Chemical composition and antioxidant activity of oil from wild Achillea setacea and A. vermicularis

ABSTRACT

The objective of this study was to determine and compare the composition of volatile components of two species, A. setacea and A. vermicularis, under the temperate climatic conditions (north of Iran) and to investigate quantification of fatty acids in the oil extracted using gas chromatography-flame ionization detector and to evaluate the antioxidant potential and the phytochemical profile in terms of phenolic acids and flavonoids content of the oils obtained from the plants. Gas chromatography-mass spectrometry analysis of the essential oils showed that the major compounds of A. setacea were nerolidol (20%) and α-cubebene (14%), while in A. vermicularis were camphor (15%) and borneol (13%). Oil analysis revealed that the major components were palmitic and myristic acids. Chromatographic separation of their phenolic compounds (high-performance liquid chromatography) demonstrated that sinapic, gallic, caffeic, vanillic, syringic, and ferulic acids were present in the two oils of the plants, but in different amounts. These results confirmed A. setacea and A. vermicularis as important sources of bioactive metabolites.

KEYWORDS:

• Achillea setacea
• Achillea vermicularis
• Essential oil
• GC-MS
• HPLC
• GC

Introduction

In a society increasingly concerned with health and nutrition, medicinal plants emerge as alternative to synthetic products, used not only in traditional medicine but also in a number of food and pharmaceutical products due to their nutritional properties and bioactivity.^[1] Achillea (yarrow) species and their oil play an important role in a wide range of substances of natural origin, used in our everyday life. In fact, flowering tops and mainly their inflorescences are a rich source of active substances (essential oils, phenolic compounds, flavonoids, tannins) which are considered as valuable secondary metabolites for the food and pharmaceutical industries.^[2] The genus Achillea grows in temperate climates in dry or semi-dry habitats. It belongs to the family Asteraceae (Compositae) and tribe Anthemideae, in flowering plants, has one of the largest families; and includes more than 115 species distributed in the Europe, North Africa, and temperate areas of Asia.^[2,3] According to distribution of genus Achillea, two main centers of diversity occur in southeast Europe and southwest Asia.^[4] Nineteen Achillea species have been recognized and distributed in different geographical and ecological regions of Iran.^[5] The same genus is represented in Turkey by 46 taxa that 25 of them are endemic.^[6] Species of the genus Achillea are widely used for numerous pharmacological properties, such as antimicrobial, anti-inflammatory, antiallergic, and antioxidant activities.^[3,7,8] Also in Iran, Achillea species are used as diuretic and menstrual regularity agents for wound healing, diarrhea, flatulence, and abdominal pain.^[9,10] Phenolic compounds, such as flavonoids and phenolcarbonic acids, constitute one of the most important groups of pharmacologically active components in yarrow.^[8,11,12] Recent studies have demonstrated that the Achillea genus possesses antioxidant activity which is pertained to its total phenolic and flavonoid contents.^[11–13] Also the morphologic and chemical composition of Achillea species was affected by environmental conditions such as plant genetic type, seasonality, cultivar, geographic location, and developmental stage, because of a chemically polymorphic and perennial.^[14] Hence, there is currently a global approach to research about them. In Iran, most studies have focused on the chemodiversity of volatile oils from the aromatic plants.^[11] A. vermicularis is distributed in the mountainous regions in the north, northwest, west, and central parts of Iran.^[5] Some researchers have reported the major constituent in the oil of the plant as camphor (20–35%), 1,8-cineole (15–30%), germacrene D (10–15%), and piperitone (5–10%) from Iran.^[11,15] Recently, antioxidant properties of A. vermicularis have been reported in methanolic and ethanolic extracts.^[11,16] In spite of several studies undertaken on the herb, there are still several unknown aspects that need more attention. The plant A. setacea investigated in this report is a wild species of Iran which finds habitat in north and west-north of Iran.^[5] A survey of the literature revealed that no chemical study (such as volatile content, fatty acid composition) has been carried out previously on A. setacea grown in Iran. However, since essential oil yield and their components is related to environmental factors and in addition, composition of the essential oil of A. setacea has not been investigated from Iran, therefore this study was carried out to determine and compare the composition of volatile components of two species, A. setacea and A. vermicularis, under the temperate climatic conditions (north of Iran); to investigate quantification of fatty acids in the oil extracted using gas chromatography; to evaluate the antioxidant potential and the phytochemical profile in terms of phenolic acids and flavonoids content of the oils obtained from the plants.

