International Journal of Food Microbiology 168–169 (2014) 1–7
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Antifungal and antiaflatoxigenic properties of Cuminum cyminum (L.) seed essential oil and its efficacy as a preservative in stored commodities Akash Kedia, Bhanu Prakash, Prashant K. Mishra, N.K. Dubey ⁎ Laboratory of Herbal Pesticides, Centre of Advanced Study in Botany, Banaras Hindu University, Varanasi 221005, UP, India
a r t i c l e
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Article history: Received 7 June 2013 Received in revised form 1 October 2013 Accepted 15 October 2013 Available online 23 October 2013 Keywords: Aspergillus flavus Cumin seed essential oil Antifungal Antioxidant Aflatoxin Shelf life
a b s t r a c t The study reports potential of Cuminum cyminum (cumin) seed essential oil (EO) as a plant based shelf life enhancer against fungal and aflatoxin contamination and lipid peroxidation. The EO showed efficacy as a preservative in food systems (stored wheat and chickpeas). A total of 1230 fungal isolates were obtained from food samples, with Aspergillus flavus LHP(C)-D6 identified as the highest aflatoxin producer. Cumin seed EO was chemically characterized through GC–MS where cymene (47.08%) was found as the major component. The minimum inhibitory concentration and minimum aflatoxin inhibitory concentration of EO were 0.6 and 0.5 μl/ml respectively. The EO showed toxicity against a broad spectrum of food borne fungi. The antifungal action of EO on ergosterol content in the plasma membrane of A. flavus was determined. The EO showed strong antioxidant potential having IC50 0.092μl/ml. As a fumigant in food systems, the EO provided sufficient protection of food samples against fungal association without affecting seed germination. In view of the antifungal and antiaflatoxigenic nature, free radical scavenging potential and efficacy in food system, cumin seed EO may be able to provide protection of food commodities against quantitative and qualitative losses, thereby enhancing their shelf life. The present investigation comprises the first report on antifungal mode of action of cumin seed EO and its efficacy as fumigant in food systems. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Fungal and aflatoxin contamination and lipid peroxidation cause quantitative and qualitative losses of food commodities and adversely affect their shelf life. Fungi represent one of the major threats for biodeterioration of cereals and pulses during storage causing economic losses to growers by increasing the free fatty acid content of seeds and decreasing germination ability (Dhingra et al., 2001). In addition, the mycotoxins secreted by different food borne molds cause qualitative losses of commodities, potentially inducing various health problems in consumers. Some species of Aspergillus are highly aflatoxigenic, particularly in tropical and subtropical countries, secreting high level of aflatoxins. Furthermore, aflatoxin is classified as group 1 human carcinogen by the International Agency for Research on Cancer (Mishra et al., 2013). Approximately 25% of cereals consumed all over the world are contaminated by mycotoxins (Devegowda et al., 1998). Moreover, lipid peroxidation of food items during storage due to the chain reactions of free radical oxidation is another major problem for qualitative losses of foods (Prakash et al., 2012). The use of synthetic preservatives is currently discouraged because of potential undesirable biological effects in animals and humans and reports of carcinogenic risks (Osman and Abdulrahman, 2003). Therefore, in recent years the development of safe plant based preservatives ⁎ Corresponding author. Tel.: +91 9415295765. E-mail addresses: [email protected], [email protected] (N.K. Dubey). 0168-1605/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.ijfoodmicro.2013.10.008
has been targeted as a viable strategy to enhance shelf life of food items against microbial deterioration, mycotoxin problems and lipid peroxidation. Some plant essential oils have proved their efficacy as antifungal agents against a wide range of fungi (Prakash et al., 2012; Soliman and Badeaa, 2002), as antiaflatoxigenic (Farag et al., 2006; Jaya et al., 2011) and antioxidant agents (Tomaino et al., 2005), and many of them are generally recognized as safe (GRAS). Currently some essential oil-based preservatives such as ‘DMC Base Natural’ comprising 50% EO from rosemary, sage and citrus and 50% glycerol are already commercially available as a safe food additive (Burt, 2004). Carvone has been introduced under the trade name TALENT in The Netherlands (Tripathi and Dubey, 2004). Formulations of some EOs viz., thyme oil (Thymus capitatus), jojoba oil (Simmondsia californica) and rosemary oil (Rosemarinus officianalis) are also frequently used as plant-based safe preservatives (Dayan et al., 2009). Cuminum cyminum L. (cumin), the second most popular spice in the world after black pepper, is cultivated mainly in India, China, Arabia and in the countries adjoining the Mediterranean Sea (Hajlaoui et al., 2010). The medicinal applications of cumin include use as a stimulant, carminative, an astringent, against indigestion, flatulence and diarrhea (Norman, 1990). There are reports on antimicrobial efficacy of cumin EO against a number of food pathogenic microorganisms (Hajlaoui et al., 2010; Khosravi et al., 2011a,b; Mohammadpour et al., 2012; Singh et al., 2002), however, the practical efficacy of EO in food system as preservative is lacking. Hence, the aim of the present study was to investigate the antifungal, antiaflatoxin, antioxidant and phytotoxic
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effect of Cuminum cyminum seed essential oil. We also assessed its in vivo efficacy as a fumigant in a model food system (wheat and chickpea stored in plastic containers) to explore the possibility of its recommendation as a plant based preservative to enhance the shelf life of food items. In the present investigation wheat and chickpea were selected as a food system to assess the practical efficacy of cumin seed EO as a preservative as these commodities are often naturally contaminated by storage fungi and aflatoxins, particularly in tropical and subtropical countries (Ahmad and Singh, 1991; Giray et al., 2007). The calculated probable daily intake of aflatoxin B1 through consumption of contaminated wheat for the population in some regions of India has been reported to be much higher than that of the suggested provisional maximum tolerable daily intake (Toteja et al., 2006).
2. Materials and methods 2.1. Chemicals and equipment Acetone, chloroform, methanol, sodium sulfate, tween-80, toluene, isoamyl alcohol, potassium hydroxide, absolute ethanol, n-heptane, Potato Dextrose Agar (PDA) medium (Potato, 200 g; Dextrose, 20 g; Agar, 15 g and distilled water 1000 ml, pH 5.6 ± 0.2), SMKY medium; (Sucrose, 200 g; MgSO4.7H2O, 0.5 g; KNO3, 0.3 g and yeast extract, 7 g; 1000 ml distilled water), Potato Dextrose Broth (PDB) medium and 2,2-diphenyl-1-picrylhydrazil (DPPH) were procured from HiMedia Laboratories Pty Ltd., Mumbai, India. Hydro-distillation apparatus (Merck Specialities Pty Ltd., Mumbai, India), centrifuge, UV transilluminator (Zenith Engineers, Agra, India) and spectrophotometer (Systronics India Ltd., Mumbai, India) were the major equipment used in the present investigation.
2.2. Moisture content and pH To calculate moisture content of three varieties of wheat (Triticum aestivum) viz. Malviya HUW 234, K68 and K65, and three varieties of chickpea (Cicer arietinum) viz. Samrat, Pusa 256 and Kabuli; 50 g of each seed sample was dried at 100 °C in hot air oven for 24 h and difference with the fresh weight was calculated (Mandeel, 2005). One gram of each sample was finely ground using a sterilized mixer grinder. A 1:10 (sample:distilled water) (w/v) suspension of each sample was prepared and stirred for 24 h. The pH of the suspension was recorded using an electronic pH meter.
2.3. Mycobiota analysis Analysis of the mycobiota of three varieties of wheat viz. Malviya HUW 234, K65 and K68 and chickpea viz. Samrat, Pusa 256 and Kabuli was performed by the direct plating method as described by Shukla et al. (2009) and also by the serial dilution technique (Prakash et al., 2010) using PDA medium. For direct plating, seeds were surface sterilized with 1% solution of sodium hypochlorite and rinsed in three changes of sterile distilled water. Five seeds were then placed in each Petri plate equidistantly containing 10 ml medium. For serial dilution test, 10 g of each powdered sample was homogenized in 90 ml sterile distilled water in an Erlenmeyer flask (250 ml). Five fold serial dilutions were prepared and 1 ml of aliquot (10− 4) of each sample was inoculated in each Petri dish containing 10 ml medium. Ten replicates of each treatment and sample were prepared and incubated at 27 ± 2 °C for seven days. Different fungal colonies were counted and species were identified following Domsch et al. (1980), Pitt (1979) and Raper and Fennell (1977). The developing fungal colonies were isolated and maintained on PDA at 4 °C.
2.4. Selection of the most toxigenic strain of Aspergillus flavus Fourteen strains of A. flavus from each variety of wheat and chickpea samples were randomly selected to test their aflatoxin B1 (AFB1) producing potential by thin layer chromatography (TLC) following Mishra et al. (2012). The isolates were aseptically inoculated in 25 ml SMKY medium in 100 ml flask and incubated for 10 days (27 ± 2 °C). The content of each flask was filtered through Whatman no. 1 filter paper and extracted with 20 ml chloroform using separating funnel. The extract was evaporated to dryness in a water bath and redissolved in 1 ml chloroform. An aliquot (50 μl) of chloroform extract was spotted on TLC plates and developed in solvent toluene:isoamyl alcohol: methanol (90:32:2; v/v/v). The plates were air dried and AFB1 was observed in UV transilluminator (360 nm). The blue spots were scraped from the TLC plate, dissolved in methanol (5 ml) and centrifuged at 3000 rpm (5 min). Absorbance of the supernatant was recorded at 360 nm and AFB1 was estimated as: AFB1 content (μg/l) = D × M/E × L × 1000 where, D = absorbance, M = molecular weight (312), E = molar extinction coefficient AFB1 (21,800), and L = path length (1 cm). A. flavus strain LHP(C)-D6 was found to be the most toxigenic strain and was selected as test fungus for further investigations. 2.5. Isolation of essential oil Essential oil (EO) was isolated from seeds of C. cyminum collected from the Botanical Garden of Banaras Hindu University, Varanasi, India. The identification and authentication of the plant were carried out with the help of authentic flora (Dubey, 2004) and voucher specimen was submitted to the herbarium of the laboratory of herbal pesticides, Department of Botany, Banaras Hindu University, Varanasi. Five hundred grams of seeds was thoroughly washed twice with distilled water and subjected to hydro distillation in Clevenger's apparatus for 3 h. The volatile fraction (EO) was separated, traces of water removed by passing through anhydrous sodium sulfate, then stored in a clean dark glass vial and kept at 4 °C until use. 2.6. Chemical characterization of cumin seed essential oil Cumin seed EO was subjected to gas chromatography (PerkinElmer Auto XL GC, MA, USA) equipped with a flame ionization detector and the GC condition were: EQUITY-5 column (60 m × 0.32 mm × 0.25 μm); the carrier gas was H2; column head pressure 10 psi; oven temperature program isotherm 2 min at 70 °C, 3 °C/min gradient to 250 °C, isotherm10 min; injection temperature, 250 °C; detector temperature 280 °C. GC–MS analysis was performed using PerkinElmer Turbomass GC–MS. The GC column was EQUITY-5 (60 m × 0.32 mm × 0.25 μm) fused silica capillary column. The GC conditions were: injection temperature, 250 °C; column temperature, isothermal at 70 °C for 2 min, then programmed to 250 °C at 37 °C/min and held at this temperature for 10 min; ion source temperature, 250 °C. Helium was the carrier gas. The effluent of the GC column was introduced directly into the source of MS and spectra obtained in the EI mode with 70 eV ionization energy. The sector mass analyzer was set to scan from 40 to 500 amu for 2 s. The identification of individual compounds was based on their retention times relative to those of authentic samples and matching spectral peaks available with Wiley, NIST and NBS mass spectral libraries or with the published data (Adams, 2007). 2.7. Evaluation of toxicity of cumin seed EO against A. flavus strain LHP(C)-D6 The toxicity of cumin seed EO against A. flavus strain LHP(C)-D6 was measured in terms of minimal inhibitory concentration (MIC) and minimal fungicidal concentration (MFC) by the poisoned food assay.
