Food Chemistry 130 (2012) 520–527
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The control of Aspergillus ﬂavus with Cinnamomum jensenianum Hand.-Mazz essential oil and its potential use as a food preservative Jun Tian, Bo Huang, Xiuli Luo, Hong Zeng, Xiaoquan Ban, Jingsheng He, Youwei Wang ⇑ Key Laboratory of Combinatorial Biosynthesis and Drug Discovery (Wuhan University), Ministry of Education, and Institute of Traditional Chinese Medicine & Natural Products, Wuhan University School of Pharmaceutical Sciences, Wuhan 430071, PR China
a r t i c l e
i n f o
Article history: Received 9 February 2011 Received in revised form 16 May 2011 Accepted 18 July 2011 Available online 22 July 2011 Keywords: Essential oil Cinnamomum jensenianum Hand.-Mazz Chemical composition Antifungal activity Aﬂatoxin Ultrastructure Ergosterol
a b s t r a c t The essential oil extracted from the bark of Cinnamomum jensenianum Hand.-Mazz was tested for antifungal activity against Aspergillus ﬂavus. Fifty-ﬁve components accounting for 96.66% of the total oil composition were identiﬁed by GC–MS. The major components were eucalyptol (17.26%) and a-terpineol (12.52%). Mycelial growth and spore germination was inhibited by the oil in a dose-dependent manner. The oil also exhibited a noticeable inhibition on the dry mycelium weight and the synthesis of aﬂatoxin B1 (AFB1) by A. ﬂavus, completely restraining AFB1 production at 6 ll/ml. The possible mode of action of the oil against A. ﬂavus is discussed based on changes in the mycelial ultrastructure. To conﬁrm the target of the oil in the plasma membrane, studies on the inhibition of ergosterol synthesis were performed. Results show that the oil caused a considerable reduction in the ergosterol quantity. Thus, the essential oil from C. jensenianum can be used as a potential source for food preservative. Ó 2011 Elsevier Ltd. All rights reserved.
1. Introduction Fungal contamination is a serious problem during storage in food industries. Food is susceptible to be affected by various microorganisms such as Aspergillus species resulting in severe economic losses. Furthermore, some species represent a very serious risk for consumers because of their production of dangerous secondary metabolites. Fungi, especially Aspergillus ﬂavus, are often mainly responsible for the spoilage of many foods and production of aﬂatoxins, a group of extremely hazardous and common secondary metabolites. Aﬂatoxin has carcinogenic, teratogenic, hepatotoxic, mutagenic, and immunosuppressive properties and can inhibit several metabolic systems (Joseph, Jayaprakasha, Selvi, Jena, & Sakariah, 2005). About 4.5 billion people are affected by uncontrolled amounts of aﬂatoxin in developing countries and aﬂatoxicosis is ranked 6th among the 10 most important health risks identiﬁed by Williams et al. (2004). In this context, investigators are looking for new sources of materials to control spoilage caused by fungi in food. However, the application of synthetic preservatives has led to a number of environmental and health problems because they are themselves carcinogenic, teratogenic, and highly toxic with long degradation periods (Lingk, 1991). Accordingly, the public demands more ⇑ Corresponding author. Tel.: +86 27 68759323; fax: +86 27 68759010. E-mail address: [email protected]
(Y. Wang). 0308-8146/$ - see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodchem.2011.07.061
acceptable preservatives that are biodegradable and safe to humans and the environment. Different plant products have been recognized and used for microbial inactivation in food industries because of their antifungal and antitoxigenic activities (Murthy, Ramalakshmi, & Srinivas, 2009). Among the different groups of plant products, essential oils are especially considered as one of the most promising groups of natural products for the formulation of safer antifungal agents (Varma & Dubey, 2001). The majority of the essential oils are classiﬁed as Generally Recognized As Safe (GRAS) and are have low risk for developing resistance to pathogenic microorganisms (Cardile et al., 2009). Cinnamomum jensenianum Hand.-Mazz, an endemic plant in China, belongs to the Lauraceae family. It is mainly grown in the provinces of Hubei, Hunan, Guizhou, Sichuan, and Jiangxi of China. It is traditionally used in the treatment of gastritis, abdominal pain, anemofrigid-damp arthralgia, and traumatic injury (Editorial Committee of National Chinese Medical Manage Bureau, 1999). However, to our knowledge and according to literature survey, there are no available reports on the chemical composition, antifungal and anti-aﬂatoxin properties of the essential oil from C. jensenianum (CJEO). In this study, the chemical composition of the essential oil extracted from the bark of C. jensenianum was investigated and evaluated for its effects on the mycelial growth, spore germination, mycelium weight, and AFB1 content in A. ﬂavus. In addition, the possible mode of action of CJEO and its effect on the morphological
