International Journal of Drug Development and Research

Key words

Artemisia minor Jacq. ex Bess., chemotype, GCMS analysis, anti-microbial activity, Trilokinath (3020m), Lahaul & Spiti (Cold Desert).

Introduction

The family Asteraceae comprises of many aromatic and medicinal plants. The genus Artemisia known as “wormwood” belongs to the tribe Anthemideae and is one of the largest genera of the Asteraceae family. This family includes more than 800 species widely distributed throughout the world, especially, in South-West of Asia and Central Europe [1]. Several Artemisia species have been found to grow above 2600m. Out of 34 species of the genus Artemisia known from India, 15 species have been documented in the flora of Lahaul & Spiti [2]. The genus has always been of great interest to botanical, pharmaceutical and food industries [3]. Artemisia species are reported to possess anti-diabetic effect and have been used in many countries of middle east and Iran as a herbal medicine for treatment of diabetes, high blood pressure, anti-migraine, anti-fungal activity, useful as tonic, stomachic, anti-bacterial and anti-septic [4-5]. The essential oils from Artemisia genus are also used for various purposes such as flavoring, fragrances, rodents, mite repellents and for antispasmodic, anti-pyretic, anti-inflammatory and abortifacient activities. The Artemisia species have been known as a folk medicine resource, which is used for the treatment of epidemic hepatitis [6]. Because of application of Artemisia in traditional medicine, many species of this genus have been surveyed by phytochemists and pharmacologists [7-8]. Sesquiterpene lactones and acetylenes have been reported from some species of Artemisia such as A. assoana, A. lantana and A. pedemontana [9] and artemisia ketone, 1, 8-cineole, davanone, camphor, thujone, myrcene and germacrene-D have also been reported in essential oil of A. absinthium, A. scoparis and A. vulgaris [10-12]. The essential oil from various species of the genus are used in soaps, detergents, cosmetics, perfumes, as aromatherapy claims [13-14]. Chemical analysis of oils from different Artemisia species such as: A. afra, A. annua, A. arborescene, A. elegantissima, A. maritima, A. myriantha, A. nilagirica, A. scoparia, A. vulgaris, etc. have already been done by many workers from India and abroad [15- 19]. In spite of many studies on the genus Artemisia, there are still many problems in systematic interpretations. The quality and quantity of the essential oil varies a lot with the genetic makeup of the taxa and the prevalent environmental factors. As there is no report on the chemical analysis of essential oil and anti-microbial activity in A. minor Jacq. ex Bess. from this study area, the aim of this work is to provide the first report on the GCMS analysis of essential oil along with anti-microbial activity in the population of A. minor Jacq. ex Bess., a naturally growing species, from Trilokinath (3020m) - a cold desert region of India.

Materials and Methods

Plant materials

Aerial parts of A. minor at full flowering and fruiting stage were collected from Trilokinath, altitude range of (3020m) in the month of July, 2007 (Table 1). Plants specimens were identified by the Botanical Survey of India (BSI, Northern Circle), Dehradoon and specimens were deposited in the Herbarium, Department of Botany, Punjabi University, Patiala (Punjab), India.

Volatile oil distillation

One hundred grams fresh aerial parts of leaves and fine stems were separated and then immersed in water in a round bottom flask and hydrodistilled for 3h in a full glass Clevenger-type apparatus, giving transparent light yellow oil. The oil was decanted to be used as essential oil. To improve the recovery and analysis, the essential oil was taken in n-pentane, dried over anhydrous sodium sulphate until the last traces of water were removed and then stored in a dark glass bottle at 4oC prior to GC-MS analysis [20].

