Food Research International 49 (2012) 201–208 Contents lists available at SciVerse ScienceDirect Food Research International journal homepage: www.elsevier.com/locate/foodres Assessment of some essential oils as food preservatives based on antifungal, antiaﬂatoxin, antioxidant activities and in vivo efﬁcacy in food system Bhanu Prakash, Priyanka Singh, Akash Kedia, N.K. Dubey ⁎ Laboratory of Herbal Pesticides, Centre of Advanced Study in Botany, Banaras Hindu University, Varanasi-221005, India a r t i c l e i n f o Article history: Received 15 June 2012 Accepted 22 August 2012 Available online 31 August 2012 Keywords: Aﬂatoxin B1 Antifungal Antioxidant Essential oils In vivo Seed germination a b s t r a c t The present study strengthens the food preservative potential of plant essential oils (EOs) viz. Origanum majorana L., Coriandrum sativum L., Hedychium spicatum Ham. ex Smith, Commiphora myrrha (Nees) Engl., and Cananga odorata Hook.f. & Thomson based on their antifungal, antiaﬂatoxin, and antioxidant efﬁcacy. The essential oils were found more efﬁcacious than some prevalent organic preservatives viz. salicylic acid, BHT, ascorbic acid and gallic acid as they inhibited the growth and aﬂatoxin secretion of the aﬂatoxigenic strain Aspergillus ﬂavus (LHP-6) at lower concentrations. The minimum inhibitory concentration of EOs against the toxigenic strain of A. ﬂavus ranged between 2.0 μl/ml and 3.0 μl/ml. In addition, the EOs exhibited broad fungitoxic spectrum against nine food borne molds and strong antioxidant activity. The IC50 value of the oils ranged between 1.3 and 21.67 μl/ml while their total phenolic content ranged between 2.90 and 33.33 μg/mg. All the EOs except C. odorata showed non-phytotoxic nature on germination of chickpea seeds. During in vivo investigations in food system all the ﬁve essential oils showed above 50% protection of chickpea seed from A. ﬂavus infestation showing their potential as plant based preservatives for enhancement of shelf life of food items. © 2012 Published by Elsevier Ltd. 1. Introduction In view of the rapid increase in world population many countries experience perpetual food shortage resulting in chronic and often widespread hunger among signiﬁcant number of people. Loss of food commodities due to storage pests is a major reason of food crisis particularly in tropical countries. According to FAO, food borne molds and their toxic metabolites render quantitative and qualitative losses of nearly 25% of agricultural food items throughout the world (Pittet, 1998; Singh et al., 2010a). Mold infestation results in reduction of grain quality, change in colour and texture, increase in free fatty acids, reduction in nutritional value and germination ability (Dhingra, Mizubuti, Napoleao, & Jham, 2001; Shukla, Kumar, Singh, & Dubey, 2009). Mycotoxins produced by some mold strains have many times resulted to famine even in developed countries (Wagacha & Muthomi, 2008). Among different mold species contamination of food items by the Aspergillus species, particularly A. ﬂavus, has drawn signiﬁcant attention to the scientiﬁc community as some of these species have the capability to excrete aﬂatoxins in affected food items on onset of favourable environmental conditions. Aﬂatoxins are well known for their possible role as carcinogen affecting different human systems (Pier, 1992; Sindhu, Chempakam, Leela, & Bhai, 2011). Because of the cosmopolitan nature of A. ﬂavus, aﬂatoxin contamination has been regarded as an unavoidable contaminant of food items by WHO ⁎ Corresponding author. Tel.: +91 9415295765; fax: +91 5422368174. E-mail address: [email protected] (N.K. Dubey). 0963-9969/$ – see front matter © 2012 Published by Elsevier Ltd. http://dx.doi.org/10.1016/j.foodres.2012.08.020 (Wagacha & Muthomi, 2008). Hence, the presence of molds and aﬂatoxin in stored food commodities is a potential health threat to humans and livestock. In addition to fungal and mycotoxin contamination, lipid peroxidation due to the chain reaction of free radical oxidation in food items is another major problem for food industries because the oxidised lipid imposes an undesirable inﬂuence on humans (Deba, Xuan, Yasuda, & Tawata, 2008). Aﬂatoxins have been also reported to enhance the generation of free radicals (ROS) (Choy, 1993; Prakash, Singh, Mishra, & Dubey, 2012). Hence, prevention of fungal growth, aﬂatoxin secretion as well as lipid peroxidation by using a single measure will be a novel and economical strategy to combat food losses during storage and transit. Some synthetic organic formulations have been recommended to control storage losses of food items. However, in view of some serious drawbacks related to environmental issues, safety concerns, development of resistant races to pests and residual toxicities (Shukla et al., 2009), there is a need for