Chemical Composition, Antimicrobial, and Antioxidant Activities of Essential Oil and Ethanol Extract of Coriandrum sativum L. Leaves from Turkey

Abstract

This study reports the chemical composition, antioxidant, and antimicrobial activity of the essential oil and ethanol extract of Coriandrum sativum L. leaves. Gas chromatography/mass spectrometry analysis identified 19 compounds representing 95.30% of the oil. (E)-2-decenal (29.87%), linalool (21.61%), (E)-2-dodecenal (7.03%), dodecanal (5.78%), (E)-2-undecenal (3.84%), (E)-2-tridecenal (3.56%), (E)-2-hexadecenal (2.47%), tetradecenal (2.35%), and α-pinene (1.64%) were the main components identified in the essential oil. The samples were screened for their antioxidant activities using 2,2-diphenyl-1-picrylhydrazyl radical scavenging and β-caroten bleaching assay. IC₅₀ value for ethanol extract of C. sativum was determined as 74.87 ± 0.03 μg/mL. Total antioxidant activity value for C. sativum ethanol extract was 85.85 ± 0.04%. Total phenolic content for ethanol extract of the plant was determined as 14.97 ± 0.05 mg gallic acid equivalents/g dry weight. The essential oil and ethanol extract were also tested for antimicrobial activity against 28 different foodborne microorganisms, including 19 bacteria, 7 fungi, and 2 yeast species. The ethanol extract of the plant showed weak antimicrobial activities against microbial strains in both disc diffusion and minimal inhibition concentration tests. This study suggested that Coriandrum sativum L. leaves may be used as a potential source of food flavoring, and for their antioxidants and antimicrobial properties.

Keywords:

• Antimicrobial activity
• Antioxidant activity
• Coriandrum sativum L.
• Essential oil
• Gas chromatography/mass spectrometry (GC/MS)

INTRODUCTION

Turkey is a country located in the temperate zone with remarkable plant diversity. This plant diversity is due to varied geographical features and climate diversity, the country’s position as a natural bridge between two continents and also as a midpoint of three different geographical plant zones. Each of the plants which have been classified based on their color, texture, shape, and the chemicals they contain, could be the subject of extensive researches. The story that starts with a seed and ends with a fruit and the mid-products that form in this process give a different meaning to each plant. The relationships that make plants indispensable show themselves in every area. From medical to nutrition uses or from art to decoration, the mark of plants and flowers can be found in all elements.

Coriandrum sativum L., known as coriander (seeds of C. sativum L.) or cilantro (leaves of C. sativum L.), is commonly utilized for its fresh leaves and the dry powder of its fruits which have organoleptic and flavoring properties. It is among the most widely used medicinal plants, possessing nutritional as well as medicinal properties.^[1,2] Hippocrates (ca. 460–377 B.C.) used coriander in traditional Greek medicine. The seeds of coriander have been discovered in the ancient Egyptian tomb of Rameses the Second. This herb had been called the “spice of happiness” by the Egyptians because they considered it an aphrodisiac. Coriander was also used to flavor wine and as a medicine by the Greeks and Romans.^[3] The coriander plant is mainly used for making sauces, salads, and soups. The fresh, green leaves possess a unique aroma and they are frequently used as an important ingredient in Thai and Vietnamese cuisine.^[2] Furthermore, it is commonly found in Turkish cuisine and in particular the plant’s fresh leaves are added to a soup named “ayran aşı” which is a natural product that is made from yogurt, with high nutritional value. Thanks to addition of cilantro, this soup has a special aroma and atypical taste from other soups cooked in the same way.

Microorganisms, including Gram positive and Gram negative bacteria, in addition to fungi, have been recognized as the main causers of various human infections and play a critical role in food spoilage. There is a worldwide trend of exploring new alternatives to control foodborne diseases. This trend has necessitated the development of new antimicrobial drugs. Aromatic and medicinal plants are sources of active molecules and produce a wide variety of volatile aliphatic and cyclic hydrocarbons. Bioactive phytochemicals from these plants are often recovered as “essential oils” (EOs) by hydrodistillation of whole tissues or seed.^[4,5]