Materials and methods

Plant materials

Plants of Achillea setacea (code no. 5537) and Achillea vermicularis (code no. 5536) were collected at the beginning of flowering in June 2015 from Guilan province (north of Iran). Their locations were marked by a global positioning system (GPS) system. One of the regions was road of Kelishom to Jirandeh (A. vermicularis: N:36º 41^/52^//, E:49º 42^/59^//and 1738 m above sea level-soil: sandy loam). Another was road of Asalem to Khalkhal near Kerman village (A. setacea: N:37º 36^/13.4^//, E:48º 41^/26.5^//and 2091 m above sea level-soil: sandy loam). The Achillea specimens were authenticated and stored in the herbarium of the Guilan Agricultural Research Center (GARC).

Essential and fatty acids extraction

Aerial parts of Achillea samples were dried at room temperature (20–25°C) in a shaded place for 5 days. Flowers and leaves were separated after drying. All samples were hydro-distilled for essential oil by grossly pulverized powdered plant (100 g) using a Clevenger type apparatus for 180 min.^[17] The oils was filtered over anhydrous sodium sulfate and stored in closed sterilized glass vials at +4°C in dark until being tested and analyzed.

To determine fatty acids, shade dried plant materials (flowers and leaves: 10 g) were powdered and passed through a sieve of 2 mm mesh size. Oil was extracted in Soxhlet apparatus using diethyl ether (40–60°C) as solvent. The lipid extracts were collected in a flask and subsequently treated with sodium sulfate to remove traces of water. At the end of the extraction, the extraction solvent was evaporated in a rotary evaporator to get the dry extract. Then, the methyl esters were prepared by dissolving 0.5 g of the oil in about 5 mL of n-hexane to which 5 mL the methanolic solution of sodium hydroxide (2 mol L⁻¹) were added. The mixture was sealed and heated in boiling water bath for 20 min. The hexane extract was filtered through 0.45 μm membrane filter.^[18]

Essential oil analysis

Analysis was performed on an Agilent 5973 gas chromatograph equipped with an ion-trap mass spectrometer detector (Varian Saturn 2100), using a HP-5MS (5 % of phenyl dimethylpolysiloxane), fused-silica capillary column (30 m × 0.25 mm i.d., 0.25 μm film thickness). Helium was used as a carrier gas. The injection volume was 1 μL. The column temperature was 120°C, with a 5 min initial hold and then it was increased to 260°C at 10°C min⁻¹ rate. The injector and detector temperatures were 250 and 200°C, respectively, and manifold at 70°C with line transfer at 240°C. The capillary column was coupled to a mass selective detector; the ionization energy voltage was 70 eV, electron multiplier voltage was 3000 v and ion resource temperature 200°C. Mass spectra were scanned in the range of 30–300 amu.

The constituents of the essential oils were identified by comparison of their Kovats retention indices (RI). Identification of individual compounds was made by comparison of their mass spectra with those of the internal reference mass spectra library (Wiley7n.1) or with authentic compounds and confirmed by comparison of their RI with authentic compounds or with those of reported in the literature.^[19]

Fatty acids analysis

Methylated samples (1 μL) were injected into a gas chromatograph (GC, Beifen SP-3420, Beifen, China) equipped with a flame ionization detector (FID) at 280°C and the fatty acid methyl esters were separated using a DB-FFAP(polyethyleneglycol acid modified in Agilent GC-Columns) column (30 m × 0.25 mm i.d and 0.25 μm of thickness) which treated with polyethylene glycol and nitrogen was used as carrier gas at a flow rate of 1.8 mL min⁻¹. The GC oven temperature was programmed as follows: 100°C (hold 10 min) and then kept at 230°C at 35 min. an injector temperature of 250°C. The peaks were identified by measuring the retention time of the samples and comparing the same standards analyzed under the same conditions.^[20]

Determination of phenolic acids in the oils by high-performance liquid chromatography-ultraviolet (HPLC-UV)