eco-friendly, biodegradable and safer alternatives to control biodeterioration and biodegradation of food items. At the same time, western society also appears to favour a trend of ‘green consumerism’ (Smid & Gorris, 1999) desiring fewer synthetic food additives and products with a smaller impact on the environment. Recently, EOs of aromatic plants are the thrust area of interest of researchers throughout the world in view of their potential as natural source of antimicrobial and antioxidant compounds (Prakash et al., 2012; Viuda-Martos et al., 2011). Many essential oil products are on the ‘Generally Recognised as Safe’ (GRAS) list fully approved by the Food and Drug Administration (FDA) and Environment 202 B. Prakash et al. / Food Research International 49 (2012) 201–208 Protection Agency (EPA) in the USA for food and beverage consumption and their vapour activity as fumigant strengthens their application on food items with wide coverage (Burt, 2004; Viuda-Martos et al., 2011). The aims of the present study were to investigate the antifungal, antiaﬂatoxin, antioxidant, total phenolic content (TPC) and phytotoxic effect of ﬁve plant essential oils viz. Origanum majorana L., Coriandrum sativum L., Hedychium spicatum Ham. ex Smith, Commiphora myrrha (Nees) Engl, and Cananga odorata Hook.f. & Thomson and to assess their in vivo efﬁcacy as plant based fumigants in food system (chickpea) stored in plastic containers so as to explore the possibility of their recommendation in enhancing the shelf life of food items. 2. Materials and methods 2.1. Chemicals and equipments All the chemicals used in the present investigation were procured from Hi Media Laboratories Pvt. Ltd., Mumbai. The major equipments used were hydro-distillation apparatus (Merck Specialities Pvt. Ltd., Mumbai, India), centrifuge, UV transilluminator (Zenith Engineers, Agra, India) and spectrophotometer (Systronics India Ltd., Mumbai, India). 2.2. Plant essential oils Selection of plants (family) and their part viz., H. spicatum (Zingiberaceae) (rhizome), C. sativum (Apiaceae) (fruits), C. myrrha (Burseraceae) (oleoresin-gum), O. majorana (Lamiaceae) (leaf), and C. odorata (Annonaceae) (ﬂower) was done on the basis of ethnobotanical literature and traditional use for the treatment of several diseases (Prajapati, Purohit, Sharma, & Kumar, 2003). Desired plant parts were collected from the campus of Banaras Hindu University, Varanasi, India, during August and October, 2010; while EO of C. odorata was procured from Central Institute of Medicinal and Aromatic Plants (CIMAP), Lucknow, India. The plants were identiﬁed with the help of relevant taxonomic literature/ﬂora (Duthie, 1960; Dubey, 2004) and their voucher specimens (Zing./Hed-024/2010; Api./Cor-014/2010; Bur./Com-03/2010/; Lab./Ori-095/2010) were deposited in the herbarium of the Laboratory of Herbal Pesticides, Department of Botany, BHU, Varanasi. The plant parts were thoroughly washed twice with distilled water prior to hydrodistillation and then subjected to hydrodistillation (4 h) in Clevenger's apparatus (Prakash et al., 2010). The EO was separated and collected in sterilised glass vial. Water traces from the essential oil were removed by adding anhydrous sodium sulphate. The EO was stored at 4 °C in the dark for further experiments. 2.3. Fungal strains and their culture conditions A total of nine molds viz., Aspergillus niger, Aspergillus candidus, Aspergillus terreus, Aspergillus fumigatus, Alternaria alternata, Cladosporium cladosporioides, Fusarium nivale, Penicillium italicum including AFB1 producing strain of A. ﬂavus LHP-6 previously isolated from spices (Prakash et al., 2011), were used for the present investigation. Their cultures were routinely maintained on Czapek-Dox Agar (CDA) (NaNO3, 2 g; K2HPO4, 1 g; MgSO4, 0.5 g; KCl, 0.5 g; FeSO4, 0.01 g; sucrose, 30 g; agar, 15 g; 1 l distilled water, pH 6.8 ± 0.2) medium on slant at 4 °C. Prior to testing, the mold isolates were re-inoculated to freshly prepared CDA medium and incubated for seven days at 27 ± 2 °C to ensure viability and purity of strains. Spore suspension of the fungal strain (LHP-6) was prepared in 0.1% Tween-80, and the spore density was adjusted to ~ 10 6 spore/ml with the help of a hemocytometer. 