Subscribe Now Subscribe Today
Abstract
Fulltext PDF
References
Research Article
 

Mycobiota and Mycotoxins of Egyptian Peanut (Arachis hypogeae L.) Seeds



M.S. Youssef, O.M.O. El-Maghraby and Y.M. Ibrahim
 
ABSTRACT

Sixty-three species in addition to 3 varieties of 21 genera were isolated from 20 samples of each of untreated (51 species + 3 varieties of 21 genera and 51.24x103 cfu g-1 dry weight seeds), roasted (28 + 2 of 12 and 11.5x103 cfu) and roasted with salt (28 + 2 of 7 and 7.5x103 cfu) on dextrose-Czapek’s agar at 28 °C using dilution-plate method. The dominant fungal genera with their respective species on three types of seeds were Aspergillus (A. niger, A. flavus and A. ficuum), Penicillium (P. citrinum) and Fusarium (F. oxysporum). Based on biological, TLC, spectrophotometeric and ELISA assays, fourteen samples (23.3%) out of 60 tested proved to be toxic with different mycotoxins; aflatoxins, sterigmatocystin, ochratoxins, diacetoxyscirpenol and zearalenone. Also, mycoflora and myctoxins of six cultivars, widely cultivated in Upper Egypt were studied as pre-storage and post-storage in normal store for 3, 6, 12 and 24 months. A total of 28 species belonging to 14 genera were identified on dextrose-Czapek’s agar medium (25 species of 12 genera) and cellulose-Czapek’s agar medium (24 of 12) using dilution-plate method at 28 °C. Aspergillus (A. niger, A. flavus and A. fumigatus), Fusarium (F. oxysporum) and Penicillium (P. citrinum) were the most prevalent fungal genera and species and their counts increased with lengthening of storage period. Cultivars were non-toxic, but toxins production appeared after 12 and 24 months of storage on two and three cultivars, respectively.

Services
Related Articles in ASCI
Similar Articles in this Journal
Search in Google Scholar
View Citation
Report Citation

 
  How to cite this article:

M.S. Youssef, O.M.O. El-Maghraby and Y.M. Ibrahim, 2008. Mycobiota and Mycotoxins of Egyptian Peanut (Arachis hypogeae L.) Seeds. International Journal of Botany, 4: 349-360.

DOI: 10.3923/ijb.2008.349.360

URL: https://scialert.net/abstract/?doi=ijb.2008.349.360

INTRODUCTION

Peanuts are unique among cultivated crops in that produce seed-bearing pods below the soil surface. Pods are in direct contact with soil populations and the seeds are frequently invaded by soil fungi before harvest (Horn and Greene, 1995).

Infection of peanut by Aspergillus, especially members of A. flavus group and A. niger group occurs under both pre-harvest and post-harvest conditions. Pre-harvest infection by A. flavus and A. parasiticus and consequent aflatoxin contamination is a major problem in the semi-arid tropic environment. These fungi are widespread in light sandy soils most suitable for peanut cultivation. Peanut pods when in direct contact with spores of A. flavus in soil are frequently invaded before harvest. The mode and extent of invasion by A. flavus depend on soil population density of A. flavus, soil moisture content and soil temperature during the pod development to maturity period (Horn et al., 1995). Post-harvest contamination may occur when stored products are not maintained at a safe moisture level. Also, these fungi can invade and produce toxins in peanut kernels before harvest, during drying and in storage. Owing to the toxicity and carcinogenicity of aflatoxins contaminated commodities destined for human or animal consumption pose a serious health hazards and are therefore, closely monitored and regulated. Apart from its effect on health, aflatoxins contamination also impacts the agricultural economy, through the loss of produce and thus time and costs involved in monitoring and decontamination (Craufurd et al., 2006; Kumar et al., 2008).

The average aflatoxin concentration accepted by PAC (aflatoxin testing program) in the United States depends on the conditions of peanut crop after shelling (rejected lots with over 25 ppb and accepted lots with 25 ppb aflatoxin or less). In the European Union, the aflatoxin B and the total aflatoxins level in peanut products are regulated with Maximum Residue Levels (MRLs) that cannot be greater than 2 and 4 ng g-1, respectively (EU Commission Regulation, 2002).

Egypt exports shelled nuts at about 7 millions US$ and in shell nuts at about 4.4 millions US$, yearly (FAO, 2006). The major mycotoxins found in Egyptian peanuts were aflatoxins (El-Maghraby and El-Maraghy, 1987). Aflatoxin concentration is the most important quality problem in peanut worldwide with serious health implications for human as well as livestock (Sinha et al., 1999; D’Mello, 2003). The climatic conditions as well as the food production chains are characteristic in most parts of Africa and the largest mycotoxin-poisoning epidemic in a decade was reported in Africa during the last 5 years (Wagacha and Muthomi, 2008).

The present investigation was designed to study the mycoflora and mycotoxins of 20 samples of each of untreated, roasted and roasted with salt peanut seeds prepared for human consumption, collected from different localities in Egypt. Also, fungal contamination and natural occurrence of mycotoxins on six peanut cultivars, widely cultivated in Upper Egypt were studied as post-harvest (1-3 months) and stored in normal store for 3, 6, 12 and 24 months.

