Invasive Species Compendium

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Aspergillus flavus
(Aspergillus ear rot)

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Datasheet

Aspergillus flavus (Aspergillus ear rot)

Summary

  • Last modified
  • 18 December 2021
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Natural Enemy
  • Preferred Scientific Name
  • Aspergillus flavus
  • Preferred Common Name
  • Aspergillus ear rot
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Fungi
  •     Phylum: Ascomycota
  •       Subphylum: Pezizomycotina
  •         Class: Eurotiomycetes

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Pictures

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PictureTitleCaptionCopyright
A. flavus ear rot (Iowa, USA). Mould is evident; seeds are discoloured and rotten.
TitleSymptoms on maize ear
CaptionA. flavus ear rot (Iowa, USA). Mould is evident; seeds are discoloured and rotten.
CopyrightDenis C. McGee/Iowa State University
A. flavus ear rot (Iowa, USA). Mould is evident; seeds are discoloured and rotten.
Symptoms on maize earA. flavus ear rot (Iowa, USA). Mould is evident; seeds are discoloured and rotten. Denis C. McGee/Iowa State University
Maize ear infected with A. flavus.
Contaminated seeds exhibit blue-green-yellow fluorescence under UV light
TitleUV fluorescence of infected seeds
CaptionMaize ear infected with A. flavus. Contaminated seeds exhibit blue-green-yellow fluorescence under UV light
CopyrightDenis C. McGee/Iowa State University
Maize ear infected with A. flavus.
Contaminated seeds exhibit blue-green-yellow fluorescence under UV light
UV fluorescence of infected seedsMaize ear infected with A. flavus. Contaminated seeds exhibit blue-green-yellow fluorescence under UV lightDenis C. McGee/Iowa State University
Seeds damaged by A. flavus.
TitleSymptoms on seeds
CaptionSeeds damaged by A. flavus.
CopyrightICRISAT
Seeds damaged by A. flavus.
Symptoms on seedsSeeds damaged by A. flavus.ICRISAT
Seeds damaged by A. flavus (right) are disoloured and rotten. Healthy seeds (left).
TitleSymptoms on seeds
CaptionSeeds damaged by A. flavus (right) are disoloured and rotten. Healthy seeds (left).
CopyrightICRISAT
Seeds damaged by A. flavus (right) are disoloured and rotten. Healthy seeds (left).
Symptoms on seedsSeeds damaged by A. flavus (right) are disoloured and rotten. Healthy seeds (left).ICRISAT
Comparison of healthy seeds (left) with seeds damaged by A. flavus (right).
TitleSymptoms on seeds
CaptionComparison of healthy seeds (left) with seeds damaged by A. flavus (right).
CopyrightICRISAT
Comparison of healthy seeds (left) with seeds damaged by A. flavus (right).
Symptoms on seedsComparison of healthy seeds (left) with seeds damaged by A. flavus (right).ICRISAT
Groundnut seed infected by A. flavus.
TitleSymptoms on seeds
CaptionGroundnut seed infected by A. flavus.
CopyrightICRISAT
Groundnut seed infected by A. flavus.
Symptoms on seedsGroundnut seed infected by A. flavus.ICRISAT
A. flavus conidiophores emerging from infected maize kernels.
TitleA. flavus conidiophores
CaptionA. flavus conidiophores emerging from infected maize kernels.
CopyrightDenis C. McGee/Iowa State University
A. flavus conidiophores emerging from infected maize kernels.
A. flavus conidiophoresA. flavus conidiophores emerging from infected maize kernels.Denis C. McGee/Iowa State University
Aflaroot - root damaged by A. flavus.
TitleSymptoms on root
CaptionAflaroot - root damaged by A. flavus.
CopyrightICRISAT
Aflaroot - root damaged by A. flavus.
Symptoms on rootAflaroot - root damaged by A. flavus.ICRISAT
A. flavus symptoms on seedling leaf.
TitleSymptoms on seedling leaf
CaptionA. flavus symptoms on seedling leaf.
CopyrightICRISAT
A. flavus symptoms on seedling leaf.
Symptoms on seedling leafA. flavus symptoms on seedling leaf.ICRISAT

