Trachyspermum ammi L. essential oil as plant based preservative in food system

Trachyspermum ammi L. essential oil as plant based preservative in food system

Industrial Crops and Products 69 (2015) 104–109 Contents lists available at ScienceDirect Industrial Crops and Products journal homepage: www.elsevi...

1MB Sizes 0 Downloads 13 Views

Recommend Documents

No documents
Industrial Crops and Products 69 (2015) 104–109

Contents lists available at ScienceDirect

Industrial Crops and Products journal homepage:

Trachyspermum ammi L. essential oil as plant based preservative in food system Akash Kedia, Bhanu Prakash, Prashant K. Mishra, Abhishek K. Dwivedy, N.K. Dubey ∗ Laboratory of Herbal Pesticides, Centre of Advanced Study in Botany, Banaras Hindu University, Varanasi 221005, India

a r t i c l e

i n f o

Article history: Received 25 October 2014 Received in revised form 14 January 2015 Accepted 10 February 2015 Keywords: Ajowan Essential oil Aspergillus flavus Aflatoxin B1 Preservative

a b s t r a c t The study reports efficacy of Trachyspermum ammi fruit essential oil (EO) as plant based preservative in food system. GC–MS analysis of EO identified cymene as chief component (76.27%) followed by thymol (13.30%), dl-limonene (3.23%), 1,8-cineole (2.58%) and ␥-terpinene (1.68%). The minimum inhibitory (MIC) and minimum aflatoxin inhibitory concentrations (MAIC) of EO were 0.8 and 0.6 ␮L/mL, respectively, against Aspergillus flavus LHP(C)-D6. Among EO components, thymol showed the highest antifungal and antiaflatoxigenic activity suggesting synergistic mode of action of EO components. Microscopic observations at sub lethal and lethal concentrations of EO using SEM and TEM revealed its mode of action on fungal plasma membrane causing considerable reduction in ergosterol quantity and increased leakages of Ca+2 , K+ and Mg+2 from EO treated A. flavus cells. The EO completely inhibited aflatoxin B1 production in wheat and chickpea as analyzed through HPLC and caused no toxicity in seed viability. The LD50 of EO was 6620.43 ␮L/kg for oral-toxicity on mice and IC50 was 0.467 ␮L/mL through DPPH analysis. The ajowan EO may thus be formulated as safe plant based preservative for enhancement of shelf life of food items. © 2015 Elsevier B.V. All rights reserved.

1. Introduction In spite of the advancements in food production technologies, fungal and mycotoxin contamination on food commodities is still one of the major threats for their spoilage. Among food spoiling fungi, the dominance of Aspergilli, particularly Aspergillus flavus, is a matter of great concern in view of secretion of aflatoxins which are hidden killers to consumers as they are unavoidable contaminant of food items (Williams et al., 2004). Aflatoxin contamination in food items was found to be responsible for Indian childhood cirrhosis, a clinical condition mainly occur in the Indian subcontinent (Reddy and Raghavender, 2007). Aflatoxin B1 (AFB1 )-8,9-exoepoxide can also accelerate reactive oxygen species formation leading to lipid peroxidation which is an another major problem for qualitative losses of food items (Prakash et al., 2012a). Most of the synthetics preservatives used for food preservation cause multiple side effects to human health due to residual toxicity, provoking the application of some safer preservatives. In this prospect, due to their volatility, plant-based essential oils (EOs) are gaining interest as they have long been used in traditional medicine and pharmaceutical preparations and many of them are generally recognized as safe (GRAS) (Prakash et al., 2012b). The antibacterial, antifungal, anti-

∗ Corresponding author. Tel.: +91 9415295765. E-mail address: [email protected] (N.K. Dubey). 0926-6690/© 2015 Elsevier B.V. All rights reserved.

cancer, antimutagenic, antidiabetic, antiviral, antiinflammatory, and antiprotozoal properties of essential oils are well documented (Raut and Karuppayil, 2014). Trachyspermum ammi L., commonly known as ajowan, is a well known traditionally used spice. Essential oil of T. ammi fruits has been previously investigated for its antimicrobial and antioxidant activity (Gandomi et al., 2013). Some preliminary reports on its antiaflatoxigenic (Gemeda et al., 2014) and antitermitic (Seo et al., 2009) nature also suggests its broad efficacy. However, investigations on mode of action of ajowan EO against aflatoxin secreting strains of A. flavus and efficacy in food system as preservative against fungal and aflatoxin contaminations are lacking. Therefore, the aim of the present study was to practically standardize the fungitoxic dose of chemically characterized ajowan EO and its major compounds, mechanism of action against food spoiling toxigenic A. flavus, efficacy in food system and safety profile so as to recommend it as plant based preservative to enhance the shelf life of food items. 2. Materials and methods 2.1. Extraction of ajowan EO The freshly harvested ajowan fruits (1–2 days after harvest, moisture content 8–10%) from 3 months old plants were collected from Botanical garden, Banaras Hindu University, Varanasi, India