2.4. Determination of minimum inhibitory concentration (MIC) and minimum fungicidal concentration (MFC) of EOs and organic preservatives against aﬂatoxigenic strain of A. ﬂavus The minimal inhibitory concentration (MIC) and minimal fungicidal concentration (MFC) of the EOs and some prevalent organic preservatives viz. salicylic acid, butylated hydroxytoluene (BHT), butylated hydroxyanisole (BHA), ascorbic acid and gallic acid against the toxigenic strain of A. ﬂavus LHP-6 were determined by the broth dilution method as reported earlier by Shukla et al. (2009). Requisite amounts of the EOs and organic preservatives were dissolved separately in 0.5 ml DMSO and incorporated to 9.5 ml PDB (potato dextrose broth) tubes to achieve different concentrations at 0.25 to 10.0 μl/ml and 0.1 to 10.0 mg/ml respectively. PDB tubes containing DMSO (0.5 ml) only served as control. Thereafter, 50 μl spore suspension (containing 106 spores/ml) of toxigenic strain of A. ﬂavus LHP-6 was added to the control as well as sets treated with EOs and organic preservatives. The tubes were kept in BOD incubator for 10 days incubation period (27± 2 °C). The lowest concentration of EOs/organic preservatives that inhibited the complete growth of test molds was taken as MIC. The minimum fungicidal concentration (MFC) of EOs/organic preservatives was determined following Shukla et al. (2009). Five hundred microliters of medium from the test tube showing no visible growth was subcultured on freshly prepared treatment-free PDA plates to determine the MFC of EOs. MFC is the lowest concentration of EOs/preservatives at which there was no revival of growth of the inhibited fungal inoculum on treatment-free PDA plates because of permanent inhibition. 2.5. Efﬁcacy of plant essential oils and organic preservatives in inhibition of aﬂatoxin B1 secretion The aﬂatoxin inhibitory efﬁcacy of plant essential oils and organic preservatives was investigated following Kumar, Mishra, Dubey, and Tripathi (2007). Different amounts of essential oils/organic preservatives were dissolved separately in 0.5 ml DMSO and then added to 24.5 ml SMKY broth (sucrose, 200 g; MgSO4·7H2O, 0.5 g; KNO3, 0.3 g and yeast extract, 7 g; 1 l distilled water) medium to achieve ﬁnal concentration of 0.5 to 10.0 μl/ml and 0.1 to 10.0 mg/ml respectively. Streptomycin (300 mg/l) was added to the medium for controlling bacterial growth. A spore suspension (50 μl) of toxigenic strain of A. ﬂavus (LHP-6) containing 10 6 spores/ml prepared in 0.1% (v/v) Tween-80 was inoculated in each ﬂask and was kept for 10 days incubation period at 27 ± 2 °C in BOD incubator. SMKY broth containing only 50 μl of toxigenic strain of A. ﬂavus (LHP-6) suspension was kept as control. Thereafter, the content of each ﬂask (25 ml) was ﬁltered (Whatman No. 1) and the ﬂasks with biomass of ﬁltered mycelium were autoclaved at 121 °C for 15 min to kill the hazardous spores. The autoclaved ﬁltered fungal mycelium was harvested and dried at 80 °C (12 h) and weighed. The content of each ﬂask was ﬁltered and extracted with 20 ml chloroform. The extract was evaporated to dryness on a water bath and re-dissolved in 1 ml chloroform. Fifty microliters of chloroform extract was spotted on TLC plates along with the standard of AFB1 and developed in toluene:isoamylalcohol:methanol (90:32:2; v/v/v). The plate was air dried and AFB1 was observed in UV-transilluminator (360 nm). For the quantiﬁcation of AFB1 the blue spots on TLC plates were scratched, dissolved in methanol (5 ml) and centrifuged at 5000 rpm (5 min). Absorbance of the supernatant was recorded at 360 nm and AFB1 was calculated following Kumar et al. (2007). AFB1 contentðμg=lÞ ¼ DM 1000 EL D = absorbance, M = molecular weight (312), E = molar extinction coefﬁcient AFB1 (21,800), L = path length (1 cm). B. Prakash et al. / Food Research International 49 (2012) 201–208 In addition, percent inhibition of AFB1 was calculated as follows: %inhibition ¼ ðC−T=CÞ 100: Where; T = mean concentration of AFB1 in the treatment, C = mean concentration of AFB1 in the control. 2.6. Fungitoxic spectrum of plant essential oils against food borne molds Fungitoxic spectrum of plant essential oils against nine food borne mold species including the toxigenic strain of A. ﬂavus (LHP-6) was evaluated by poisoned food technique as described earlier by Shukla et al. (2009) with slight modiﬁcation. A 5 mm diameter disc of each fungal species was cut from the periphery of seven day old colony and was inoculated aseptically on the centre of the PDA Petri dish amended with test oils (0.25 to 10.0 μl/ml). The PDA plates without EOs inoculated with the respective fungal species served as controls. Both the treatment and control sets were incubated for ten days at 27 ± 2 °C in BOD incubator. The colony diameters of fungal species in treatment and control sets were measured. The percent mycelia growth inhibition (MGI) was calculated with the following formula %MGI ¼ ðDc−DtÞ=Dc 100: Where, Dc = fungal mycelial growth diameter in control sets, Dt = fungal mycelial growth diameter in treated sets. The lowest concentration of EOs that inhibited the complete growth of test molds was taken as MIC. 2.7. Free radical scavenging activity of essential oils and comparative efﬁcacy with some prevalent antioxidants 2.7.1. DPPH free radical scavenging The preliminary antioxidant activity of plant EOs was assayed through TLC method following Tepe, Daferera, Sokmen, Sokmen, and