Identity

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Preferred Scientific Name

  • Aspergillus flavus Link

Preferred Common Name

  • Aspergillus ear rot

Other Scientific Names

  • Aspergillus fasciculatum Batista & H. Maia
  • Aspergillus flavus f. magnasporus (Sakag. & G. Yamada) Nehira
  • Aspergillus flavus var. wehmeri (Costantin & Lucet) Blochwitz
  • Aspergillus humus E. V. Abbott
  • Aspergillus luteus (Tiegh.) C. W. Dodge
  • Aspergillus parasiticus Speare
  • Aspergillus wehmeri Costantin & Lucet

International Common Names

  • English: aflatoxin producing mould; Aspergillus boll rot of cotton; storage rot of groundnut, maize and rice; yellow mould of peanut

Local Common Names

  • Germany: Lagerfäule: Erdnuss; Lagerfäule: Mais; Lagerfäule: Reis; Mykotoxinbildner (u. a. Aflatoxin)

EPPO code

  • ASPEFL (Aspergillus flavus)

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Fungi
  •         Phylum: Ascomycota
  •             Subphylum: Pezizomycotina
  •                 Class: Eurotiomycetes
  •                     Subclass: Eurotiomycetidae
  •                         Order: Eurotiales
  •                             Family: Trichocomaceae
  •                                 Genus: Aspergillus
  •                                     Species: Aspergillus flavus

Notes on Taxonomy and Nomenclature

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Aspergillus flavus and A. parasiticus are closely related fungi that contaminate seeds and plant debris of many crops in the field and in storage. A. flavus produces aflatoxin B1 and B2, whereas A. parasiticus produces G1, G2, and M1. Researchers rarely distinguish between the two in their work (Diener et al., 1987).

A novel genetic approach for classifying species of Aspergillus section flavi is described, using polymerase chain reaction (PCR) amplification of the 5.8S ribosomal DNA-intervening internal transcribed spacer regions (ITS I-5.8S-ITS II) with universal primers and analysis of the PCR product by the principle of single-strand conformation polymorphism (SSCP). Non-radiolabelled PCR-SSCP analysis is inexpensive and practical to perform without special apparatus or skill and should assist in fungal morphological identification (Kumeda and Asao, 1996).

Description

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Conidial heads are radiate, with both metulae and phialides. Conidia are globose to subglobose, echinulate, usually 3-6 µm diameter. Sclerotia are dark-red to black, and 400-700 µm diameter.

Distribution

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A. flavus is a member of the storage fungi that are distributed throughout the world on decaying seeds and grains (Christensen and Meronuck, 1986).

Distribution Table

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The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Last updated: 03 Aug 2022
Continent/Country/Region Distribution Last Reported Origin First Reported Invasive Reference Notes

Africa

BeninPresent
BotswanaPresent
Burkina FasoPresent
CameroonPresent
ChadPresent
EgyptPresent
EthiopiaPresent
GhanaPresent
KenyaPresent
LibyaPresent
MoroccoPresent
NigerPresent
NigeriaPresent
SenegalPresent
SudanPresent
TanzaniaPresent
UgandaPresent
ZambiaPresent

Asia

ArmeniaPresent
BahrainPresent
BangladeshPresent
ChinaPresent
-AnhuiPresent
-JiangsuPresent
IndiaPresent
-Andhra PradeshPresent
-AssamPresent
-BiharPresent
-ChhattisgarhPresent
-GujaratPresent
-HaryanaPresent
-Himachal PradeshPresent
-Jammu and KashmirPresent
-KarnatakaPresent
-KeralaPresent
-Madhya PradeshPresent
-MaharashtraPresent
-ManipurPresent
-OdishaPresent
-PunjabPresent
-RajasthanPresent
-Tamil NaduPresent
-Uttar PradeshPresent
-UttarakhandPresent
-West BengalPresent
IndonesiaPresent
-JavaPresent
IranPresent
IraqPresent
IsraelPresent
JapanPresent
-HonshuPresent
-KyushuPresent
LebanonPresent
NepalPresent
OmanPresent
PakistanPresent
PhilippinesPresent
Saudi ArabiaPresent
TaiwanPresent
ThailandPresent
TurkeyPresent
VietnamPresent
YemenPresent