A. Kedia et al. / Industrial Crops and Products 69 (2015) 104–109

(25◦ 20 N/83◦ 00 E) in the month of February, 2011. The EO was isolated from 500 g sample through hydro-distillation in Clevenger’s apparatus (Merck Specialities Pvt., Ltd., Mumbai, India). The volatile fraction (EO) was separated and traces of water removed by passing through anhydrous Na2 SO4 . The EO was collected in a clean dark glass vial and kept at 4 ◦ C until use. 2.2. GC and GC–MS analysis of ajowan EO The ajowan EO was subjected to gas chromatography (PerkineElmer Auto XL GC, MA, USA) with flame ionization detector having conditions: EQUITY-5 column (60 m × 0.32 mm × 0.25 ␮m); carrier gas H2 ; column head pressure 10 psi; oven temperature program isotherm 2 min at 70 ◦ C, 3 ◦ C/min gradient to 250 ◦ C, isotherm 10 min; injection temperature, 250 ◦ C; detector temperature 280 ◦ C. Analysis was also accomplished with a PerkinElmer Turbomass GC–MS following Kedia et al. (2014a). The identification of individual compounds was done with comparison of their relative retention times respect to control samples on capillary columns and through matching of their mass spectra of peaks with available mass spectral libraries of Wiley, NIST and NBS or with published data. From this study, cymene, thymol, dl-limonene, 1,8-cineole and ␥-terpinene (procured from Ozone International, Mumbai, India; purity >99%) (>1% contribution in EO composition) were selected to test antifungal and antiaflatoxigenic efficacy against A. flavus along with EO. 2.3. Culture conditions of test fungi Toxigenic strain of A. flavus strain LHP(C)-D6 along with 19 additional food spoiling fungi viz. Absidia ramosa, Alternaria alternata, Aspergillus fumigatus, Aspergillus glaucus, Aspergillus niger, Aspergillus terreus, Aspergillus unguis, Cladosporium cladosporioides, Curvularia lunata, Fusarium oxysporum, Mucor sp., Mycelia sterilia, Penicillium citrinum, Penicillium italicum, Penicillium luteum, Penicillium purpurogenum, Penicillium sp., Rhizopus stolonifer and Spondylocladium australe isolated previously from chickpea and wheat during mycobiota analysis (Kedia et al., 2014a) were selected in this study. Their cultures were routinely preserved on PDA slants at 4 ◦ C. 2.4. Evaluation of fungitoxicity of ajowan EO and the components against A. flavus Requisite amounts of test chemicals were dissolved separately in Petri plates containing 0.5 mL Tween-20 (5%) and then 9.5 mL PDA was added so as to obtain different concentrations (0.1–1.0 ␮L/mL). 5 mm diameter disk of seven days old culture of A. flavus was inoculated onto prepared Petri dishes. The control sets contained PDA with Tween-20 but no test chemicals. The Petri dishes were incubated at 27 ± 2 ◦ C for 7 days. Fungal colony diameters of treatment and control sets were measured. The lowest concentration resulting in no growth was noted. 2.5. Evaluation of ajowan EO and the components against aflatoxin B1 production Requisite amounts of test chemicals were dissolved separately in 0.5 mL Tween-20 (5%) and then to 24.5 mL SMKY medium (Sucrose, 200 g; MgSO4 ·7H2 O, 0.5 g; KNO3 , 0.3 g and yeast extract, 7 g; 1000 mL distilled water), so as to achieve different concentrations ranging from 0.1 to 1.0 ␮L/mL. The medium was then inoculated with a 5 mm diameter disk of 7-day-old culture of A. flavus. The control sets were run without adding the test chemicals.


