The part of the plants examined and the drying method used are presented in Table 1. Dried samples were purchased from the local market or collected from different sites in Greece. All samples were analyzed within three months of collection. Gallic acid, gentisic acid, p-coumaric acid, vanillic acid, ferulic acid, syringic acid, (+)-catechin, quercetin, apigenin, naringenin, myricetin, were purchased from Sigma-Aldrich (Steinheim, Germany). Luteolin was from Roth (Karlsruhe, Germany). Caffeic acid was from Merck (Darmstadt, Germany). (−)-Epicatechin was from Fluka AG (Buchs, Switzerland). Rutin was from Alexis Biochemicals (Lausen, Switzerland). Ascorbic acid, p-hydroxybenzoic acid, and BHT (butylated hydroxytoluene) were a kind donation from the National Agricultural Research Foundation (N.AG.RE.F, Athens, Greece). Quantification was done via calibration with standards (external standard method). All standards were prepared as stock solutions in methanol. Methanol and sodium carbonate (Na₂CO₃) were purchased from Carlo Erba-SDS. Working standards were made by diluting stock solutions in 62.5% aqueous methanol containing BHT 1 g/L, and 6mol/L HCL to yield concentrations ranging between 0.5 and 25 mg/L. Stock/working solutions of the standards were stored in darkness at −180 °C. All solvents and reagents from various suppliers were of the highest purity needed for each application. The Folin-Ciocalteu reagent was purchased from Merck (Darmstadt, Germany). 2,2-diphenyl-1-picrylhydrazyl (DPPH) was obtained from Sigma-Aldrich (Steinheim, Germany). The ABTS reagent (2,2′-azinobis-(3-ethylbenzothiaziline-6-sulfonate) was from MP Biomedicals, (Eschwege, Germany).

The extraction method used for dried samples had as follows: 50 mL of methanol were added to 1 g of dried sample. The extraction mixture was refluxed in a water bath at 70 °C for 1 h. The mixture was then filtered and placed in a test tube in the fridge.

The essential oils were prepared by hydrodistillation, 30 g of plant materials were added in 300 mL water, for 3 h using a Clevenger-type apparatus. The supernatant was separated by decantation, dried over anhydrous Na₂CO₃ and kept in sterile flasks.

To prevent oxidation, all steps were carried out in dark with the flasks covered with aluminum foil. Moreover, it is essential that the waste (due to evaporation) of the solvent was kept at the lowest minimum.

The extraction method for the HPLC analysis had as follows: 40 mL of 60% aqueous methanol containing BHT (1 g/L) to prevent oxidation of phenolics extractedwas added to 0.5 g of dried sample. Then 10 mL of 6 mol/L HCl were added (separate aglycones from glycosides). The mixture was stirred carefully. In each sample nitrogen was bubbled for ca. 40–60 s. The extraction mixture was then sonicated for 15 min and refluxed in a water bath at 90 °C for 2 h. The mixture was then filtered and made up to 100 mL with methanol furthermore filtered quickly through a 0.45 μm membrane filter (Millex-HV) and injected to HPLC.

The total phenol content (TPC) was measured using the Folin-Ciocalteu assay [3]. A volume of 0.2 mL of the extract was introduced into test tubes followed by 0.5 mL Folin-Ciocalteu’s reagent (diluted 10 times with water). The solution was then kept at dark for 5 min and then 1 mL sodium carbonate (7.5% w/v) was added. The tubes were covered with parafilm and kept again in the dark for 1 h. Absorption at 765 nm was measured with a spectrophotometer UV-vis (Jasco V-530) and compared to a gallic acid calibration curve. The results were expressed as mg gallic acid/g dried sample. Each assay was carried out in triplicate. The determination of the total phenol content of the essential oils was done as follows: a volume of 3 mL of the methanol solution of each essential oil was introduced into test tubes followed by 1 mL Folin-Ciocalteu’s reagent (diluted 10 times with water) and 1 mL sodium carbonate (7.5% w/v).

The antioxidant activity was measured in terms of hydrogen donating or radical scavenging ability using the stable radical DPPH. Experiments were carried out according to the method of Blois [7] with a slight modification [8]. The reduction of the radical is followed by a decrease in the absorbance at 517 nm. A volume of 2 mL of a methanolic or aqueous methanolic stock solution of the extracts was put into test tubes and 2 mL of 1 mM DPPH solution was added. The tubes were covered with parafilm and kept again in the dark for 1 h. Absorbance at 517 nm was measured with a spectrophotometer UV-vis (Jasco V-530) and compared to an ascorbic acid calibration curve. The results were expressed as mg ascorbic acid/g dried sample. Each assay was carried out in triplicate. The percentage inhibition of the DPPH radical was calculated using the following formula: (1) where I = DPPH inhibition (%), A₀ = absorbance of control sample (t = 0 h) and A = absorbance of a tested sample at the end of the reaction (t = 1 h).