Gas chromatography-massspectrometry

GCMS (70ev) data were measured on GCMS (QP 2010 series Shimadzu, Tokyo, Japan) equipped with AOC 20i autosampler and BP20 Capillary column (SGC International Ringwood, Australia) of 30m length, 0.25mm i.d. and 0.25µm film thickness. Temperature was programmed from 70- 220ºC at a rate of 4ºC/min, held isothermally at 70ºC and 220ºC for 4 and 5 min, respectively. Mass spectrometer source temperature, 200ºC; interface temperature, 220ºC; injector temperature, 220ºC. Sample injection volume 2µL (diluted 5µL oil in 2mL dichloromethane, HPLC grade); split ratio, 1:50 and mass scan, 50-600 amu. Helium was used as a carrier gas with 1.1mL/min flow rate.

Identification of components

The retention index was calculated for all volatile constituents using a homolog4us series of n-alkanes. The components of oil were identified by matching their massspectra with those stored in the computer library such as Wiley, New York mass spectral (MS) library, National Institute of Standards and Technology (NIST) [21] and their retention indices (RI) either with authentic compounds or with published data in the literature [22-25] based on retention indices of components on same phases of polar columns such as: BP-20, CW-20M, HP-20M and Supelcowax-10.

Microbial strains for anti-microbial activity

The microorganism strains used in the agar disc diffusion method were supplied by the Institute of Microbial Technology, Chandigarh, India. Gram-positive bacteria: Bacillus subtilis (MTCC-2451), Staphylococcus aureus (MTCC-740), Staphylococcus epidermis (MTCC-435), Gram-negative bacteria: Escherichia coli (MTCC-443), Salmonella typhimurium (MTCC-1251), Pseudomonas fluorescence (MTCC-664) and Acenetobactor calcoaceticus (MTCC-127).

Anti-microbial screening

In vitro anti-bacterial activity of the A. minor essential oil was studied against seven bacterial strains by the agar well diffusion method as described by Perez and co-workers [25] with certain modifications. Nutrient agar (Hi Media, India) was used as the bacteriological medium. The antibacterial activity of essential oils was taken at different concentrations (5, 10, 20, 40 and 80µL/well). The nutrient agar was melted and cooled to 48-50oC and a standardized inoculum of 1 × 106 CFU/mL, (0.5 McFarland) was then added aseptically to the molten agar and poured into sterile petri dishes to give a solid plate. Wells were prepared in the seeded agar plates. The test compound was introduced in the well (8.5 mm). The plates were incubated overnight at 37oC. The anti-microbial spectrum of the oils was determined for the bacterial species in terms of zone sizes around each well. The diameters of zone of inhibition produced by the agent were compared with those produced by the commercial control antibiotics, 20µL each of amoxicillin and ciprofloxacin (5mg/mL of autoclaved distilled water). These are commonly used anti-biotics to treat infections caused by several Gram-positive and Gram-negative bacteria. For each bacterial strain positive controls were maintained. The experiment was performed three times to minimize the error and the mean values are presented.

Minimal inhibition concentration

The essential oils that exhibited considerable activity were diluted with nutrient broth (1:1) in a series of seven sets of three test tubes for different microorganisms [26]. An aliquot of 1mL of the bacterial suspension (1x106) was inoculated into each tube. The control tubes were inoculated with same quantity of sterile distilled water and 75% ethanol. All tubes were incubated at 37oC for 24hrs. The lowest concentration that did not permit any visible growth when compared with the control was considered as the minimum inhibitory concentration. The contents of all tubes that showed no visible growth were cultured on nutrient agar, incubated at 37oC for 24hrs. The minimum bactericidal concentration was considered as the lowest concentration that could not produce a single bacterial colony.