A. Kedia et al. / International Journal of Food Microbiology 168–169 (2014) 1–7
Requisite amounts of the EO were dissolved separately in Petri plates containing 0.5 ml acetone and then 9.5 ml PDA added to obtain different concentrations (0.1 to 1 μl/ml). A fungal disk (5 mm diameter) of seven days old culture of A. flavus strain LHP(C)-D6 was inoculated onto prepared Petri dishes. The control sets contained PDA with acetone but no EO. The inoculated Petri dishes were incubated at 27 ± 2 °C for 7 days. Fungal colony diameters of treatment and control sets were measured. The lowest concentration of the EO resulting in no growth of the A. flavus was taken as the MIC. For determination of MFC, the inhibited fungal disks of oil treated sets were re-inoculated on fresh medium after washing with distilled water and revival of their growth (fungistatic/fungicidal) was observed after 7 days. The lowest concentration preventing revival of fungal growth was taken as the MFC (Kumar et al., 2008). 2.8. Evaluation of cumin seed EO as aflatoxin B1 suppressor For aflatoxin B1 suppressor activity of cumin seed EO, the requisite amount of EO was dissolved separately in 0.5 ml acetone and added to 24.5 ml SMKY to achieve concentrations from 0.1 to 1.0 μl/ml. The medium was inoculated with a 5 mm diameter disk of seven days old culture of A. flavus strain LHP(C)-D6. The control sets contained acetone but no EO. AFB1 was detected by thin layer chromatography as described in Section 2.4. The fungal mats were dried at 80 °C (12 h) to determine the net mycelial dry weight. 2.9. Effect of cumin seed EO on ergosterol content in plasma membrane of A. flavus strain LHP(C)-D6 The ergosterol content in the plasma membrane of A. flavus was assessed following Tian et al. (2012) with slight modifications. A 100 μl aliquot of spore suspension of A. flavus strain LHP(C)-D6 (106 spores/ml) in 0.1% Tween-80 was inoculated in PDB medium containing 0, 0.15, 0.3, 0.45 and 0.6 μl/ml cumin seed EO. After 4 days of incubation at 27 ± 2 °C, mycelia was harvested and washed twice with distilled water. The net wet weight of the cell pellet was measured. Five ml of 25% alcoholic potassium hydroxide solution (25 g KOH and 35 ml sterile distilled water, brought to 100 ml with 100% ethanol) was added to each cell pellet and vortex mixed for 2 min, followed by incubation in an 85 °C water bath for 4 h. Sterols were then extracted from each sample by adding a mixture of 2 ml sterile distilled water and 5 ml n-heptane followed by sufficient vortex mixing for 2 min. After allowing the layers to separate for 1 h at room temperature, the n-heptane layer was separated and analyzed by scanned spectrophotometry between 230 and 300 nm. The presence of ergosterol and the late sterol intermediate 24(28) dehydroergosterol in the n-heptane layer resulted in a characteristic four peaked curve. The absence of ergosterol in samples was indicated by a flat line. The ergosterol amount was calculated as a percentage of the wet weight of the cells as follows:% ergosterol + % 24(28) dehydroergosterol = (A282/290)/pellet weight; % 24(28) dehydroergosterol = (A230/518)/pellet weight; and % ergosterol = (% ergosterol + % 24(28) dehydroergosterol) — % 24(28) dehydroergosterol, where 290 and 518 are the E values (in percentages per cm) determined for crystalline ergosterol and 24(28) dehydroergosterol, respectively, and pellet weight is the net wet weight (g). 2.10. Spectrum of toxicity of cumin seed EO against food borne molds isolated from wheat and chickpea mycoflora analysis
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only acetone (0.5 ml) and 9.5 ml PDA inoculated with the test fungi served as control. The plates of both treatment and control sets were incubated at 27 ± 2 °C for seven days. The percent inhibition of fungal growth was determined. The plates showing no visible fungal growth were sub-cultured on EO-free PDA plates to determine if the inhibition was reversible (fungistatic/fungicidal). 2.11. Antioxidant activity of cumin seed EO through DPPH free radical analysis Free radical scavenging activity of the cumin seed EO was measured by recording the extent of bleaching of a DPPH solution from purple to yellow following Prakash et al. (2012). Different concentrations (0.02 to 0.2 μl/ml) of the EO were added to 0.004% DPPH solution in methanol (5 ml). After 30 min incubation at room temperature (25 ± 2 °C), the absorbance was measured against a blank at 517 nm using a spectrophotometer. The antioxidant activity was measured through scavenging of DPPH free radical with reduction in absorbance of the sample. The IC50 of the test compounds, which represented the concentration that caused 50% neutralization of DPPH radicals, was measured from the graph plotting percentage inhibition against concentration. 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. 2.12. In vivo investigation with cumin seed EO against fungal deterioration of wheat and chickpea during storage Experiments were designed to fumigate the wheat and chickpea samples with cumin seed EO in airtight containers following Varma and Dubey (2001). In one set (uninoculated treatment), 1.2 ml of the EO, soaked in a 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 0.6 μl/ml air. In another set (inoculated treatment), the wheat and chickpea samples, prior to treatment with the oil, were inoculated with 2 ml spore suspension of A. flavus strain LHP(C)-D6 prepared in 0.1% Tween 80. The control set also contained two sets — the uninoculated control and the inoculated control. After 12 months of storage at laboratory conditions (temperature 10–46 °C and RH 30–90%), food samples of both treatment and control sets were analyzed for fungi using the direct plating and serial dilution methods as mentioned in Section 2.3. The percent protection in the uninoculated and inoculated treatments was observed by following formula. Percent protection = Dc − Dt / Dc × 100 where, Dc = percent occurrence of total fungi of control set and Dt = percent occurrence of total fungi in treatment set. 2.13. Effect of cumin seed EO on germination of fumigated seeds The germination of wheat and chickpea seeds, treated with cumin seed EO was tested after 12 months of storage by placing 50 randomly selected seeds in Petri plates containing moistened blotting paper. The seeds germinated within a week were recorded as viable. The per cent germination was calculated with respect to control sets. 2.14. Statistical analysis
The toxicity of cumin seed EO for 19 food borne molds, isolated from wheat and chickpea during mycoflora analysis, was evaluated by the poisoned food technique. The oil at its MIC (viz. 0.6 μl/ml) was added separately to Petri plates containing 0.5 ml acetone and 9.5 ml molten PDA. A 5 mm disk from a seven day old colony from each fungus was separately placed on the center of the prepared plates. Plates containing
All experiments except mycoflora analysis 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 b 0.05). The analysis of data was performed with the SPSS program version 16.0.
A. Kedia et al. / International Journal of Food Microbiology 168–169 (2014) 1–7
28 54 – – – – – 1 – – – – – – – – – – – Total 72 70 – – – – – – – – – – – – – – – – – 36 26 – – – – – 1 2 7 9 – – – – – – – –
Samrat
DP
MalwiyaHUW234
SD
38 5 16 38 – – 8 12 – – – – – 6 – – – – –
DP
34 33 4 – – – – 1 – 2 – 2 – – – – – – –
50 44 – – – – – – 13 7 – – – – – 2 – – –
DP SD DP
SD
Total Kabuli Pusa 256 Chickpea mycoflora analysis
K68 Wheat mycoflora analysis
Serial no.
Compound
Retention time (min.)
%
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
Alpha-pinene Camphene Benzaldehyde Laevo beta pinene Cymene L-Limonene Gamma-terpinene Para-alpha-dimethyl styrene Cherry propanol Alpha-terpineol Myrtenol P-Allylanisole Cuminaldehyde Carvotanacetone (E)-Cinnamaldehyde Eugenol
9.126 9.650 10.00 10.60 12.38 12.50 13.70 15.001 19.351 19.651 19.951 20.001 22.08 22.326 23.40 27.376
0.99 0.14 0.02 11.50 47.08 1.05 19.36 0.65 0.02 0.22 0.04 0.12 14.92 0.17 0.84 0.33 Total 97.45
70 50 – – – – – – – 2 – – – – – – – – – DP = Direct plating method; SD = Serial dilution method
DP
23 16 – – 8 – – 5 3 6 8 – 1 – 3 1 – – – 25 – – – 13 7 15 1 – – – 9 9 3 – – – – –
SD DP
22 19 15 5 9 3 – 9 4 – 1 4 – – – – – 2 1 42 – 39 28 22 13 10 1 – – – – – – – 3 – –
SD
The moisture content of wheat samples varied from 12 to 14% whereas that of chickpea samples varied from 11 to 16%. The pH values were slightly acidic, for wheat samples the pH ranged from 6.2 to 6.67 whereas for chickpea samples it was in the range of 6.0 to 6.14. During mycological survey, a total of 1230 isolates belonging to 11 different genera and 19 species were isolated. Aspergillus. was the dominant genus. A. flavus and A. niger were found in all the investigated samples and A. flavus was found to be the dominant species (Table 1). In assessing the toxigenicity of A. flavus isolates from the commodities, 20 isolates from wheat and 19 isolates from chickpea were found to be aflatoxigenic with blue spots on TLC plates at 360 nm. The toxigenic isolate A. flavus LHP(C)-D6, from Chickpea var. Samrat was selected as test fungus for detailed study as it produced maximum aflatoxin B1 (2199.84 μg/l). The EO obtained from cumin seed was pale yellow in color with a spicy odor. The yield ranged between 0.9 and 1.2% on fresh weight basis through hydrodistillation. Chemical characterization by the GC–MS analysis of the cumin seed oil revealed the presence of 16 compounds comprising 97.45% of total EO. Their retention time and area percentage are summarized in Table 2. The major components of EO were cymene (47.08%), gamma-terpinene (19.36%), cuminaldehyde (14.92%) and laevo beta pinene (11.50%). The MIC of cumin seed EO for complete inhibition of growth of A. flavus strain LHP(C)-D6 was found at 0.6 μl/ml. The fungicidal concentration (MFC value) of the EO was recorded at 0.9 μl/ml. The EO inhibited AFB1 production as well as
40 7 1 – 6 16 – – 8 – 1 2 – – – – – – – Aspergillus flavus Aspergillus niger Curvularia lunata Aspergillus glaucus Alternaria alternata Penicillium citrinum Aspergillus unguis Aspergillus terreus Rhizopus stolonifer Aspergillus nidulans Mucor sp. Mycelia sterilia Penicillium italicum Cladosporium cladosporioides Penicillium purpurogenum Penicillium luteum Fusarium oxysporum Absidia ramosa Spondylocladium australe 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19.
SD
3. Results
K65 No. of fungal isolates Fungus isolated Sr no.
Table 1 Fungi isolated from wheat and chickpea samples.
Table 2 GC–MS analysis of Cuminum cyminum seed essential oil.