After 10 days incubation at 27 ± 2 ◦ C, the content of each set was filtered through Whatman no. 1 filter paper and extracted with 20 mL chloroform in a separating funnel. The extract was evaporated to dryness on a water bath and re-dissolved in 1 mL chloroform. Fifty microliters of chloroform extract was spotted on TLC plate along with a standard of AFB1 (Hi-Media Laboratories Pvt., Ltd., Mumbai, India) and developed in toluene:isoamyl-alcohol:methanol (90:32:2; v/v/v). The plate was air-dried and observed in UV transilluminator (360 nm). The blue spots were scrapped, dissolved in methanol (5 mL) and centrifuged at 3000 rpm (5 min). The supernatant was collected and absorbance was recorded in spectrophotometer at 360 nm and AFB1 was estimated as: AFB1 content (␮g/L) = D × M/E × L × 1000, where D = absorbance, M = molecular weight of AFB1 (312), E = molar extinction coefficient (21,800), and L = path length (1 cm cell was used) (Kedia et al., 2014b). 2.6. Fungitoxic spectrum of EO against 19 food spoiling molds The EO at its MIC (0.8 ␮L/mL) was added separately to Petridishes containing 0.5 mL Tween-20 (5%) and then added 9.5 mL molten PDA to it. A 5 mm fungal disk from a seven day old culture from each fungus was placed separately on the center of the prepared Petri-plates. Plates containing only Tween-20 and PDA inoculated with the test fungi served as controls. The plates were incubated at 27 ± 2 ◦ C. After 7 days, fungal colony diameters of treatment and control sets were measured and the percent inhibition of fungal growth was determined. 2.7. Mode of action of EO against aflatoxigenic strain of A. flavus 2.7.1. Electron microscopic observations For electron microscopic study, 5 days old culture of A. flavus grown on PDA plates was exposed to 0.4 and 0.8 ␮L/mL of EO while control sets were without EO. After 7 days, 2 × 2 mm segments from the margin were excised and placed in vials containing 2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer at 4 ◦ C and fixed overnight. The samples were rinsed thrice with the same buffer and post-fixed in 1% osmium tetraoxide for 1 h at 4 ◦ C. The specimens were then dehydrated in a graded series of acetone upto absolute for a period of 20 min in each dilution. For scanning electron microscopy (SEM), the samples were finally sputter-coated with gold and viewed under scanning electron microscope (Quanta 200F, the Netherlands). For transmission electron microscopy (TEM), the samples were cleaned with toluene and then polymerized in araldite CY212 (TAAB, UK) in an oven at 50 ◦ C for overnight. Ultrathin sections (60–70 nm) of the specimen blocks were hand trimmed with an ultra-microtome (Leica EMUC6) and placed on 300 mesh copper grids. The sections were stained with 12.5% alcoholic uranyl acetate for 20 min and then with lead–citrate. After washing with doubledistilled water for 1 min, the samples were dried and viewed under transmission electron microscope (Morgagni 268D) (Fei Company, the Netherlands) at an operating voltage 80 kV. Images were digitally acquired by a CCD camera (Megaview III, Fei Company) using iTEM software (Sift Imaging System, Münster, Germany) attached to the microscope. 2.7.2. Effect of EO on ergosterol content in plasma membrane of A. flavus To estimate ergosterol content in the plasma membrane, 100 ␮L aliquot of spore suspension of A. flavus (106 spores/mL) prepared in 0.1% Tween-20 (Tian et al., 2012) was inoculated in SMKY bearing 0, 0.2, 0.4, 0.6 and 0.8 ␮L/mL EO and incubated at 27 ± 2 ◦ C for 4 days. The mycelia were filtered, washed twice with distilled water and the net wet weight of each cell pellet was measured. To


A. Kedia et al. / Industrial Crops and Products 69 (2015) 104–109

each cell pellet, 5 mL of 25% alcoholic potassium hydroxide solution was added and vortex-mixed for 2 min, followed by incubation in a water bath (85 ◦ C) for 4 h. From each sample, sterols were extracted by adding a mixture of 2 mL sterile distilled water and 5 mL n-heptane followed by sufficient vortex mixing for 2 min. After 1 h, the n-heptane layer was separated and examined by scanned spectrophotometry between 230 and 300 nm. A characteristic four peaked curve depicted the presence of ergosterol and the late sterol intermediate 24(28) dehydroergosterol in the n-heptane layer. A flat line indicated the absence of ergosterol. The amount of ergosterol was calculated following Tian et al. (2012). 2.7.3. Determination of ions leakage The biomass of A. flavus grown on PDB medium for 5 days, was filtered, washed with sterile distilled water thrice, suspended in 20 mL 0.85% saline solution and fumigated with 0.4 and 0.8 ␮L/mL EO for 12 h. Control sets received no EO. The biomass was filtered again and the supernatant was analyzed using atomic absorption spectrometry (PerkinElmer, AAnalyst 800, USA) for Ca+2 , K+ and Mg+2 ions (Helal et al., 2007) 2.8. Efficacy of EO as food preservative 2.8.1. In vivo investigations with ajowan EO on fungal and aflatoxin contamination of wheat and chickpea samples during storage Experiments were designed to fumigate the wheat and chickpea seed samples with ajowan EO in airtight containers following Prakash et al. (2012b). In one set (uninoculated treatment), EO soaked cotton swab was introduced in airtight closed plastic container (2 L), containing 1 kg of wheat (var. K68, moisture content 12–14%) and 1 kg of chickpea (var. Samrat, moisture content 15–16%) separately to achieve a concentration 0.8 ␮L/mL air (similar to MIC). In another set (inoculated treatment), prior to treatment, the samples were inoculated with 3 mL spore suspension of A. flavus. The control set also contained the uninoculated control and the inoculated control. After 12 months of storage at 10–46 ◦ C and RH 30–90%, food samples were subjected to mycoflora analysis to check A. flavus contamination using the direct plating and serial dilution methods (Kedia et al., 2014a). To estimate AFB1 content in each set, HPLC analysis with photo-diode-array (PDA) detector (Waters, Bangalore, India) was performed. The samples for HPLC were prepared by homogeneous liquid–liquid extraction (HLLE) method following SheijooniFumani et al. (2011). All aqueous solutions were prepared using double-distilled–deionised water developed by a Milipore-DQ3 system (Synergy, Germany). Five g well milled sample was mixed with 10 mL methanol/water (8:10) and shaken at 400 rpm for 30 min on a mechanical shaker. The sample was centrifuged at 3000 rpm for 5 min. Four milliliter supernatant was then mixed with 300 ␮L chloroform and 6 mL water containing 3% potassium bromide. After 5 min centrifugation at 3000 rpm, the settled phase was collected in a screw cap vial and dried in water bath under steam of nitrogen. To this, 50 ␮L methanol (HPLC grade) was added and finally injected to HPLC. A calibration curve of standard solution of aflatoxin B1 was prepared in a range of 12.5–500 ng/50 ␮L. Separation was carried out on a C-18 reverse phase column (250 mm × 4.6 mm i.d × 5 ␮m) and the mobile phase was methanol–acetonitrile–water (17:19:64 v/v) at a flow rate at 1.2 mL/min. Aflatoxin B1 was detected at 365 nm (Rohman and Triwahyudi, 2008). 2.8.2. Effect on germination of fumigated seeds The germination of wheat and chickpea seeds, fumigated with EO upto 12 months of storage was tested by placing 50 randomly selected seeds along with fresh seeds in Petri plates containing