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and physiological changes of A. ﬂavus were explored using transmission electron microscopy (TEM). The effect of CJEO on ergosterol synthesis was also assessed. 2. Materials and methods 2.1. Plant material The barks of C. jensenianum plants were harvested from the Wuhan Botanical Garden, in the Hubei Province, in China, in March 2010. The identiﬁcation of the plant material was initially done based on its morphological features and was ﬁnally conﬁrmed by Prof. Youwei Wang of the College of Pharmacy in Wuhan University. A voucher specimen number (No. 626) has been deposited in the herbarium of the Institute of the Traditional Chinese Medicine and Nature Products, College of Pharmacy, Wuhan University. 2.2. Isolation of the essential oil A total of 200 g of the powdered barks (800 mesh size) were hydrodistiled for approximately 5 h using a Clevenger-type apparatus (SS85-1000, Shenshi Chemical Engineering Co., Ltd., Wuhan, China). The yield of the essential oil was 0.35% (v/w). The oil was dried over anhydrous sodium sulphate. After ﬁltration, the oil was stored in air-tight sealed glass vials covered with aluminium foil at approximately 4 °C for further testing and chemical analysis. 2.3. Gas chromatography–mass spectrometry (GC–MS) analysis The chemical composition of the essential oil was analyzed using GC–MS. The essential oil (10 ll) was dissolved in acetone (50 ll), and 1 ll of the solution was injected into a GC–MS (QP-2010, Shimadzu Co., Kyoto, Japan). The capillary column was Rtx-5MS (length = 30 m, i.d. = 0.25 mm, thickness = 0.25 lm). Helium was used as the carrier gas at a ﬂow rate of 1.00 ml/min. The column inlet pressure was 49.7 kPa. The GC column oven temperature was increased from 40 to 280 °C at a rate of 10 °C /min with a ﬁnal hold time of 6 min. The injector and detector temperatures were maintained at 280 °C. The EI mode was at 70 eV, while the mass spectra were recorded in the 45–400 amu range with the ion source-temperature set to 200 °C. The relative percentage of the oil constituents was expressed as percentages, obtained by peak area normalization. The identiﬁcation of the compounds was performed by comparing their retention indices based on a homologous series of n-alkane indices on the Rtx-5MS capillary column, referring to known compounds from the literature (Adams, 2001) and also by comparing their mass spectra with those stored in the spectrometer database using NIST05.LIB and NIST05s.LIB (National Institute of Standards and Technology) and whenever possible, by co-injection with authentic compounds. 2.4. Fungal strains used Aspergillus ﬂavus CCAM 080001 was obtained from the Culture Collection of State Key Laboratory of Agricultural Microbiology (CCAM), China. The fungal strain cultures were maintained on a potato dextrose agar (PDA) slant at 4 °C. The old cultures were transferred to fresh slant every two months in order to avoid a decline in strain viability.
temperature of 45–50 °C to procure the required concentrations of 1, 2, 4, 6, and 8 ll/ml. The control plates (without essential oil) were inoculated following the same procedure. A fungal disc (9 mm in diameter) of mycelial material, cut from the periphery of a ﬁve-day-old culture using a cork borer, was inoculated aseptically into the center of each petri dish. The plates were sealed with polyethylene ﬁlm and incubated at a temperature of 28 ± 2 °C (Murthy et al., 2009). The efﬁcacy of the treatment was evaluated each day for 9 days by measuring the average of two perpendicular diameters of each colony. All tests were performed in triplicate. The percentage inhibition of the radial growth of the tested strain by the oils, compared with the control, was calculated at day 9, using the following formula (Albuquerque et al., 2006):
Percentage mycelial inhibition ¼ ½ðdc dtÞ=dc 100 where dc (cm) is the mean colony diameter for the control sets and dt (cm) is the mean colony diameter for the treatment sets. The lowest concentration that completely inhibited the growth of the fungus was considered the minimum inhibitory concentration (MIC). 2.6. Spore germination assay Fungal spore germination and growth kinetics were tested using the slightly modiﬁed method of Bajpai, Shukla, and Kang (2008). CJEO was added to the glass tube containing 1 ml 0.1 (v/v) Tween-20 to obtain ﬁnal concentrations of 1, 2, 4, 6 and 8 ll/ml. A spore suspensions of A. ﬂavus was obtained from its 3 days old cultures, harvested by adding 5 ml sterile water containing 0.1% (v/v) Tween-20 to each petri dish, and gently scraping the mycelial surface three times with a sterile L-shaped spreader to free spores. The homogenous spore suspension of A. ﬂavus containing 107 spores/ml was then inoculated into each of the above tubes. From this, 10 ll aliquots of the spore suspension were incubated into fresh PDA medium in separate depression slides. Depression slides containing the spores were assembled with the cover slip and then incubated in a moisture chamber at 28 °C for 20 h in three replicates. For each treatment, 200 spores were examined and the extent of spore germination was assessed by looking for the emergence of germ tubes. The number of germinated spores was scored and reported as a percentage of spore germination. A. ﬂavus was also tested by a kinetic study to assess its antifungal activity. A spore suspension (10 ll) of A. ﬂavus containing 108 spores/ml prepared in 0.1% (v/v) Tween-20 was inoculated to different concentrations (2, 4, and 8 ll/ml) of 5 ml CJEO solution, and a homogenous suspension (about 2 105 spore/ml) was mixed vigorously by vortex (TS-1, Kylin-Bell Lab Instruments Co., Ltd., Shanghai, China) for 1 min. Samples without any oil treatment were considered as controls. After speciﬁc time intervals, i.e., 30, 60, 90, 120, 150, and 180 min, the reaction mixtures were collected and centrifuged at 6000g (TGL-16C, Anting Scientiﬁc Instrument Factory, Shanghai, China) for 5 min at room temperature. The supernatant was discarded and the remainder was resuspended in 10 ml sterilized distiled water. From this, 10 ll aliquots of the spore suspension were also taken to the depression slides, which were handled as described above. About 200 spores were examined and the percentage of spore germination was calculated.