EOs are aromatic liquids that usually have antimicrobial and antioxidant properties which are obtained from herbs, fruit, spices, or flowers. They can be used as food flavoring agents or preservatives and for medicinal purposes.^[2,4,6] C. sativum EO has a characteristic odor of linalool and a soft, sweet, aromatic flavor. Antimicrobial activity has been reported for the EO extracted from C. sativa seeds and leaves against different species of both Gram positive and negative bacteria, yeasts, and fungi. At the same time, its high antioxidant and antimicrobial activities have been related to the presence of phytochemicals such as linalool, camphor, geraniol, and geranyl acetate.^[1,7,8]

Turkish C. sativum leaves (cilantro) have not been studied previously with regard to EO composition. This work presents (1) the chemical composition of the EO from leaves of C. sativum, (2) in vitro antioxidant activity of the plant EO and ethanol extract using two complementary assays: β-caroten bleaching (BCB) and 2,2-diphenyl-1-picrylhydrazyl (DPPH) radical tests, (3) total phenolic contents of the plant EO and ethanol extract, and (4) antimicrobial activity of the plant EO and ethanol extract.

MATERIALS AND METHODS

Plant Collection and Authentication

Plant samples (Coriandrum sativum L. samples) commonly grown in Eastern Anatolia were purchased from local producers in Ilıca, Erzurum, Turkey. The healthy plant leaves, without damage or insects, were randomly selected and immediately transported to the laboratory. The taxonomic identification of the plant material was confirmed by a senior plant taxonomist, Meryem Sengül Köseoglu, in the Department of Biology, Atatürk University, Erzurum, Turkey.

Other Materials in the Experiment

In this study, a total of 28 food-associated microorganisms (including 19 bacteria, 7 fungi, and 2 yeast species), frequently reported in foods, were used. The microorganisms used are listed in Tables 1 and 2. Microorganisms were provided by the Food Microbiology Laboratory of the Department of Food Engineering in the Faculty of Agriculture at Ataturk University in Erzurum, Turkey. All chemicals used for the research purposes were obtained from Merck (Darmstadt, Germany) and Sigma-Aldrich Chemie (Steinheim, Germany).

TABLE 1 Antibacterial activity of the ethanol extract and essential oil of Coriandrum sativum

	Ethanol extract		Essential oil		Reference antibiotics
Bacterial strain	DD^a	MIC^b	DD^c	MIC^b	DD^d	MIC^e
Acinetobacter lwoffi BC 2819	–	500 ± 0.00	8 ± 0.50	500 ± 0.00	17 ± 0.58 (AMC30)	7.8 ± 0.00
Bacillus cereus BC 6830	–	500 ± 0.00	9 ± 0.87	500 ± 0.00	12 ± 0.00 (CC2)	7.8 ± 0.00
Bacillus subtilis BC 5211	–	500 ± 0.00	11 ± 0.87	500 ± 0.00	36 ± 0.00 (AMC30)	7.8 ± 0.00
Flavobacterium indologenes BC 1520	8 ± 0.40	500 ± 0.00	13 ± 0.21	500 ± 0.00	27 ± 0.00 (AZM15)	7.8 ± 0.00
Enterococcus faecalis	10 ± 0.42	500 ± 0.00	16 ± 0.68	250 ± 0.00	18 ± 0.64 (SCF105)	31.3 ± 0.00
Escherichia coli BC 1402	–	–	11 ± 0.60	500 ± 0.00	22 ± 0.58 (OFX10)	15.6 ± 0.00
Escherichia coli BC 2326	–	–	10 ± 0.75	500 ± 0.00	26 ± 0.58 (AZM15)	7.8 ± 0.00
Klebsiella pneumoniae ozaenae BC32	10 ± 0.00	500 ± 0.00	18 ± 0.67	250 ± 0.00	19 ± 0.30 (CC2)	31.3 ± 0.00
Klebsiella pneumoniae BC 1749	10 ± 0.30	500 ± 0.00	16 ± 0.00	250 ± 0.00	30 ± 0.00 (OFX10)	62.3 ± 0.00
Listeria monocytogenes BC 8353	–	–	25 ± 0.65	31.3 ± 0.00	22 ± 0.00 (SAM20)	7.8 ± 0.00
Proteus mirabilis BC 2644	12 ± 0.35	500 ± 0.00	19 ± 0.30	250 ± 0.00	28 ± 0.58 (OFX10)	125 ± 0.00
Providencia alcalifaciens	10 ± 0.00	500 ± 0.00	14 ± 0.47	500 ± 0.00	33 ± 0.17 (OFX10)	62.5 ± 0.00
Pseudomonas aeruginosa BC 4372	–	–	7 ± 0.50	500 ± 0.00	28 ± 0.35 (OFX10)	7.8 ± 0.00
Pseudomonas fluorescens BC 7324	–	–	9 ± 0.42	500 ± 0.00	31 ± 0.00 (OFX10)	125 ± 0.00
Pseudomonas pseudoalcaligenes BC 3445	10 ± 0.00	500 ± 0.00	15 ± 0.68	250 ± 0.00	32 ± 0.17 (OFX10)	125 ± 0.00
Salmonella Typhimurium RSSK 95091	–	–	9 ± 0.59	500 ± 0.00	12 ± 0,75 (TE30)	7.8 ± 0.00
Staphylococcus aureus ATCC 29213	–	–	21 ± 0.38	31.3 ± 0.00	34 ± 0.12 (TE30)	7.8 ± 0.00
Streptococcus pyogenes ATCC 176	13 ± 0.57	500 ± 0.00	26 ± 0.00	62.5 ± 0.00	25 ± 0.06 (OFX10)	7.8 ± 0.00
Yersinia enterocolitica BC 0184	10 ± 0.60	500 ± 0.00	13 ± 0.81	500 ± 0.00	26 ± 0.29 (AZM15)	31.3 ± 0.00