The amount of sample necessary to decrease the absorbance of DPPH by 50% (IC₅₀) was calculated graphically for the methanolic solutions of the essential oils of rosemary, thyme, oregano, and sage in six different concentrations.

The ABTS radical cation decolorization method is based on the reduction of ABTS^•+ radicals by antioxidants of the plant extracts tested. The reaction’s mechanism involves the electron-donating ability and results in the decolorization of the radical. ABTS was first dissolved in deionized water to a 7-mM concentration. Then, the solution of K₂S₂O₈ was prepared at a concentration of 2.45 mM. The two solutions were mixed with a ratio of 1:1 and kept in the dark for 24–48 h. The ABTS solution was then diluted in aqueous methanol with a ratio of 1:25. A volume of 20 μL (diluted 1:10) of aqueous methanolic plant extract was added to 2 mL of ABTS^•+ solution, and the mixture was kept at a standard temperature of 30 °C. The absorbance was measured at 734 nm in 0, 5, and 10 min after initial mixing. All solutions were used on the day of preparation, and all determinations were carried out in triplicate. The results were expressed as mg ascorbic acid/g dried sample. The percentage of inhibition of ABTS^•+ was calculated using the following formula: (2) where I = DPPH inhibition (%), A_t=0 = absorbance of control sample (t = 0 h) and A_t = absorbance of a tested sample in 5 or 10 min. Antioxidant capacity was expressed in mg ascorbic acid/g dried sample.

Samples of sunflower oil (3.5 g) containing 0.2% w/w extract were subjected to oxidation at 110 °C (air flow 20 L/h) [9]. The standard compounds (0.2% addition) were also examined. The rate of auto-oxidation of sunflower oil was estimated according to the increase in the induction period (IP). In the case of an increase in the IP, the extract displayed antioxidant ability, whereas a decrease in the IP indicated oxidative ability. The protection factors and the antioxidant activity of the samples were calculated according to the following formulas [10]: (3) where IP_x = induction period of the sample with additive, IP_k = induction period of sample without additive, IP_BHT = induction period of sample with added synthetic antioxidant BHT.

The analytical HPLC system employed consisted of a JASCO high performance liquid chromatograph coupled with a UV-vis multi-wavelength detector (MD-910 JASCO). The analytical data were evaluated using a JASCO data processing system (DP-L910/V). The separation was achieved on a Waters Spherisorb^® 5μm ODS2 4.6 × 250 mm column at ambient temperature. The mobile phase consisted of water with 1% glacial acetic acid (solvent A), water with 6% glacial acetic acid (solvent B), and water-acetonitrile (65:30, v/v) with 5% glacial acetic acid (solvent C). The gradient used was: 100% A 0–10 min, 100% B 10–30 min, 90% B/10% C 30–50 min, 80% B/20% C 50–60 min, 70% B/30% C 60–70 min, 100% C 70–105 min, 100% A 105–110; post time 10 min before next injection [11]. The flow rate was 0.5 mL/min and the injection volume was 20 μL. The monitoring wavelength was 280 nm for the phenolic acids and 320 and 370 nm (flavones, flavonoles). The identification of each compound was based on a combination of retention time and spectral matching.

The results of the determination of total phenolics are demonstrated in Table 1. Nepeta melissifolia, Phlomis lanata and Origanum vulgare demonstrated the highest total phenol content with more than 15.0 mg gallic acid/g dried sample. Geranium purpureum, Matricaria chamomilla and Lavandula vera seemed to have the lowest antioxidant capacity with less than 7 mg gallic acid/g dried sample. The results of this study were comparable to other similar ones [12,13,14]. Similar amounts in plant phenolics have also been reported from herbs and medicinal plants collected in Finland [3].

The methanolic solution of the essential oil of thyme showed the highest antioxidant capacity with 18 mg gallic acid/g dried sample (Table 2). The essential oil of rosemary was found to contain a relatively low concentration of phenols with only half that present in the essential oil of thyme. Viuda-Martos [14] demonstrated similar results. These results indicated that the phenolic compounds had a major contribution to the antioxidant capacity of herbs.