2.5. Antifungal activity assay
2.7. A. ﬂavus growth and analysis of aﬂatoxin B1
The antifungal activity of the essential oil was tested against A. ﬂavus following our previously published method (Tian et al., 2011). Aliquots of the essential oil dissolved separately in 0.5 ml of 5% (v/v) Tween-20, were pipetted aseptically onto glass petri dishes (9 1.5 cm) containing 9.5 ml PDA medium at a
The anti-aﬂatoxigenic efﬁcacy of CJEO was studied on A. ﬂavus following Kumar, Shukla, Singh, Prasad, and Dubey (2008). A spore suspension (100 ll) of A. ﬂavus containing 107 spores/ml, prepared in 0.1% (v/v) Tween-20, was added to 20 ml potato dextrose broth (PDB) medium in an Erlenmeyer ﬂask. The required amounts of
J. Tian et al. / Food Chemistry 130 (2012) 520–527
CJEO dissolved in 5% (v/v) Tween-20 were transferred to the PDB medium to obtain 1, 2, 4, 6, and 8 ll/ml concentrations. The control sets contained the medium without oil. The ﬂasks were incubated at 28 ± 2 °C for 10 days. Three replicates of each treatment were performed, and the experiment was repeated three times. After incubation, the mycelia produced in liquid cultures were ﬁltered and washed through ﬁlter paper (DX102, Xinhua Paper Co., Ltd., Hangzhou, China). The weight of the dry mycelia for each mycelium was determined after drying at 60 °C for 24 h. AFB1 in the ﬁltrate was extracted twice with 25 ml chloroform in a separating funnel. The chloroform extracts were combined, evaporated to dryness, and the residue was redissolved in chloroform up to 1 ml in a volumetric ﬂask. Silica gel-G thin layer plate was used for the AFB1 analysis. Fifty microlitres of each sample spotted onto the TLC sheets were developed in the solvent system comprising of toluene: isoamyl alcohol: methanol (90:32:2 v/v/v) (Reddy, Viswanathan, & Venkitasubramanian, 1970). The identity of AFB1 was detected under UV lamp at 365 nm and conﬁrmed chemically by spraying triﬂuoroacetic acid (Bankole & Joda, 2004). For the quantiﬁcation of AFB1, amethyst-ﬂuorescent spots of AFB1 on the TLC were scraped out, dissolved in 5 ml cold methanol, and centrifuged at 2000g (TGL-16C, Anting Scientiﬁc Instrument Factory, Shanghai, China) for 5 min. The absorbance of the supernatant was made using a UV–Visible spectrophotometer (UV-1240, Shimadzu, Tokyo, Japan) at a wavelength of 360 nm. The amount of AFB1 present in the sample was calculated according to the formula by Sinha, Sinha, and Prasad (1993):
AFB1 content ðlg=mlÞ ¼ ðD MÞ=ðE lÞ 1000
amount of 100 ll containing 107 spores/ml (the spore population was counted using a hemocytometer) of A. ﬂavus spore suspension was inoculated in PDB medium containing 0, 0.25, 0.5, 1.0, and 2.0 ll/ml of CJEO for 4 days at 28 ± 2 °C. After incubation, mycelia was harvested and washed twice with distiled water. The net wet weight of the cell pellet was determined. Five millilitres of 25% alcoholic potassium hydroxide solution was added to each sample and vortex mixed for 2 min (TS-1, Kylin-Bell Lab Instruments Co., Ltd., Shanghai, China), followed by incubation at 85 °C for 4 h. Sterols were extracted from each sample by adding a mixture of 2 ml sterile distiled water and 5 ml n-heptane. Then, the mixture was sufﬁciently mixed by vortex (TS-1, Kylin-Bell Lab Instruments Co.,Ltd., Shanghai, China) for 2 min allowing the layers to separate for 1 h at room temperature. The n-heptane layer was analyzed by scanned spectrophotometry (UV-1700, Shimadzu, Tokyo, Japan) between 230 and 300 nm. The presence of ergosterol (at 282 nm) and the late sterol intermediate 24(28) dehydroergosterol (at 230 and 282 nm) in the n-heptane layer led to a characteristic curve. The ergosterol amount was calculated as a percentage of the wet weight of the cells and was based on the absorbance and wet weight of the initial pellet. The calculated formula of the ergosterol amount is as follows:
% ergosterol þ %24ð28Þ dehydroergosterol ¼ ðA282=290Þ=pellet weight;%24ð28Þ dehydroergosterol ¼ ðA230=518Þ=pellet weight; and % ergosterol ¼ ð% ergosterol þ %24ð28Þ dehydroergosterolÞ %24ð28Þ dehydroergosterol
where D is the absorbance, M is the molecular weight of aﬂatoxin (312 g/mol), E is the molar extinction coefﬁcient (21, 800 l/ (mol cm)), and l is the path length (1 cm cell was used). In addition, AFB1 inhibition was calculated as follows:
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).