Table 1 Chemical profile of Trachispermum ammi fruit essential oil. Sr. no.


Retention time (min)


1 2 3 4 5 6 7 8

2-Isopropyloxetane 1-Heptyn-3-ol 2,6-Dimethyl-3-heptanone Cymene dl-Limonene 1,8-Cineole ␥-Terpinene Thymol

10.825 11.150 11.375 12.53 12.625 12.775 13.801 24.58

0.25 0.25 0.90 76.27 3.23 2.58 1.68 13.30 Total: 98.46

moistened blotting paper and 50 seeds in small earthen pots. The seeds germinated within a week were recorded as viable. The per cent germination was calculated with respect to control sets. 2.8.3. Determination of LD50 on mice The LD50 value, which represents the lethal dose per unit body weight for killing 50% population was calculated on mice (Mus musculus L., average weight 30 g, age 3 months) through oral toxicity (Kedia et al., 2014b). Different doses of EO (0.05–0.5 mL) were orally distributed with 0.5 mL stock solution (Tween-80 and distilled water in a 1:1 ratio) separately to each group of animals (10 male mice). In control set, only 0.5 mL stock solution was orally administrated. After 4 h, the mortality was recorded and the LD50 value was calculated through Probit analysis. 2.8.4. Antioxidant activity through DPPH free radical analysis Different concentrations of EO (0.1–1.0 ␮L/mL) were added to 5 mL 0.004% DPPH solution in methanol. The absorbance was measured at 517 nm against a blank using a spectrophotometer after 30 min incubation at room temperature (25 ± 2 ◦ C). The IC50 value, which represented the concentration that caused 50% neutralization of DPPH radicals, was measured from the graph plotting percentage inhibition against concentration. The percent inhibition was calculated from the formula, I% = (Ablank − Asample /Ablank ) × 100 where, Ablank is the absorbance of the control (without test material) and Asample is the absorbance of the test material (Mishra et al., 2012). 2.9. Statistical analysis All experiments were repeated thrice and data are the mean ± standard error subjected to one way ANOVA. Means were separated by Tukey’s multiple range tests when ANOVA was significant (p < 0.05). Probit analysis was performed to estimate lethal dose (LD50 ) with their 95% confidence limits. The analysis of data was performed with the SPSS program version 16.0. 3. Results and discussion The yield of EO ranged between 2.5 and 2.8% on fresh weight basis. GC–MS analysis depicted the presence of 8 compounds comprising 98.46% of total EO (Table 1). The major compounds were identified as cymene (76.27%) followed by thymol (13.30%), dllimonene (3.23%), 1,8-cineole (2.58%) and ␥-terpinene (1.68%). In most of the earlier reports with ajowan fruit EO, thymol was identified as the chief component (Khajeh et al., 2004; Mahboubi and Kazempour, 2011; Gandomi et al., 2013). In the present investigation, cymene was found to be the chief component. Such variation could be due to the ecological and geographical conditions, seasons, age of plant and time of harvesting (Burt, 2004). During fungitoxic assay against A. flavus only ajowan EO and thymol showed complete inhibition with MIC at 0.8 and 0.1 ␮L/mL, respectively (Table 2). Rest of the oil components exhibited fungi-

A. Kedia et al. / Industrial Crops and Products 69 (2015) 104–109


Table 2 Antifungal activity of ajowan EO and the components against A. flavus LHP(C)-D6. Conc. (␮L/mL)

Diameter of A. flavus LHP(C)-D6 (cm) EO






Control 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0

5.90 ± 0.17a 4.70 ± 0.06b 3.87 ± 0.07c 2.43 ± 0.03d 1.70 ± 0.15e 1.37 ± 0.07ef 1.10 ± 0.10f 0.73 ± 0.03g 0.00 ± 0.00h 0.00 ± 0.00h 0.00 ± 0.00h