Europe

CzechiaPresent
GreecePresent
ItalyPresent
-SicilyPresent
PolandPresent
PortugalPresent
RomaniaPresent
RussiaPresentPresent based on regional distribution.
-Russian Far EastPresent
SerbiaPresent
SlovakiaPresent
SpainPresent
SwedenPresent
Union of Soviet Socialist RepublicsPresent
United KingdomPresent

North America

CubaPresent
HondurasPresent
MexicoPresent
PanamaPresent
United StatesPresent
-AlabamaPresent
-ArizonaPresent
-CaliforniaPresent
-FloridaPresent
-GeorgiaPresent
-IllinoisPresent
-IndianaPresent
-IowaPresent
-LouisianaPresent
-MississippiPresent
-New MexicoPresent
-North CarolinaPresent
-PennsylvaniaPresent
-TexasPresent

Oceania

AustraliaPresent
-QueenslandPresent

South America

ArgentinaPresent
BoliviaPresent
BrazilPresent
-GoiasPresent
-Minas GeraisPresent
-ParaibaPresent
-ParanaPresent
-Rio de JaneiroPresent
-Rio Grande do SulPresent
-Santa CatarinaPresent
-Sao PauloPresent
ColombiaPresent
EcuadorPresent
PeruPresent
VenezuelaPresent

Risk of Introduction

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Economic importance: high
Distribution: worldwide
Seedborne incidence: moderate
Seed transmitted: not recorded from naturally infected seeds
Seed treatment: yes
Overall risk: low

A. flavus is already distributed worldwide as storage fungi in decaying seeds and grain. For this reason it represents no quarantine risk.