Polissiou (2005). Five microliters of the EOs (1:10 dilution in methanol) was applied separately on TLC plates and developed in ethyl acetate and methanol (1:1 v/v). The plates were sprayed with 0.2% DPPH solution in methanol (2,2-diphenyl‐1-picrylhydrazil) and left at room temperature for 30 min. The yellow spot that developed on the plates was due to bleaching of the purple‐coloured DPPH reagent indicating a positive antioxidant activity of EOs. After conﬁrming the radical scavenging activity of plant EOs detailed investigation was performed through spectrophotometric assay following Prakash et al. (2010). Free radical scavenging activity of the EOs was measured by recording the extent of bleaching of the purple-coloured DPPH solution to yellow. Different concentrations (1.0 to 30.0 μl/ml) for the essential oils (1.0 to 10.0 μg/ml), BHT, BHA, ascorbic acid (100 to 500 mg/ml) and salicylic acid were added to 0.004% methanolic solution of DPPH and kept in the dark at room temperature (25 ± 2 °C) for 30 min. Thereafter the absorbance was taken against a blank at 517 nm using a spectrophotometer. Scavenging of DPPH free radical with reduction in absorbance of the sample was taken as a measure of their antioxidant activity. IC50 (concentration of the EOs/organic antioxidants that caused 50% neutralisation of DPPH radicals) was calculated from the graph plotted on percentage inhibition and concentration. I% ¼ Ablank –Asample =Ablank 100 where, Ablank is the absorbance of the blank (without EOs/organic antioxidants), and Asample is the absorbance of the test EOs/organic antioxidants. 2.7.2. β-Carotene/linoleic acid bleaching assay The β-carotene/linoleic acid bleaching test was performed by the method described by Ebrahimabadi et al. (2010). In this assay, the 203 antioxidant activity of EOs was determined by measuring the efﬁcacy of EO to inhibit the conjugated diene hydroperoxide formation arising from linoleic acid and β-carotene coupled oxidation in an emulsiﬁed aqueous system. A stock solution of β-carotene and linoleic acid was prepared by dissolving 0.5 mg of β-carotene in 1 ml of chloroform, 25 μl of linoleic acid and 200 μl Tween 40. The chloroform was completely evaporated under vacuum in a rotatory evaporator at 40 °C. One hundred milliliters of distilled water was then added and the resulting mixture was vigorously stirred to form a β-carotene–linoleic acid emulsion. The samples (2 g/l) were dissolved in DMSO and 350 μl of each sample solution was added to 2.5 ml of the above mixture in test tubes. The test tubes were incubated in a water bath at 50 °C for 2 h. As the sample solution was added to β-carotene–linoleic acid emulsion in each tube, the zero time absorbance (A0) was measured at 470 nm using a spectrophotometer. Second absorbance (At) was measured after 2 h incubation period. BHT and BHA were used as a positive control. Antioxidant activity of EOs/synthetic antioxidant was calculated using the following equation: I% ¼ ðAt =A0 Þ 100 where, I%=percent inhibition, (At)=absorbance of β-carotene after 2 h, (A0)=absorbance of β-carotene at the beginning of the experiments. 2.8. Determination of total phenolic content of EOs Total phenolic contents of essential oils were determined spectrophotometrically using the Folin–Ciocalteu reagent according to the method of Gholivand, Rahimi-Nasrabadi, Batooli, and Ebrahimabadi (2010). A solution (0.1 ml) was prepared in a volumetric ﬂask containing the 1000 μg oil mixed with the 46 ml sterilised distilled water and 1 ml Folin–Ciocalteu reagent. The mixture was thoroughly mixed by shaking in an electronic shaker and allowed to react for 3 min. Three milliliters aqueous solution of 2% Na2CO3 was then added to it and left for 2 h incubation period at 25 ± 2 °C. Absorbance of each mixture was measured at 760 nm. The same procedure was also applied to the standard solutions of gallic acid and an equation was obtained by standard curve. Total phenolic contents of the oils were obtained by putting the absorbance value of oil at 760 nm to standard curve and equation expressed as μg gallic acid equivalent/mg of the oil. Absorbance ¼ 0:0012 gallicacidðμgÞ þ 0:024 2.9. Phytotoxicity assay of essential oils The phytotoxicity of the EOs was determined in terms of seed germination and seedling growth of chickpea (Cicer arietinum var. Radha) with respect to the control sets following Kordali et al. (2008). The Radha variety of chickpea purchased from the local market of Varanasi, India was surface-sterilised with sodium hypochlorite (1%) for 5 min to avoid possible microbial contamination and then rinsed in distilled water (three times). The seeds were then soaked in sterile distilled water (1 h). Empty and undeveloped seeds ﬂoating in sterile distilled water were discarded. Two layers of ﬁlter paper moistened with 10 ml of distilled water were placed on the bottom of each Petri plate (9 cm). Thereafter 5 seeds were placed equidistantly on the ﬁlter paper. Ten microliters of each EO was allowed to drip on Whatman No. 1 ﬁlter paper strip placed on the lid using a micropipette. Petri plates were sealed with paraﬁlm to prevent the essential oil vapours from escaping and kept at 23 ± 2 °C in a growth chamber. The length of radicle and plumule was monitored at 24, 48, 72, 96 and 120 h interval. 