5.90 ± 0.17a 5.93 ± 0.09a 5.57 ± 0.12a 5.33 ± 0.09ab 5.30 ± 0.15bc 5.23 ± 0.12bc 5.13 ± 0.09bc 4.97 ± 0.09c 4.93 ± 0.12c 4.90 ± 0.15c 4.77 ± 0.12c

5.90 ± 0.17a 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b

5.90 ± 0.17a 5.73 ± 0.23a 5.70 ± 0.20a 5.67 ± 0.28a 5.63 ± 0.18a 5.60 ± 0.18a 5.43 ± 0.17a 5.40 ± 0.15a 5.23 ± 0.09ab 5.03 ± 0.07ab 4.47 ± 0.15b

5.90 ± 0.17a 4.53 ± 0.15b 4.27 ± 0.17b 4.10 ± 0.12bc 3.60 ± 0.15cd 2.67 ± 0.12d 2.27 ± 0.17de 2.17 ± 0.07de 1.97 ± 0.07ef 1.80 ± 0.10ef 1.50 ± 0.15f

5.90 ± 0.17a 5.20 ± 0.15b 5.20 ± 0.10b 4.90 ± 0.12bc 4.50 ± 0.12c 3.63 ± 0.03d 3.43 ± 0.24de 3.23 ± 0.13de 3.13 ± 0.03de 2.97 ± 0.07e 2.90 ± 0.10e

Values are mean (n = 3) ± standard error. The means followed by same letter in the same column are not significantly different according to ANOVA and Tukey’s multiple comparison tests.

Table 3 Antiaflatoxigenic activity of ajowan EO and the components against A. flavus LHP(C)-D6. Conc. (␮L/mL)

Control 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0

Aflatoxin content (␮g/L) EO






1758.98 ± 35.97a 1435.67 ± 36.68b 1050.53 ± 29.88c 455.71 ± 224.79d 333.94 ± 34.40de 152.66 ± 20.79de 0.00 ± 0.00f 0.00 ± 0.00f 0.00 ± 0.00f 0.00 ± 0.00f 0.00 ± 0.00f

1758.98 ± 35.97a 1747.10 ± 21.94ab 1747.38 ± 26.16ab 1684.28 ± 14.86abc 1667.68 ± 30.76abc 1635.65 ± 25.38bc 1637.23 ± 19.91bc 1614.38 ± 19.75c 1605.56 ± 10.58c 1594.60 ± 34.04c 1566.62 ± 23.63c

1758.98 ± 35.97a 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b 0.00 ± 0.00b

1758.98 ± 35.97a 1799.86 ± 11.54a 1749.12 ± 21.12a 1732.71 ± 27.57ab 1650.16 ± 25.58bc 1637.68 ± 19.38bcd 1626.46 ± 12.97cd 1575.18 ± 24.13cde 1565.08 ± 19.20cde 1539.54 ± 21.82de 1523.01 ± 11.23e

1758.98 ± 35.97a 1633.28 ± 17.42b 1574.15 ± 23.75b 1505.19 ± 14.67bc 1401.62 ± 14.61cd 1283.15 ± 48.68d 1161.93 ± 20.99e 955.28 ± 15.81f 869.93 ± 33.48fg 795.35 ± 53.47g 589.81 ± 21.77h

1758.98 ± 35.97a 1653.49 ± 38.29ab 1628.34 ± 27.79ab 1601.12 ± 43.96b 1549.12 ± 29.91b 1337.37 ± 33.47c 1248.71 ± 27.72cd 1223.45 ± 57.14cde 1137.18 ± 23.92de 1078.36 ± 11.58e 913.80 ± 36.62f

Values are mean (n = 3) ± standard error. The means followed by same letter in the same column are not significantly different according to ANOVA and Tukey’s multiple comparison tests.

toxicity above 1 ␮L/mL. Similar results were also recorded during antiaflatoxigenic assay of ajowan EO and its components (Table 3). The findings emphasize that the overall activity of EO was due to the synergistic effects of its different components as cymene, the chief component, showed the poorest activity. Thymol was the main antimicrobial and anti-aflatoxigenic component of the ajowan EO and its strong activity is due to the presence of a phenolic OH group as suggested by Farag et al. (1989). The observation agrees with Delgado et al. (2004) that cymene has poor antimicrobial action and thymol when mixed with cymene, enhances the activity. In addition, ajowan EO also exhibited broad spectrum of fungal toxicity causing 100% growth inhibition of all 19 food spoiling fungal species at its MIC suggesting its recommendation for complete protection against food spoiling fungi. The mode of antimicrobial action of some preservative agents in food shows disruption of plasma membrane (Brul and Coote, 1999; Hazan et al., 2004). In the present investigation, through

SEM analysis, the EO fumigated hyphae were found abnormal with distorted conidiophores and conidia with conspicuous depressions when compared with control hyphae (Fig. 1). The degree of abnormality increased on high concentration. This study agrees with earlier observations of fumigation effects of lemongrass EO on A. flavus (Helal et al., 2007) and Tagetes patula EO on Botrytis cineria (Romagnoli et al., 2005). The distortion could be due to leakage of cell contents as supported by TEM observations. Fig. 2 illustrates the effect of EO on the ultrastructure of A. flavus. Control hyphae showed uniform plasma-membrane with normal cell organelles and abundant matrix. At 0.4 ␮L/mL of EO, the plasmalemma showed roughness with small lomasomes and decreased matrix. At 0.8 ␮L/mL, detachment of plasmalemma from the cell wall with abundant lomasomes was observed. Changes in the cell permeability due to destruction of plasma membrane caused loss of the normal shape and the formation of lomasomes inside the cells. This result supported the findings of Razzaghi-Abyaneh et al.