480 314 75 71 58 47 33 31 30 24 19 17 10 9 3 3 3 2 1 1230
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Table 3 Effect of different concentrations of Cuminum cyminum seed EO on growth, dry weight, aflatoxin B1 production and nature of toxicity to the A. flavus strain LHP(C)-D6. Conc. (μl/ml)
Diameter (cm)
Mycelial dry wt. (g)
AFB1 content (μg/l)
Nature of toxicity
CNT 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0
5.95 ± 0.14e 4.84 ± 0.07d 4.57 ± 0.14d 3.11 ± 0.08c 3.00 ± 0.17c 1.69 ± 0.02b 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a
0.46 ± 0.01f 0.42 ± 0.01e 0.41 ± 0.00de 0.38 ± 0.01d 0.22 ± 0.00c 0.04 ± 0.00b 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a
1517.06 ± 145.25 1245.14 ± 24.79 1173.58 ± 37.87 1030.45 ± 65.59 495. 93 ± 48.13 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00
Static Static Static Static Static Static Static Static Static Cidal Cidal
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.
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Fig. 2. Antioxidant activity of cumin seed EO through DPPH free radical analysis. Fig. 1. Effect of different concentrations of cumin seed EO on mycelial wet wt. and ergosterol content of A. flavus strain LHP(C)-D6.
mycelial dry weight in a dose dependent manner and at 0.5 μl/ml, it completely inhibited the production of AFB1 (Table 3). The effects of cumin seed EO on ergosterol content in the plasma membrane of A. flavus strain LHP(C)-D6 with respect of mycelial wet wt. are shown in Fig. 1. A dose dependent decrease in ergosterol content was observed on increasing concentration of the EO. A reduction percentage of the ergosterol content as compared with the control was found to be 28.57, 35.71, 64.29 and 100% respectively for 0.15, 0.3, 0.45 and 0.6 μl/ml concentrations. Hence, the cumin seed oil completely reduced the ergosterol content at 0.6 μl/ml concentrations which is equal to its MIC value. The oil exhibited a broad spectrum of fungal toxicity inhibiting all 19 food borne fungal species except Rhizopus stolonifer at its MIC (Table 4). The EO was found to be fungicidal against most of the fungal species except Absidia ramosa, Aspergillus glaucus, A. niger, Aspergillus terreus, Aspergillus unguis, Fusarium oxysporum and Mucor sp. The bleaching of the purple color of the DPPH to yellow confirmed the positive antioxidant activity of EO. The oil showed strong free radical scavenging activity as its IC50 value was found to be 0.092 μl/ml (Fig. 2).
Table 4 Fungitoxic spectrum and nature of toxicity of Cuminum cyminum seed EO at 0.6 μl/ml concentration against some food borne fungi. Serial no.
Fungal species
% mycelial inhibition
Nature of toxicity
1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19.
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 Spondylocladium australe
100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 88.51 ± 0.98 100.00 ± 0.00
Static Cidal Cidal Static Static Static Static Cidal Cidal Static Static Cidal Cidal Cidal Cidal Cidal Cidal – Cidal
Values are mean (n = 3) ± standard error.
The results of in vivo trials in food system are based on percent protection of wheat and chickpea from fungal association in the uninoculated and inoculated treatments. In the uninoculated samples, 61.29 and 27.27% protection of wheat and chickpea respectively against storage molds was observed for serial dilution sets, whereas for direct plating method the percent protection of wheat and chickpea was 65.85 and 75.00% respectively. Similarly in inoculated samples, 65.15 and 50.00% protection was recorded for serial dilution sets and 20.00 and 50.00% protection was recorded for direct plating method for wheat and chickpea samples respectively. The seed germination tests revealed that wheat and chickpea seeds fumigated with cumin seed EO had no loss in viability even after 12 months of treatment with 100% seeds germinating in Petri plates. Hence, the cumin seed EO was not phytotoxic. 4. Discussion The extent of mold contamination depends on various conditions such as source, harvesting period, storage practices and chemical nature of the substrate (Prakash et al., 2010). Moisture content and, to a lesser extent, pH are two other important abiotic factors that govern fungal growth and proliferation on stored food commodities (Ozkan et al., 2003). In the present study the moisture content and pH of selected commodities were found within conducive range for fungal proliferation (FAO, 1980). The results indicate that all the selected samples were heavily contaminated with the different mold species. A total of 19 fungal species were recorded from the samples and Aspergillus was found to be dominant in both serial dilution and direct plating methods. The finding supports the earlier observations where Aspergillus spp. were dominant in some stored edible pulses (Shukla et al., 2009) and wheat (Varma and Dubey, 2001). Higher incidence of Aspergilli than other molds may be due to their saprophytic nature and ability to colonize diverse substrates because of secretion of various hydrolytic enzymes (Leite de Souza et al., 2005). The relative frequency of A. flavus and A. niger was higher than that of the remaining fungal species supporting the earlier observations made by Prakash et al. (2011) and Shukla et al. (2009). The quantitative estimation of aflatoxin B1 production in the present study reveals that the samples were highly contaminated by toxigenic strains of A. flavus. Hence, the biodeterioration of the samples was qualitative as well as quantitative. In the present investigation cumin seed oil was selected for detailed investigation to assess its efficacy as plant based preservative. Prior to antimicrobial assay, an essential oil should be chemically characterized to standardize its chemical nature. The chemical profile of essential oil of a particular plant species shows
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different chemotypic variations because of ecological and geographical conditions, age of plant and time of harvesting (Prakash et al., 2010). Such chemotypic variations would definitely affect the biological activity of the essential oil. Hence, it is advisable that the percentage of the major components of the EOs should be mentioned if applied as food additive (Bagamboula et al., 2004). In the present investigation, the GC and GC–MS analysis of cumin oil revealed the presence of 16 compounds comprising 97.45% of total oil. Cymene was identified as the major compound contributing 47.08% followed by gammaterpinene (19.36%), cuminaldehyde (14.92%) and laevo beta pinene (11.50%). In earlier reports, cuminaldehyde (36–39%) was reported as the major component of cumin seed EO from China (Li and Jiang, 2004), Bulgaria (Jirovetz et al., 2005) and Tunisia (Hajlaoui et al., 2010). However, alpha-Pinene (29.2%) was reported as the major component of cumin seed EO from Iran (Mohammadpour et al., 2012). This variation in EO composition could be due to ecological and geographical conditions. In this study, the antifungal activity of cumin seed oil has been assessed. The study of MIC is important to determine the minimum dose to control fungal populations resulting in least wastage of the pesticide (Kumar et al., 2008). The cumin seed EO showed high antifungal activity against A. flavus LHP(C)-D6 as its MIC was lower than that of some earlier reported EOs viz. Origanum majorana, Coriandrum sativum, Hedychium spicatum, Commiphora myrrha and Cananga odorata (Prakash et al., 2012), Cinnamomum jensenianum (Tian et al., 2012) and some prevalent fungicides such as Nystatin and Wettasul-80 (Prakash et al., 2010) emphasizing its superiority as an antimicrobial agent at a low prescribed dose. The oil was fungicidal in nature above MIC value as its MFC against test fungus was slightly greater than its MIC. Therefore, cumin seed EO could be capable of permanent disinfection of food borne fungi from food items. In addition, the EO also exhibited marked efficacy in checking aflatoxin B1 production by A. flavus LHP(C)-D6 at a concentration lower than MIC. Hence, the EO would be acting through two different modes of action as inhibitor of both fungal growth and aflatoxin production as has been earlier reported by Prakash et al. (2012), Rasooli and Abyaneh (2004) and Tian et al. (2012). The antifungal mechanism of EO involves membrane disruption by their lipophilic compounds. The lowmolecular weight and highly lipophilic components of EO pass easily through cell membranes and cause disruption to the fungal cell organization (Chao et al., 2005). Likewise, the aflatoxin B1 production inhibition efficacy of EO may be because of inhibition of carbohydrate catabolism in A. flavus by acting on some key enzymes, reducing its ability to produce aflatoxins (Tian et al. 2011). The MIC of the cumin EO for inhibition of growth of test fungus and aflatoxin production was different from that of earlier reports (Khosravi et al., 2011a,b; Mohammadpour et al., 2012). This difference in MIC may be due to the difference in chemical profile of the oil, strain of the test fungus, technique adopted for antifungal assay and the medium used. In the present investigation the mode of action of the cumin EO has been observed which would be helpful in its formulation as preservative. Ergosterol is specific to fungi and is the major sterol component of the fungal cell membrane. It is also responsible for maintaining the cell function and integrity (Rodriguez et al., 1985). There is a previous study by Kelly et al. (1995) stating that the primary action mechanism, by which azole antifungal drugs inhibit fungal cell growth, is the disruption of normal sterol biosynthetic pathways resulting in a decrease of ergosterol biosynthesis. Some studies have shown that essential oils can also cause a considerable reduction in the quantity of ergosterol (Pinto et al., 2006, 2009). The cumin seed oil completely reduced the ergosterol content at 0.6 μl/ml concentrations which is equal to its MIC value. The reduction in mycelial weight as well as ergosterol content on increasing concentration of oil clearly suggests its action on plasma membrane of fungus supporting earlier findings of Kelly et al. (1995) and Tian et al. (2012). The present findings revealed that cumin seed EO can induce a considerable impairment of
the ergosterol biosynthesis by A. flavus and subsequent decrease in biomass on increasing concentration of EO. Hence, the plasma membrane is an important antifungal target of cumin seed EO. This emphasizes that the antimicrobial components of the essential oils cross the cell membrane, interact with the enzymes and proteins of the membrane, thus producing a flux of protons towards the cell exterior which induces disruption to the fungal cell organization and, ultimately, their death as has been supported by the transmission electron microscopy by Nogueira et al. (2010); and Tian et al. (2012). In addition, cumin seed EO also exhibited strong antioxidant activity and can enhance the shelf life of food products by controlling free radical scavenging and oxidation of unsaturated lipids. The IC50 value of essential oil was found to be better than that of the earlier reported essential oils viz. Cananga odorata and Coriandrum sativum (Prakash et al., 2012). Free radical scavenging activity of EO may be due to the presence of the phenolic compounds or synergistic effect of overall compounds (Sharififar et al., 2007). The practical applicability of cumin seed EO has been observed as fumigant in food system in plastic storage containers. The oil showed remarkable in vivo efficacy in protecting the fumigated wheat and chickpea samples, from fungal contamination during storage up to 12 months. Hence, further large scale trials are required for its recommendation as a plant based preservative. The preliminary phytotoxic test of cumin seed EO was performed in the form of seed germination test. Results on seed germination test showed 100% germination of wheat and chickpea seeds in EO treated sets. The oil treated food commodities may thus be used for sowing purpose by the growers. Moreover, since EO is volatile in nature, the vapors may be easily eliminated from the fumigated food items after sun drying. In this connection, currently microencapsulation technology is prescribed to use the essential oils as antimicrobials for preservation of stored food commodities and in food industry for flavor stabilization (Burt, 2004). Essential oils of many edible and medicinal plants are used in different pharmaceutical preparations which minimize questions regarding their safe use. Essential oils from aromatic and medicinal plants are potentially useful as antimicrobial agents and their use as medicines have long been recognized (Kim et al., 2005). The attraction of modern society towards herbal products desiring fewer synthetic ingredients in foods and recommendation of herbal products as ‘generally recognized as safe’ (GRAS) as food additives may lead scientific interest in the exploitation of essential oils as plant based food additives (Smid and Gorris, 1999). Moreover, the regulatory status of cumin and cumin oil in the USA is regarded as safe, GRAS 2340 and GRAS 2343 (Parthasarathy et al., 2008). The cumin seeds are traditionally used as safe culinary items in form of spice. Bulk of raw materials will be available for formulation of oil as preservative as the cumin plants grow luxuriantly in different tropical and subtropical countries. Because of renewable source, high yield of the oil, and efficacy at low concentrations, the oil may be economically recommended for formulation as plant based preservatives. However, standardization of dose during practical application is desirable before its recommendation to agri-firms. This is the first report on mode of antifungal action of cumin seed EO and its efficacy as fumigant in food system. In conclusion, based on efficacy as inhibitor to fungal growth and aflatoxin secretion, its free radical scavenging potential, efficacy in food system and favorable safety profile, the cumin seed oil has possibility to provide complete protection of food commodities against quantitative and qualitative losses, thereby, enhancing their shelf life.