204 B. Prakash et al. / Food Research International 49 (2012) 201–208 2.10. In vivo efﬁcacy of EOs as fumigants in food system (chickpea) stored in plastic containers In vivo fumigant efﬁcacy of EOs was assessed following Singh et al. (2010b). chickpea seeds var. Radha were procured from the local market of Varanasi and their moisture content was determined following Prakash et al. (2010). The seeds were surface-sterilised with 1% solution of sodium hypochlorite and rinsed 3–5 times with sterilised distilled water under aseptic laboratory conditions. One kilogramme of chickpea seed was kept separately in plastic containers having an aerial volume of 2 l. Three milliliters spore suspension of seven-day-old culture of toxigenic strain of A. ﬂavus (LHP-6) was inoculated in seed sample of each container through uniform spraying. Seed samples in containers were separately fumigated with the EOs at their respective MIC value with respect to aerial volume of container. The control sets were kept parallel to treatment sets without EOs. All the containers were kept airtight and stored for 6 months at 28 ± 2 °C; 70% RH. After 6 months of storage the seed samples of control and treated sets were subjected to mycobiota analysis by the serial dilution method of Aziz, Youssef, El-Fouly, and Moussa (1998). The percent protection of chickpea seed in treatments was calculated based on number of A. ﬂavus isolates in treatment and control sets as follows: Dc−Dt 100: Dc Where; % P = percent protection, Dc = total number of A. ﬂavus isolates from chickpea seed samples of control sets, Dt = total number of A. ﬂavus isolates from chickpea seed samples in EO treatment sets. A 2.11. Statistical analysis All the experiments were performed in triplicate and data analysis was done on mean ± SE subjected to one way ANOVA. Means are separated by Tukey's multiple range test when ANOVA was signiﬁcant (p b 0.05) (SPSS 10.0; Chicago, IL, USA). 3. Results The minimum inhibitory concentration (MIC) and minimum fungicidal concentration (MFC) of plant essential oils and commonly used food preservatives against the toxigenic strain of A. ﬂavus LHP-6 are summarised in Table 1. Among essential oils the lowest MIC against the toxigenic strain of A. ﬂavus was depicted in the case of C. odorata (2.0 μl/ml), C. sativum (2.5 μl/ml) followed by H. spicatum (2.5 μl/ml), C. myrrha and O. majorana (3.0 μl/ml), while in the case of synthetic % inhibition of mycelial dry weight (MDW) %P ¼ preservatives only BHA (0.15 mg/ml) and salicylic acid (3.0 mg/ml) showed pronounced activity in inhibition of growth of toxigenic strain of A. ﬂavus. BHT, ascorbic acid, and gallic acid showed poor antifungal activity up to 10.0 mg/ml. The MFC of EOs and preservatives was found to be C. odorata (5.0 μl/ml), H. spicatum (6.0 μl/ml), O. majorana (7.0 μl/ml), BHA (0.3 mg/ml) and salicylic acid (5.0 mg/ml). During antiaﬂatoxin investigation, a gradual decrease in mycelia dry weight (MDW) and aﬂatoxin B1 production by the toxigenic strain of A. ﬂavus was found on increasing the concentration of EOs (Fig. 1A and B). However, in the case of synthetic preservatives, BHT, gallic acid, and ascorbic acid did not cause complete inhibition of MDW of the toxigenic strain of A. ﬂavus in liquid medium up to 10.0 mg/ml; only BHA and salicylic acid cause complete inhibition of MDW at 0.15 mg/ml and 3.0 mg/ml respectively (Fig. 2A). BHA and salicylic acid cause complete inhibition of aﬂatoxin B1 at 0.10 mg/ml and 3.0 mg/ml respectively; while ascorbic acid and gallic acid at 10.0 mg/ml. However BHT was recorded as a poor aﬂatoxin inhibitory agent causing only 61.68% inhibition at 10 mg/ml (Fig. 2B). The result of fungitoxic spectrum against nine food borne molds depicted that all the essential oils exhibited broad antifungal activity (Table 2). The MIC of all the EOs against the tested fungal species ranges between C. odorata (1.5 to 2.25 μl/ml), C. sativum (2.0 to 3.0 μl/ml), H. spicatum (2.25 to 3.0 μl/ml), C. myrrha (2.5 to 3.5 μl/ml) and O. majorana (2.5 to 3.5 μl/ml). Discoloration of the purple colour of the DPPH radical on thin layer chromatographic (TLC) plate was assessed as a positive sign of antioxidant activity of EOs. The IC50 value and % inhibition of oxidation of linoleic acid are summarised in Table 3. The lowest IC50 value was recorded in the case of essential oil