Fig. 1. Scanning electron microscopy illustrates the effect of ajowan EO on A. flavus morphology ((A), control, (B) treatment with 0.4 ␮L/mL EO, (C) treatment with 0.8 ␮L/mL EO).


A. Kedia et al. / Industrial Crops and Products 69 (2015) 104–109

Fig. 2. Transmission electron microscopy illustrates the effect of ajowan EO on A. flavus ultrastructure ((A), control, (B) treatment with 0.4 ␮L/mL EO, (C) treatment with 0.8 ␮L/mL EO). PM = plasma membrane, N = nucleus, L = lomasomes. Table 4 Efficacy of ajowan EO as ergosterol synthesis inhibitor and ion leakages from A. flavus cells. Conc. (␮L/mL)

Ergosterol content (%)

Control 0.2 0.4 0.6 0.8

0.61 ± 0.01 0.53 ± 0.02b 0.48 ± 0.01b 0.13 ± 0.01c 0.00 ± 0.00d

Ion content (␮L/L) Ca+2



5.49 ± 0.28 – 6.28 ± 0.08b – 7.36 ± 0.21c a


108.77 ± 15.33 – 586.33 ± 103.22b – 900.00 ± 10.07c a

5.65 ± 0.24a – 7.10 ± 0.33b – 8.22 ± 0.19c

Values are mean (n = 3) ± standard error. The means followed by same letter in the same column are not significantly different according to ANOVA and Tukey’s multiple comparison tests.

(2006). Hence, plasma membrane is the important primary target of EO. Further, to verify the effects on plasma membrane, the effect of EO on the amount of ergosterol was also measured. A dose dependent decrease in ergosterol content was observed on increasing concentration of the EO (Table 4). Ergosterol is specific to fungi and is the major sterol component of the fungal cell membrane responsible for maintaining the cell function and integrity (Rodriguez et al., 1985). Thus, the antimicrobial components of the essential oils interacted with the membrane, causing misbalance in cell permeability and disruption in fungal cell organization leading to their death (Tian et al., 2012). The ion leakage test showed increased leakage from the fumigated hyphae on increasing the EO concentration as compared to control (Table 4). The increased leakage of these ions might have occurred as a consequence of cell death at sub lethal and lethal concentrations of EO as supported by TEM observations. The EO showed remarkable efficacy in protecting the food commodities from A. flavus contamination up to 12 months of storage. For wheat, the percent protection was found to be 46.15% for inoculated samples and 75.86% for uninoculated samples. Similarly, in case of chickpea, 65.22% protection was recorded for inoculated sets and 70.83% for uninoculated samples. The less protection in inoculated sets may be due to increased inoculum density. From this observation, it can be concluded that a higher concentration above MIC is required for complete protection as some amount of EO might be absorbed by the food system (Varma and Dubey, 2001). Another, significant result of the present study was the efficacy of ajowan EO in checking aflatoxin B1 contamination in food system during in vivo trials in storage containers. The aflatoxin B1 content estimated during HPLC analysis revealed a good linearity by determination coefficients (r2 ) 0.997. The calibration graph was linear (Fig. 3), Limit of detection (LOD) value was 0.286 ng/g and the recovery of AFB1 was 74.13%. The mean AFB1 levels in inoculated wheat and chickpea samples in control sets were 14.35 and 21.01 ␮g/kg, respectively. For uninoculated control sets, the values

Fig. 3. Calibration curve for aflatoxin B1 in HPLC method.

were 12.86 and 16.19 ␮g/kg, respectively. However, aflatoxin B1 was not detected in any of the EO fumigated samples. The aflatoxin inhibition may be attributed to reduced fungal growth, however, some other factors may also govern inhibition of aflatoxin production as EO did not caused complete protection from A. flavus contamination. Tian et al. (2012) stated that the EO can inhibit carbohydrate catabolism in Aspergillus species by acting on some key enzymes, reducing its ability to produce aflatoxins. Moreover, because of the lesser fungal association in EO fumigated sets, the fungal cells used most of the available energy to sustain viability, not for toxin production as has been emphasized by Burt (2004). Although, different EOs have been previously tested for their in vivo fungitoxicity of food commodities like cereals, pulses, spices, fruits and vegetables (Varma and Dubey, 2001; Tripathi and Dubey, 2004; Tripathi and Kumar, 2007; Prakash et al., 2012c) the in vivo fungitoxicity of ajowan EO against fungal as well as aflatoxin contamination in food preservation has been tested for the first time in the present study. The application of EO in food preservation would be one of the economically viable and effective stratagies due to its volatility and penetration in food systems. Further, the EO can be eliminated during sun drying. The seed germination tests revealed 100% seeds germination in EO treated sets showing its nonphytotoxic nature. The LD50 value of ajowan EO was found to be 6620.43 ␮L/kg body weight on mice which was found higher than the values for previously reported botanicals viz. pyrethrum, carvone and some commercial pesticides viz. Bavistin (Prakash et al., 2011). The high LD50 value of ajowan EO will harbor high safety profile with mammals if applied to fumigate food commodities. The oil showed strong