MATERIALS AND METHODS

Collection of peanut seed samples: Twenty fruit samples, 500 g, each were collected after harvest of 2004 production, from the Egyptian markets in different Governorates, put in a sterile polyethylene bags, transferred to the laboratory, then the pericarps were unfolded and the seeds were released under sterile conditions, put in other sterile polyethylene bags and kept in a cool place (3-5 °C). As well as, 20 samples of each of roasted and roasted with salt (500 g, each) were collected from the peanut roasters in different Governorates of Egypt, transferred to the laboratory, shelled in sterile conditions and put the seeds of each sample in a sterile polyethylene bag sealed and put in another one, which was also sealed to minimize the loss of water-content and kept in a cool place (3-5 °C) till fungal cultivation, isolation, identification and mycotoxins assay.

Also, the 6 cultivars (Ismaila1, Giza4, Giza5, Giza6, Local262 and R92) were kindly supplied by Oil Crops Department of Agriculture Research Center, Shandaweel Station for Agriculture Researches, Sohag Governorate, Upper Egypt. The seed samples (500 g, each) were put in sterile polyethylene bags, transferred to the laboratory and kept in cool (3-5 °C) conditions till different assays. Storage of cultivars for 3, 6, 12 and 24 months was carried out in oil crops store (Shandaweel Station for Agriculture Researches). The fungal flora and mycotoxins investigations of cultivars were performed pre-storage and after each storage period.

Determination of moisture content of peanut seeds: Twenty grams of each seed sample were milled and dried in an oven at 105 °C for 24 h, then cooled in a desiccator and re-weighted to a constant weight. The moisture content was calculated as percentage of the dry weight.

Determination of seed-borne fungi: Dilution-plate method as described by Johnson and Curl (1972) was used for isolation of fungi. Modified 1% dextrose-Czapek’s agar medium (g L-1; sodium nitrate 3.0, magnesium sulphate 0.5, potassium chloride 0.5, di-potassium hydrogen phosphate 1.0, iron sulphate 0.01, dextrose 10.0, agar agar 15.0-20.0, pH 6.6 ± 0.1) was used as cultivation and isolation medium. In case of seed cultivars fungal flora, modified 1% dextrose-Czapek’s agar and 2.1% cellulose-Czapek’s agar media were used. Chloramphenicol (0.5 mg mL-1) as bacteriostatic agent and rosebengal (30 ppm) to restrict for widespreading fungi because of stimulate slow growing fungi, were added to the medium (Al-Doory, 1980).

Five plates were used for each sample tested and each cultivar after every storage period, in addition to different control samples as post-harvest (pre-storage). The plates were incubated at 28 ± 2 °C for 7-15 days and the developing fungi were identified, counted and calculated per gram dry weight of each tested sample. The colonies of slow growing fungi, as well as mycelial bits were transferred to slants with special media to ensure precise counting, then to plate for identification.

Taxonomic identification of fungi (based on purely morphologically macro- and microscopic characteristics) was carried out according to (Booth, 1971, 1977; Raper and Fennell, 1977; Christensen and Raper, 1978; Pitt, 1979, 1991; Domsch et al., 1980; Moubasher, 1993; Klich, 2002; Samson et al., 2002; Summerell et al., 2003).

Sample preparation for mycotoxins analysis
Extraction procedures: Fifty grams of each sample were defatted by extraction with cyclohexane (150 mL) for 10 h using Soxhlet type extractor. The defatted residue was extracted with ethyl acetate (three times, 50 mL/each). The extracts were combined, dried over anhydrous sodium sulphate, filtered and then concentrated under vacuum to near dryness, transferred into a brown glass vial and evaporated under nitrogen stream.

Clean up of crude extracts: For cleaning up the crude extracts (purified from interfering compounds), it was suspended in 1 mL chloroform and applied to 14x0.8 cm column containing 2.5 g kiesel gel 60, 70/230 silica gel (MERCK. Germany). The washing and eluting solvents (8 mL, each) for aflatoxins (n-hexane followed by ether and 3: 97 methanol-chloroform, respectively), sterigmatocystin (n-hexane and 3: 97 methanol-chloroform, respectively), ochratoxins and citrinin (n-hexane and 5: 95 methanol-chloroform, respectively), trichothecenes (dichloromethane and 5: 95 methanol-dichloromethane, respectively), zearalenone (n-hexane and 5: 95 acetone-benzene, respectively), fusarin C (dichloromethane and 10: 90 methanol-dichloromethane, respectively) and moniliformin (chloroform and 5: 3: 2 toluene-acetone-methanol, respectively) were carried out according to AOAC (1984), Jarvis et al. (1986) and Dorner (1998).

Bioassay of mycotoxins: Three bioassay tests for mycotoxins detection were used; brine shrimps (Artemia salina L.) larvae, Chlorella vulgaris Bejerinck and Bacillus subtilis according to Korpinen (1974), Bean et al. (1992) and Földes et al. (2000), respectively.