^aDD: Diameter of inhibition zone [mm] around the disks (6 mm) impregnated with 300 µg of ethanol extract.

^bMIC: Minimal inhibitory concentration [µg/mL].

^cDD: Diameter of inhibition zone [mm] around the disks (6 mm) impregnated with 10 µL of essential oil.

^dDD: Diameter of inhibition zone [mm] around the reference standard antibiotic disks (Oxoid) AMC30 (20 µg amoxicillin + 10 µg clavulanic asit/disc), CC2 (2 µg clindamycin/disc), AZM15 (15 µg azithromycin/disc), SCF105 (30 µg sulbactam + 75 µg cefoperazone/disc), OFX10 (10 µg Ofloxacin/disc), SAM20 (10 µg sulbactam + 10 µg ampicillin/disc), TE30 (30 µg tetracycline/disc).

^eMIC: Minimal inhibitory concentration [µg/mL] of the reference antibiotic clarithromycin (Oxoid).

–: no inhibition zone.

TABLE 2 Antifungal activity of ethanol extract and essential oil of C. sativum

	Ethanol extract		Essential oil		Amphotericin B
	DD^a	MIC^b	DD^c	MIC^b	DD^d	MIC^e
Yeasts
Candida albicans ATCC 1223	9 ± 0.21	500 ± 0.00	18 ± 0.25	7.8 ± 0.00	15 ± 0.00
Candida crusei ATCC 14243	–	–	9 ± 0.44	250 ± 0.00		125 ± 0.00
Fungi
Aspergillus flavus BC 101	–	–	8 ± 0.55	500 ± 0.00	17 ± 0.00	250 ± 0.00
Aspergillus niger BC 102	–	–	14 ± 0.21	250 ± 0.00	21 ± 0.00	62.5 ± 0.00
Aspergillus niger BC 103	–	–	17 ± 0.36	250 ± 0.00	19 ± 0.29	62.5 ± 0.00
Cladosporium herbarum BC 106	–	–	22 ± 0.26	250 ± 0.00	10 ± 0.36	250 ± 0.00
Penicillium brevicompactum BC 109	–	–	19 ± 0.47	125 ± 0.00	14 ± 0.23	62.5 ± 0.00
Penicillium roquefortii BC 111	–	–	21 ± 0.00	250 ± 0.00	11 ± 0.00	250 ± 0.00
Trichothecium roseum BC 116	–	–	26 ± 0.00	125 ± 0.00	32 ± 0.49	62.5 ± 0.00

^aDD: Diameter of inhibition zone [mm] around the disks (6 mm) impregnated with 300 µg of ethanol extract.

^bMIC: Minimal inhibitory concentration [µg/mL].

^cDD: Diameter of inhibition zone [mm] around the disks (6 mm) impregnated with 10 µL of essential oil.

^dDD: Diameter of inhibition zone [mm] around the reference standard antimycotic disk (20 µg amphotericin B/disk; Sigma. MIC: Minimal inhibitory concentration [µg/mL] of the reference antimycotic amphotericin B (Sigma).