To perform this analysis, pure methanol was added to essential oil at a ratio of 1:1 (vol/vol), blended for 1 h, and centrifuged 10 min. Then, A 20 μL aliquot of the extracted solution (essential oil) was separated using a HPLC Knauer system equipped with ultraviolet-visible (UV-Vis) detector and a Eurospher 100-5 C-18 column (25 cm × 4.6 mm; 5 μm) according to the method proposed the authors.^[21] The mobile phase consisted of HPLC grade water with 2% A: acetic acid, and B: acetonitrile. Solvent gradient was used as follows: from 0 to 5 min isocratic 85% A flow, from 5 to 19 min (14 min) a linear gradient of A: 85% to B: 100%. After termination of the cycle, 15 min of column equilibration (85% A) were allowed prior next injection. Phenolic compounds were detected at a wavelength of 280 nm and identified by comparing their relative retention times and UV spectra with authentic compounds; they were detected using an external standard method.

Estimation of total phenols contents

The total phenolic content (TPC) was determined by the spectrophotometric method^[22] with a little modification. In brief, a 20 µL of the samples (essential oil) was mixed with 1 mL of Folin-Ciocalteu’s phenol reagent. After 5 min, 0.8 mL of a 7.5% Na₂CO₃ solution was added to the mixture. The mixture was kept in the dark for 30 min at 23ºC, after which the absorbance was read at 765 nm. The TPC was determined from extrapolation of calibration curve which was made by preparing gallic acid solution. The estimation of the phenolic compounds was carried out in triplicate. The TPC was expressed as milligrams of gallic acid equivalents (GAE) per gram of dried sample (The calibration equation for Gallic acid: y = 0.0421 x – 0.0232, R² = 0.998).

Estimation of total flavonoid content (TFC)

TFC was determined following a method by Youngjae et al.^[23] with a little modification. Briefly, a 20 µL of samples (essential oil) was kept in 1ml of deionized water. Then 75 µL of 5% NaNO₂ was added to this mixture, which was allowed stand for 5 min at room temperature, and 0.15 mL of 10% AlCl₃.6H₂O was added. The mixture was allowed to stand for 6 min at room temperature, and 0.5 mL of 1 mol L⁻¹ NaOH was added, and the total volume was made up to 3 mL with deionized water. The absorbance of the solution was measured immediately at 510 nm (WPA Biowave S2100). TFC were expressed in terms of g quercetin equivalents/100 g powder (the calibration equation for quercetin: y = 0.0779 x – 0.0136, R² = 0.9979).

DPPH radical scavenging assay

The measurement of DPPH radical scavenging activity was carried out according to the method of Hatano et al.^[24] A total of 10 µL of each of the samples was added to 2 mL of methanolic DPPH (2,2-diphenyl-1-picrylhydrazyl) (0.0023 mol L⁻¹) solution. The mixture was incubated in room temperature for 30 min before the change in absorbance at 520 nm was measured. The radical scavenging activity (D) was calculated as in Eq. (1):

(1)

where A0 is the absorbance of the DPPH solution and A1 is the absorbance of the sample.

Data analysis

To evaluate the antioxidant potential of essential oils using different systems of oxidation, all data represent an average of three replicates. Mean values and standard deviation (SD; n = 3) were calculated from the results. Data analyses were performed using SPSS software version 17. Comparison between groups was performed by one-way analysis of variance (ANOVA) and P < 0.05 was considered statistically significant.

Results and discussion

Chemical composition of essential oils by gas chromatography-mass spectrometry (GC-MS)