Thin Layer Chromatography (TLC): For qualitative detection of mycotoxins, thin layer chromatography technique was employed using precoated silica gel plates type 60 F254 TLC (E, MERCK, Germany). Aflatoxins B1, B2, G1 and G2, ochratoxins A and B, sterigmatocystin, citrinin, diacetoxyscirpenol (DAS), T-2 toxin, zearalenone, moniliformin and fusarin C were used as standard references (Sigma). The plates were developed using the following solvent systems; methanol-chloroform (v/v, 3/97) for aflatoxins, ochratoxins, sterigmatocystin and citrinin, ethyl alcohol-chloroform (v/v, 5/95) for zearalenone, ethyl acetate-n-hexane (v/v, 70/30) for trichothecenes and toluene-acetone-methanol (v/v/v, 50/30/20) for moniliformin and fusarin C. The developed plates were viewed under short and/or long wave length UV (254 and/or 366 nm) light and sprayed with reagents according to Gimeno (1976), Takitani et al. (1979), AOAC (1984), Farber and Sanders (1986), Vesonder (1986) and Dorner (1998).

For quantitative determination of mycotoxins, spectrophotometeric (Cecil, model 703) technique was used at molecular coefficient of 21800 at 260 and/or 366 nm UV light according to the method described by Bean et al. (1972).

Enzyme Linked Immuno-Sorbent Assay (ELISA): For quantitative determination of aflatoxin B1 (AFB1), ELISA technique was employed according to Gathumbi et al. (2001) and Rodriguez et al. (2003) because World Health Organization (WHO, 2006) has cited aflatoxins as the most potent naturally occurring carcinogens, as well as International Agency for Research on Cancer (IARC) placed aflatoxin B1 on the list of human carcinogens (Wu, 2004) and presence of aflatoxin B1 kits.

RESULTS AND DISCUSSION

The moisture content of untreated, roasted and roasted with salt peanut seed samples (on oven dry basis) was low and ranged between 3.03-4.35, 0.81-1.75 and 1.03-1.63, respectively.

Mycological analysis of peanut seed samples tested based on dilution-plate method using 1% dextrose-Czapek’s agar medium at 28 ± 2 °C revealed that 63 species in addition to 3 varieties belonging to 21 genera were isolated and identified from untreated (51 species + 3 varieties of 19 genera), roasted (28 + 2 of 12) and roasted with salt (28 + 2 of 7) peanut seeds. The gross total viable count of fungi as well as the mean of fungal contamination of untreated seeds (72.9% of general total fungi, 51.24x103 and 2.56x103 cfu g-1 dry peanut seed sample) was remarkably high in comparison of roasted (16.4%, 11.5x103 and 0.6x103 cfu) and roasted with salt (10.7%, 7.5x103 and 0.4x103 cfu) as recorded in Table 1. These results are in harmony with that obtained by El-Maghraby and El-Maraghy (1987, 1988), who isolated (64 species + 2 varieties of 19 genera) on 1% dextrose-Czapek’s agar and (43 species + 1 variety of 16 genera) on 2.1% cellulose-Czapek’s agar medium at 28 °C from untreated peanut seeds, respectively.

Generally, the most dominant fungal genera on the three seed types tested were Aspergillus (20 species + 2 varieties, 57 cases out of 60 tested, 95% of total samples and 38.03% of general total fungi) and Penicillium (17 species, 34 cases, 56.7 and 20.4%), respectively. A. niger (22 cases out of 60 tested and 36.7% of total samples), A. flavus (20 and 33.3%), A. ficuum (16 and 26.7%), A. oryzae (11 and 18.3%), A. parasiticus (11 and 18.3%) and P. citrinum (9 and 15%) were the most prevalent species. Also, the high degrees of occurrence of Aspergillus and Penicillium were detected on different seed types tested; that on untreated (90 and 75% of the samples and 20.9 and 24.3% of total fungi), roasted (95, 35, 85.3 and 7.3%) and roasted with salt (100, 60, 81.4 and 13.9%) peanut seed samples, respectively. On the other hand, Fusarium (4 species, 12 cases, 20 and 12.2%) isolated from untreated (3 species, 50 and 16.4%) and roasted (2 species, 10 and 1.2%) seed samples only, respectively and disappeared on roasted with salt samples as recorded in Table 1.

Also, the order of occurrence frequency of isolated species varied on different seed types, that on untreated seeds A. flavus (40% of the samples and 4.9% of total fungi), followed by A. aculeatus (40 and 4.7%), A. niger (35 and 1.3%), A. flavus var. columnaris (25 and 1.1%), P. citrinum (30 and 15.1%) and F. oxysporum (30 and 0.7%) were the most dominant, respectively. While, on roasted seeds, A. ficuum (45 and 27.8%), A. niger (35 and 30.8%), A. parasiticus (30 and 10.4%), A. flavus (25 and 4.3%) and P. chrysogenum (15 and 2.3%), whereas, on roasted seeds with salt, A. niger (40 and 28.1%), A. flavus (35 and 4.9%), A. oryzae (25 and 18.1%), A. ficuum (25 and 5.4%) and P. roquefortii (20 and 4.3%) were the most frequent, respectively as shown in Table 1.

These obtained results are in full agreement with those previously recorded by El-Maghraby and El-Maraghy (1988), and Gonçales et al. (2008) that Aspergillus and Penicillium were the most dominant genera in Egyptian peanut, chickpea seeds and Brazilian peanut seeds, respectively. Also, Christensen (1991) reported that Aspergillus and Penicillium species are the main components of storage fungi play an important role in seed deterioration and they have competitive ability against other fungi due to heavy spores and mycotoxins production of their diversity species. Creppy (2002) reported that seeds and grains are more liable to fungal infection particularly Aspergillus, Penicillium and Fusarium species in tropical and sub-tropical regions dependent on high levels of moisture content. Costa and Scussel (2002) reported that the problem of food contamination with mycotoxins has led to an increasing concern of toxigenic fungi contamination, mainly Aspergillus, Penicillium and Fusarium genera.