Host Plants and Other Plants Affected

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Plant nameFamilyContextReferences
Abelmoschus esculentus (okra)MalvaceaeOther
Acacia nilotica (gum arabic tree)FabaceaeMain
Allium cepa (onion)LiliaceaeHabitat/association
Allium sativum (garlic)LiliaceaeUnknown
Althaea officinalis (Marsh-mallow)MalvaceaeUnknown
Arachis hypogaea (groundnut)FabaceaeMain
Bertholletia excelsa (Brazil nut)LecythidaceaeMain
Brassica napusBrassicaceaeUnknown
Brassica napus var. napus (rape)BrassicaceaeOther
Brassica rapa (field mustard)BrassicaceaeUnknown
Capsicum (peppers)SolanaceaeOther
Capsicum annuum (bell pepper)SolanaceaeUnknown
Capsicum frutescens (chilli)SolanaceaeOther
Carthamus tinctorius (safflower)AsteraceaeOther
Cicer arietinum (chickpea)FabaceaeOther
CitrusRutaceaeUnknown
Corylus avellana (hazel)BetulaceaeHabitat/association
Cucumis sativus (cucumber)CucurbitaceaeUnknown
Cucurbita moschata (pumpkin)CucurbitaceaeUnknown
Cucurbita pepo (marrow)CucurbitaceaeOther
Cydonia oblonga (quince)RosaceaeOther
Daucus carota (carrot)ApiaceaeOther
Delonix regia (flamboyant)FabaceaeHabitat/association
Dendrocalamus hamiltoniiPoaceaeUnknown
Echinacea purpurea (purple coneflower)AsteraceaeUnknown
Elaeis guineensis (African oil palm)ArecaceaeOther
Ficus carica (common fig)MoraceaeOther
Glycine max (soyabean)FabaceaeOther
Gossypium (cotton)MalvaceaeMain
Helianthus annuus (sunflower)AsteraceaeOther
Hordeum vulgare (barley)PoaceaeOther
Hypericum perforatum (St John's wort)ClusiaceaeUnknown
Ipomoea batatas (sweet potato)ConvolvulaceaeUnknown
Jatropha curcas (jatropha)EuphorbiaceaeHabitat/association
Lagenaria siceraria (bottle gourd)CucurbitaceaeUnknown
Lathyrus sativus (grass pea)FabaceaeOther
Lens culinarisUnknown
Lens culinaris subsp. culinaris (lentil)FabaceaeOther
Leucaena leucocephala (leucaena)FabaceaeHabitat/association
Lupinus albus (white lupine)FabaceaeUnknown
Malus domestica (apple)RosaceaeOther
Momordica charantia (bitter gourd)CucurbitaceaeUnknown
Musa (banana)MusaceaeOther
Oryza sativa (rice)PoaceaeOther
Parthenium hysterophorus (parthenium weed)AsteraceaeHabitat/association
Peltophorum pterocarpum (copperpod)FabaceaeHabitat/association
Phaseolus vulgaris (common bean)FabaceaeOther
Phoenix dactylifera (date-palm)ArecaceaeOther
Piper betle (betel pepper)PiperaceaeUnknown
Pistacia vera (pistachio)AnacardiaceaeOther
Pisum sativum (pea)FabaceaeOther
Plumeria (frangipani)ApocynaceaeUnknown
Prunus domestica (plum)RosaceaeOther
Prunus dulcis (almond)RosaceaeOther
Prunus persica (peach)RosaceaeOther
Prunus salicina (Japanese plum)RosaceaeOther
Quercus griffithiiFagaceaeUnknown
Quercus serrata (glandbearing oak)FagaceaeUnknown
Raphanus sativus (radish)BrassicaceaeUnknown
Salvia officinalis (common sage)LamiaceaeUnknown
Sapindus trifoliatusSapindaceaeUnknown
Sesamum indicum (sesame)PedaliaceaeOther
Solanum lycopersicum (tomato)SolanaceaeOther
Solanum melongena (aubergine)SolanaceaeUnknown
Solanum tuberosum (potato)SolanaceaeUnknown
Sorghum bicolor (sorghum)PoaceaeOther
Theobroma cacao (cocoa)MalvaceaeOther
Torreya taxifoliaTaxaceaeUnknown
Trigonella foenum-graecum (fenugreek)FabaceaeOther
Triticum (wheat)PoaceaeOther
Triticum aestivum (wheat)PoaceaeOther
Triticum aestivum subsp. speltaUnknown
Unonopsis guatterioidesAnnonaceaeUnknown
Vaccinium (blueberries)EricaceaeHabitat/association
Vigna unguiculata (cowpea)FabaceaeOther
Vitis vinifera (grapevine)VitaceaeUnknown
wheat (stored grain)Other
Zea mays (maize)PoaceaeMain
Zingiber officinale (ginger)ZingiberaceaeUnknown
Ziziphus jujuba (common jujube)RhamnaceaeOther
Ziziphus mauritiana (jujube)RhamnaceaeMain

Growth Stages

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Flowering stage, Fruiting stage, Post-harvest, Seedling stage, Vegetative growing stage

Symptoms

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Seeds are discoloured and rotten. A greenish-yellow mould may be evident.

List of Symptoms/Signs

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SignLife StagesType
Fruit / extensive mould
Fruit / lesions: black or brown
Leaves / abnormal colours
Leaves / fungal growth
Leaves / necrotic areas
Roots / necrotic streaks or lesions
Seeds / discolorations
Seeds / rot
Stems / discoloration of bark
Stems / mould growth on lesion
Whole plant / dwarfing

Biology and Ecology

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A. flavus is widespread in soils, crop residues and air throughout the world (Diener et al., 1987). Inoculum densities of A. flavus ranged from 1-2 to 110-120 infested debris propagules per litre of soil in six cotton fields in California, USA (Kiyomoto and Ashworth, 1974). In groundnut field plots in Taiwan, populations of A. flavus ranged from 786 to 3467 propagules per gram of dry soil (Lee and Chuang, 1992). The frequency of aflatoxin-producing strains in the soil in Taiwan varied with the growth stage of groundnut. Sandy loam and soil of 5-10 cm depth yielded a greater frequency of aflatoxin-producing strains. Of the total A. flavus population in the soil, 45-72% were aflatoxin-producing strains, with some seasonal fluctuation. Significantly more aflatoxin-producing strains were isolated from the soil than from pod shells and kernels (Lee and Chuang, 1997).