Acknowledgments This work was financially supported by University Grant Commission (UGC), New Delhi, India.
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References Adams, R.P., 2007. Identification of Essential Oil Components by Gas Chromatography/ Mass Spectrometry. Allured Publishing Corporation, Carol Stream, IL. Ahmad, S.K., Singh, P.L., 1991. Mycofloral changes and aflatoxin contamination in stored chickpea seeds. Food Addit. Contam. 8, 723–730. Bagamboula, C.F., Uyttendaele, M., Debevere, J., 2004. Inhibitory effect of thyme and basil essential oils, carvacrol, thymol, estragol, linalool and p-cymene towards Shigella sonnei and S. flexneri. Food Microbiol. 21, 33–42. Burt, S., 2004. Essential oils: their antibacterial properties and potential applications in foods — a review. Int. J. Food Microbiol. 94, 223–253. Chao, L.K., Hua, K.F., Hsu, H.Y., Cheng, S.S., Lin, J.Y., Chang, S.T., 2005. Study on the antiinflammatory activity of essential oil from leaves of Cinnamomum osmophloeum. J. Agric. Food Chem. 53, 7274–7278. Dayan, F.E., Cantrell, C.L., Duke, S.O., 2009. Natural products in crop protection. Bioorg. Med. Chem. 17, 4022–4034. Devegowda, G., Raju, M.V.L.N., Swang, H.V.L.N., 1998. Mycotoxins: novel solutions for their counteraction. Feedstuffs 70, 12–15. Dhingra, O.D., Mizubuti, E.S.G., Napoleao, I.T., Jham, G., 2001. Free fatty acid accumulation and quality loss of stored soybean seeds invaded by Aspergillus ruber. Seed Sci. Technol. 29, 193–203. Domsch, K.H., Gams, W., Anderson, T.H., 1980. Compendium of Soil Fungi, vol. 1 and 2. Academic Press, London. Dubey, N.K., 2004. Flora of BHU Campus. Banaras Hindu University Press, Varanasi, India. FAO (Food and Agriculture Organisation), 1980. On-farm maize drying and storage in the humid tropics. FAO Agric. Serv. Bull. 40, 1–54. Farag, R.S., Daw, Z.Y., Abo-Raya, S.H., 2006. 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. Giray, B., Girgin, G., Engin, A.B.., Aydın, S., Sahin, G., 2007. Aflatoxin levels in wheat samples consumed in some regions of Turkey. Food Control 18, 23–29. Hajlaoui, H., Mighri, H., Noumi, E., Snoussi, M., Trabelsi, N., Ksouri, R., Bakhrouf, A., 2010. Chemical composition and biological activities of Tunisian Cuminum cyminum L. essential oil: a high effectiveness against Vibrio spp. strains. Food Chem. Toxicol. 48, 2186–2192. Jaya, Prakash, B., Dubey, N.K., 2011. Evaluation of chemically characterised essential oils of Coleus aromaticus, Hyptis suaveolens and Ageratum conyzoides against storage fungi and aflatoxin contamination of food commodities. Int. J. Food Sci. Technol. 46, 754–760. Jirovetz, L., Buchbauer, G., Stoyanova, A.S., Georgiev, E.V., Damianova, S.T., 2005. Composition, quality control and antimicrobial activity of the essential oil of cumin (Cuminum cyminum L.) seeds from Bulgaria that had been stored for up to 36 years. Int. J. Food Sci. Technol. 40, 305–310. Kelly, S.L., Lamb, D.C., Corran, A.J., Baldwin, B.C., Kelly, D.E., 1995. Mode of action and resistance to azole antifungals associated with the formation of 14α-methylergosta8, 24(28)-dien-3β, 6α-diol. Biochem. Biophys. Res. Commun. 207, 910–915. Khosravi, A.R., Minooeianhaghighi, M.H., Shokri, H., Emami, S.A., Alavi, S.M., Asili, J., 2011a. The potential inhibitory effect of Cuminum cyminum, Ziziphora clinopodioides and Nigella sativa essential oils on the growth of Aspergillus fumigatus and Aspergillus flavus. Braz. J. Microbiol. 42, 216–224. Khosravi, A.R., Shokri, H., Minooeianhaghighi, M., 2011b. Inhibition of aflatoxin production and growth of Aspergillus parasiticus by Cuminum cyminum, Ziziphora clinopodioides and Nigella sativa essential oils. Foodborne Pathog. Dis. 8 (12), 1275–1280. Kim, J.H., Campbell, B.C., Yu, J., Mahoney, N., Chan, K.L., Molyneux, R.J., Bhatnagar, D., Cleveland, T.E., 2005. Examination of fungal stress response genes using Saccharomyces cerevisiae as a model system: targeting genes affecting aflatoxin biosynthesis by Aspergillus flavus Link. Appl. Microbiol. Biotechnol. 67, 807–815. Kumar, A., Shukla, R., Singh, P., Prasad, C.S., Dubey, N.K., 2008. Assessment of Thymus vulgaris L. essential oil as a safe botanical preservative against post harvest fungal infestation of food commodities. Innov. Food Sci. Emerg. Technol. 9, 575–580. Leite de Souza, E., Antonio da Silva, C., Paiva de Sousa, C., 2005. Bacteriocins: molecules of fundamental impact on the microbial ecology and potential food biopreservatives. Braz. Arch. Biol. Technol. 48 (4), 559–566. Li, R., Jiang, Z., 2004. Chemical composition of the essential oil of Cuminum cyminum L. from China. Flavour Fragr. J. 19, 311–313. Mandeel, Q.A., 2005. Fungal contamination of some imported species. Mycopatholgia 159, 291–298. Mishra, P.K., Shukla, R., Singh, P., Prakash, B., Kedia, A., Dubey, N.K., 2012. Antifungal, antiaflatoxigenic, and antioxidant efficacy of Jamrosa essential oil for preservation of herbal raw materials. Int. Biodeterior. Biodegrad. 74, 11–16.
7
Mishra, P.K., Singh, P., Prakash, B., Kedia, A., Dubey, N.K., Chanotiya, C.S., 2013. Assessing essential oil components as plant-based preservatives against fungi that deteriorate herbal raw materials. Int. Biodeterior. Biodegrad. 80, 16–21. Mohammadpour, H., Moghimipour, E., Rasooli, I., Fakoor, M.H., Astaneh, S.A., Moosaie, S.S., Jalili, Z., 2012. Chemical composition and antifungal activity of Cuminum cyminum L. essential oil from Alborz mountain against Aspergillus species. Jundishapur. J. Nat. Pharm. Prod. 7 (2), 50–55. Nogueira, J.H., Gonçalez, E., Galleti, S.R., Facanali, R., Marques, M.O., Felício, J.D., 2010. Ageratum conyzoides essential oil as aflatoxin suppressor of Aspergillus flavus. Int. J. Food Microbiol. 137, 55–60. Norman, J., 1990. The Complete Book on Spices. Doerling Kindersley, London. Osman, K.A., Abdulrahman, H.T., 2003. Risk assessment of pesticide to human and the environment. Saudi J. Biol. Sci. 10, 81–106. Ozkan, M., Kirca, A., Cemeroglu, B., 2003. Effect of moisture content on CIE color values in dried apricots. Eur. Food Res. Technol. 216, 2170–2219. Parthasarathy, V.A., Chempakam, B., Zachariah, T.J., 2008. Chemistry of Spices. CABI, Oxfordsire, United Kingdom (U.K.). Pinto, E., Pina-Vaz, C., Salgueiro, L., Goncalves, M.J., Costa-de-Oliveira, S., Cavaleiro, C., et al., 2006. Antifungal activity of the essential oil of Thymus pulegioides on Candida, Aspergillus and dermatophyte species. J. Med. Microbiol. 55 (10), 1367–1373. Pinto, E., Vale-Silva, L., Cavaleiro, C., Salgueiro, L., 2009. Antifungal activity of the clove essential oil from Syzygium aromaticum on Candida, Aspergillus and dermatophyte species. J. Med. Microbiol. 58 (11), 1454–1462. Pitt, J.I., 1979. The Genus Penicillum. Academic Press, Sydney, Australia. Prakash, B., Shukla, R., Singh, P., Kumar, A., Mishra, P.K., Dubey, N.K., 2010. Efficacy of chemically characterized Piper betle L. essential oil against fungal and aflatoxin contamination of some edible commodities and its antioxidant activity. Int. J. Food Microbiol. 142, 114–119. 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., 2012. 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, 201–208. Raper, K.B., Fennell, D.L., 1977. The Genus Aspergillus. Krieger publishing Corporation, Huntington, New York. Rasooli, I., Abyaneh, M.R., 2004. Inhibitory effects of Thyme oils on growth and aflatoxin production by Aspergillus. parasiticus. Food Control 15, 479–483. 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. Sharififar, F., Moshafi, M.H., Mansouri, S.H., Khodashenas, M., Khoshnoodi, M., 2007. In vitro evaluation of antibacterial and antioxidant activities of the essential oil and methanol extract of endemic Zataria multiflora Boiss. Food Control 18, 800–805. Shukla, R., Kumar, A., Singh, P., Dubey, N.K., 2009. Efficacy of Lippia alba (Mill.) N.E. Brown essential oil and its monoterpene aldehyde constituents against fungi isolated from some edible legume seeds and aflatoxin B1 production. Int. J. Food Microbiol. 135, 165–170. Singh, G., Kapoor, I.P.S., Pandey, S.K., Singh, U.K., Singh, R.K., 2002. Antibacterial activity of volatile oils of some spices. Phytother. Res. 16 (7), 680–682. Smid, E.J., Gorris, L.G.M., 1999. Natural Antimicrobials for Food Preservation. In: Rahman, M.S. (Ed.), Handbook of Food Preservation. Marcel Dekker, New York, pp. 285–308. Soliman, K.M., Badeaa, R.I., 2002. Effect of oil extracted from some medicinal plants on different mycotoxigenic fungi. Food Chem. Toxicol. 40, 1669–1675. Tian, J., Ban, X., Zeng, H., He, J., Huang, B., Youwei, W., 2011. Chemical composition and antifungal activity of essential oil from Cicuta virosa L. var. latisecta Celak. Int. J. Food Microbiol. 145, 464–470. 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. Tomaino, A., Cimino, F., Zimbalatti, V., Venuti, V., Sulfaro, V., De Pasquale, A., Saija, A., 2005. Influence of heating on antioxidant activity and the chemical composition of some spice essential oils. Food Chem. 89, 549–554. Toteja, G.S., Mukherjee, A., Diwakar, S., Singh, P., Saxena, B.N., Sinha, K.K., et al., 2006. Aflatoxin B1 contamination in wheat grain samples collected from different geographical regions of India: a multicenter study. J. Food Prot. 69 (6), 1463–1467. 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. 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.