Table 1:
Total count (TC, calculated per g dry weight peanut seeds) in each peanut sample, percentage (TC%, per total count of fungi), number of cases of isolation (NCI, out of 20 samples) of fungal genera and species isolated from peanut sample types (untreated, roasted and roasted with salt seeds) on 1% dextrose-Czapek’s agar medium at 28 ± 2 °C using dilution-plate method
- : No fungus isolated, OR: Occurrence remarks; H: High occurrence, (more than 10 samples out of 20 tested), M: Moderate occurrence, (6-10 samples), L: Low occurrence (3- 5 samples), R: Rare occurrence, (less than 3 samples)

The toxicity test using three biological assays (Artemia salina L. larvae, Chlorella vulgaris Bejerinck and Bacillus subtilis) revealed that 14 samples (23.3%) out of 60 peanut seed samples tested proved to be toxic. Based on thin layer chromatography (TLC), spectrophotometeric and ELISA analyses, aflatoxins B1 and B2 or B1, B2, G1 and G2 were detected in 20, 10 and 15% of untreated, roasted and roasted with salt samples, respectively. These toxic samples were heavily contaminated with many members of Aspergillus flavus group (A. flavus, A. parasiticus, A. flavus var. columnaris, A. oryzae and A. flavo-furcatis) as aflatoxins-producers as shown in Table 2.

Similarly, 42.5% of Egyptian peanut, 35% of soybean and 32% of Brazilian peanut seed samples were proved to be contaminated by aflatoxins B1 and B2 or B1, B2, G1 and G2 (El-Maghraby and El-Maraghy, 1987; El-Kady and Youssef, 1993; Gonçales et al., 2008), respectively. Aflatoxins are potent hepatotoxic, carcinogenic metabolites produced by A. flavus Link, A. parasiticus Spear and A. nomius Kurtzman, Hort and Hesseltine as reported by Kurtzman et al. (1987) and Richard (2007). Species of A. flavus group have been responsible for production of carcinogenic aflatoxins in peanuts (Arachis hypogeae L.) in soils worldwide. The mode and extent of invasion by A. flavus depend on high levels of A. flavus group colonization of peanut fruit in soil, moisture content and temperature of soil during the pod development to maturity period. These fungi can also, invade and produce toxins in peanut kernels before harvest, during drying and in storage. Aflatoxins contamination of peanut does not affect yield only, but also, causes serious health risks to human and cattle (Craufurd et al., 2006; Kumar et al., 2008).

Sterigmatocystin was recorded in 15% of roasted seeds and 5% of roasted with salt seed samples. These samples were contaminated by Emericella nidulans var. acristata as sterigmatocystin-producing fungus as stated in Table 2. This toxin was naturally reported in 8.5% of Egyptian sunflower seeds and 8% of peanut seeds (El-Maraghy and El-Maghraby, 1986; El-Maghraby and El-Maraghy, 1987), respectively. This toxin was also detected by Egyptian thermotolerant fungal isolates including Emericella nidulans. E. nidulans var. lata and E. quadrilineata (El-Maraghy and El-Maghraby, 1986).

Ochratoxins A and B occurred naturally in 5% of each of untreated and roasted peanut seed samples.

Table 2: Biological assay and natural occurrence of mycotoxins in the toxic peanut (untreated, roasted and roasted with salt) seed samples
SN = Sample No, % MC = Percentage moisture content, M = Moderate toxicity, 50-75% of dead larvae, L = Low toxicity, 25-50% of dead larvae

These samples were rich in A. ochraceus (A. alutaceus), A. niger and A. carbonarius as ochratoxins-producing fungi (Table 2). Ochratoxin A (OA), a phenylalanyl derivative of a substituted isocoumarin with nephrotoxic, nephrocarcinogenic, teratogenic and immunosuppressive properties, is mainly produced by Aspergillus ochraceus, also by A. niger, A. carbonarius and some species of Penicillium (Serra et al., 2003; Wagacha and Muthomi, 2008). This toxin was recorded as a contaminant in plant products worldwide (Creppy, 2002; Ghitakou et al., 2006; Wagacha and Muthomi, 2008; Kumar et al., 2008). The toxin was also found frequently and in high average concentrations in blood samples obtained from people living in Balkan countries and affected by Balkan Endemic Nephropathy disease (Pfohl-Leszkowicz et al., 2002).

Zearalenone combined with diacetoxyscirpenol (simple microcyclic trichothecene) were detected in one untreated seed sample (5% of samples tested). This sample heavily contaminated by Fusarium species (F. oxysporum, F. sporotrichioides and F. sulphureum) as diacetoxyscirpenol and zeralenone-producers as recorded in Table 2. Zearalenone can co-occur with simple microcyclic trichothecenes synthesized by Fusarium species (El-Maghraby, 1996; Schollenberger et al., 2006; Kumar et al., 2008). These toxins were detected in Egyptian oil seeds (El-Maraghy and El-Maghraby, 1986; El-Maghraby and El-Maraghy, 1987) and cereal grains (El-Maghraby et al., 1995; El-Maghraby, 1996; Al-Abssy, 2002).