Inhibition ð%Þ ¼ ð1 X=YÞ 100; where X (lg/ml) is the mean concentration of AFB1 in the treatment and Y (lg/ml) is the mean concentration of AFB1 in the control. 2.8. TEM observations Five-day-old fungal materials of A. ﬂavus on PDA exposed to 2 and 4 ll/ml of CJEO and control without oil were used for TEM observations to study the mode of action of essential oil (Nogueira et al., 2010). The small segments measuring 5 5 mm were excised at the margin of the colony from the cultures growing on the PDA plates. Then, the segments were promptly placed in vials containing 2.5% glutaraldehyde in 0.1 M phosphate buffer saline (PBS) (pH 7.2) at 4 °C and ﬁxed overnight. The ﬁxed samples were rinsed with the same buffer three times for 10 min each. Afterwards, the samples were dehydrated in a graded series of ethanol (70%, 80%, 90%, 95%, and 100%) for a period of 20 min in each alcohol dilution. The last step was performed for 30 min three times. The dehydrated specimens were then embedded and polymerized in Spurr’s resin at 65 °C for 72 h. Ultrathin sections (approximately 50 nm in thickness) were hand trimmed with a diamond knife using an LKB-V Ultratome for TEM observations (Tecnai G2 20 S-Twin, FEI Company, Hillsboro, USA). 2.9. Measurement of the ergosterol content in the plasma membrane of A. ﬂavus The ergosterol content in the plasma membrane of A. ﬂavus was detected by a previously described method with slight modiﬁcations (Arthington-Skaggs, Jradi, Desai, & Morrison, 1999). An
2.10. Statistical analysis All data are reported as means ± standard deviations. The significant differences between mean values were determined by Duncan’s Multiple Range test (p < 0.01), following one-way ANOVA. The statistical analysis was performed using a statistical software (SPSS, 13.0; Chicago, USA).
3. Results 3.1. Chemical composition of essential oil A total of 55 different components of the essential oil, accounting for 96.66% of the total oil composition, were identiﬁed by GC–MS analyses. The identiﬁed chemical composition, retention time, and percentage composition are given in Table 1. The oil mainly contained a complex mixture of oxygenated monoterpenes (46.63%) and sesquiterpene hydrocarbons (36.13%). The most abundant components of the essential oil were eucalyptol (17.26%) and a-terpineol (12.52%). Several other components such as ()-terpinen-4-ol (7.60%), d-cadinene (6.59%), a-copaene (4.50%), b-elemene (4.27%), caryophyllene (4.25%), a-muurolene (3.38%), bornyl acetate (3.28%), a-cadinol (2.85%), borneol (2.78%), T-muurolol (2.35%), a-guaiene (2.13%), and a-gurjunene (2.00%) were in less amounts. However, monoterpene hydrocarbons, oxygenated sesquiterpenes, aldehydes, and others were also found as trace or minor components.
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Table 1 Chemical composition of the essential oil isolated by hydrodistillation from Cinnamomum jensenianum Hand.-Mazz.
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55
842 924 931 939 951 976 981 991 1005 1014 1026 1035 1040 1046 1062 1073 1089 1098 1114 1133 1142 1148 1165 1177 1190 1265 1285 1339 1350 1355 1361 1367 1378 1392 1401 1406 1411 1418 1433 1451 1477 1484 1500 1508 1522 1542 1564 1571 1579 1580 1630 1646 1654 1689 1697
Diacetone alcohol Tricyclene a-Thujene a-Pinene Camphene Sabinene b-Pinene b-Myrcene a-Phellandrene a-Terpinene p-Cymene Eucalyptol cis-b-Ocimene cis-b-Terpineol c-Terpinene trans-Linalool oxide Terpinolene Linalool Fenchyl alcohol Isopulegol ()-Camphor Camphene hydrate Borneol ()-Terpinen-4-ol a-Terpineol trans-Citral Bornyl acetate d-elemene a-Terpinyl acetate a-Cubebene Acetyleugenol Ylangene a-Copaene b-elemene Lauraldehyde a-Gurjunene a-Cedrene Caryophyllene a-Guaiene a-Humulene c-Muurolene a-Amorphene a-Muurolene b-Bisabolene d-Cadinene a-Calacorene trans-Nerolidol Caryophyllenyl alcohol Caryophyllene oxide n-Myristaldehyde a-Muurolol T-Muurolol a-Cadinol Longifolene-(V4) trans-Farnesol
0.22 0.01 0.08 1.78 0.82 0.07 0.80 0.25 0.06 0.38 1.28 17.26 0.03 0.02 0.60 0.02 0.26 1.44 0.07 0.11 0.04 0.10 2.78 7.60 12.52 0.06 3.28 0.29 0.08 0.50 0.23 0.25 4.50 4.27 0.21 2.00 1.39 4.25 2.13 1.10 1.44 0.62 3.38 0.48 6.59 1.07 0.51 0.33 0.50 0.33 0.35 2.35 2.85 1.87 0.66
Retention indices relative to a series of n-alkanes on Rtx-5 capillary column. The relative proportions of the essential oil constituents.
3.2. Antifungal activity assay The growth of A. ﬂavus during the nine days is shown in Fig. 1. Results indicate that mycelia growth was considerably reduced with the increasing concentration of CJEO but increased with incubation time. The mycelial growth was delayed by three days for A. ﬂavus at 6 ll/ml concentration and an MIC of 8 ll/ml was obtained after nine days of incubation. The mycelial growth inhibition percentage was determined at day 9. The oil produced a signiﬁcant reduction in mycelium growth with the four fungi species at 1, 2, 4, and 6 ll/ml concentrations with reduction percentages of 27.6%, 59.7%, 81.4%, and 87.0%, respectively.
Fig. 1. Effects of the different concentrations of CJEO on the colony diameter (cm) growth of Aspergillus ﬂavus in PDA. The plates were incubated at a temperature of 28 ± 2 °C for 9 days. Values are means (n = 3) ± standard deviations.
Fig. 2. Effect of CJEO on spore germination of A. ﬂavus. (a) Effect of different concentrations of CJEO on spore germination of the tested fungi. (b) Growth kinetics of the inhibition of the tested fungi by CJEO.