A. Kedia et al. / Industrial Crops and Products 69 (2015) 104–109

free-radical-scavenging activity in a dose-dependent manner and its IC50 value was recorded as 0.467 ␮L/mL which was found better than some of the earlier reported essential oils viz. Origanum majorana, Coriandrum sativum, Hedychium spicatum, Commiphora myrrha and Cananga odorata and synthetic preservative salicylic acid (Prakash et al., 2012b). Ajowan is a common plant growing abundantly in many parts of the world including India and have several applications in ethnomedicine and as flavoring agent. Because of renewable source, high yield of EO, and efficacy at low concentrations, the ajowan EO may be economically recommended for formulation as safe plant based preservative for food safety and enhancement of shelf life of food items. 4. Conclusion In view of strong antifungal, antiaflatoxigenic, antioxidant efficacy and favorable safety limits, the ajowan EO may be formulated as plant based food preservative by agro-industries for enhancement of shelf life of food items from fungal and aflatoxin contamination as well as their deterioration due to lipid peroxidation. Its efficacy in food system strengthens to carry out further large scale experiments for its future recommendation as a better alternative of synthetic preservatives. Acknowledgements This work was supported by University Grant Commission, New Delhi, India. Authors are thankful to SAIF, AIIMS, New Delhi and IIT, BHU, Varanasi, India for TEM and SEM facility. References Brul, S., Coote, P., 1999. Preservative agents in foods: mode of action and microbial resistance mechanisms. Int. J. Food Microbiol. 50 (1), 1–17. Burt, S., 2004. Essential oils: their antibacterial properties and potential applications in foods – a review. Int. J. Food Microbiol. 94, 223–253. Delgado, B., Fernandez, P.S., Palop, A., Periago, P.M., 2004. Effect of thymol and cymene on Bacillus cereus vegetative cells evaluated through the use of frequency distributions. Food Microbiol. 21, 327–334. Farag, R.S., Daw, Z.Y., Abo-Raya, S.H., 1989. Influence of some spice essential oils on Aspergillus parasiticus growth and production of aflatoxins in a synthetic medium. J. Food Sci. 54 (1), 74–76. Gandomi, H., Abbaszadeh, S., Jebellijavan, A., Sharifzadeh, A., 2013. Chemical constituents, antimicrobial and antioxidative effects of Trachyspermum ammi essential oil. J. Food Process. Preserv. 38, 1690–1695. Gemeda, N., Woldeamanuel, Y., Asrat, D., Debella, A., 2014. Effect of Cymbopogon martinii, Foeniculum vulgare and Trachyspermum ammi essential oils on the growth and mycotoxins production by Aspergillus species. Int. J. Food Sci. 2014, 1–9. Hazan, R., Levine, A., Abeliovich, H., 2004. Benzoic acid, a weak organic acid food preservative, exerts specific effects on intracellular membrane trafficking pathways in Saccharomyces cerevisiae. Appl. Environ. Microbiol. 70 (8), 4449–4457. Helal, G.A., Sarhan, M.M., Abu Shahla, A.N.K., Abou El-Khair, E.K., 2007. Effects of Cymbopogon citratus L. essential oil on the growth: morphogenesis and aflatoxin production of Aspergillus flavus ML2-strain. J. Basic Microbiol. 47, 5–15. Kedia, A., Prakash, B., Mishra, P.K., Chanotiya, C.S., Dubey, N.K., 2014b. Antifungal antiaflatoxigenic, and insecticidal efficacy of spearmint (Mentha spicata L.) essential oil. Int. Biodeterior. Biodegrad. 89, 29–36.