Zearalenone has been implicated in numerous mycotoxicoses in farm animals, causing infertility and reproductive problems such as abortions, false heat, recycling, reabsorption of fetuses and mummies and vulval uterine prolapse (Schollenberger et al., 2006). Doses of ZEN that are much greater than concentrations, which have hormonal effect may have genotoxic and carcinogenic effects (Mitterbauer et al., 2003). In blood, zearalenone and its metabolite, zearalenol bind to human sex hormone-binding globulin to some extent (Eriksen and Alexander, 1998). As well as, trichothecenes are well-known strong irritants and have been associated with naturally occurring outbreaks of vomiting feed refusal and possibly gastric ulcers when consumed. Also, trichothecenes inhibit protein synthesis which followed by a secondary disruption of DNA and RNA synthesis. They affect the actively dividing cells and can decrease antibody levels, immunoglobulins and certain other humoral factors such as cytokines (Eriksen and Pettersson, 2004; Richard, 2007).

Effect of storage on mycoflora and mycotoxins of peanut cultivars: Based on dilution-plate method, 28 species belonging to 14 genera were isolated and identified on 1% dextrose-Czapek’s (25 species of 12 genera) and 2.1% cellulose-Czapek’s agar media (24 species of 12 genera) from six peanut seed cultivars (Ismailia1, Giza4, Giza5, Giza6, Local262 and R92) in pre-storage and after storage for 3, 6, 12 and 24 months. The most dominant fungal genera and their respective species were; Aspergillus (A. niger, A. flavus and A. fumigatus), Fusarium (F. oxysporum) and Penicillium (P. citrinum) as recorded in Table 3. These results are in harmony with those obtained by El-Maghraby and El-Maraghy (1988), who surveyed 40 groundnut seed samples collected throughout Egypt and isolated 43 fungal species belonging to 16 genera. The most dominant genera with respective species were Aspergillus (A. fumigatus matching in 62% of the samples, A. flavus, 57%, A. niger, 55%), Fusarium (F. oxysporum, 55%) and Penicillium (P. chrysogenum, 52%). Also, several reports on mycoflora of peanut seeds worldwide are in agreement with the previous results (Chisholm and Coates-Beckford, 1997; Gonçales et al., 2008).

Table 3:
Total count (TC, calculated per g dry weight peanut seeds), number of cases of isolation (NCI, out of 6 samples) of fungal genera and species isolated from 6 peanut seed cultivars, in pre-storage and stored for 3, 6, 12 and 24 months on 1% dextrose- and 2.1% cellulose-Czapek’s agar media at 28 ± 2 °C
- = No fungus isolated

With regarding the results obtained, the gross fungal count of the stored peanut six cultivars under normal conditions, faintly increased than control samples in pre-storage on both of dextrose- and cellulose-Czapek’s agar media (103.7 and 102.7%), respectively after 3 months of storage, followed by relatively increased (119.1 and 110.3%) after 6 months, sharply increased (347.7 and 306.2%) after 12 months and extremely increased (1151.7 and 1131.5%) after 24 months of storage period. The sharply increase after 12 months is up to promotion of Fusarium (F. oxysporum) by 72.4% on dextrose-Czapek’s and Aspergillus and Fusarium counts (A. flavus, A. fumigatus and F. oxysporum) by totally 65.8% on cellulose-Czapek’s agar. As well as, the extremely increase after 24 months of storage is due to flourishing of Aspergillus (79.1 and 73.7%) on the two media used, respectively in addition to appearing of Trichoderma viride, Apocrea chrysosperma and Rhizopus stolonifer (collectively, 15.4 and 14.8%) on the two isolation media, respectively (Table 3).

These results are in harmony with those obtained by Youssef (1987), who studied the effect of different relative humidity (~0, ~75, ~80, ~85, ~92 and ~100%) levels for 1, 2, 3, 4 and 5 months of incubation periods at 20 °C on mycoflora of soybean seeds at 28 and 45 °C, as well as Abdel-Hafez et al. (1992) studied the effect of different moisture content (11.5, 17, 22.7 and 28%) levels at 8, 18 and 28 °C on paddy grain fungi. They reported that an increase in moisture content of seeds and grains, increase in incubation temperature in addition to lengthening of storage period led to flourishing of some fungi especially storage fungi which, consequently led to extremely in gross total fungal count.

Concerning mycotoxins, the different cultivars ((Ismailia1, Giza4, Giza5, Giza6, Local262 and R92) tested proved to be non-toxic and no mycotoxins could be detected in control samples (pre-storage) and after 3 and 6 months of storage period. But, after 12 months of storage, two cultivars had low toxicity with producing aflatoxins B1, B2, G1 and G2 on (Giza5) cultivar and diacetoxyscirpenol combined with zearalenone on (Local262) cultivar. Also, after 24 months of storage, three cultivars proved to be toxic; two cultivars (Giza4 and Giza5) had low toxicity with producing aflatoxins B1, B2, G1 and G2 and the third one (Local262) recorded high toxicity with producing diacetoxyscirpenol and zearalenone (Table 4). Similarly, USDA, RMA (2005) stated that, post-harvest aflatoxin contamination can increase during storage and if crop drying is delayed. Insects or rodent infestations may also facilitate mould invasion of some stored commodities. Corn, peanuts and cotton seeds are the commodities with the high risk of contamination during storage.