3.3. The effect of the essential oil on spore germination The efﬁcacies of CJEO on the spore germination of A. ﬂavus are shown in Fig. 2. Fig. 2a indicates that the percentage of spore germination was signiﬁcantly (p < 0.01) inhibited by the different concentrations of essential oil. All A. ﬂavus spores germinated after 20 h of incubation at 28 °C in PDB without the essential oil. However, CJEO completely inhibited the germination of spores at 8 ll/ml. Observations show an inhibitory effect on the spore germination of A. ﬂavus within the range of 12.0–82.3% at concentrations ranging from 1 to 6 ll/ml. The inhibition growth kinetics of A. ﬂavus by the essential oil is shown in Fig. 2b. Exposure of the A. ﬂavus spores to different concentrations of the essential oil for a period of 0–180 min caused varying degrees of spore germination inhibition. Results indicate that spore germination was reduced with increasing exposure time and CJEO concentration. The essential oil at 2 and 4 ll/ml exhibited antifungal activity but not rapid killing, and about 40–50% inhibition was observed at an exposure time of 90 min. However, there was a visible increase in the killing rate at 8 ll/ml after 30 min
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Table 2 Efﬁcacy of the different concentrations of CJEO on dry mycelium weight and aﬂatoxin B1 synthesis by A. ﬂavus. Oil concentration (ll/ml)
Inhibition of AFB1 synthesis (%)
Control 1 2 4 6 8
404.0 ± 6.5a 307.4 ± 4.9b 229.1 ± 5.5c 147.6 ± 5.5d 65.3 ± 3.0e 0.0 ± 0.0f
387.8 ± 6.9a 265.0 ± 12.0b 181.6 ± 13.9c 58.9 ± 12.0d 0.0 ± 0.0e 0.0 ± 0.0e
0.0 31.6 53.2 84.8 100.0 100.0
DMW, dry mycelium weight (mg); AFB1, aﬂatoxin B1 content (lg/ml). Values are mean (n = 3) ± standard deviations. Values followed by the same letter in each column are not signiﬁcantly different in ANOVA and Duncan Multiple Range Test (p < 0.01).
of exposure, and complete inhibition of spore germination was observed at 180 min of exposure. 3.4. Efﬁcacy of the essential oil on dry mycelium weight and aﬂatoxin B1 content The efﬁcacy of the essential oil on A. ﬂavus aﬂatoxin production and dry mycelium weight in PDB medium is shown in Table 2. The ﬁve different concentrations of essential oil efﬁciently caused different degrees of inhibition in terms of dry mycelium weight and AFB1 synthesis (p < 0.01). No mycelium was recorded and the oil completely inhibited mycelial production at 8 ll/ml. However, mycelial growth was observed at 6 ll/ml although AFB1 production was completely inhibited. The AFB1 content was reduced to about half as compared with that of the control at 2 ll/ml. 3.5. TEM observations To investigate the mode of CJEO against A. ﬂavus, changes in the ultrastructure of A. ﬂavus observed by TEM are presented in Fig. 3. In the control hyphae (Fig. 3A and B), the cell wall was uniform and thoroughly surrounded by an intact ﬁbrillar layer. The plasma membrane was uniform with a smooth surface. The cytoplasmic matrix was abundant. The main organelles, such as mitochondria, vacuoles, nucleus, and electron dense granules, have normal and uniform structures. By contrast, in the CJEO treated hyphae (Fig. 3C–F), ultrastructural alterations were conspicuous in the plasma membrane, ﬁbrillar layer, and cytoplasm. At 2 ll/ml of CJEO, the plasmalemma became rough with continuous folding into the cytoplasm and festooned with small lomasomes. A decreased cytoplasmic matrix was also observed (Fig. 3C). Some mitochondria suffered an extensive disruption of the internal structure with a decrease in the mitochondrial cristae (Fig. 3D). The cell ultrastructure damage was aggravated in the presence of 4 ll/ml concentration of CJEO (Fig. 3E and F). The major alterations were observed by some signs including massive vacuolation of cytoplasm with vacuole fusion, appearance of numerous lomasomes with folding, and detachment of plasma membrane from the cell wall (Fig. 3E). As shown in Fig. 3F, the ﬁbrillar layers have gradually lost their integrity, becoming thinner, and eventually failing to deposit on the cell wall. The plasma membrane was also folded at many sites. The cytoplasmic matrix and some cytoplasmic organelles, such as the electron dense granules, were absent. In addition, the mitochondria suffered a severe disruption of the internal structure with complete lysis. 3.6. Measurement of plasma membrane ergosterol content The efﬁcacies of CJEO on the ergosterol content in the plasma membrane of A. ﬂavus are shown in Fig. 4. The total ergosterol con-
tent was determined at 0, 0.25, 0.5, 1.0, and 2.0 ll/ml concentrations of CJEO with a value of 0.415 ± 0.004%, 0.179 ± 0.002%, 0.126 ± 0.002%, 0.061 ± 0.000%, and 0.012 ± 0.000%, respectively (data not presented). The results demonstrate that the ergosterol content (at 282 nm) in the plasma membrane of A. ﬂavus was signiﬁcantly inhibited by the different concentrations of essential oil. A dose-dependent decrease in ergosterol production was observed when isolates were grown in the presence of CJEO. After incubation of A. ﬂavus at 0.25, 0.5, and 1.0 ll/ml concentrations of CJEO, a reduction percentage of the ergosterol content in the plasma membrane as compared with the control was observed at 56.9% for 0.25 ll/ml, 69.8% for 0.5 ll/ml, and 85.2% for 1.0 ll/ml. The A. ﬂavus cells growing in the presence of 2.0 ll/ml concentrations of the CJEO showed the highest inhibition to ergosterol with a value of 97.2% compared with the control.