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International Journal of Food Microbiology journal homepage: www.elsevier.com/locate/ijfoodmicro
Antifungal and antiaflatoxigenic properties of Cuminum cyminum (L.) seed essential oil and its efficacy as a preservative in stored commodities Akash Kedia, Bhanu Prakash, Prashant K. Mishra, N.K. Dubey ⁎ Laboratory of Herbal Pesticides, Centre of Advanced Study in Botany, Banaras Hindu University, Varanasi 221005, UP, India
a r t i c l e
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Article history: Received 7 June 2013 Received in revised form 1 October 2013 Accepted 15 October 2013 Available online 23 October 2013 Keywords: Aspergillus flavus Cumin seed essential oil Antifungal Antioxidant Aflatoxin Shelf life
a b s t r a c t The study reports potential of Cuminum cyminum (cumin) seed essential oil (EO) as a plant based shelf life enhancer against fungal and aflatoxin contamination and lipid peroxidation. The EO showed efficacy as a preservative in food systems (stored wheat and chickpeas). A total of 1230 fungal isolates were obtained from food samples, with Aspergillus flavus LHP(C)-D6 identified as the highest aflatoxin producer. Cumin seed EO was chemically characterized through GC–MS where cymene (47.08%) was found as the major component. The minimum inhibitory concentration and minimum aflatoxin inhibitory concentration of EO were 0.6 and 0.5 μl/ml respectively. The EO showed toxicity against a broad spectrum of food borne fungi. The antifungal action of EO on ergosterol content in the plasma membrane of A. flavus was determined. The EO showed strong antioxidant potential having IC50 0.092μl/ml. As a fumigant in food systems, the EO provided sufficient protection of food samples against fungal association without affecting seed germination. In view of the antifungal and antiaflatoxigenic nature, free radical scavenging potential and efficacy in food system, cumin seed EO may be able to provide protection of food commodities against quantitative and qualitative losses, thereby enhancing their shelf life. The present investigation comprises the first report on antifungal mode of action of cumin seed EO and its efficacy as fumigant in food systems. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Fungal and aflatoxin contamination and lipid peroxidation cause quantitative and qualitative losses of food commodities and adversely affect their shelf life. Fungi represent one of the major threats for biodeterioration of cereals and pulses during storage causing economic losses to growers by increasing the free fatty acid content of seeds and decreasing germination ability (Dhingra et al., 2001). In addition, the mycotoxins secreted by different food borne molds cause qualitative losses of commodities, potentially inducing various health problems in consumers. Some species of Aspergillus are highly aflatoxigenic, particularly in tropical and subtropical countries, secreting high level of aflatoxins. Furthermore, aflatoxin is classified as group 1 human carcinogen by the International Agency for Research on Cancer (Mishra et al., 2013). Approximately 25% of cereals consumed all over the world are contaminated by mycotoxins (Devegowda et al., 1998). Moreover, lipid peroxidation of food items during storage due to the chain reactions of free radical oxidation is another major problem for qualitative losses of foods (Prakash et al., 2012). The use of synthetic preservatives is currently discouraged because of potential undesirable biological effects in animals and humans and reports of carcinogenic risks (Osman and Abdulrahman, 2003). Therefore, in recent years the development of safe plant based preservatives ⁎ Corresponding author. Tel.: +91 9415295765. E-mail addresses: [email protected], [email protected] (N.K. Dubey). 0168-1605/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.ijfoodmicro.2013.10.008
has been targeted as a viable strategy to enhance shelf life of food items against microbial deterioration, mycotoxin problems and lipid peroxidation. Some plant essential oils have proved their efficacy as antifungal agents against a wide range of fungi (Prakash et al., 2012; Soliman and Badeaa, 2002), as antiaflatoxigenic (Farag et al., 2006; Jaya et al., 2011) and antioxidant agents (Tomaino et al., 2005), and many of them are generally recognized as safe (GRAS). Currently some essential oil-based preservatives such as ‘DMC Base Natural’ comprising 50% EO from rosemary, sage and citrus and 50% glycerol are already commercially available as a safe food additive (Burt, 2004). Carvone has been introduced under the trade name TALENT in The Netherlands (Tripathi and Dubey, 2004). Formulations of some EOs viz., thyme oil (Thymus capitatus), jojoba oil (Simmondsia californica) and rosemary oil (Rosemarinus officianalis) are also frequently used as plant-based safe preservatives (Dayan et al., 2009). Cuminum cyminum L. (cumin), the second most popular spice in the world after black pepper, is cultivated mainly in India, China, Arabia and in the countries adjoining the Mediterranean Sea (Hajlaoui et al., 2010). The medicinal applications of cumin include use as a stimulant, carminative, an astringent, against indigestion, flatulence and diarrhea (Norman, 1990). There are reports on antimicrobial efficacy of cumin EO against a number of food pathogenic microorganisms (Hajlaoui et al., 2010; Khosravi et al., 2011a,b; Mohammadpour et al., 2012; Singh et al., 2002), however, the practical efficacy of EO in food system as preservative is lacking. Hence, the aim of the present study was to investigate the antifungal, antiaflatoxin, antioxidant and phytotoxic
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effect of Cuminum cyminum seed essential oil. We also assessed its in vivo efficacy as a fumigant in a model food system (wheat and chickpea stored in plastic containers) to explore the possibility of its recommendation as a plant based preservative to enhance the shelf life of food items. In the present investigation wheat and chickpea were selected as a food system to assess the practical efficacy of cumin seed EO as a preservative as these commodities are often naturally contaminated by storage fungi and aflatoxins, particularly in tropical and subtropical countries (Ahmad and Singh, 1991; Giray et al., 2007). The calculated probable daily intake of aflatoxin B1 through consumption of contaminated wheat for the population in some regions of India has been reported to be much higher than that of the suggested provisional maximum tolerable daily intake (Toteja et al., 2006).
2. Materials and methods 2.1. Chemicals and equipment Acetone, chloroform, methanol, sodium sulfate, tween-80, toluene, isoamyl alcohol, potassium hydroxide, absolute ethanol, n-heptane, Potato Dextrose Agar (PDA) medium (Potato, 200 g; Dextrose, 20 g; Agar, 15 g and distilled water 1000 ml, pH 5.6 ± 0.2), SMKY medium; (Sucrose, 200 g; MgSO4.7H2O, 0.5 g; KNO3, 0.3 g and yeast extract, 7 g; 1000 ml distilled water), Potato Dextrose Broth (PDB) medium and 2,2-diphenyl-1-picrylhydrazil (DPPH) were procured from HiMedia Laboratories Pty Ltd., Mumbai, India. Hydro-distillation apparatus (Merck Specialities Pty Ltd., Mumbai, India), centrifuge, UV transilluminator (Zenith Engineers, Agra, India) and spectrophotometer (Systronics India Ltd., Mumbai, India) were the major equipment used in the present investigation.
2.2. Moisture content and pH To calculate moisture content of three varieties of wheat (Triticum aestivum) viz. Malviya HUW 234, K68 and K65, and three varieties of chickpea (Cicer arietinum) viz. Samrat, Pusa 256 and Kabuli; 50 g of each seed sample was dried at 100 °C in hot air oven for 24 h and difference with the fresh weight was calculated (Mandeel, 2005). One gram of each sample was finely ground using a sterilized mixer grinder. A 1:10 (sample:distilled water) (w/v) suspension of each sample was prepared and stirred for 24 h. The pH of the suspension was recorded using an electronic pH meter.
2.3. Mycobiota analysis Analysis of the mycobiota of three varieties of wheat viz. Malviya HUW 234, K65 and K68 and chickpea viz. Samrat, Pusa 256 and Kabuli was performed by the direct plating method as described by Shukla et al. (2009) and also by the serial dilution technique (Prakash et al., 2010) using PDA medium. For direct plating, seeds were surface sterilized with 1% solution of sodium hypochlorite and rinsed in three changes of sterile distilled water. Five seeds were then placed in each Petri plate equidistantly containing 10 ml medium. For serial dilution test, 10 g of each powdered sample was homogenized in 90 ml sterile distilled water in an Erlenmeyer flask (250 ml). Five fold serial dilutions were prepared and 1 ml of aliquot (10− 4) of each sample was inoculated in each Petri dish containing 10 ml medium. Ten replicates of each treatment and sample were prepared and incubated at 27 ± 2 °C for seven days. Different fungal colonies were counted and species were identified following Domsch et al. (1980), Pitt (1979) and Raper and Fennell (1977). The developing fungal colonies were isolated and maintained on PDA at 4 °C.
2.4. Selection of the most toxigenic strain of Aspergillus flavus Fourteen strains of A. flavus from each variety of wheat and chickpea samples were randomly selected to test their aflatoxin B1 (AFB1) producing potential by thin layer chromatography (TLC) following Mishra et al. (2012). The isolates were aseptically inoculated in 25 ml SMKY medium in 100 ml flask and incubated for 10 days (27 ± 2 °C). The content of each flask was filtered through Whatman no. 1 filter paper and extracted with 20 ml chloroform using separating funnel. The extract was evaporated to dryness in a water bath and redissolved in 1 ml chloroform. An aliquot (50 μl) of chloroform extract was spotted on TLC plates and developed in solvent toluene:isoamyl alcohol: methanol (90:32:2; v/v/v). The plates were air dried and AFB1 was observed in UV transilluminator (360 nm). The blue spots were scraped from the TLC plate, dissolved in methanol (5 ml) and centrifuged at 3000 rpm (5 min). Absorbance of the supernatant was recorded at 360 nm and AFB1 was estimated as: AFB1 content (μg/l) = D × M/E × L × 1000 where, D = absorbance, M = molecular weight (312), E = molar extinction coefficient AFB1 (21,800), and L = path length (1 cm). A. flavus strain LHP(C)-D6 was found to be the most toxigenic strain and was selected as test fungus for further investigations. 2.5. Isolation of essential oil Essential oil (EO) was isolated from seeds of C. cyminum collected from the Botanical Garden of Banaras Hindu University, Varanasi, India. The identification and authentication of the plant were carried out with the help of authentic flora (Dubey, 2004) and voucher specimen was submitted to the herbarium of the laboratory of herbal pesticides, Department of Botany, Banaras Hindu University, Varanasi. Five hundred grams of seeds was thoroughly washed twice with distilled water and subjected to hydro distillation in Clevenger's apparatus for 3 h. The volatile fraction (EO) was separated, traces of water removed by passing through anhydrous sodium sulfate, then stored in a clean dark glass vial and kept at 4 °C until use. 2.6. Chemical characterization of cumin seed essential oil Cumin seed EO was subjected to gas chromatography (PerkinElmer Auto XL GC, MA, USA) equipped with a flame ionization detector and the GC condition were: EQUITY-5 column (60 m × 0.32 mm × 0.25 μm); the carrier gas was H2; column head pressure 10 psi; oven temperature program isotherm 2 min at 70 °C, 3 °C/min gradient to 250 °C, isotherm10 min; injection temperature, 250 °C; detector temperature 280 °C. GC–MS analysis was performed using PerkinElmer Turbomass GC–MS. The GC column was EQUITY-5 (60 m × 0.32 mm × 0.25 μm) fused silica capillary column. The GC conditions were: injection temperature, 250 °C; column temperature, isothermal at 70 °C for 2 min, then programmed to 250 °C at 37 °C/min and held at this temperature for 10 min; ion source temperature, 250 °C. Helium was the carrier gas. The effluent of the GC column was introduced directly into the source of MS and spectra obtained in the EI mode with 70 eV ionization energy. The sector mass analyzer was set to scan from 40 to 500 amu for 2 s. The identification of individual compounds was based on their retention times relative to those of authentic samples and matching spectral peaks available with Wiley, NIST and NBS mass spectral libraries or with the published data (Adams, 2007). 2.7. Evaluation of toxicity of cumin seed EO against A. flavus strain LHP(C)-D6 The toxicity of cumin seed EO against A. flavus strain LHP(C)-D6 was measured in terms of minimal inhibitory concentration (MIC) and minimal fungicidal concentration (MFC) by the poisoned food assay.