Results and Discussion

The essential oil obtained by hydrodistillation from the aerial parts of wild growing A. minor, with a yield of 0.40%, was pale yellowish and possessed a strong odour. By gas chromatography mass spectroscopy, 18 out of 22 components were identified, representing 65.37% of the composition of essential oil. The major constituents recorded from essential oil of A. minor were: 1, 8-cineole (22.30%), camphor (12.64%), davanone (12.33%), ascaridole (11.11%) and á-phellandrene (5.23%). The different constituents of essential oil of A. minor are well represented with their relative area percentage (Table 2). In the previous published reports on other species of the genus Artemisia, cineol, thujone and monoterpenes were reported to be the major constituents in A. maritima [27] and á-thujone (63.25%); sabinene (7.83%); 1, 8-cineole (6.54%) and germacrene-D (2.22%) were also considered as major components in A. annua. In one recent report on populations of A. maritima from Lahaul & Spiti, 1, 8–cineole, chrysanthenone, germacrene-D and borneol were recorded as major components, with small percentages of rare artedouglasia oxides was also recorded in one population [28].

In spite of many studies on the genus Artemisia, there is no report on essential oil composition of A. minor from the study area. Therefore, it is interesting to find such a high percentage of major constituents, 1, 8-cineole (22.30%), camphor (12.64%), davanone (12.33%), ascaridole (11.11%), á- phellandrene (5.23%) and one minor but rare constituent i.e. artedouglasia oxide-C (0.72%) first time in essential oil of the species in the population of A. minor from Trilokinath of Lahaul & Spiti (Cold Desert) region of North Indian higher altitude Himalayas. This observation about the presence of artedouglasia oxide-C in A. minor has not been reported earlier in the same species, but reported only in the oil of A. maritima [28] and A. laciniata [29], from which it is concluded that, A. minor oil from Trilokinath (3020m) is a new chemotype.

Anti-microbial activity showed that, the inhibition zones were found increased considerably when the concentration rate increased. Therefore it can be said that quantity of the oil was important for inhibition effect. Among all Gram-positive bacterial growths, the maximum zone of inhibition was recorded against Bacillus subtilis (MTCC-2451) i.e. 33mm, followed by Staphylococcus epidermis (MTCC-435) i.e. 27mm and 25mm zone of inhibition against Staphylococcus aureus (MTCC- 740). On the other hand four different Gram-negative bacterial strains were tested and among these microorganisms Pseudomonas fluorescence (MTCC-664) showed maximum zone of inhibition i.e. 32mm, followed by Acenetobactor calcoaceticus (MTCC- 127) i.e. 23mm. The minimum zone of inhibition was recorded against the Escherichia coli (MTCC-443) strain i.e. 4mm (Table 3). The minimal inhibition concentration (MIC) was 1µL recorded in Gram-negative strain Pseudomonas fluorescence (MTCC-664) followed by a Gram-positive strains Bacillus subtilis (MTCC-2451) and Staphylococcus aureus (MTCC-740), both showed 5µL of minimal inhibition concentration (MIC) (Table 4).

From these it is concluded that the essential oil showed maximum zone of inhibition and minimal inhibition concentration against Bacillus subtilis (MTCC-2451) and Pseudomonas fluorescence (MTCC-664) bacterial strains, which indicate that A. minor Jacq. ex Bess. essential oil has capacity to inhibit the growth of both Grampositive and Gram-negative bacterial strains when used in a higher amount. Further it can be concluded on the basis of previous studies on Artemisia genus and present results that A. minor Jacq. ex Bess. a higher altitude medicinal and aromatic plant act as an important anti-microbial agent against many Gram-positive and Gram-negative bacterial strains and a new chemotype with higher percentage of some most important chemical constituents, which was previously undescribed from this study area of “Cold Desert.”

Acknowledgments

The authors are grateful to His Holiness Baba Iqbal Singh Ji President, The Kalgidhar Trust & Founder Chancellor of Eternal University (H.P.), Hon’ble Vice- Chancellor Dr. Manmohan S. Atwal, Eternal University (H.P.) India, the Director, IHBT (CSIR), Palampur (H.P.) and Head, Department of Botany, Punjabi University Patiala (Punjab), India for providing necessary facilities and support.

Conflict of Interest

NIL

Source of Support

NONE

Tables at a glance

Table 1 Table 2 Table 3 Table 4

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