A. flavus was the dominant species in soil samples taken across western and central Texas, Georgia/Alabama, and Virginia/North Carolina; the S strain of this species, characterized by production of numerous small sclerotia of <400 µm diameter, was present primarily in the cotton-growing regions of eastern-central Texas and Louisiana. Among the groundnut-growing regions, Georgia and Alabama had the highest and western Texas the lowest soil densities of species from section Flavi. Variability in the density of A. flavus and A. parasiticus in soil may result from regional differences in the frequency of drought and in soil temperature, and the influence of these factors on the susceptibility of groundnut seeds to fungal invasion (Horn and Dorner, 1998).

In maize fields in Iowa the population recovered declined from an estimated 1231 colonies per gram of dry soil in autumn 1988, immediately after an aflatoxin epidemic, to 396 colonies per gram of dry soil in autumn 1990 (Shearer et al., 1992). Aspergillus was found in 67-100% of stored wheat samples taken from seven farms in Slovakia (Trancinova et al., 2001). Sclerotia of A. flavus were detected in samples of insect-damaged and mouldy maize ears from a field that was left unharvested following the 1981 growing season, and their dispersal into soil from the combine harvester was demonstrated during harvest (Wicklow et al., 1984). Sclerotia, overwintered in field soil at two sites in southern Georgia, survived in greater numbers than those overwintered at three sites in central Illinois (Wicklow, 1987). Also, sclerotia placed on the soil surface germinated 8 days prior to the maize silking date (Wicklow and Wilson, 1986). However, the role of sclerotia as inoculum sources for A. flavus infection of maize has not been determined.

McGee et al. (1996) showed that, in years not favourable for aflatoxin development, populations of A. flavus in the soil and on maize crop residues in Iowa changed very little from year to year. Unpublished work by the same authors (DC McGee, OM Olanya, GM Hoyos and LH Tiffany, Iowa State University, Iowa, USA, personal communication, 1996) showed that soil temperatures in the range 30-39°C greatly increased soil populations of A. flavus within 1 week of incubation and that this increase corresponded with increased release of conidia of A. flavus into the air. In groundnut fields in Taiwan, the population of A. flavus in field plots varied within season, increasing irregularly in June or July and decreasing in November and December of the same year or February and March of the following year. The number of fungal propagules was lower in irrigated than in non-irrigated fields, but fluctuations in the soil population were the same in both field types. The most important factors governing the fungal population were the average RH per day and average highest temperature per day in the 3 weeks before the collection of soil.. Temperature was, however, the most important factor affecting seasonal variation of A. flavus in soils (Lee and Chuang, 1992).

Maize samples in Costa Rica were frequently contaminated with A. flavus; about 80% of samples contained >20 ng/g of aflatoxins. Samples kept on the cob after harvest contained almost no aflatoxin whereas shelled samples were often highly contaminated. A. flavus was isolated more frequently from maize shelled immediately after harvest than from maize dried on the cob. Samples harvested with a water content of at least 18% often contained >70% A. flavus-infected (Mora and Lacey, 1997).

Imported and local groundnut samples collected from wholesale markets, storage and shops in Aden, Hodida and Sana'a in January/February, 1998, showed A. flavus in 37.4% of the samples. Mycotoxins were detected in 52% of samples at levels of <10 to 160 µg/kg (Al-Nahdi, 2000).

More than 75% soil samples and more than 70% groundnut kernel samples collected from farmers' fields in Karnataka, India, were contaminated by A. flavus. A positive correlation was found between A. flavus population density and kernel infection level in about 25% samples (Kumar et al., 2001).