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Requisite amounts of the EO were dissolved separately in Petri plates containing 0.5 ml acetone and then 9.5 ml PDA added to obtain different concentrations (0.1 to 1 μl/ml). A fungal disk (5 mm diameter) of seven days old culture of A. flavus strain LHP(C)-D6 was inoculated onto prepared Petri dishes. The control sets contained PDA with acetone but no EO. The inoculated Petri dishes were incubated at 27 ± 2 °C for 7 days. Fungal colony diameters of treatment and control sets were measured. The lowest concentration of the EO resulting in no growth of the A. flavus was taken as the MIC. For determination of MFC, the inhibited fungal disks of oil treated sets were re-inoculated on fresh medium after washing with distilled water and revival of their growth (fungistatic/fungicidal) was observed after 7 days. The lowest concentration preventing revival of fungal growth was taken as the MFC (Kumar et al., 2008). 2.8. Evaluation of cumin seed EO as aflatoxin B1 suppressor For aflatoxin B1 suppressor activity of cumin seed EO, the requisite amount of EO was dissolved separately in 0.5 ml acetone and added to 24.5 ml SMKY to achieve concentrations from 0.1 to 1.0 μl/ml. The medium was inoculated with a 5 mm diameter disk of seven days old culture of A. flavus strain LHP(C)-D6. The control sets contained acetone but no EO. AFB1 was detected by thin layer chromatography as described in Section 2.4. The fungal mats were dried at 80 °C (12 h) to determine the net mycelial dry weight. 2.9. Effect of cumin seed EO on ergosterol content in plasma membrane of A. flavus strain LHP(C)-D6 The ergosterol content in the plasma membrane of A. flavus was assessed following Tian et al. (2012) with slight modifications. A 100 μl aliquot of spore suspension of A. flavus strain LHP(C)-D6 (106 spores/ml) in 0.1% Tween-80 was inoculated in PDB medium containing 0, 0.15, 0.3, 0.45 and 0.6 μl/ml cumin seed EO. After 4 days of incubation at 27 ± 2 °C, mycelia was harvested and washed twice with distilled water. The net wet weight of the cell pellet was measured. Five ml of 25% alcoholic potassium hydroxide solution (25 g KOH and 35 ml sterile distilled water, brought to 100 ml with 100% ethanol) was added to each cell pellet and vortex mixed for 2 min, followed by incubation in an 85 °C water bath for 4 h. Sterols were then extracted from each sample by adding a mixture of 2 ml sterile distilled water and 5 ml n-heptane followed by sufficient vortex mixing for 2 min. After allowing the layers to separate for 1 h at room temperature, the n-heptane layer was separated and analyzed by scanned spectrophotometry between 230 and 300 nm. The presence of ergosterol and the late sterol intermediate 24(28) dehydroergosterol in the n-heptane layer resulted in a characteristic four peaked curve. The absence of ergosterol in samples was indicated by a flat line. The ergosterol amount was calculated as a percentage of the wet weight of the cells as follows:% ergosterol + % 24(28) dehydroergosterol = (A282/290)/pellet weight; % 24(28) dehydroergosterol = (A230/518)/pellet weight; and % ergosterol = (% ergosterol + % 24(28) dehydroergosterol) — % 24(28) dehydroergosterol, where 290 and 518 are the E values (in percentages per cm) determined for crystalline ergosterol and 24(28) dehydroergosterol, respectively, and pellet weight is the net wet weight (g). 2.10. Spectrum of toxicity of cumin seed EO against food borne molds isolated from wheat and chickpea mycoflora analysis
3
only acetone (0.5 ml) and 9.5 ml PDA inoculated with the test fungi served as control. The plates of both treatment and control sets were incubated at 27 ± 2 °C for seven days. The percent inhibition of fungal growth was determined. The plates showing no visible fungal growth were sub-cultured on EO-free PDA plates to determine if the inhibition was reversible (fungistatic/fungicidal). 2.11. Antioxidant activity of cumin seed EO through DPPH free radical analysis Free radical scavenging activity of the cumin seed EO was measured by recording the extent of bleaching of a DPPH solution from purple to yellow following Prakash et al. (2012). Different concentrations (0.02 to 0.2 μl/ml) of the EO were added to 0.004% DPPH solution in methanol (5 ml). After 30 min incubation at room temperature (25 ± 2 °C), the absorbance was measured against a blank at 517 nm using a spectrophotometer. The antioxidant activity was measured through scavenging of DPPH free radical with reduction in absorbance of the sample. The IC50 of the test compounds, which represented the concentration that caused 50% neutralization of DPPH radicals, was measured from the graph plotting percentage inhibition against concentration. 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. 2.12. In vivo investigation with cumin seed EO against fungal deterioration of wheat and chickpea during storage Experiments were designed to fumigate the wheat and chickpea samples with cumin seed EO in airtight containers following Varma and Dubey (2001). In one set (uninoculated treatment), 1.2 ml of the EO, soaked in a 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 0.6 μl/ml air. In another set (inoculated treatment), the wheat and chickpea samples, prior to treatment with the oil, were inoculated with 2 ml spore suspension of A. flavus strain LHP(C)-D6 prepared in 0.1% Tween 80. The control set also contained two sets — the uninoculated control and the inoculated control. After 12 months of storage at laboratory conditions (temperature 10–46 °C and RH 30–90%), food samples of both treatment and control sets were analyzed for fungi using the direct plating and serial dilution methods as mentioned in Section 2.3. The percent protection in the uninoculated and inoculated treatments was observed by following formula. Percent protection = Dc − Dt / Dc × 100 where, Dc = percent occurrence of total fungi of control set and Dt = percent occurrence of total fungi in treatment set. 2.13. Effect of cumin seed EO on germination of fumigated seeds The germination of wheat and chickpea seeds, treated with cumin seed EO was tested after 12 months of storage by placing 50 randomly selected seeds in Petri plates containing moistened blotting paper. The seeds germinated within a week were recorded as viable. The per cent germination was calculated with respect to control sets. 2.14. Statistical analysis
The toxicity of cumin seed EO for 19 food borne molds, isolated from wheat and chickpea during mycoflora analysis, was evaluated by the poisoned food technique. The oil at its MIC (viz. 0.6 μl/ml) was added separately to Petri plates containing 0.5 ml acetone and 9.5 ml molten PDA. A 5 mm disk from a seven day old colony from each fungus was separately placed on the center of the prepared plates. Plates containing
All experiments except mycoflora analysis 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 b 0.05). The analysis of data was performed with the SPSS program version 16.0.
A. Kedia et al. / International Journal of Food Microbiology 168–169 (2014) 1–7
28 54 – – – – – 1 – – – – – – – – – – – Total 72 70 – – – – – – – – – – – – – – – – – 36 26 – – – – – 1 2 7 9 – – – – – – – –
Samrat
DP
MalwiyaHUW234
SD
38 5 16 38 – – 8 12 – – – – – 6 – – – – –
DP
34 33 4 – – – – 1 – 2 – 2 – – – – – – –
50 44 – – – – – – 13 7 – – – – – 2 – – –
DP SD DP
SD
Total Kabuli Pusa 256 Chickpea mycoflora analysis
K68 Wheat mycoflora analysis
Serial no.
Compound
Retention time (min.)
%
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
Alpha-pinene Camphene Benzaldehyde Laevo beta pinene Cymene L-Limonene Gamma-terpinene Para-alpha-dimethyl styrene Cherry propanol Alpha-terpineol Myrtenol P-Allylanisole Cuminaldehyde Carvotanacetone (E)-Cinnamaldehyde Eugenol
9.126 9.650 10.00 10.60 12.38 12.50 13.70 15.001 19.351 19.651 19.951 20.001 22.08 22.326 23.40 27.376
0.99 0.14 0.02 11.50 47.08 1.05 19.36 0.65 0.02 0.22 0.04 0.12 14.92 0.17 0.84 0.33 Total 97.45
70 50 – – – – – – – 2 – – – – – – – – – DP = Direct plating method; SD = Serial dilution method
DP
23 16 – – 8 – – 5 3 6 8 – 1 – 3 1 – – – 25 – – – 13 7 15 1 – – – 9 9 3 – – – – –
SD DP
22 19 15 5 9 3 – 9 4 – 1 4 – – – – – 2 1 42 – 39 28 22 13 10 1 – – – – – – – 3 – –
SD
The moisture content of wheat samples varied from 12 to 14% whereas that of chickpea samples varied from 11 to 16%. The pH values were slightly acidic, for wheat samples the pH ranged from 6.2 to 6.67 whereas for chickpea samples it was in the range of 6.0 to 6.14. During mycological survey, a total of 1230 isolates belonging to 11 different genera and 19 species were isolated. Aspergillus. was the dominant genus. A. flavus and A. niger were found in all the investigated samples and A. flavus was found to be the dominant species (Table 1). In assessing the toxigenicity of A. flavus isolates from the commodities, 20 isolates from wheat and 19 isolates from chickpea were found to be aflatoxigenic with blue spots on TLC plates at 360 nm. The toxigenic isolate A. flavus LHP(C)-D6, from Chickpea var. Samrat was selected as test fungus for detailed study as it produced maximum aflatoxin B1 (2199.84 μg/l). The EO obtained from cumin seed was pale yellow in color with a spicy odor. The yield ranged between 0.9 and 1.2% on fresh weight basis through hydrodistillation. Chemical characterization by the GC–MS analysis of the cumin seed oil revealed the presence of 16 compounds comprising 97.45% of total EO. Their retention time and area percentage are summarized in Table 2. The major components of EO were cymene (47.08%), gamma-terpinene (19.36%), cuminaldehyde (14.92%) and laevo beta pinene (11.50%). The MIC of cumin seed EO for complete inhibition of growth of A. flavus strain LHP(C)-D6 was found at 0.6 μl/ml. The fungicidal concentration (MFC value) of the EO was recorded at 0.9 μl/ml. The EO inhibited AFB1 production as well as
40 7 1 – 6 16 – – 8 – 1 2 – – – – – – – Aspergillus flavus Aspergillus niger Curvularia lunata Aspergillus glaucus Alternaria alternata Penicillium citrinum Aspergillus unguis Aspergillus terreus Rhizopus stolonifer Aspergillus nidulans Mucor sp. Mycelia sterilia Penicillium italicum Cladosporium cladosporioides Penicillium purpurogenum Penicillium luteum Fusarium oxysporum Absidia ramosa Spondylocladium australe 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19.
SD
3. Results
K65 No. of fungal isolates Fungus isolated Sr no.
Table 1 Fungi isolated from wheat and chickpea samples.
Table 2 GC–MS analysis of Cuminum cyminum seed essential oil.