In this research, the amount of the essential oil obtained from A. setacea growing wild was 1.2% v/w dry and in A. vermicularis it was 0.6% v/w dry matter. Forty-five major components of the essential oil were identified using GC-MS analysis which some constituents were observed in both species, but some constituents were identified only in one species. In the oil of A. setacea, a sum of 33 components were characterized representing 90.61% of the total oil, with nerolidol (20.01%) and α-cubebene (14.02%) as the main constituents. Other compounds with low concentrations were β-cadinene (6.36%) and germacrene D (5.53%), spathulenol (5.16%), lavandulyl acetate (5.12%), β-selinenol (4.87%), β-selinene (4.41%), and 1,8-cineol (4.08%). The oil obtained of this plant was found monoterpene (20.07%) and sesquiterpene (70.54%; Table 1). In the oil of A. vermicularis, 25 components were characterized, representing 85.95% of the total oil. GC-MS analysis of the essential oil of this plant resulted in identification of different types of volatiles including mono (55.12%) and sesquiterpenoids (30.83%) as their major constituents. Among the characterized compounds, the oxygenated monoterpenes, camphor (18.8%), borneol (13.08%), and terpinen-4-ol (6.72%) were the major constituents, respectively. The essential oil of A. vermicularis was rich also in sesquiterpenoids: α-chamigrene (5.13%) and caryophyllene oxide (4.12%), respectively (Table 1).

Table 1. Main components of the essential oils from A. setacea and A. vermicularis.

	Percent composition (%)
Compounds	RI*	A. setacea	A. vermicularis
Santolina triene	909	0.52	nd
α-Pinene	935	0.61	nd
Sabinen	972	0.31	nd
β-Pinene	977	1.46	nd
p-Cymene	1027	nd	0.28
1, 8-cineol	1034	4.08	2.28
γ-Terpinene	1060	nd	0.29
Artemisia ketone	1063	2.2	nd
Cis-sabinene hydrate	1066	nd	1.60
Trans-sabinene hydrate	1093	nd	1.28
Linalool	1099	0.71	nd
Chrysanthenone	1125	nd	1.12
Cis-verbenol	1140	nd	0.63
Camphor	1145	0.61	18.8
Trans-chrysanthemol	1163	0.56	nd
Borneol	1167	1	13.08
Terpinen-4-ol	1176	nd	6.72
α-Terpineol	1188	3.03	3.69
Fragranol	1212	nd	2.09
Trans-carveol	1218	nd	0.68
Pulegone	1232	nd	0.21
Lavandulyl acetate	1279	5.12	nd
Verbenol acetate	1282	0.38	nd
Thymol	1289	1.51	1.87
Carvacrol	1299	nd	0.38
Piperitenone	1344	1.21	nd
α-Cubebene	1351	14.02	1.33
α-Copaene	1376	0.29	nd
α-Gurgujene	1407	0.42	nd
β-Caryophyllene	1418	0.98	0.59
α-Humulene	1455	0.27	nd
Germacrene D	1483	5.53	3.14
β-Selinene	1486	4.41	nd
Valencene	1491	2.46	1.6
Bicyclogermacrene	1492	0.47	nd
α-Chamigrene	1503	nd	5.13
(E,E)-α-Farnesene	1505	0.8	nd
β-Bisabolene	1510	0.68	1.31
γ-Cadinene	1515	046	nd
δ-Cadinene	1525	6.36	nd
(E)-Nerolidol	1564	20.01	1.44
Spathulenol	1576	5.16	2.96
Epiglobulol	1578	nd	1.95
Caryophyllene oxide	1582	2.95	4.12
β-Selinenol	1644	4.87	nd
(Z)-α-Bisabolene epoxide	1675	nd	1
Farnesol	1712	0.15	nd
Monoterpene hydrocarbons		2.9	0.57
Oxygenated monoterpenes		17.17	54.55
Sesquiterpene hydrocarbons	37.36		14.61
Oxygenated sesquiterpenes	33.18		16.22
Total	90.61	85.95

nd: not found; RI*: retention indices as determined on HP-5MS column.