Table 4: Effect of storage periods (3, 6, 12 and 24 months) on mycotoxins production on different peanut cultivars tested
Brine shrimp test: H = High toxicity; > 75% dead larvae, M = Moderate toxicity; 50-75% dead larvae, L = Low toxicity; 25-50% dead larvae, NT = Non toxic; < 25% dead larvae, Mycotoxins detected: DAS = Diacetoxyscirpenol, ZEN = Zearalenone, UTF = Unidentified toxin factor, ND = None detected

CONCLUSION

In conclusion, it is clearly evident that peanut seeds is a good substrate for mold infection and production of dangerous mycotoxins; aflatoxins, ochratoxins, sterigmatocystin, trichothecenes and zearalenone with potentially hazards to the health of both humans and animals. So, for human public health, peanuts and their products in different stages of production chain from the field to the final consumer must be subjected to quality control and good testing protocol for molds and mycotoxins contamination to ensure a food supply free of toxic levels of mycotoxins. Also, genetic modification resistant cultivars to fungal invasion and mycotoxin production is effective means beside to, biological control and others in prevention or minimizing mycotoxin synthesis on agricultural commodities in field, post-harvest and in storage. Therefore cultivation of suitable and selective resistant cultivars seeds and grains in different climatic and environmental conditions must be applied all over the world.

REFERENCES
AOAC., 1984. Association of Official Analytical Chemists. Official Methods of Analysis. 13th Edn. Washington DC., pp: 429.

Abdel-Hafez, S.I.I., I.A. El-Kady, M.B. Mazen and O.M.O. El-Maghraby, 1992. Effect of temperature and moisture content on germination capacity and paddy grain-borne fungi from Egypt. Abhath Al-Yarmouk Pure Sci. Eng., 1: 91-105.

Al-Abssy, A.A., 2002. Fungal contamination of wheat grains during storage and its effect on mycotoxins production and grain quality. M.Sc. Thesis, Assiut University, Egypt.

Al-Doory, Y., 1980. Laboratory Medical Mycology. 1st Edn., Lea and Febiger, Philadelphia Kimpton Publishers, London, pp: 410.

Bean, G.A., B.B. Jarvis and M.B. Aboul-Nasr, 1992. Biological assay for the detection of Myrothecium species produced macrocyclic trichothecenes. Mycopathologia, 119: 175-180.
CrossRef  |  Direct Link  |  

Bean, G.A., J.A. Schilinger and W.I. Klaman, 1972. Occurrence of aflatoxins and aflatoxin-producing strains of Aspergillus species in soybean. Applied Microbiol., 24: 437-439.
Direct Link  |  

Booth, C., 1971. The Genus Fusarium. Commonwealth Mycological Institute, Kew, Surrey, England, Pages: 237.

Booth, C., 1977. Fusarium Laboratory Guide to the Identification of the Major Species. 1st Edn., Commenwealth Mycological Institute, England, pp: 58.

Chisholm, F.V. and P.L. Coates-Beckford, 1997. Fungi associated with seeds of three legume species in Jamaica and seed germination at harvest and after storage. Trop. Agric. (Trinidad), 74: 121-127.
Direct Link  |  

Christensen, C.M. and K.P. Raper, 1978. Synoptic key to Aspergillus nidulans group species and related Emericella species. Trans. Br. Mycol. Soc., 71: 177-191.
CrossRef  |  

Christensen, C.M., 1991. Fungi and Seed Quality in Handbook of Applied Mycology. In: Food and Feeds, Arora, D.K., K.G. Mukerji and E.H. March (Eds.). Marcel Dekker, New York, pp: 99.

Costa, L.L.F. and V.M. Scussel, 2002. Toxigenic fungi in beans (Phaseolus vulgaris L.) classes black and color cultivated in the state of Santa Catarina, Brazil. Braz. J. Microbiol., 33: 138-144.
Direct Link  |  

Craufurd, P.Q., P.V.V. Prasad, F. Waliyar and A. Taheri, 2006. Drough, pod yield, pre-harvest Aspergillus infection and aflatoxin contamination on peanut in Niger. Field Crops Res., 98: 20-29.
CrossRef  |  Direct Link  |  

Creppy, E.E., 2002. Update of survey, regulation and toxic effects of mycotoxins in Europe. Toxicol. Lett., 127: 19-28.
CrossRef  |  Direct Link  |  

D'Mello, J.P.F., 2003. Mycotoxins in Cereal Grains, Nuts and other Plant Products. In: Food and Safety: Contaminant and Toxins, D'Mello, J.P.F. (Ed.). CABI, Wallingford, UK., ISBN: 9780851996073, pp: 65-90.

Domsch, K.W., W. Gams and T.H. Anderson, 1980. Compendium of Soil Fungi. 1st Edn., Academic Press, London, ISBN-10: 0122204018, pp: 22-23.

Dorner, J.W., 1998. Chromatographic Analysis of Mycotoxins. In: Chromatographic of Environmental and Food Toxicants. Shibamato, T. (Ed.). Marcel Dekker, Inc., New York, Hong Kong.