4. Discussion In recent years, consumer demand for effective and safe natural products to control food spoilage without chemical residues has increased. Essential oils, aromatic volatile products of the secondary metabolism of plants, have formed the basis of many applications in food ﬂavouring and preservation industries (Rahman & Kang, 2009). C. jensenianum is a common green and economic tree in China thus, it is an advantage to develop it into a food preservative due to its abundance. In general, most chemical components of essential oils are terpenoids, including monoterpenes, sesquiterpenes, and their oxygenated derivatives. Terpenes are the active antimicrobial compounds of essential oils. The action mechanism of this class of compounds is not fully understood, but it is speculated to involve membrane disruption by these lipophilic compounds (Cowan, 1999). These low molecular weight, highly lipophilic compounds of essential oils easily pass through cell membranes to induce biological responses (Chao et al., 2005). CJEO exhibits antifungal activities, which may be attributed to the presence of eucalyptol and a-terpineol. Eucalyptol and a-terpineol have been shown to possess antifungal properties to control food contamination (Adegoke, Iwahashi, Komatsu, Obuchi, & Iwahashi, 2000; Sokovic, Brkic, Dzamic, Ristic, & Marin, 2009; Sokovic, Tzakou, Pitarokili, & Couladis, 2002). However, whole essential oil has greater antifungal activity, which may be attributed to some minor components that has a synergistic effect with the major components. In this study, CJEO showed a pronounced antifungal efﬁcacy against the tested fungi. The mycelium growth was recorded to change with increasing concentrations of the oil and incubation time. The colony diameter expanded along with the increase in the number of incubation days. The essential oil also showed a remarkable effect in restraining the germination of A. ﬂavus. As the concentration of the essential oil increased, a pronounced reduction in the percentage of spore germination was observed on A. ﬂavus. Some studies focused on the effects of the compounds on fungal spore germination. Yenjit, Issarakraisila, Intana, and Chantrapromma (2010) found that fernenol, arundoin, and the mixture of stigmasterol and b-sitosterol greatly inhibit spore germination and germ tube elongation in Colletotrichum gloeosporioides with EC50 values of 45.8, 62.3, and 86.9 mg/l. The kinetic study of A. ﬂavus exhibited that the exposure time of the essential oil had a little effect on the antifungal activity at the concentration of 2 ll/ml. However, the spore germination was greatly inhibited at the concentration of 6 and 8 ll/ml. Similar types of results were also reported by Bajpai et al. (2008). A. ﬂavus can produce AFB1, the most dangerous toxic metabolite among all classes of aﬂatoxin. Thus, an examination on the content of aﬂatoxin B1 was performed in the present study. Our results ex-
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Fig. 3. Transmission electron microscopy illustrates the effect of CJEO on the ultrastructural aspects of A. ﬂavus. (A and B) Control mycelia are homogenous and their cells contain abundant cytoplasmic matrix. The cell wall (CW), plasma membrane (PM), ﬁbrillar layer (FL), and intracellular organelles including the mitochondria (M), vesicles (V), electron dense granules (EDG), and nucleus (N) have usual and uniform structures. (C) Longitudinal section of A. ﬂavus hypha exposed to 2 ll/ml of essential oil. Plasma membrane (PM) appears to be the primary target of essential oil. It becomes irregular with the continuous folding into the cytoplasm. The change of the plasma membrane (PM) led to the reduction of cytoplasmic matrix and formation of small lomasomes (L). (D) Cross section of A. ﬂavus hypha exposed to 2 ll/ml of essential oil. Mitochondria (M) show some abnormalities, such as the disruption of the mitochondrial structures and reduction of cristae. (E and F) Longitudinal sections of A. ﬂavus hyphae exposed to 4 ll/ml of essential oil. Complete cell depression and disorganization signs, such as extensive destruction, massive vacuolation of cytoplasm with vacuole fusion, formation of massive lomasomes (L), () dissociation of plasma membrane (PM) on the cell wall (CW), thinner ﬁbrillar layer (FL), severe absence of cytoplasmic matrix, and lysis of hyphae membranous organelles. Bar = 2 lm.
hibit that CJEO can efﬁcaciously inhibit dry mycelium weight and the AFB1 synthesis of A. ﬂavus. The essential oil exhibited antiaﬂatoxigenic properties at concentrations lower than its fungitoxic concentration, and similar types of results were also reported by Rasooli et al. (2008) as well as Shukla, Kumar, Singh, and Dubey (2009). Some studies have shown that there is a direct correlation between fungal growth and AFB1 production (Kumar, Shukla, Singh, & Dubey, 2010; Kumar et al., 2008). However, the inhibition of AFB1 production cannot be completely attributed to the insufﬁcient fungal growth, but instead, it can be attributed to the inhibition of carbohydrate catabolism in A. ﬂavus by acting on some key enzymes leading to the reduction of its ability to produce AFB1
(Tatsadjieu, Dongmo, Ngassoum, Etoa, & Mbofung, 2009). The inhibition mechanism of AFB1 production is not very clear. CJEO may interfere with some steps in the metabolic pathways of the A. ﬂavus, which controls the biosynthesis of AFB1. Hence, determining the AFB1 suppression mechanism requires further investigation on CJEO. The results from our studies indicate that the TEM of CJEO-treated A. ﬂavus in comparison with untreated samples appeared to be dose-dependent pathologic changes of fungal cells especially on membranous structures. Different concentrations of the CJEO pass not only in the cell wall but also in the plasma membrane and then interact with the membranous structures of cytoplasmic organ-
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in vivo against food contamination microorganisms for their possible application as natural food preservatives. Acknowledgements This work was supported by the National Mega Project on Major Drug Development (2009ZX09301-14-1), the Commonweal Specialized Research Fund of China Agriculture (201103016), the Key Program of Natural Science Foundation of Hubei Province of China (2010CBB02301), and the Fundamental Research Funds for the Central Universities (20103010101000185). Fig. 4. UV spectrophotometric sterol proﬁles of A. ﬂavus treated with CJEO and compared with those of an untreated control.
