive bacteria and from 386 ± 8.3 μg/mL to 1225.2 ± 100.9 μg/mL gram-negative bacteria, respectively. The most potent activity (386 ± 8.3 μg/mL) was observed against E. coli.