Isolation of the EO

Plant leaves were lyophilized by freeze-drying (Xianou-12N, Nanjing, China). The lyophilized plant material was ground with a blender, stored in glass containers, kept in a refrigerator and used within a few days. Four portions (75 g each) of the ground plant material were separately subjected to hydrodistillation for 240 min using a Clevenger-type apparatus. The EO was collected, dried under anhydrous sodium sulfate, and stored at low temperature (4°C) for further analysis.

Preparation of the Ethanol Extract

The ground plant material (200 g) was extracted in a Soxhlet (number of the exchanges in Soxhlet extraction is six) with ethanol for 72 h. The extracts were filtered using Whatman filter paper (No: 1) and then concentrated in vacuo at 40°C using a rotary evaporator. The residues obtained were stored in a freezer at −80°C before further tests. All volatile components of the extract, including the EO and trace amount of the solvent (ethanol) used in the extraction process, were exactly removed using a vacuum oven. The extracts were stored in vials in the refrigerator and were used for analysis.

Gas Chromatography/Mass Spectrometry (GC/MS) Analysis

GC/MS analysis of the EO was carried out on an Agilent-7890B gas chromatography equipped with an Agilent 5975C mass selective detector and HP-5MS cap. Column (30 m × 0.32 mm, 0.25 μm film thickness). The oven temperature was programmed from 60 to 240° at 3°/min and injector temperature of 220°; the carrier gas was He (1 mL/min). Identification of components of EO was based on the comparison of the Wiley 275.L and Wiley/n.L library data of the GC/MS system and literature data.^[9,10]

Antioxidant Activity

DPPH radical scavenging assay

Antioxidant activity of the plant was measured by DPPH free radical scavenging method as described.^[11,12] In the assay, 0.1 mM solution of DPPH was prepared in ethanol and 0.5 mL of this solution was added to 1.5 mL of extract solution in ethanol at different concentrations (20–100 μg/mL). These solutions were vortexed in depth and incubated in the dark at 25°C for 30 min. Thirty minutes later, the absorbance was measured at 517 nm against blank samples without the scavenger. A standard curve was prepared using different concentrations of DPPH. The capability to scavenge the DPPH free radical was calculated using the following equation:

where, A_c is the absorbance at 517 nm of the control reaction (containing DPPH solution without C. sativum extract), and A_s is the absorbance of the test sample.

β-caroten bleaching (BCB) assay

The antioxidant activity of the plant extract was determined according to the BCB method described by Kaur and Kapoor^[13] with some modifications. In the BCB assay, antioxidant capacity is determined by measuring the inhibition of the volatile organic compounds and the conjugated hydroperoxides arising from linoleic acid oxidation. Briefly, 4 mL of β-carotene solution (0.1 mg in 1 mL chloroform), 40 mg of linoleic acid and 400 mg of Tween 40 were transferred to a round-bottom flask. The mixture was then evaporated at 50°C by means of a rotary evaporator to remove chloroform. Then, 100 mL of oxygenated distilled water was slowly added to the residue and vigorously agitated to give a stable emulsion. Then, 800 μL of extract was added to 3 mL of β-carotene/linoleic acid emulsion. As soon as the emulsion was added to each tube, the zero time absorbance was measured at 470 nm using a visible spectrophotometer (T60V; PG Instruments Ltd.). The mixtures were incubated at 50°C for 100 min, and measurement was carried out at 10 min intervals for 100 min. Water, instead of plant extract, was used as control. A blank, devoid of β-carotene, was prepared for background subtraction. Butylated hydroxyanisole (BHA) and butylated hydroxytoluene (BHT) were used as a standard. All samples were assayed in triplicate. The degradation rate (DR) was calculated according to first-order kinetics using the following equation:

where ln is natural log, a is the initial absorbance (470 nm) at time 0, b is the absorbance (470 nm) at 100 min, and t is time. The antioxidant activity (AA) was expressed as percent of inhibition relative to the control, using the following formula:

Determination of total phenolic content

Total phenolic constituent of the plant extract was performed employing the literature methods involving Folin–Ciocalteu reagent and gallic acid as standard.^[14] Extract solution (0.1 mL) containing 1000 μg extract was taken in a volumetric flask, 46 mL distilled water and 1 mL Folin–Ciocalteu reagent were added and shaken vigorously. After 3 min, 3 mL of solution 2% sodium carbonate was added and the mixture was allowed to stand for 120 min with intermittent shaking. Absorbance was measured at 760 nm using a visible spectrophotometer (T60V; PG Instruments Ltd., Leicestershire, UK). Measurements were carried out in triplicate. The same procedure was repeated to all standard gallic acid solutions (0–1000 mg, 0.1 mL-L) and standard curve was obtained.