elles. However, the results clearly presented that CJEO exerts its effect directly on the plasma membrane without any obvious damage on the cell wall. Changes in the cell permeability due to the break down of the plasma membrane at variable intervals led to the loss of the normal shape of fungal mycelia and the formation of membrane bound vesicles inside the cells. Our results are in agreement with the previous ﬁndings of Razzaghi-Abyaneh et al. (2006). Additionally, in a previous work, Nogueira et al. (2010) reported morphological alterations in A. ﬂavus by TEM observations. They found that ultrastructural changes, which were concentration dependent of the essential oil of Ageratum conyzoides, were more evident in the endomembrane system, affecting mainly the mitochondria. Degradation was also observed in the surrounding ﬁbrils. Thus, based on our study, the plasma membrane and the organelles may be the important targets of the essential oil main components, and the main components of CJEO may affect the above targets leading to an imbalance in the intracellular osmotic pressure, blockage of enzymatic reactions, leakage of cytoplasmic contents, and so on, eventually resulting to the death of the cell. The plasma membrane appears to be the main target of the essential oil based on TEM data. In order to ensure the CJEO target in the plasma membrane, the effect of CJEO on the amount of ergosterol was assessed. 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, Low, Bottema, & Parks, 1985). As mentioned earlier, there is a previous study by Kelly, Lamb, Corran, Baldwin, and Kelly (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 oil can cause a considerable reduction in the quantity of ergosterol (Pinto, Vale-Silva, Cavaleiro, & Salgueiro, 2009; Pinto et al., 2006). In our study, the ergosterol content was determined by previously described methods (Arthington-Skaggs et al., 1999), and it is also an absolute measurement. This sterol quantitation method is indicative of the ergosterol and 24(28) dehydroergosterol contents based on the exclusive spectral absorption pattern produced between 230 and 300 nm by extracted sterols. Hence, the important target in the plasma membrane was determined by quantitating the total intracellular ergosterol production in cells grown in increasing concentrations of CJEO. Our observations reveal that CJEO can induce a considerable impairment of the ergosterol biosynthesis by A. ﬂavus. Thus, the plasma membrane is an important antifungal target of CJEO. In summary, the promising results of the antifungal activity and studies on its components indicate that employing CJEO as a food preservative must be encouraged. Further studies are being undertaken for the individual components and their antifungal activity
References Adams, R. P. (2001). Identiﬁcation of essential oil by gas chromatography/quadrupole mass spectroscopy. Illinois: Allured Publishing Corporation. Adegoke, G. O., Iwahashi, H., Komatsu, Y., Obuchi, K., & Iwahashi, Y. (2000). Inhibition of food spoilage yeasts and aﬂatoxigenic moulds by monoterpenes of the spice Aframomum danielli. Flavour and Fragrance Journal, 15(3), 147–150. Albuquerque, C. C., Camara, T. R., Marian, R. D. R., Willadino, L., Marcelino, C., & Ulisses, C. (2006). Antimicrobial action of the essential oil of Lippia gracilis Schauer. Brazilian Archives of Biology and Technology, 49(4), 527–535. Arthington-Skaggs, B. A., Jradi, H., Desai, T., & Morrison, C. J. (1999). Quantitation of ergosterol content: Novel method for determination of ﬂuconazole susceptibility of Candida albicans. Journal of Clinical Microbiology, 37(10), 3332–3337. Bajpai, V. K., Shukla, S., & Kang, S. C. (2008). Chemical composition and antifungal activity of essential oil and various extract of Silene armeria L. Bioresource Technology, 99(18), 8903–8908. Bankole, S. A., & Joda, A. O. (2004). Effect of lemon grass (Cymbopgon citratus Stapf.) powder and essential oil on mould deterioration and aﬂatoxin contamination of melon seeds (Colocynthis citrullus L.). African Journal of Biotechnology, 3, 52–59. Cardile, V., Russo, A., Formisano, C., Rigano, D., Senatore, F., Arnold, N. A., et al. (2009). Essential oils of Salvia bracteata and Salvia rubifolia from Lebanon: Chemical composition, antimicrobial activity and inhibitory effect on human melanoma cells. Journal of Ethnopharmacology, 126(2), 265–272. Chao, L. K., Hua, K. F., Hsu, H. Y., Cheng, S. S., Liu, J. Y., & Chang, S. T. (2005). Study on the antiinﬂammatory activity of essential oil from leaves of Cinnamomum osmophloeum. Journal of Agricultural and Food Chemistry, 53(18), 7274–7278. Cowan, M. M. (1999). Plant products as antimicrobial agents. Clinical Microbiology Reviews, 12(4), 564–582. Editorial Committee of National Chinese Medical Manage Bureau (1999). Chinese Herb, vol. 3. Shanghai: Shanghai Scientiﬁc & Technical Publishers [pp. 349]. Joseph, G. S., Jayaprakasha, G. K., Selvi, A. T., Jena, B. S., & Sakariah, K. K. (2005). Antiaﬂatoxigenic and antioxidant activities of Garcinia extracts. International Journal of Food Microbiology, 101(2), 153–160. 