The DPPH method was evidently introduced nearly 50 years ago by Blois [7] and it is widely used to test the ability of compounds to act as free radical scavengers or hydrogen donors, and to evaluate antioxidant capacity. The parameter IC₅₀ (“efficient concentration” value), is used for the interpretation of the results from the DPPH method and is defined as the concentration of substrate that causes 50% loss of the DPPH activity (color). Some plant extracts and essential oils were examined in relation to their IC₅₀ value, while others were tested for their antioxidant capacity. IC₅₀ values of Nepeta melissifolia, Mentha pulegium and Phlomis lanata (5.1 ± 0.4, 13.5 ± 0.5, 23.9 ± 0.4 μg/mL) were found to be similar to BHT and ascorbic acid (18.5 ± 0.4, 3.9 ± 0.3 μg/mL). The IC₅₀ values of the examined plant extracts and essential oils are presented in Figure 1 and Figure 2, respectively.

As the IC₅₀ concentration and the antioxidant capacity have inversely proportional values, Salvia officinalis was established to have the lowest antioxidant capacity while Origanum vulgare was found to be the richest of all. Similar studies in plant extracts [15] exhibited higher hydrogen-donating capacity.

For the majority of the plants, as shown in Table 3, the DPPH method showed low antioxidant capacity. Only Sideritis cretica and Geranium purpureum were found to be the richest in antioxidants, approaching 1 mg ascorbic acid/g dried sample.

The results of the determination of antioxidant activity are demonstrated in Table 4. Origanum vulgare, Origanum dictamnus and Thymus vulgaris L. indicated the highest % inhibition for 0–5 min. In most cases as time elapsed, the % inhibition decreased due to the fact that the antioxidants of the aromatic plants scavenge the cation radical ABTS^•+. The values found in the present study were also comparable to similar research work [16,17].

The outcome of the Rancimat test as shown in Table 5 supports the hypothesis that aromatic plants are good sources of natural antioxidants such as phenolic compounds. When working accurately, this method offers an efficient, simple and automated assay. When ground material was added to sunflower oil, protection factors were slightly higher compared to the addition of methanol extracts. Similar PF values for ethanol and acetone extracts of plants of Greek origin have been reported [9].

Based on the table above, most aromatic plants indicated low protection against oil oxidation. Yet, the ground material of Nepeta melissifolia demonstrated very high protection while Sideritis cretica was found to have pro-oxidative action. Similar values were also established by Kintzios et al. [18] for the aromatic plants of Labiatae family. Aruoma et al. [19] found that Rosmarinus officinalis had medium protection (PF: 2.36) against oil oxidation in contrast with Thymus vulgaris L. and Origanum vulgare that indicated low protection.

The present method is simple, easy to use, and effective enough for the identification and quantification of major phenolic compounds in aromatic plants. This has been reported by other authors, who have used the same method of extraction and analysis of major flavonoid aglycones [1,20,21]. Generally, reversed phase HPLC with C₁₈ columns is the most popular technique for the analysis of polyphenols in different foods, despite the fact that the separation of procyanidins is not satisfactory with these phases [21]. Spherisorb C₁₈ stationary phase, which was used in this study to separate phenolic acids and flavonoids in aromatic plants, produced quite good results. After extraction and acid hydrolysis, the content of phenolic substances was determined. After extraction and acid hydrolysis the content of phenolic substances was determined. Quantification was done via a calibration with standards (external standard method). The amount of phenolic acids detected in the analyzed samples is shown in Table 6. Additionally, the content of flavonoids identified in the same plant extracts is shown in Table 7. Results are expressed in mg/ 100g dry sample. (+)-Catechin, quercetin and rutin were the most abundant flavonoids. Quercetin has been reported to be present in a large number of aromatic plants [1]. Rutin (quercetin 3-o-rhamnose glycoside) was present along with 22 other flavones, flavonoles and their glycosides in tea leaves [22]. Rutin in some cases can be hydrolyzed to quercetin (aglycone). This could have happened in the examined samples. Caffeic acid was detected in all the examined plant extracts. Ferulic acid was also detected in all the methanolic extracts, except from P. lanata, in rather high concentration. Phenolic acids are found in nature as esters and rarely as glycosides or in free form [23]. General comments have been published [12,24,25]. Thus, hydrolysis was needed for the identification and quantitative determination of phenolic acids. The data presented in Table 6 and Table 7 are considered as indicative of phenolic content of these aromatic plants. Papers about most of the examined plant extracts are very scarce in the literature. Among others reasons, time of harvest and storing conditions are considered responsible for the observed variations in the phenolic contents.