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
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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.
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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.
In conclusion, in view of their potential as inhibitory of fungal
growth and aflatoxin secretion and efficacy as fumigant in food system
in controlling fungal infestation as well as their free radical scavenging
activity, the essential oils may be recommended for formulation of
plant based preservatives for enhancement of shelf life of food items
during post harvest processing.
Acknowledgement
Bhanu Prakash is thankful to the Council of Scientific and Industrial
Research (CSIR), New Delhi, India for financial assistance as senior
research fellow (SRF).
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