480 314 75 71 58 47 33 31 30 24 19 17 10 9 3 3 3 2 1 1230
4
Table 3 Effect of different concentrations of Cuminum cyminum seed EO on growth, dry weight, aflatoxin B1 production and nature of toxicity to the A. flavus strain LHP(C)-D6. Conc. (μl/ml)
Diameter (cm)
Mycelial dry wt. (g)
AFB1 content (μg/l)
Nature of toxicity
CNT 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0
5.95 ± 0.14e 4.84 ± 0.07d 4.57 ± 0.14d 3.11 ± 0.08c 3.00 ± 0.17c 1.69 ± 0.02b 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a
0.46 ± 0.01f 0.42 ± 0.01e 0.41 ± 0.00de 0.38 ± 0.01d 0.22 ± 0.00c 0.04 ± 0.00b 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a 0.00 ± 0.00a
1517.06 ± 145.25 1245.14 ± 24.79 1173.58 ± 37.87 1030.45 ± 65.59 495. 93 ± 48.13 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00
Static Static Static Static Static Static Static Static Static Cidal Cidal
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.
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5
Fig. 2. Antioxidant activity of cumin seed EO through DPPH free radical analysis. Fig. 1. Effect of different concentrations of cumin seed EO on mycelial wet wt. and ergosterol content of A. flavus strain LHP(C)-D6.
mycelial dry weight in a dose dependent manner and at 0.5 μl/ml, it completely inhibited the production of AFB1 (Table 3). The effects of cumin seed EO on ergosterol content in the plasma membrane of A. flavus strain LHP(C)-D6 with respect of mycelial wet wt. are shown in Fig. 1. A dose dependent decrease in ergosterol content was observed on increasing concentration of the EO. A reduction percentage of the ergosterol content as compared with the control was found to be 28.57, 35.71, 64.29 and 100% respectively for 0.15, 0.3, 0.45 and 0.6 μl/ml concentrations. Hence, the cumin seed oil completely reduced the ergosterol content at 0.6 μl/ml concentrations which is equal to its MIC value. The oil exhibited a broad spectrum of fungal toxicity inhibiting all 19 food borne fungal species except Rhizopus stolonifer at its MIC (Table 4). The EO was found to be fungicidal against most of the fungal species except Absidia ramosa, Aspergillus glaucus, A. niger, Aspergillus terreus, Aspergillus unguis, Fusarium oxysporum and Mucor sp. The bleaching of the purple color of the DPPH to yellow confirmed the positive antioxidant activity of EO. The oil showed strong free radical scavenging activity as its IC50 value was found to be 0.092 μl/ml (Fig. 2).
Table 4 Fungitoxic spectrum and nature of toxicity of Cuminum cyminum seed EO at 0.6 μl/ml concentration against some food borne fungi. Serial no.
Fungal species
% mycelial inhibition
Nature of toxicity
1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19.
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 Spondylocladium australe
100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 100.00 ± 0.00 88.51 ± 0.98 100.00 ± 0.00
Static Cidal Cidal Static Static Static Static Cidal Cidal Static Static Cidal Cidal Cidal Cidal Cidal Cidal – Cidal
Values are mean (n = 3) ± standard error.
The results of in vivo trials in food system are based on percent protection of wheat and chickpea from fungal association in the uninoculated and inoculated treatments. In the uninoculated samples, 61.29 and 27.27% protection of wheat and chickpea respectively against storage molds was observed for serial dilution sets, whereas for direct plating method the percent protection of wheat and chickpea was 65.85 and 75.00% respectively. Similarly in inoculated samples, 65.15 and 50.00% protection was recorded for serial dilution sets and 20.00 and 50.00% protection was recorded for direct plating method for wheat and chickpea samples respectively. The seed germination tests revealed that wheat and chickpea seeds fumigated with cumin seed EO had no loss in viability even after 12 months of treatment with 100% seeds germinating in Petri plates. Hence, the cumin seed EO was not phytotoxic. 4. Discussion The extent of mold contamination depends on various conditions such as source, harvesting period, storage practices and chemical nature of the substrate (Prakash et al., 2010). Moisture content and, to a lesser extent, pH are two other important abiotic factors that govern fungal growth and proliferation on stored food commodities (Ozkan et al., 2003). In the present study the moisture content and pH of selected commodities were found within conducive range for fungal proliferation (FAO, 1980). The results indicate that all the selected samples were heavily contaminated with the different mold species. A total of 19 fungal species were recorded from the samples and Aspergillus was found to be dominant in both serial dilution and direct plating methods. The finding supports the earlier observations where Aspergillus spp. were dominant in some stored edible pulses (Shukla et al., 2009) and wheat (Varma and Dubey, 2001). Higher incidence of Aspergilli than other molds may be due to their saprophytic nature and ability to colonize diverse substrates because of secretion of various hydrolytic enzymes (Leite de Souza et al., 2005). The relative frequency of A. flavus and A. niger was higher than that of the remaining fungal species supporting the earlier observations made by Prakash et al. (2011) and Shukla et al. (2009). The quantitative estimation of aflatoxin B1 production in the present study reveals that the samples were highly contaminated by toxigenic strains of A. flavus. Hence, the biodeterioration of the samples was qualitative as well as quantitative. In the present investigation cumin seed oil was selected for detailed investigation to assess its efficacy as plant based preservative. Prior to antimicrobial assay, an essential oil should be chemically characterized to standardize its chemical nature. The chemical profile of essential oil of a particular plant species shows
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different chemotypic variations because of ecological and geographical conditions, age of plant and time of harvesting (Prakash et al., 2010). Such chemotypic variations would definitely affect the biological activity of the essential oil. Hence, it is advisable that the percentage of the major components of the EOs should be mentioned if applied as food additive (Bagamboula et al., 2004). In the present investigation, the GC and GC–MS analysis of cumin oil revealed the presence of 16 compounds comprising 97.45% of total oil. Cymene was identified as the major compound contributing 47.08% followed by gammaterpinene (19.36%), cuminaldehyde (14.92%) and laevo beta pinene (11.50%). In earlier reports, cuminaldehyde (36–39%) was reported as the major component of cumin seed EO from China (Li and Jiang, 2004), Bulgaria (Jirovetz et al., 2005) and Tunisia (Hajlaoui et al., 2010). However, alpha-Pinene (29.2%) was reported as the major component of cumin seed EO from Iran (Mohammadpour et al., 2012). This variation in EO composition could be due to ecological and geographical conditions. In this study, the antifungal activity of cumin seed oil has been assessed. The study of MIC is important to determine the minimum dose to control fungal populations resulting in least wastage of the pesticide (Kumar et al., 2008). The cumin seed EO showed high antifungal activity against A. flavus LHP(C)-D6 as its MIC was lower than that of some earlier reported EOs viz. Origanum majorana, Coriandrum sativum, Hedychium spicatum, Commiphora myrrha and Cananga odorata (Prakash et al., 2012), Cinnamomum jensenianum (Tian et al., 2012) and some prevalent fungicides such as Nystatin and Wettasul-80 (Prakash et al., 2010) emphasizing its superiority as an antimicrobial agent at a low prescribed dose. The oil was fungicidal in nature above MIC value as its MFC against test fungus was slightly greater than its MIC. Therefore, cumin seed EO could be capable of permanent disinfection of food borne fungi from food items. In addition, the EO also exhibited marked efficacy in checking aflatoxin B1 production by A. flavus LHP(C)-D6 at a concentration lower than MIC. Hence, the EO would be acting through two different modes of action as inhibitor of both fungal growth and aflatoxin production as has been earlier reported by Prakash et al. (2012), Rasooli and Abyaneh (2004) and Tian et al. (2012). The antifungal mechanism of EO involves membrane disruption by their lipophilic compounds. The lowmolecular weight and highly lipophilic components of EO pass easily through cell membranes and cause disruption to the fungal cell organization (Chao et al., 2005). Likewise, the aflatoxin B1 production inhibition efficacy of EO may be because of inhibition of carbohydrate catabolism in A. flavus by acting on some key enzymes, reducing its ability to produce aflatoxins (Tian et al. 2011). The MIC of the cumin EO for inhibition of growth of test fungus and aflatoxin production was different from that of earlier reports (Khosravi et al., 2011a,b; Mohammadpour et al., 2012). This difference in MIC may be due to the difference in chemical profile of the oil, strain of the test fungus, technique adopted for antifungal assay and the medium used. In the present investigation the mode of action of the cumin EO has been observed which would be helpful in its formulation as preservative. Ergosterol is specific to fungi and is the major sterol component of the fungal cell membrane. It is also responsible for maintaining the cell function and integrity (Rodriguez et al., 1985). There is a previous study by Kelly et al. (1995) stating that the primary action mechanism, by which azole antifungal drugs inhibit fungal cell growth, is the disruption of normal sterol biosynthetic pathways resulting in a decrease of ergosterol biosynthesis. Some studies have shown that essential oils can also cause a considerable reduction in the quantity of ergosterol (Pinto et al., 2006, 2009). The cumin seed oil completely reduced the ergosterol content at 0.6 μl/ml concentrations which is equal to its MIC value. The reduction in mycelial weight as well as ergosterol content on increasing concentration of oil clearly suggests its action on plasma membrane of fungus supporting earlier findings of Kelly et al. (1995) and Tian et al. (2012). The present findings revealed that cumin seed EO can induce a considerable impairment of
the ergosterol biosynthesis by A. flavus and subsequent decrease in biomass on increasing concentration of EO. Hence, the plasma membrane is an important antifungal target of cumin seed EO. This emphasizes that the antimicrobial components of the essential oils cross the cell membrane, interact with the enzymes and proteins of the membrane, thus producing a flux of protons towards the cell exterior which induces disruption to the fungal cell organization and, ultimately, their death as has been supported by the transmission electron microscopy by Nogueira et al. (2010); and Tian et al. (2012). In addition, cumin seed EO also exhibited strong antioxidant activity and can enhance the shelf life of food products by controlling free radical scavenging and oxidation of unsaturated lipids. The IC50 value of essential oil was found to be better than that of the earlier reported essential oils viz. Cananga odorata and Coriandrum sativum (Prakash et al., 2012). Free radical scavenging activity of EO may be due to the presence of the phenolic compounds or synergistic effect of overall compounds (Sharififar et al., 2007). The practical applicability of cumin seed EO has been observed as fumigant in food system in plastic storage containers. The oil showed remarkable in vivo efficacy in protecting the fumigated wheat and chickpea samples, from fungal contamination during storage up to 12 months. Hence, further large scale trials are required for its recommendation as a plant based preservative. The preliminary phytotoxic test of cumin seed EO was performed in the form of seed germination test. Results on seed germination test showed 100% germination of wheat and chickpea seeds in EO treated sets. The oil treated food commodities may thus be used for sowing purpose by the growers. Moreover, since EO is volatile in nature, the vapors may be easily eliminated from the fumigated food items after sun drying. In this connection, currently microencapsulation technology is prescribed to use the essential oils as antimicrobials for preservation of stored food commodities and in food industry for flavor stabilization (Burt, 2004). Essential oils of many edible and medicinal plants are used in different pharmaceutical preparations which minimize questions regarding their safe use. Essential oils from aromatic and medicinal plants are potentially useful as antimicrobial agents and their use as medicines have long been recognized (Kim et al., 2005). The attraction of modern society towards herbal products desiring fewer synthetic ingredients in foods and recommendation of herbal products as ‘generally recognized as safe’ (GRAS) as food additives may lead scientific interest in the exploitation of essential oils as plant based food additives (Smid and Gorris, 1999). Moreover, the regulatory status of cumin and cumin oil in the USA is regarded as safe, GRAS 2340 and GRAS 2343 (Parthasarathy et al., 2008). The cumin seeds are traditionally used as safe culinary items in form of spice. Bulk of raw materials will be available for formulation of oil as preservative as the cumin plants grow luxuriantly in different tropical and subtropical countries. Because of renewable source, high yield of the oil, and efficacy at low concentrations, the oil may be economically recommended for formulation as plant based preservatives. However, standardization of dose during practical application is desirable before its recommendation to agri-firms. This is the first report on mode of antifungal action of cumin seed EO and its efficacy as fumigant in food system. In conclusion, based on efficacy as inhibitor to fungal growth and aflatoxin secretion, its free radical scavenging potential, efficacy in food system and favorable safety profile, the cumin seed oil has possibility to provide complete protection of food commodities against quantitative and qualitative losses, thereby, enhancing their shelf life.