EU, Commission Regulation, 2002. The European Scientific Committee for Food. March, 2002.

El-Kady, I.A. and M.S. Youssef, 1993. Survey of mycoflora and mycotoxins in Egyptian soybean seeds. J. Basic Microbiol., 33: 371-378.
CrossRef  |  

El-Maghraby, O.M.O. and S.S.M. El-Maraghy, 1987. Mycoflora and mycotoxins of peanut (Arachis hypogeae L.) seeds in Egypt. I-Sugar fungi and natural occurrence of mycotoxins. Mycopathologia, 98: 165-170.
PubMed  |  Direct Link  |  

El-Maghraby, O.M.O. and S.S.M. El-Maraghy, 1988. Mycoflora and mycotoxins of peanut (Arachis hypogeae L.) seeds in Egypt. II-Cellulose-decomposing and mycotoxins-producing fungi. Mycopathologia, 104: 19-24.
Direct Link  |  

El-Maghraby, O.M.O., 1996. Mycotoxins and mycoflora of rice in Egypt with special reference to trichothecenes production and control. J. Nat. Toxins, 5: 49-59.
Direct Link  |  

El-Maghraby, O.M.O., I.A. El-Kady and S.A. Soliman, 1995. Mycoflora and Fusarium toxins of three types of corn grains in Egypt with special reference to production of trichothecene-toxins. Microbiol. Res., 150: 225-232.
Direct Link  |  

El-Maraghy, S.S.M. and O.M.O. El-Maghraby, 1986. Mycoflora and mycotoxins of sunflower (Helianthus annus L.) seeds in Egypt. II- Sterigmatocystin production by thermophilic (or thermotolerant) fungi. Pak. J. Biochem., 20: 1-9.

Eriksen, G.S. and H. Pettersson, 2004. Toxicological evaluation of trichothecenes in animal feed. Anim. Feed Sci. Technol., 114: 205-234.
Direct Link  |  

Eriksen, G.S. and J. Alexander, 1998. Fusarium toxins in cereals-A risk assessment. Nordic Council of Ministers, Tema Nord Copenhagen, 502: 7-27.

FAO, 2006. Food and nutrition paper, perspective on mycotoxins. Food and Agriculture Organization of the United Nations, Rome.

Farber, J.M. and G.W. Sanders, 1986. Fusarin C production by North American isolates of Fusarium moniliforme. Applied Environ. Microbiol., 51: 381-384.
PubMed  |  Direct Link  |  

Foldes,T., I. Banhegyi, Z. Herpai, I. Varga and J. Szigeti, 2000. Isolation of Bacillus subtilis strains from rhizosphere of cereals and in vitro screening for antagonism against phytopathogenic, food-borne pathogenic and spoilage micro-organisms. J. Applied Microbiol., 89: 840-846.
Direct Link  |  

Gathumbi, J.K., E. Usleber and E. Mrtlbauer, 2001. Production of ultrasensitive antibodies against aflatoxin B1. Lett. Applied Microbiol., 32: 349-351.
CrossRef  |  

Ghitakou, S., K. Koutras, E. Kanellou and P. Markaki, 2006. Study of aflatoxin B1 and ochratoxin A production by natural microflora and Aspergillus parasiticus in black and green olives of Greek origin. Food Microbiol., 23: 612-621.
CrossRef  |  

Gimeno, A., 1976. Thin layer chromatographic determination of aflatoxins, ochratoxins, sterigmatocystin, zearalenone, citrinin, T-2 toxin, diacetoxyscirpenol, penicillic acid, patulin and penitrem A. J. Assoc. Official Anal. Chem., 62: 579-585.
PubMed  |  

Goncalez, E., J.H.C. Nogueira, H. Fonseca, J.D. Felicio, F.A. Pino and B. Correa, 2008. Mycobiota and mycotoxins in Brazilian peanut kernels from sowing to harvest. Int. J. Food. Microbiol., 123: 184-190.
Direct Link  |  

Horn, B.W. and R.L. Greene, 1995. Vegetative compatibility within population of Aspergillus flavus, A. parasiticus and A. tamarii from peanut field. Mycologia, 85: 324-332.

Horn, B.W., R.L. Greene and J.W. Dorner, 1995. Effect of corn and peanut cultivation on soil populations of Aspergillus flavus and A. parasiticus in Southwestern Georgia. Applied Environ. Microbiol., 61: 2472-2475.
Direct Link  |  

Jarvis, B.B., Y.W. Lee, S.N. Comezoglu and C.S. Yatawara, 1986. Trichothecenes produced by Stachybotrys atra from Eastern Europe. Applied Environ. Microbiol., 51: 915-918.
Direct Link  |  

Johnson, L.F. and E.A. Curl, 1972. Methods for Research on Ecology of Soil-Borne Pathogens. 1st Edn., Burgess Publ. Co., Minneapolis, pp: 187.

Klich, M.A., 2002. Identification of Common Aspergillus species. 1st Edn. Centraalbureau Voor Schimmelcultures, Utrecht, Netherlands, pp: 122.