of C. odorata (1.3 μl/ml) while Essential oils C. odorata C. myrrha H. spicatum C. sativum O. majorana Preservatives Salicylic acida BHAa BHTa Ascorbic acida Gallic acida a mg/ml; nf not found. MIC (μl/ml) 2.0 3.0 2.5 2.5 3.0 MFC (μl/ml) 5.00 nf 6.0 nf 7.0 C. myrrha C. odorata H. spicatum C. sativum O. majorana 90 80 70 b bc bc b e d c 20 c c c 30 c d c 40 b b cd 50 aaaaa aa a a b c 60 c d 10 0.0 0.5 1.0 1.5 2.0 2.5 3.0 aaaaa a aaaa 2.5 3.0 3.5 Concentration (µl/ml) C. myrrha C. odorata H. spicatum C. sativum O. majorana 110 100 % inhibition of Aflatoxin B1 Samples a 100 0 B Table 1 Minimum inhibitory concentration (MIC) and minimum fungicidal concentration (MFC) of plant essential oils and preservatives against the toxigenic strain of A. ﬂavus LHP-6. 110 90 80 aa b b b b c c 70 b c cd 60 c d 50 d 40 e 30 e d d e 20 10 3.00 0.15 >10.0 >10.0 >10.0 5.0 0.30 nf nf nf 0 0.0 0.5 1.0 1.5 2.0 3.5 Concentration (µl/ml) Fig. 1. A and B: Effect of essential oils on mycelial biomass and aﬂatoxin B1 production by A. ﬂavus (LHP-6) in SMKY medium. B. Prakash et al. / Food Research International 49 (2012) 201–208 % inhibition of mycelial dry weight (MDW) A Table 3 Antioxidant activity of plant essential oils and preservatives. Salicylic acid BHT Ascorbic acid Gallic acid 120 a 100 a a a a 80 b 60 c b c c b 40 d c a a b b a a de e f d d 4.5 6.0 d DPPH (IC50) (μl/ml) β‐Carotene/linoleic acid inhibition (%) Total phenolic content (μg/mg) C. odorata C. sativum H. spicatum O. majorana C. myrrha BHT BHA Vit. C Salicylic acid 1.30 ± 0.03e 2.90 ± 0.06d 21.67 ± 0.22b 7.2 ± 0.12c 1.90 ± 0.06e 7.40 ± 0.21f* 4.65 ± 0.05g* 3.34 ± 0.03h* 216.0 ± 1.15a† 51.28 ± 0.34c 25.19 ± 0.63e 8.30 ± 0.44g 22.10 ± 0.72f 47.25 ± 0.17d 70.74 ± 0.53b 82.65 ± 0.57a nd nd 10.84 ± 1.66b 4.15 ± 0.85bc 2.90 ± 0.45c 10.00 ± 0.84b 33.33 ± 2.5a nd nd nd nd ⁎(μg/ml); †(mg/ml); nd not determined. The means followed by the same letter in the same column are not signiﬁcantly different according to ANOVA and Tukey's multiple comparison tests. Values are mean (n = 3) ± SE. e e Test samples a a 20 205 f 0 0.0 1.5 3.0 7.5 9.0 10.5 12.0 Concentration (mg/ml) Salicylic acid BHT Ascorbic acid Gallic acid B 110 a % inhibition of aflatoxin B1 100 a a a b 90 a d d 70 d a a b b a aa c c 80 b c a 60 b 50 e 40 30 e e d e c d c 4. Discussion f 20 10 0 0.0 of seeds was recorded following 48 h in all the EO treated sets including control. The EO of C. odorata showed a 100% phytotoxic effect as there was no emergence of radicle or plumule up to 172 h incubation. However, the length of radicles in O. majorana EO treated seeds was higher than the control (Table 4). During in vivo investigations of the essential oils on food system as fumigants in plastic containers, the oils were found to be inhibitory on fungal contamination of chickpea seeds from A. ﬂavus infestation. The order of percent protection of seed sample of chickpea with the A. ﬂavus infestation was C. odorata (77.38%) > H. Spicatum (72.02%) > O. majorana (67.86%) > C. sativum (65.48%) > C. myrrha (55.36%) (Table 5). Moisture content of chickpea seeds was recorded to be 14.86 ± 1.23%. 1.5 3.0 4.5 6.0 7.5 9.0 10.5 12.0 Concentration (mg/ml) Fig. 2. A and B: Effect of organic preservative on mycelial biomass and aﬂatoxin B1 production by A. ﬂavus (LHP-6) in SMKY medium. highest in H. spicatum (21.67 μl/ml) and for preservatives the lowest IC50 value was recorded in the case of ascorbic acid (3.34 μg/ml) and highest in salicylic acid (216 mg/ml). Oxidation of linoleic acid was moderately inhibited by the EO ranges between 8.3 and 51.28% compared to the positive control of BHA (82.65%) and BHT (70.74%). The total phenolic content (TPC) of the essential oils is summarised in Table 3. The lowest phenolic content was recorded for H. spicatum (2.90 μg/mg) while highest in the case of C. myrrha (33.33 μg/mg). During phytotoxic assay, the EOs (except for C. odorata) did not show adverse effect on germination of chickpea seeds. A 100% germination Table 2 Fungitoxic spectrum of plant essential oils against nine food borne molds. Fungal species Aspergillus ﬂavus Aspergillus niger Aspergillus terreus Aspergillus candidus Aspergillus fumigatus Alternaria alternate Cladosporium cladosporioides Fusarium nivale Penicillium italicum Minimum inhibitory concentration (MIC) (μl/ml) C. odorata C. sativum H. spicatum O. majorana C. myrrha 2.0 2.0 2.0 2.0 2.0 2.25 2.25 2.5 3.0 2.5 2.5 2.75 2.5 3.0 2.5 2.5 2.5 2.5 3.0 2.5 3.0 3.0 3.5 3.25 3.0 3.0 3.25 3.5 3.0 3.5 3.0 2.5 3.0 3.0 3.5 1.5 1.5 2.0 2.25 2.25 2.5 2.75 2.5 2.5 3.0 The ﬁndings of the present investigation reveal the EOs of O. majorana, C. sativum, H. spicatum, C. myrrha and C. odorata as suitable candidate for formulation of plant based food additives/ preservatives against biodeterioration of food items from the storage molds and lipid peroxidation