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 14amethylergosta-8, 24(28)-dien-3b, 6a-diol. Biochemical and Biophysical Research Communications, 207, 910–915. Kumar, A., Shukla, R., Singh, P., & Dubey, N. K. (2010). Chemical composition, antifungal and antiaﬂatoxigenic activities of Ocimum sanctum L. Essential oil and its safety assessment as plant based antimicrobial. Food and Chemical Toxicology, 48(2), 539–543. 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. Innovative Food Science & Emerging Technologies, 9(4), 575–580. Lingk, W. (1991). Health risk evaluation of pesticide contamination in drinking water. Gesunde Pﬂangen, 43, 21–25. Murthy, P. S., Ramalakshmi, K., & Srinivas, P. (2009). Fungitoxic activity of Indian borage (Plectranthus amboinicus) volatiles. Food Chemistry, 114(3), 1014–1018. Nogueira, J. H. C., Goncalez, E., Galleti, S. R., Facanali, R., Marques, M. O. M., & Felicio, J. D. (2010). Ageratum conyzoides essential oil as aﬂatoxin suppressor of Aspergillus ﬂavus. International Journal of Food Microbiology, 137(1), 55–60. 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. Journal of Medical Microbiology, 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. Journal of Medical Microbiology, 58(11), 1454–1462. Rahman, A., & Kang, S. C. (2009). In vitro control of food-borne and food spoilage bacteria by essential oil and ethanol extracts of Lonicera japonica Thunb. Food Chemistry, 116(3), 670–675. Rasooli, I., Fakoor, M. H., Yadegarinia, D., Gachkar, L., Allameh, A., & Rezaei, M. B. (2008). Antimycotoxigenic characteristics of Rosmarinus ofﬁcinalis and Trachyspermum copticum L. essential oils. International Journal of Food Microbiology, 122(1–2), 135–139. Razzaghi-Abyaneh, M., Shams-Ghahfarokhi, M., Kawachi, M., Eslamifar, A., Schmidt, O. J., Schmidt, A., et al. (2006). Ultrastructural evidences of growth inhibitory
J. Tian et al. / Food Chemistry 130 (2012) 520–527 effects of a novel biocide, AkacidÒplus, on an aﬂatoxigenic Aspergillus parasiticus. Toxicon, 48(8), 1075–1082. Reddy, T. V., Viswanathan, L., & Venkitasubramanian, T. A. (1970). Thin layer chromatography of aﬂatoxins. Analytical Biochemistry, 38(2), 568–571. Rodriguez, R. J., Low, C., Bottema, C. D., & Parks, L. W. (1985). Multiple functions for sterols in Saccharomyces cerevisiae. Biochimica et Biophysica Acta, 837, 336–343. Shukla, R., Kumar, A., Singh, P., & Dubey, N. K. (2009). Efﬁcacy of Lippia alba (Mill.) NE Brown essential oil and its monoterpene aldehyde constituents against fungi isolated from some edible legume seeds and aﬂatoxin B1 production. International Journal of Food Microbiology, 135(2), 165–170. Sinha, K. K., Sinha, A. K., & Prasad, G. (1993). The effect of clove and cinnamon oils on growth and aﬂatoxin production by Aspergillus ﬂavus. Letters in Applied Microbiology, 16, 114–117. Sokovic, M. D., Brkic, D. D., Dzamic, A. M., Ristic, M. S., & Marin, P. D. (2009). Chemical composition and antifungal activity of Salvia desoleana Atzei & Picci essential oil and its major components. Flavour and Fragrance Journal, 24(2), 83–87. Sokovic, M., Tzakou, O., Pitarokili, D., & Couladis, M. (2002). Antifungal activities of selected aromatic plants growing wild in Greece. Nahrung/Food, 46(5), 317–320.
Tatsadjieu, N. L., Dongmo, P. M. J., Ngassoum, M. B., Etoa, F. X., & Mbofung, C. M. F. (2009). Investigations on the essential oil of Lippia rugosa from Cameroon for its potential use as antifungal agent against Aspergillus ﬂavus Link ex. fries. Food Control, 20(2), 161–166. Tian, J., Ban, X. Q., Zeng, H., He, J. S., Huang, B., & Wang, Y. W. (2011). Chemical composition and antifungal activity of essential oil from Cicuta virosa L. var. latisecta Celak. International Journal of Food Microbiology, 145, 464–470. 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. International Journal of Food Microbiology, 68(3), 207–210. Williams, J. H., Phillips, T. D., Jolly, P. E., Stiles, J. K., Jolly, C. M., & Aggarwal, D. (2004). Human aﬂatoxicosis in developing countries: a review of toxicology, exposure, potential health consequences, and interventions. American Journal of Clinical Nutrition, 80(5), 1106–1122. Yenjit, P., Issarakraisila, M., Intana, W., & Chantrapromma, K. (2010). Fungicidal activity of compounds extracted from the pericarp of Areca catechu against Colletotrichum gloeosporioides in vitro and in mango fruit. Postharvest Biology and Technology, 55(2), 129–132.