Airborne inoculum of A. flavus has been detected in several states of the USA (Bothast et al., 1978; Holtmeyer and Wallin, 1981; McGee et al., 1996). A distinct gradient in spore density of A. flavus was found in relation to distance from piles of waste maize in the vicinity of maize storage sites at three locations in Iowa. The spore density gradient also correlated well with leaf, silk and kernel infection at the same distances in adjacent maize fields (Olanya et al., 1995). Large numbers of airborne conidia of A. flavus exist in aflatoxin-contaminated cotton fields in Arizona, with the number of propagules in soil, air and on cotton leaves highest in mid- to late August (Diener et al., 1987).

In field and greenhouse studies, inoculation of external maize silks that were yellow-brown resulted in more extensive colonization of the silks and a greater number of infected kernels than inoculation of brown silks. The pattern of tissue colonization was very similar in silk-inoculated ears and in those that were not inoculated; growth generally proceeded from the ear tip towards the base, colonizing the silks first, then the glumes and (by the late milk stage) the kernel surfaces, but rarely penetrating the cob pith. Silk senescence and the subsequent growth of A. flavus down the silks was rapid in environment chambers at 30-34°C; the fungus reached the base of some ears in 4 days. Equivalent progress took 4-13 days in the field. The mycelium of A. flavus spread quickly from the silks onto the kernel surfaces, forming a clustered distribution. The percentage of kernels colonized within an ear half was correlated with the extent of contamination of the associated silks by A. flavus, but showed no relationship to the location or extent of visible insect damage. Internal infection of kernels did not appear until the early dent stage (Marah and Payne, 1984).

Silks of three physiological ages (green unpollinated, yellow-brown and brown) of preharvest maize were examined by SEM at 4, 8 and 24 h after inoculation. The few conidia that germinated on unpollinated silks failed to colonize. Conidia on yellow-brown silks germinated in 4-8 hours and colonized the silks extensively, especially near pollen grains, where thick hyphal mats produced numerous conidiophores. Indirect and direct penetration of silk was observed. Conidia germinated on brown silks but hyphal growth was sparse. SEM observation of split kernels from ears inoculated with A. flavus showed early hyphal growth localized in the tip cap (Marsh and Payne, 1984). Inoculation of maize ears with A. flavus 6 days after midsilk resulted in as many, or more, infected grains than inoculation of ears 12 and 18 days after midsilk. Multiple inoculations did not increase incidence of grain infection or aflatoxin contamination (Scott and Zummo, 1994).

Evidences in support of soil invasion of groundnuts by A. flavus as opposed to aerial invasion include a higher percentage invasion of kernels rather than flower or aerial pegs and preliminary data from two air samplings showed an absence of propagules of A. flavus or A. parasiticus in air around the experimental facility (Cole et al., 1986). However, infection of groundnut flowers does occur, as indicated by recovery of A. flavus from field-collected flowers (Griffin and Garren, 1976a).

In greenhouse and field studies, cotton flowers were inoculated with A. flavus at the involucral nectaries. Bolls developing from early-season flowers had significantly higher percentages of A. flavus-infected seed than bolls from flowers formed later in the season. Seeds from bolls inoculated 2 weeks after anthesis had the same infection levels as those from flowers inoculated at anthesis. These results indicate that early-season flowers are predisposed to A. flavus infection and that the degree of susceptibility at anthesis is retained through early boll development (Klich, 1990).

The occurrence of fluorescent fibres from cottonseed harvested from fields in Arizona indicated that seeds had been infected by A. flavus during development. Hyphae, conidial heads and conidia were identified readily in differentially-stained cotyledon tissue processed for light microscopy. Hyphae were located throughout the cotyledon and in the non-lignified layers of the seed coat. The identification of hyphae in cross sections of vessel elements within the seed coat provided ultrastructural evidence supporting the hypothesis that A. flavus may enter seeds via the vascular tissue. The observations demonstrated that the hyphae localized within fluorescent seeds had features characteristic of A. flavus and that fungal-like structures do not occur within uninfected seeds (Huizar et al., 1990).