The essential oil of A. setacea species was analyzed for the first time from Iran. The main constituents in the volatile oil showed some differences from those in the essential oils derived of A. setacea from Turkey and Middle Europe. For example, 1,8-cineole (18.5%), sabinene (10.8%), α-bisabolon oxide A (27%), and borneol (20%) were previously reported as major components of the oil of A. setacea.^[25–27] Whereas our studies showed that nerolidol (20.01%), α-cubebene (14.02%), and β-cadinene (6.36%) were main constituents of the oil. Also, sesquiterpene hydrocarbons (37.36%) were dominating compounds in this species. Noteworthy, was α-nerolidol (20.01%) and α-cubebene (14.02%), were not reported as major components previously. According to the report on A. vermicularis growing in Turkey, the major components of essential oil were camphor (41.3%) and 15-hexadecanolide (19.6%).^[26] In another study, 1,8-cineole (15–30%), camphor (20–35%), and germacrene D (12–15%) were previously reported as main components in A. vermicularis from other parts of Iran,^[11,15] whereas, in our study, camphor (18.8%), borneol (13.08%), and terpinen-4-ol (6.72%) were the major constituents. In addition to these findings, our results indicated that terpinen-4-ol, α-chamigrene, epiglobulol, and β-bisabolene were reported for the first time in A. vermicularis essential oil from the country. On the other hand, according to previous researches, Achillea has a particular pattern of accumulation of essential oils during its development. There is a reverse relation between chamazulene and spathulenol amounts in yarrow.^[28] In this study, the same result was obtained and no chamazulene was observed. In conclusion, the variations between the essential oils of these plants growing in different areas are may be attributed mainly to the differences in their chemical polymorphic structure and geographical conditions.^[2,14,28]

Determination of fatty acids content

Table 2 summarizes the results obtained from GC analysis showed the fatty acid composition saturated fatty acids and unsaturated fatty acids were detected and quantified. The oil content of the aerial parts A. setacea was 0.31%, which is higher than the oil content of A. vermicuraris (0.26%). In A. setacea palmitic acid (33.45%) was found in the sample in highest concentration. Among the other fatty acid with concentration are myristic acid (13.36%), arachidic acid (11.15%), and capric acid (10.3%). The most abundant fatty acid was in A. vermicularis palmitic acid with concentration 38.86% followed by myristic acid (18.62%), arachidic acid (8.86%), and capric acid (8.71%). Evidence suggested that polyunsaturated fatty acids (PUFAs) especially linoleic acid (LA) and linolenic acid (LN) are essential to human health. However, due to incapability of human body to synthesize these fatty acids, they are provided in the diet. Therefore, more studies were carried out to nutritional and pharmaceutical values.^[29] However, no study has been carried out on oil content (fatty acid) of yarrow aerial parts and its quality in Iran. Goli et al.^[30] reported, the seed oil of A. tenuifolia growing in Tehran province (Iran) contained mainly linoleic (69.4%), oleic (14.5%) acids, and LN (1.7%). According to our research, in A. setacea LA (ω-6) and LN (ω-3) were obtained in good quantities as 8.04 and 1.5%, respectively. Also in the oil from aerial parts of A. vermicularis were found LA (5%) and LA (1.28%). According to tests done in A. setacea were not observed erucic acid (toxic) or oleic acid in A. vermicularis. The saturated fatty acid composition of examined species showed low intraspecific variability while it was significant in unsaturated fatty acid composition (Table 2). Also, the differences in individual contents of fatty acids when compared to the bibliographic references, may be due to genotypes of different or environmental factors.^[30] However, the presence of these important fatty acids in appreciable amounts can make the oils beneficial for health, which can be used in the preparation of phytopharmaceutical or pharmaceutical preparations.

Table 2. The fatty acids composition (%) of A. setacea and A. vermicularis by GC.

			Concentration (%)
Fatty acid	Chemical formula	Acronym	A. setacea	A. vermicularis
Capric acid	C10H20O2	C10:0	10.3	8.71
Lauric acid	C12H24O2	C12:0	3.38	11.7
Myristic acid	C14H28O2	C14:0	13.36	18.62
Palmitic acid	C16H32O2	C16:0	33.45	38.86
Palmitoleic acid	C16H30O2	C16:1	4.24	1.71
Stearic acid	C18H36O2	C18:0	4.24	0.58
Oleic acid	C18H34O2	C18:1	2.4	nd
Linoleic acid (ω-6)	C18H32O2	C18:2	8.04	5
Linolenic acid(ω-3)	C18H30O2	C18:3	1.5	1.28
Arachidic acid	C20H40O2	C20:0	11.15	8.86
Behenic acid	C22H44O2	C22:0	1.78	nd
Erucic acid	C22H42O2	C22:1	nd	1.66
Lignoceric acid	C24H48O2	C24:0	5.74	3.02
Saturated fatty acids			83.4	90.35
Monounsaturated fatty acids			6.64	3.37
Polyunsaturated fatty acids			9.9	6.24

nd: not detected.