Antimicrobial Activity

Disc-diffusion assay

Ethanol extract was dissolved in dimethyl sulfoxide (DMSO; Sigma) to produce solutions of concentration 30 mg/mL and sterilized by filtration through 0.45 μm Millipore filters (Schleicher & Schuell, Microscience, Dassel, Germany). The antimicrobial activity of the ethanol extract and EO was measured by disc diffusion method, using 100 μL of suspension, containing 1 × 10⁸ colony forming unit (CFU)/mL of bacteria, 1 × 10⁶ CFU/ml of yeast and 1 × 10⁴ spores/mL of fungi. Bacteria were spread on Nutrient Agar (NA), yeast were spread on sabouraud dextrose agar (SDA) and fungi were spread on potato dextrose agar (PDA). Sterile filter paper discs (6 mm in diameter) infused with 300 μg of ethanol extract and 10 μL of EO, were placed on the inoculated agar. SAM20 (10 μg sulbactam + 10 μg ampicillin/disc), OFX10 (10 μg ofloxacin/disc), AMC30 (20 μg amoxicillin + 10 μg clavulanic asit/disc), KF30 (30 μg cephalothin/disc), TE30 (30 μg tetracycline/disc), and AZM15 (15 μg azithromycin/disc) were used as positive reference standards to determine the sensitivity of one strain/isolate in each microbial species tested. The microbial cultures were incubated for 24 h at 37°C for mesophilic bacterial strains, 20°C for 48 h for psychrotrophic bacterial strains, at 30°C for 48 h for the yeasts and at room temperature for 72 h for fungi isolates.^[15] Antimicrobial activity was evaluated by measuring the disk diffusion (DD) zone of inhibition against the test organisms. Each assay was performed in triplicate in the aseptic environment for the reproducibility of the result.

Microwell dilution assay

Inoculates of the strains were prepared from 12 h broth cultures, and suspensions were adjusted to 0.5 McFarland standard turbidity. The EO and ethanol extract of C. sativum were dissolved in DMSO, diluted to the highest concentration tested (500 μg/mL) with distilled, sterile H₂O. Then serial two-fold dilutions were made to obtain a concentration range from 7.8 to 500 μg/mL in 10-mL sterile test tubes containing Nutrient Sabouraud Dextrose Broth. Bacterial and yeast strains sensitive to the plant extract and EO in disc diffusion assay were studied for their minimal inhibition concentration (MIC) values using the micro-well dilution assay method.^[16]

MIC agar dilution assay

The agar dilution method was used to determine the MIC values of the fungus isolates.^[17] The EO and ethanol extract of plant were added aseptically to sterile molten PDA medium containing Tween 20 (0.5% v/v, Sigma), at the appropriate volume to produce a concentration range of 7.8–500 μg/mL. The prepared PDA solutions were immediately poured into Petri plates after vortexing. The inoculated plates were incubated at room temperature for 72 h. At the end of the incubation period, the plates were evaluated for the presence or absence of growth. Each test was repeated at least three times.

RESULTS AND DISCUSSION

GC/MS Analysis of C. sativum Leaves EO

The EO of C. sativum obtained by hydro-distillation was analyzed for its chemical composition. Nineteen components, representing 95.30% of the oil, were identified (Table 3). The major constituents identified in the EO of the coriander leaves were (E)-2-decenal (29.87%), linalool (21.61%), (E)-2-dodecenal (7.03%), dodecanal (5.78%), (E)-2-undecenal (3.84%), (E)-2-tridecenal (3.56%), (E)-2-hexadecenal (2.47%), tetradecenal (2.35%), and α-pinene (1.64%).

The previous studies about C. sativum EO composition showed that seed, fruit, and leaf oils of C. sativum contained linalool, (E)-2-decenal, dodecanal, α-pinene, camphor, cymene, myrcene, geranyl acetate, borneol, terpinen-4-ol, geraniol, and tetradecenal as predominant compounds.^{[2,6,8,18–20]} The results of C. sativum leaf EO composition reported by Shahwar et al.^[8] were in quite good accordance with our findings. Linalool and (E)-2-decenal were found in the EO of C. sativum as major components. However, the percentage of linalool (13.97%) was quite low compared to our result (21.61%). In addition to these, decenal, 1-decanol, undecanal, (E)-2-tridecanal, tetradecenal, dodecanal, tridecenal, and carvone values found in our work were higher than those found by Shahwar et al.^[8] These qualitative and quantitative differences in the EO composition of C. sativum may be attributed to climatic and environmental factors.