as well as their contamination from aﬂatoxin. To the best of our knowledge, so far there has not been any relevant study on the effectiveness of these essential oils against aﬂatoxin inhibitory role and their in vivo practical efﬁcacy on food system. Detailed in vitro investigations were done with essential oils/organic preservatives in order to standardise their fungitoxic parameters prior to in vivo experiments. The essential oils possessed high efﬁcacy as antifungal and aﬂatoxin inhibitors than the prevalent preservatives. The MIC and MFC of essential oils and preservatives were determined against the toxigenic strain of A. ﬂavus LHP-6 by the broth dilution method using PDB broth medium. This method offers better opportunity to essential oils to come in close contact with fungal spores as both of them are homogenously distributed inside the medium as has been earlier suggested by Kalemba and Kunicka (2003). Based on lower MIC value, the EOs were found superior in efﬁcacy than the prevalent organic preservatives as well as many earlier reported essential oils viz. Cicuta virosa (Tian et al., 2011), Nigella sativa (El-Nagerabi, Al-Bahry, Elshaﬁe, & AlHilali, 2012), Curcuma longa L. (Sindhu et al., 2011) and Cinnamomum jensenianum (Tian et al., 2012). Hence, their low doses will be required for inhibition of the fungal infestation of food items. In addition, the broad fungitoxic spectrum of EOs exhibited in the present investigation strengthens their recommendation against a wide number of molds contaminating different food items. A positive correlation between the subsequent decrease in mycelium growth and aﬂatoxin B1 production with increasing concentrations was found in all treatment sets with essential oils. This may be due to the effect of different bioactive volatile components on mycelial development and the perception/transduction of signals involved in the switch from vegetative to reproductive development Table 5 Percent protection of chickpea seed samples after six month of storage fumigated with essential oils. Essential oils R radicle; P plumule; nf not found. The means followed by the same letters in the same row (separately for radicle and plumule) are not signiﬁcantly different according to ANOVA and Tukey's multiple comparison tests. Values are mean (n = 3) ± SE. P nf nf nf nf nf nf nf nf nf nf nf nf 1.33 ± 0.22a 1.4 ±0.02b 2.83 ± 0.20a 0.49 ± 0.03a 2.43 ± 0.22a 3.60 ± 0.25a 4.70 ± 0.09a 5.97 ± 0.64a nf nf 0.37 ± 0.09b 0.5 ± 0.10c 0.60 ± 0.12c 0.13 ± 0.03b 0.33 ± 0.03c 0.93 ± 0.19c 1.40 ± 0.20c 1.47 ± 0.23c nf nf 0.53 ± 0.03b 1.17 ± 0.08b 1.77 ± 0.15b 0.17 ± 0.05b 1.23 ± 0.35b 2.00 ± 0.23b 3.30 ± 0.38b 3.87 ± 0.24b nf nf 0.60 ± 0.06b 0.81 ± 0.05c 1.23 ± 0.22bc nf nf 1.34 ± 0.24a 1.83 ± 0.35a 3.2 ± 0.51a 0.50 ± 0.03a 2.40 ± 0.10a 3.50 ± 0.53a 4.50 ± 0.29a 5.60 ± 0.35a 24 48 72 96 120 R R 0.17 ± 0.03b 0.77 ± 0.23bc 1.81 ± 0.15b 2.43 ± 0.28b 3.0 ± 0.29b O. majorana C. myrrha H. spicatum C. sativum P Negative control Mean length of seedlings (cm) Duration of exposure (h) Table 4 Effect of essential oils on seed germination and seedling growth of chickpea seeds. P R P R P R P C. odorata B. Prakash et al. / Food Research International 49 (2012) 201–208 R 206 C. odorata C. sativum H. spicatum O. majorana C. myrrha a Concentration (μl/ml) 2 2.5 2.5 3 3 No. of A. ﬂavus isolates Controla Treatment 168 168 168 168 168 38 58 47 54 75 % protection 77.38 65.48 72.02 67.86 55.36 Control for all EO treated sets. as has been reported by Tzortzakis and Economakis (2007). The lowmolecular weight and highly lipophilic compounds of EOs pass easily through cell membranes and disrupt the fungal cell organisation (Chao et al., 2005; Shukla, Singh, Prakash, & Dubey, 2012). The aﬂatoxin inhibitory concentration of the EOs was found to be lower than the inhibitory concentration required for mycelial dry weight supporting the earlier ﬁndings of some workers (Shukla et al., 2009; Tian et al., 2011; Shukla et al., 2012; Tian et al., 2012). This suggests a different mode of EOs against fungal growth and toxin secretion as has been reported by Prakash et al. (2010) and Shukla et al. (2012). Hence, the oils act with two different modes of action against fungal growth and aﬂatoxin secretion. This fact should be kept in mind during their recommendation for their practical application as food preservatives. The antioxidative properties of EOs in the present investigation were measured by two commonly used methods. The low IC50 values of the essential oils strengthen their application as natural antioxidant food preservatives to reduce oxidative burden of food items. Although, as antioxidant, the EOs were not as efﬁcacious as some of the organic preservatives viz. BHT, BHA and ascorbic acid but the toxic and abnormal effects of synthetic antioxidants including their carcinogenic