Evaluation of phenolic compounds by HPLC-UV

The applicability of the proposed analytical method and the qualitative and quantitative determination of the standard phenolic compounds have already been verified.^[31] Due to their importance in plants and human health, it would be useful to know the concentration of the polyphenolic compounds and biological activities that could indicate their potentials as therapeutic agents, and also for predicting and controlling the quality of medicinal herbs.^[32] But no scientific data in the field (HPLC analysis) on Achillea is not available in Iran. Eight compounds were identified in the oils of A. setacea and A. vermicularis (Table 3). In the essential oil of A. setacea, sinapic acid ˃ caffeic acid ˃ ferulic acid ˃ were the compound found in the largest amount (31.33 ± 0.6 mg g⁻¹, 3.12 ± 0.06 mg g⁻¹, and 2.94 ± 0.02 mg g⁻¹, respectively). Also p-coumaric acid was dominant in the oil of A. vermicularis (8.59 ± 0.5 mg g⁻¹), followed by syringic acid (5.37 ± 0.05 mg g⁻¹) and ferulic acid (3.66 ± 0.3 mg g⁻¹), respectively. Both investigated plants rutin (flavonoid glycoside) was not separated following the same method. Our results revealed that the plants contain hydroxycinnamic acids (such as sinapic and p-coumaric acids) rather than hydroxybenzoic acids (such as gallic and vanillic acids) which is in good agreement with some previously reported data.^[33] The phenolic acids found in this study for these plants are known to have many biological activities due to their hydroxyl groups, which can thus, be correlated with the popular use of these plants.^[13]

Table 3. Content of phenolic compounds in essential oil of Achillea species.

		Plant material (mg g^−1 DW)
Compound	RT	A. setacea	A. vermicularis
Rutin	9.4	nd	nd
Galic acid	13.2	1.22 ± 0.09	1.17 ± 0.09
Caffeic acid	32.2	3.12 ± 0.06	2.66 ± 0.08
4-Hydroxy benzoic acid	37.52	2.91 ± 0.3	2.87 ± 0.2
Vanillic acid	38.92	0.24 ± 0.09	2.85 ± 0.2
p-coumaric acid	39.38	1.41 ± 0.3	8.59 ± 0.5
Syringic acid	40.12	2.89 ± 0.01	5.37 ± 0.05
Ferulic acid	41.84	2.94 ± 0.02	3.66 ± 0.3
Sinapic acid	42.35	31.33 ± 0.6	2.99 ± 0.01

RT: retention time (min).

Each value is presented as the mean ± SD (n = 3).

nd: not detected; DW: dry weight basis.

Determination of total polyphenolic and flavonoid contents

Among the natural compounds participating against free radicals, phenolic compounds, particularly flavonoids, constitute one of the major groups of herbal compounds acting as radical scavengers and antioxidants. Therefore, these substances have been proposed as health-promoting natural products.^[34] Significantly, the highest amount of TPC was determined in the oil of A. setacea (158.67 ± 0.93 mg g⁻¹, p < 0.05) followed by A. vermicularis oil (154.97 ± 1.06 mg g⁻¹, p < 0.05). Also concerning the content of flavonoids (TFC), the oil of A. vermicularis (88.75 ± 1.02 mg g⁻¹, p < 0.05), was richer in flavonoids, than the oil of A. setacea (70.72 ± 0.61 mg g⁻¹, p < 0.05; Table 4). To the best of our knowledge, there is no previous report on the antioxidant properties (TPC, TFC, DPPH) of Achillea essential oils in Iran. Studies have been conducted for antioxidant activities of the extracts (methanolic, ethanolic, etc.) from different Achillea species.^[11] These factors can only be directly compared with chemically similar oils, composing at least the same major compounds. However, apart from chemical variations of essential oils, differences in methods applied and diversity of plants used, make results incomparable. In a study in Iran on several of Achillea species methanol extracts was found that TPC varied from 13.56 to 188.66 (mg tannic acid g⁻¹ dry weight [dw]) while, TFC ranged from 19.23 to 79.16 (mg quercetin g⁻¹ dw).^[11] Also the author Nemeth and Bernath state 0.6% content of flavonoids and 1.48% of phenolic acids.^[3] Thus, the comparative study showed that these Achillea species seem to be a rich source of total polyphenolic and flavonoid contents that can be considered as a promising source of natural antioxidants. However, plant polyphenols such as flavonoid and phenolics are biosynthesized via several routes and constitute a heterogeneous group from the metabolic point-of-view.^[35]

Table 4. The content of total polyphenols, flavonoids, and antioxidant capacity parameters in essential oil of Achillea species.