Acknowledgments This work was financially supported by University Grant Commission (UGC), New Delhi, India.
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References Adams, R.P., 2007. Identification of Essential Oil Components by Gas Chromatography/ Mass Spectrometry. Allured Publishing Corporation, Carol Stream, IL. Ahmad, S.K., Singh, P.L., 1991. Mycofloral changes and aflatoxin contamination in stored chickpea seeds. Food Addit. Contam. 8, 723–730. Bagamboula, C.F., Uyttendaele, M., Debevere, J., 2004. Inhibitory effect of thyme and basil essential oils, carvacrol, thymol, estragol, linalool and p-cymene towards Shigella sonnei and S. flexneri. Food Microbiol. 21, 33–42. Burt, S., 2004. Essential oils: their antibacterial properties and potential applications in foods — a review. Int. J. Food Microbiol. 94, 223–253. Chao, L.K., Hua, K.F., Hsu, H.Y., Cheng, S.S., Lin, J.Y., Chang, S.T., 2005. Study on the antiinflammatory activity of essential oil from leaves of Cinnamomum osmophloeum. J. Agric. Food Chem. 53, 7274–7278. Dayan, F.E., Cantrell, C.L., Duke, S.O., 2009. Natural products in crop protection. Bioorg. Med. Chem. 17, 4022–4034. Devegowda, G., Raju, M.V.L.N., Swang, H.V.L.N., 1998. Mycotoxins: novel solutions for their counteraction. Feedstuffs 70, 12–15. Dhingra, O.D., Mizubuti, E.S.G., Napoleao, I.T., Jham, G., 2001. Free fatty acid accumulation and quality loss of stored soybean seeds invaded by Aspergillus ruber. Seed Sci. Technol. 29, 193–203. Domsch, K.H., Gams, W., Anderson, T.H., 1980. Compendium of Soil Fungi, vol. 1 and 2. Academic Press, London. Dubey, N.K., 2004. Flora of BHU Campus. Banaras Hindu University Press, Varanasi, India. FAO (Food and Agriculture Organisation), 1980. On-farm maize drying and storage in the humid tropics. FAO Agric. Serv. Bull. 40, 1–54. Farag, R.S., Daw, Z.Y., Abo-Raya, S.H., 2006. 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. Giray, B., Girgin, G., Engin, A.B.., Aydın, S., Sahin, G., 2007. Aflatoxin levels in wheat samples consumed in some regions of Turkey. Food Control 18, 23–29. Hajlaoui, H., Mighri, H., Noumi, E., Snoussi, M., Trabelsi, N., Ksouri, R., Bakhrouf, A., 2010. Chemical composition and biological activities of Tunisian Cuminum cyminum L. essential oil: a high effectiveness against Vibrio spp. strains. Food Chem. Toxicol. 48, 2186–2192. Jaya, Prakash, B., Dubey, N.K., 2011. Evaluation of chemically characterised essential oils of Coleus aromaticus, Hyptis suaveolens and Ageratum conyzoides against storage fungi and aflatoxin contamination of food commodities. Int. J. Food Sci. Technol. 46, 754–760. Jirovetz, L., Buchbauer, G., Stoyanova, A.S., Georgiev, E.V., Damianova, S.T., 2005. Composition, quality control and antimicrobial activity of the essential oil of cumin (Cuminum cyminum L.) seeds from Bulgaria that had been stored for up to 36 years. Int. J. Food Sci. Technol. 40, 305–310. Kelly, S.L., Lamb, D.C., Corran, A.J., Baldwin, B.C., Kelly, D.E., 1995. Mode of action and resistance to azole antifungals associated with the formation of 14α-methylergosta8, 24(28)-dien-3β, 6α-diol. Biochem. Biophys. Res. Commun. 207, 910–915. Khosravi, A.R., Minooeianhaghighi, M.H., Shokri, H., Emami, S.A., Alavi, S.M., Asili, J., 2011a. The potential inhibitory effect of Cuminum cyminum, Ziziphora clinopodioides and Nigella sativa essential oils on the growth of Aspergillus fumigatus and Aspergillus flavus. Braz. J. Microbiol. 42, 216–224. Khosravi, A.R., Shokri, H., Minooeianhaghighi, M., 2011b. Inhibition of aflatoxin production and growth of Aspergillus parasiticus by Cuminum cyminum, Ziziphora clinopodioides and Nigella sativa essential oils. Foodborne Pathog. Dis. 8 (12), 1275–1280. Kim, J.H., Campbell, B.C., Yu, J., Mahoney, N., Chan, K.L., Molyneux, R.J., Bhatnagar, D., Cleveland, T.E., 2005. Examination of fungal stress response genes using Saccharomyces cerevisiae as a model system: targeting genes affecting aflatoxin biosynthesis by Aspergillus flavus Link. Appl. Microbiol. Biotechnol. 67, 807–815. Kumar, A., Shukla, R., Singh, P., Prasad, C.S., Dubey, N.K., 2008. Assessment of Thymus vulgaris L. essential oil as a safe botanical preservative against post harvest fungal infestation of food commodities. Innov. Food Sci. Emerg. Technol. 9, 575–580. Leite de Souza, E., Antonio da Silva, C., Paiva de Sousa, C., 2005. Bacteriocins: molecules of fundamental impact on the microbial ecology and potential food biopreservatives. Braz. Arch. Biol. Technol. 48 (4), 559–566. Li, R., Jiang, Z., 2004. Chemical composition of the essential oil of Cuminum cyminum L. from China. Flavour Fragr. J. 19, 311–313. Mandeel, Q.A., 2005. Fungal contamination of some imported species. Mycopatholgia 159, 291–298. Mishra, P.K., Shukla, R., Singh, P., Prakash, B., Kedia, A., Dubey, N.K., 2012. Antifungal, antiaflatoxigenic, and antioxidant efficacy of Jamrosa essential oil for preservation of herbal raw materials. Int. Biodeterior. Biodegrad. 74, 11–16.
7
Mishra, P.K., Singh, P., Prakash, B., Kedia, A., Dubey, N.K., Chanotiya, C.S., 2013. Assessing essential oil components as plant-based preservatives against fungi that deteriorate herbal raw materials. Int. Biodeterior. Biodegrad. 80, 16–21. Mohammadpour, H., Moghimipour, E., Rasooli, I., Fakoor, M.H., Astaneh, S.A., Moosaie, S.S., Jalili, Z., 2012. Chemical composition and antifungal activity of Cuminum cyminum L. essential oil from Alborz mountain against Aspergillus species. Jundishapur. J. Nat. Pharm. Prod. 7 (2), 50–55. Nogueira, J.H., Gonçalez, E., Galleti, S.R., Facanali, R., Marques, M.O., Felício, J.D., 2010. Ageratum conyzoides essential oil as aflatoxin suppressor of Aspergillus flavus. Int. J. Food Microbiol. 137, 55–60. Norman, J., 1990. The Complete Book on Spices. Doerling Kindersley, London. Osman, K.A., Abdulrahman, H.T., 2003. Risk assessment of pesticide to human and the environment. Saudi J. Biol. Sci. 10, 81–106. Ozkan, M., Kirca, A., Cemeroglu, B., 2003. Effect of moisture content on CIE color values in dried apricots. Eur. Food Res. Technol. 216, 2170–2219. Parthasarathy, V.A., Chempakam, B., Zachariah, T.J., 2008. Chemistry of Spices. CABI, Oxfordsire, United Kingdom (U.K.). Pinto, E., Pina-Vaz, C., Salgueiro, L., Goncalves, M.J., Costa-de-Oliveira, S., Cavaleiro, C., et al., 2006. Antifungal activity of the essential oil of Thymus pulegioides on Candida, Aspergillus and dermatophyte species. J. Med. Microbiol. 55 (10), 1367–1373. Pinto, E., Vale-Silva, L., Cavaleiro, C., Salgueiro, L., 2009. Antifungal activity of the clove essential oil from Syzygium aromaticum on Candida, Aspergillus and dermatophyte species. J. Med. Microbiol. 58 (11), 1454–1462. Pitt, J.I., 1979. The Genus Penicillum. Academic Press, Sydney, Australia. Prakash, B., Shukla, R., Singh, P., Kumar, A., Mishra, P.K., Dubey, N.K., 2010. Efficacy of chemically characterized Piper betle L. essential oil against fungal and aflatoxin contamination of some edible commodities and its antioxidant activity. Int. J. Food Microbiol. 142, 114–119. 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., 2012. 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, 201–208. Raper, K.B., Fennell, D.L., 1977. The Genus Aspergillus. Krieger publishing Corporation, Huntington, New York. Rasooli, I., Abyaneh, M.R., 2004. Inhibitory effects of Thyme oils on growth and aflatoxin production by Aspergillus. parasiticus. Food Control 15, 479–483. 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. Sharififar, F., Moshafi, M.H., Mansouri, S.H., Khodashenas, M., Khoshnoodi, M., 2007. In vitro evaluation of antibacterial and antioxidant activities of the essential oil and methanol extract of endemic Zataria multiflora Boiss. Food Control 18, 800–805. Shukla, R., Kumar, A., Singh, P., Dubey, N.K., 2009. Efficacy of Lippia alba (Mill.) N.E. Brown essential oil and its monoterpene aldehyde constituents against fungi isolated from some edible legume seeds and aflatoxin B1 production. Int. J. Food Microbiol. 135, 165–170. Singh, G., Kapoor, I.P.S., Pandey, S.K., Singh, U.K., Singh, R.K., 2002. Antibacterial activity of volatile oils of some spices. Phytother. Res. 16 (7), 680–682. Smid, E.J., Gorris, L.G.M., 1999. Natural Antimicrobials for Food Preservation. In: Rahman, M.S. (Ed.), Handbook of Food Preservation. Marcel Dekker, New York, pp. 285–308. Soliman, K.M., Badeaa, R.I., 2002. Effect of oil extracted from some medicinal plants on different mycotoxigenic fungi. Food Chem. Toxicol. 40, 1669–1675. Tian, J., Ban, X., Zeng, H., He, J., Huang, B., Youwei, W., 2011. Chemical composition and antifungal activity of essential oil from Cicuta virosa L. var. latisecta Celak. Int. J. Food Microbiol. 145, 464–470. 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. Tomaino, A., Cimino, F., Zimbalatti, V., Venuti, V., Sulfaro, V., De Pasquale, A., Saija, A., 2005. Influence of heating on antioxidant activity and the chemical composition of some spice essential oils. Food Chem. 89, 549–554. Toteja, G.S., Mukherjee, A., Diwakar, S., Singh, P., Saxena, B.N., Sinha, K.K., et al., 2006. Aflatoxin B1 contamination in wheat grain samples collected from different geographical regions of India: a multicenter study. J. Food Prot. 69 (6), 1463–1467. 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. 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.