Total Phenolic Content

The total phenolic contents of the EO and ethanol extract determined by Folin–Ciocalteu method are reported to be gallic acid equivalents (GAE; Table 4). The amount of total phenolic compounds in the ethanol extract of plant material was determined as 14.973 mg GAE/g dry weight (DW), but phenolic compounds were not detected in the EO. Similar results were also obtained by Sriti et al.^[21] These authors reported that total phenolic contents found in coriander fruit methanolic extracts were 15.16 mg GAE/g DW. Shahwar et al.^[8] found that the methanol extract of coriander leaves is rich in total phenolic contents, which contain 30.25 mg/g.

Antioxidant Activity

In the present study, the antioxidant activity, determined by two different methods, namely DPPH and β-carotene-linoleic acid, is presented in Table 4. The vital role of the antioxidants is their interaction with oxidative free radicals. The DPPH method with the stable organic radical DPPH is used for the determination of free radical scavenging activity, usually expressed as IC₅₀. The results are expressed as IC₅₀, the amount of antioxidant necessary to decrease the initial concentration of DPPH by 50%. The lower IC₅₀ values indicate a higher antioxidant activity

In DPPH assay, IC₅₀ values were 74.87 ± 0.03 μg/mL and 26.20 ± 0.01 μg/mL for ethanol extract of C. sativum and Trolox, respectively. Sriti et al. ^[21] have showed a higher free radical scavenging activity (IC₅₀ of 32 ± 0.78 μg/mL) in comparison with our result (IC₅₀ of 74.87 ± 0.03 μL/mL). Shahwar et al.^[8] found that methanol extract of coriander leaves has significant radical scavenging activity (72.19%) at a concentration of 500 μg. Our results reveal that the EO did not show any activity with DPPH. In a study conducted by Sriti et al.,^[21] the two methanolic extracts of coriander fruit (IC₅₀ = 32 µg/mL for Tunisian coriander, IC₅₀ = 36 µg/mL for Canadian coriander) showed higher scavenging ability on DPPH radicals when compared to those reported for EOs of two fruit used in research (IC₅₀ = 61,000 µg/mL for Tunisian coriander, IC₅₀ = 60,000 µg/mL for Canadian coriander). Similar results were obtained also by Bamoniri et al.^[22] In fact, EOs contain high concentration of monoterpenic compounds that were reported to be almost ineffective. This result is in agreement with the poor performance given by the oils with similar patterns and by single monoterpenic hydrocarbons. Therefore, antioxidant activity may vary, based on the variations in the chemical composition.^[23]

In most cases, the antioxidant activity of some plants is due to their polar extracts rather than other extracts or EOs. For instance, Gulluce et al.^[16] showed that the methanolic extract from Micromeria fruticosa ssp. serpyllifolia exhibited important antioxidant activity evaluated by scavenging of the free radical DPPH and inhibition of linoleic acid oxidation, while the EO of this plant exhibited no activity or very weak activity. The results here presented are in agreement with many previously published results which showed that polar solvent extracts has more higher antioxidant activity than lower polarity solvents.^[24]

The antioxidant activity of cilantro ethanolic extract was also evaluated by the BCB method. In the BCB assay, linoleic acid produces hydroperoxides during incubation at 50°C. The presence of hydroperoxides causes rapid discoloration of β-carotene. However, hydroperoxides formed in this system can be neutralized by the antioxidants from the extracts. Table 4 shows the mean total antioxidant activity of ethanol extract of C. sativum. The means of total antioxidant activity for C. sativum ethanol extract was 85.85 ± 0.04%. BHA, used as the standard, had a higher antioxidant activity (93.20 ± 0.01%) than C. sativum ethanol extract. The EO of the plant did not show any activity with β-carotene bleaching assay.