potential restrict their application in food items (Adamez, Samino, Sanchez, & Gonzalez-Gomez, 2012; Singh et al., 2012). The poor radical scavenging and inhibition of linoleic acid activity by the H. spicatum in comparison to other tested essential oils in the present investigation may be due to its low phenolic content in comparison to other oils supporting the earlier view that the phenolic compounds play major roles in antioxidant activity (Ebrahimabadi et al., 2010; Prakash et al., 2011). In addition, aﬂatoxin inhibitory effect of phenolic compounds through mediation of oxidative stress levels in the fungi is also reported by Kim, Mahoney, Chan, Molyneux, and Campbell (2006). Kim et al. (2006) have reported that growth inhibition of A. ﬂavus by the phenolics salicylic acid, thymol, vanillyl acetone, vanillin and cinnamic acid is via targeting the mitochondrial oxidative stress defense system. Since the mitochondria are responsible for providing acetyl-CoA, a main precursor for AF biosynthesis, disruption of mitochondrial respiration chain may account in part for the inhibitory effects of antifungal phenolics on aﬂatoxin production. Hence, these plant essential oils are used as markers for the elucidation of antioxidant-based inhibition of aﬂatoxin biosynthesis. However, the high antioxidant activity of these essential oils than the earlier reported plant essential oils viz., Semenovia tragioides (Bamoniri et al., 2010); Lavandula ofﬁcinalis, Petroselinum crispum, Foeniculum vulgare (Viuda-Martos et al., 2011); Rosmarinus ofﬁcinalis L. var. typicus (Zaouali, Bouzaine, & Boussaid, 2010); Cistus ladanifer, Citrus latifolia, Cupressus lusitanica, Eucalyptus gunnii (Guimaraes, Sousa, & Ferreira, 2010), Rhizoma Homalomenae (Zeng, Zhang, Luo, & Zhu, 2011) and Artemisia annua (Cavar, Maksimovic, Vidic, & Paric, 2012) strengthens their application for possible use as antioxidants. During phytotoxic assay the EOs (except C. odorata) did not exhibit any adverse effect on germination of chickpea seeds showing their non‐phytotoxic nature for treatment of food commodities stored even for sowing purpose. The C. odorata oil may be only recommended B. Prakash et al. / Food Research International 49 (2012) 201–208 for food commodities stored for consumption purpose. However, the safety proﬁle of the oils should be recorded on animal system, although the plants from which the oils have been isolated are well known medicinal plants in the Indian system of medicine (Prajapati et al., 2003). The oils also exhibited pronounced efﬁcacy in food system providing up to > 75% protection of fumigated chickpea seeds from A. ﬂavus infestation. Hence, it may be concluded that the essential oils require higher concentrations in food system than their in vitro concentration as has been also reported by Farbood, MacNeil, and Ostovar (1976) and Tian et al. (2011). The nature of food system, fungal inoculum density, storage conditions and their moisture content should also be considered during prescribing the in vivo concentration of the oils against food infestation by molds. The most signiﬁcant ﬁnding of the present investigation was the efﬁcacy of essential oils as growth inhibitory of food infesting molds and aﬂatoxin secretion as well as their free radical scavenging activity. The prevalent organic preservatives could show only antioxidant activity being very poor as fungal growth inhibitor and aﬂatoxin suppressor. Moreover, there are also reports on enhancement of aﬂatoxin secretion by many synthetic preservatives (Badii & Moss, 1988). Plant essential oils are expected to be more advantageous over synthetic preservatives because of their biodegradable nature. The essential oils are a mixture of different major and minor components which act together for their biological activities. Because of this there would be less chance of development of resistant races of fungal strains as has been reported in the case of many synthetic preservatives (Ishii, 2006). Currently many essential oil formulations such as Sporan-TM (Rosemary oil), Promox-TM (Thyme), and DMC base natural (rosemary, sage, citrus oil combination) (Shukla et al., 2009) are in the market as food preservatives. The recent encapsulation technology is employed to retain their aroma in food systems with controlled release of vapours (Donsi, Annunziata, Sessa, & Ferrari, 2011). The abundance of raw materials because of luxuriant growth of the plants and their renewable nature makes use of essential oils economical for practical application. The ﬁndings of the present investigation warrant future research for large scale application of the essential oils as fumigants in food systems for enhancement of their shelf life by controlling their losses from fungal infestation and lipid peroxidation. 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