Kedia, A., Prakash, B., Mishra, P.K., Dubey, N.K., 2014a. Antifungal and antiaflatoxigenic properties of Cuminum cyminum (L.) seed essential oil and its efficacy as a preservative in stored commodities. Int. J. Food Microbiol. 168–169, 1–7. Khajeh, M., Yamini, Y., Sefidkon, F., Bahramifar, N., 2004. Comparison of essential oil composition of Carum copticum obtained by supercritical carbon dioxide extraction and hydrodistillation methods. Food Chem. 86, 587–591. Mahboubi, M., Kazempour, N., 2011. Chemical composition and antimicrobial activity of Satureja hortensis and Trachyspermum copticum essential oil. Iran. J. Microbiol. 3 (4), 194–200. Mishra, P.K., Shukla, R., Singh, P., Prakash, B., Dubey, N.K., 2012. Antifungal and antiaflatoxigenic efficacy of Caesulia axillaris Roxb. essential oil against fungi deteriorating some herbal raw materials and its antioxidant activity. Ind. Crop. Prod. 36, 74–80. Prakash, B., Shukla, R., Singh, P., Mishra, P.K., Dubey, N.K., Kharwar, R.N., 2011. Efficacy of chemically characterized Ocimum gratissimum L. essential oil as an antioxidant and a safe plant based antimicrobial against fungal and aflatoxin B1 contamination of spices. Food Res. Int. 44, 385–390. Prakash, B., Singh, P., Kedia, A., Dubey, N.K., 2012b. Assessment of some essential oils as food preservatives based on antifungal, antiaflatoxin, antioxidant activities and in vivo efficacy in food system. Food Res. Int. 49 (1), 201–208. Prakash, B., Singh, P., Kedia, A., Singh, A., Dubey, N.K., 2012a. Efficacy of essential oil combination of Curcuma longa L. and Zingiber officinale Rosc. as a postharvest fungitoxicant aflatoxin inhibitor and antioxidant agent. J. Food Saf. 32, 279–288. Prakash, B., Singh, P., Mishra, P.K., Dubey, N.K., 2012c. Safety assessment of Zanthoxylum alatum Roxb. essential oil, its antifungal, antiaflatoxin, antioxidant activity and efficacy as antimicrobial in preservation of Piper nigrum L. fruits. Int. J. Food Microbiol. 153 (1), 183–191. Raut, J.S., Karuppayil, S.M., 2014. A status review on the medicinal properties of essential oils. Ind. Crop. Prod. 62, 250–264. Razzaghi-Abyaneh, M., Shams-Ghahfarokhi, M., Kawachi, M., Eslamifar, A., Schmidt, O.J., Schmidt, A., et al., 2006. Ultrastructural evidences of growth inhibitory effects of a novel biocide, Akacid® plus, on an aflatoxigenic Aspergillus parasiticus. Toxicon 48 (8), 1075–1082. Reddy, B.N., Raghavender, C.R., 2007. Outbreaks of aflatoxicoses in India. Afr. J. Food Agric. Nutr. Dev. 7 (5), 1–15. Rodriguez, R.J., Low, C., Bottema, C.D., Parks, L.W., 1985. Multiple functions for sterols in Saccharomyces cerevisiae. Biochim. Biophys. Acta 837, 336–343. Rohman, A., Triwahyudi, 2008. Simultaneous determination of aflatoxin B1 , B2 , G1 and G2 using HPLC with photodiode-array (PDA) detector in some foods obtained from Yogyakarta, Indonesia. Agritech 28 (3), 109–112. Romagnoli, C., Bruni, R., Andreotti, E., Rai, M.K., Vicentini, C.B., Mares, D., 2005. Chemical characterization and antifungal activity of essential oil of capitula from wild Indian Tagetes patula L. Protoplasma 225, 57–65. Seo, S.M., Kim, J., Lee, S.G., Shin, C.H., Shin, S.C., Park, I.K., 2009. Fumigant antitermitic activity of plant essential oils and components from ajowan (Trachyspermum ammi), allspice (Pimenta dioica), caraway (Carum carvi), dill (Anethum graveolens), geranium (Pelargonium graveolens), and litsea (Litsea cubeba) oils against Japanese termite (Reticulitermes speratus Kolbe). J. Agric. Food Chem. 57 (15), 6596–6602. Sheijooni-Fumani, N., Hassan, J., Yousefi, S.R., 2011. Determination of aflatoxin B1 in cereals by homogeneous liquid–liquid extraction coupled to high performance liquid chromatography-fluorescence detection. J. Sep. Sci. 34, 1333–1337. Tian, J., Huang, B., Luo, X., Zeng, H., Ban, X., He, J., Wang, Y., 2012. The control of Aspergillus flavus with Cinnamomum jensenianum Hand.-Mazz essential oil and its potential use as a food preservative. Food Chem. 130, 520–527. Tripathi, P., Dubey, N.K., 2004. Exploitation of natural products as an alternative strategy to control postharvest fungal rotting of fruit and vegetables. Postharvest Biol. Technol. 32, 235–245. Tripathi, N.N., Kumar, N., 2007. Putranjiva roxburghii oil—a potential herbal preservative for peanuts during storage. J. Stored Prod. Res. 43 (4), 435–442. Varma, J., Dubey, N.K., 2001. Efficacy of essential oils of Caesulia axillaris and Mentha arvensis against some storage pests causing biodeterioration of food commodities. Int. J. Food Microbiol. 68, 207–210. Williams, H.J., Phillips, T.D., Jolly, E.P., Stiles, K.J., Jolly, M.C., Aggrwal, D., 2004. Human aflatoxicosis in developing countries: a review of toxicology, exposure, potential health consequences, and interventions. Am. J. Clin. Nutr. 80, 1106–1122.