Antimicrobial Activity

C. sativum EO has been reported to inhibit a broad spectrum of and has proven its efficacy as an antibacterial agent.^[20,25,26] The antimicrobial activities of the EO and ethanol extract of C. sativum leaves against foodborne microorganisms were determined by evaluating the presence of inhibition zones, zone diameter and MIC values. Their potencies were assessed both qualitatively and quantitatively. The results are given in Table 1 and Table 2. The plant EO showed significant antimicrobial activities against many microbial strains. The ethanol extract of the plant showed weak antimicrobial activities against microbial strains in both disc diffusion and MIC tests. Generally, the EO of C. sativum possessed a stronger and broader spectrum of antimicrobial activity than the ethanol extract. The ethanol extract of the plant showed antibacterial activities against Flavobacterium indologenes, Enterococcus faecalis, Klebsiella pneumoniae ozaenae, Klebsiella pneumoniae, Proteus mirabilis, Providencia alcalifaciens, Pseudomonas pseudoalcaligenes, Streptococcus pyogenes, and Yersinia enterocolitica. On the other hand, the EO of the plant showed antibacterial activities against all microorganisms used in this study.

TABLE 3 Essential oil composition of Coriandrum sativum

Compound	Percentage of total (%)	RI^a
Nonane	0.18	867
Alpha-pinene	1.64	935
Beta-pinene	0.87	978
Linalool	21.61	1096
Camphor	0.53	1172
(E)-2-Decenal	29.87	1269
Decanal	2.85	1211
Carvone	1.75	1249
1-Decanol	3.97	1274
3-Decanol	0.21	1289
Undecanal	3.78	1302
Undecanol	0.95	1349
Tetradecenal	2.35	1392
Dodecanal	5.78	1412
(E)-2-Undecanal	3.84	1421
(E)-2-Dodecanal	7.03	1456
(E)-2-Tridecanal	3.56	1597
Tridecenal	2.06	1614
(E)-2-Hexadecanal	2.47	1859
Total identified	95.30

^aRetention index on HP-5 column.

TABLE 4 Antioxidant activity and total phenolic content of the essential oil and ethanol extract of C. sativum

Sample	DPPH IC50 (μg/mL)	β- caroten/linoleic acid assay (%)	Total phenolic content (mgGAE/g DW)
Ethanol extract	74.87 ± 0.03	85.85 ± 0.04	14.97 ± 0.05
Essential oil	nd	nd	nd
Trolox	26.20 ± 0.01	–	–
BHA	–	93.20 ± 0.01	–

nd: not determined.

In the current study, it is also important to note that the most important microorganisms in food security were tested. As seen in Table 1, Listeria monocytogenes and Staphylococcus aureus were bacteria influenced by the plant oil. Silva et al.^[6] reported that the potent antibacterial activity of coriander EO against Gram positive and Gram negative bacteria is due to membrane permeability. Moreover, cilantro EO was particularly effective against Listeria monocytogenes, likely due to the presence of alcohols and aldehydes.^[1]

All Candida strains (Candida albicans and Candida krusei) studied were inhibited by plant EO (Table 2). But the ethanol extract of the plant showed only weak inhibitory activitiy against Candida albicans. It is obvious from Table 2 that the MIC value of C. albicans of the cilantro oil is higher than the positive control, amphotericin B. Owing to its broad spectrum of antifungal effect and low toxicity, cilantro EO is a promising source in the search for new antifungal drugs and could be considered useful in treatment or prevention of Candida yeast infections.^[27] Silva et al.^[6] reported that coriander EO and its main component, linalool, significantly inhibited growth of Candida.

The oil inhibited all fungi tested (Aspergillus flavus, two different strains of Aspergillus niger, Cladosporium herbarum, Penicillium brevicompactum, Penicillium roquefortii, and Trichothecium roseum). But the ethanol extract of the plant did not show antifungal activities against fungal strains (Table 2). Darughe et al.^[28] stated that coriander EO with a range of 0.15% could inhibit the growth of fungus in the cake.

CONCLUSION

In recent years, the EOs of plants have drawn great interest as sources of natural products. EOs, possessing antimicrobial and antioxidant properties as well as several biological activities, can be used instead of synthetic antioxidants and antibiotics. In conclusion, the chemical compositions of C. sativum leaves’ EO was revealed, and also the antioxidant and antimicrobial activities and total phenolic content of C. sativum leaves’ EO and ethanol extract was evaluated. Thus, the knowledge of the oil composition and the antimicrobial and antioxidant activities of samples (EO and ethanol extract) can provide important benefits for the food and medical industries. Further studies are needed to determine the toxicity and safety of C. sativum.

ACKNOWLEDGMENTS

The author is grateful to Bulent Cetin in the Department of Food Engineering at Ataturk University in Erzurum, Turkey for his help and support.