Korpinen, E.L., 1974. Studies on Stachybotrys alternans: I- Comparison of rabbit skin, mouse fibroblast culture and brine shrimp tests as detectors of Stachybotrys toxins. Acta Pathol. Microbiol. Scan. Sect. Bul., 82: 465-469.
PubMed  |  Direct Link  |  

Kumar, V., M.S. Basu and T.P. Rajendran, 2008. Mycotoxin research and mycoflora in some commercially important agricultural commodities. Crop Protect., 27: 891-905.
CrossRef  |  Direct Link  |  

Kurtzman, C.P., B.W. Horn and C.W. Hesseltine, 1987. Aspergillus nomius, a new aflatoxin-producing species related to Aspergillus flavus and Aspergillus tamarii. Antonie Van Leeuwenhoek, 53: 147-158.
PubMed  |  Direct Link  |  

Mitterbauer, R., H. Weindrofer, N. Safaie, R. Krska and M. Lemmens et al., 2003. A sensitive and inexpensive yeast bioassay for the mycotoxin zearalenone and other compounds with estrogenic activity. Applied Environ. Microbiol., 69: 805-811.
CrossRef  |  Direct Link  |  

Moubasher, A.H., 1993. Soil Fungi in Qatar and other Arab Countries. 1st Edn., Center of Scientific and Applied Research, University of Qatar, Doha, Qatar, ISBN-13: 9992121025, Pages: 566.

Pfohl-Leszkowicz, A., T. Petkova-Bocharova, I.N. Chernozemsky and M. Castegnaro, 2002. A review on etiological causes and the potential role of mycotoxins. Food Addit. Contamin., 19: 282-302.

Pitt, J.I., 1979. The Genus Penicillium and its Teleomorphic States, Eupenicillium and Talaromyces. 1st Edn., Academic Press, Ltd. London, Pages: 634.

Pitt, J.I., 1991. A Laboratory Guide to Common Penicillium Species. Common. Sci. Ind. Res. Org., Div. Food Processing, North Ryde, NS.W. Australia, Academic Press Inc. Ltd. London, Pages: 187.

Raper, K.B. and D.I. Fennel, 1977. The Genus Aspergillus. Krieger R.E. Publishing Co., Huntington, New York, USA., Pages: 686.

Richard, J.L., 2007. Some major mycotoxins and their mycotoxicosis: An overvie. Int. J. Food Microbiol., 119: 3-10.
Direct Link  |  

Rodriguez, V.M.L., D.M.M. Calonge and E.D. Ordonez, 2003. ELISA and HPLC determination of the occurrence of aflatoxin M1 in raw cow's milk. J. Food Addit. Contam., 20: 276-280.
Direct Link  |  

Samson, R.A., V.R. Hoeckstra, J.C. Frisvad and O. Filtenborg, 2002. Introduction to Food-Borne Fungi. 6th Edn. Centraalbureau voor Schimmelcultures, Baarn Delft, The Netherlands, pp: 389.

Schollenberger, M., H.M. Muller, M. Rufle, S. Suchy, S. Plank and W. Drochner, 2006. Natural occurrence of 16 Fusarium toxins in grains and feedstuffs of plant origin from Germany. Mycopathologia, 161: 43-52.
CrossRef  |  Direct Link  |  

Serra, R., L. Abrunhosa, Z. Kozakiewicz and A. Venancio, 2003. Black Aspergillus species as ochratoxin A producers in Portuguese wine grapes. Int. J. Food Microbiol., 88: 63-68.
CrossRef  |  Direct Link  |  

Sinha, B.K., K.S. Rajan and T.N. Panday, 1999. Aflatoxin contamination of animal feed in Bihar. Indian J. Vet. Res., 8: 31-38.

Summerell, B.A., B. Salleh and J.F. Leslie, 2003. A utilitarian approach to Fusarium identification. Am. Phytopathol. Soc. Plant Dis., 87: 117-128.
CrossRef  |  Direct Link  |  

Takitani, S., Y. Asaba, T. Kato, M. Suzuki and Y. Ueno, 1979. Spectrodensitometric determination of trichothecene mycotoxins with 4-(P-nitrobenzyl) pyridine on silica gel thin layer chromatograms. J. Chromatogr., 172: 335-342.
PubMed  |  Direct Link  |  

USDA, RMA, 2005. United State Department of Agriculture, Risk Management Agency. Loss Adjustment Procedures for Aflatoxin. Washington DC., USA.

Vesonder, R.F., 1986. Moniliformin produced by cultures of Fusarium moniliforme var. subglutinans isolated from swine feed. Mycopathologia, 95: 149-152.
CrossRef  |  PubMed  |  Direct Link  |  

WHO, 2006. Mycotoxins in African foods: Implications to food safety and health. AFRO Food Safety Newsletter, World Health Organization Food Safety, Issue No. July, 2006. http://www.afro.who.int/des.

Wagacha, J.M. and J.W. Muthomi, 2008. Mycotoxin problem in Africa: Current status, implications to food safety and health and possible management strategies. Int. J. Food Microbiol., 124: 1-12.
CrossRef  |  Direct Link  |  

Wu, F., 2004. Mycotoxin risk assessment for the purpose of setting international regulatory standards. Environ. Sci. Technol., 38: 4049-4055.
Direct Link  |  

Youssef, M.S., 1987. Mycoflora and mycotoxins of soybean seeds in Egypt. M.Sc. Thesis, Sohag, Assiut Univ., Egypt.

©  2019 Science Alert. All Rights Reserved
Fulltext PDF References Abstract