Samples	TPC (mg GAE g^−1	TFC (mg QUE g^−1)	DPPH (%)	IC50 (µg mL^−1)
A. setacea	158.67 ± 0.93	70.72 ± 0.61	75.82 ± 0.06	28.87 ± 0.8
A. vermicularis	154.97 ± 1.06	88.75 ± 1.02	73.32 ± 1.24	30.63 ± 1.04

Each value is the mean ± SD of three independent measurements.

GAE: gallic acid equivalents; QUE: rutin equivalents.

DPPH radical scavenging assay

The DPPH test is a very convenient method for screening small antioxidant molecules (such as flavonoids) because the reaction intensity can be analyzed by the spectrophotometric method.^[13] Radical scavenger activity of the extracts was expressed as the amount of antioxidants necessary to decrease the initial DPPH absorbance by 50% (median effective concentration value, EC₅₀) and results were given in Table 4. The scavenging activity toward DPPH radical of the EO of A. setacea (75.82 ± 0.06%, IC₅₀ = 28.87 ± 0.8 µg mL⁻¹) was significantly higher (p < 0.05) than EO of A. vermicularis (73.32 ± 1.24%, IC₅₀ = 30.63 ± 1.04 µg mL⁻¹) in all samples. This is in good agreement with the TPC values listed in Table 4. A significant positive correlation (p < 0.05) was found between TPC and DPPH-scavenging activity in both samples. Recently Gharibi et al.^[11] tested DPPH scavenging activity of eight Achillea (A. vermicularis, A. nobilis, A. wilhelmsii, A. millefolium, A. filipendulina, A. tenuifolia, A. biebersteinii, and A. eriophora) species and observed highest activity with A. eriophora methanolic extract. This antioxidant capacity was attributed to the flavonoid and phenol contents. Our results showed an agreement with previous studies that determined by Polatoglu et al. in Turkey.^[8] They reported that essential oils of A. teretifolia and A. vermicularis showed considerable DPPH scavenging activity. This high activities of the oils was attributed to the content of monoterpene and sesquiterpene alcohols. It is well-known that antioxidants react with DPPH free radical and convert it to the stable form.^[8,13,25] Therefore, two Achillea species oils, seem to contain powerful inhibitor compounds, which may act as primary antioxidants that react with free radicals as efficient hydrogen donors.

Conclusion

This is the first report on the essential oil composition of A. setacea from Iran. The results obtained clearly showed that the oil was characterized by its richness in sesquiterpene hydrocarbons. Our results confirmed variations in the qualitative and quantitative composition in the oils obtained from the aerial parts of two distinct Achillea species. These differences can probably be attributed to the genetic differences or different geographic or environmental conditions of the plant materials. The fatty acid profile plays an important role to the chemical properties, therefore, this is useful knowledge for further researches. This study revealed that the aerial parts of these plants are attractive sources of oily components, especially the essential ones, as well as of effective natural sources of unsaturated fatty acids. They are also a non-negligible source of polyphenolic and essential oil compounds with bioactive properties which are very good therapeutic agents that could orientate toward the use of these plants in pharmaceutical and cosmetic industries.

Conflict of interest

No potential conflict of interest was reported by the authors

Acknowledgments

The authors wish to thank Dr. Morady (taxonomist) in herbarium of Guilan agriculture and natural resources researches center, for identifying the plant material.

Funding

We also thank the research councils of Urmia University for financial supports of the present study with a research grant (996/2015/D30). No potential conflict of interest was reported by the authors.

Additional information

Funding

We also thank the research councils of Urmia University for financial supports of the present study with a research grant (996/2015/D30). No potential conflict of interest was reported by the authors.