Experiments in which cotton plant parts were inoculated with A. flavus demonstrated that the fungus could enter seedlings, flowers and developing bolls early in the season, and infect bolls maturing in August and September (Klich, 1988). Another study showed that the critical period for aflatoxin formation was a 30-45-day interval commencing about the date of initial boll opening as a result of A. flavus infection (Russell et al., 1987).

The incidence of A. flavus and aflatoxin levels were higher in groundnut seeds than in castor, sesame and cotton. Amongst oil cakes, A. flavus was more prevalent on cotton followed by groundnut, castor and sesame. Analysis of oil samples revealed that aflatoxin contamination was highest in groundnut followed by sesame, cotton and castor (Verma et al., 1997).

In general, production of aflatoxin B1 by A. flavus in barley grains was less in the presence of Penicillium verrucosum, Fusarium sporotrichioides and Hyphopichia burtonii (Ramakrishna et al., 1996).

The development of spores, cleistothecia and sclerotia in A. flavus and A. parasiticus is affected by linoleic acid and light. The specific morphological effects of linoleic acid include the induction of precocious and increased asexual spore development in A. flavus and A. parasiticus strains. Spore development was induced by hydroperoxylinoleic acids, which are linoleic acid derivatives produced during fungal colonization of seeds. Light treatments also increased the production of asexual spores. The sporogenic effects of light and linoleic acid may be significant environmental signals for fungal development (Calvo et al., 1999).

The specific activity of kojic acid formation of Aspergillus strains was maximal during the exponential growth phase. Carbohydrates such as glucose, saccharose, maltose and galactose were the most utilized. A comparatively high yield of kojic acid (8.5-9.5 g/kg) was obtained by solid-phase fermentation of grain and grain-forage with a high amount of proteins and carbohydrates (maize, oats, rye and barley). The toxicity of the preparation in chickens produced a number of clinical and pathological symptoms characteristic of kojitoxicosis (Kharchenko and Kuts, 1999).

The aflatoxin content and incidence of fungi in boiled, baked and raw arecanut samples from different commercial outlets in South Africa contained aflatoxins B1, B2, G1 and G2. In countries such as India and Taiwan, chewers of raw areca may be exposed to concentrations of aflatoxins which may enhance the carcinogenic effects of these nuts on human tissues (Bijl et al., 1996).


Epidemiology

A number of environmental and edaphic factors influence infection, development and the spread of A. flavus and subsequent aflatoxin production. These factors influence preharvest, harvest and transportation and postharvest stages of aflatoxin development.

Aflatoxin contamination from A. flavus was greatest in groundnuts grown under drought stress with a mean geocarposphere temperature of 29.6°C. It was concluded that groundnuts grown under drought stress may not be contaminated with aflatoxin unless drought is accompanied by elevated geocarposphere temperature during the latter part of the growing cycle (Cole et al., 1985). In plot tests on Florunner groundnuts, the incidence of A. flavus in drought-stressed, unshelled, sound mature kernels decreased with decreases in the mean 5 cm deep soil temperature. The mean threshold geocarposphere temperature required for aflatoxin development during the later growth stages of the plant was 26.3-29.6°C; 31.3°C was too high for contamination to occur (Cole et al., 1984). Incidence of A. flavus was greatest in edible groundnuts from the 30, 40 and 50 days stress treatments. A threshold stress period for preharvest contamination of groundnuts by A. flavus was more than 20 days and possibly less than 30 days (Sanders et al., 1985).

Temperature is considered to be a key factor in high aflatoxin contamination in cotton grown in the western USA, where extended periods of temperatures above 32°C occur during boll development and opening (Kiyomoto and Ashworth, 1974). This was corroborated in a growth chamber study, where excised cotton bolls were inoculated with an aflatoxin producing strain of A. flavus and exposed to 7 or 3 days with short (2-hour) or long (10-hour) diurnal maximum temperature cycles. The maximum and minimum temperatures were 30-32°C and 16-18°C. The percentage bright-greenish-yellow fluorescence of locks and seeds, and also seed infection of bright-greenish-yellow fluorescent seeds, increased as the duration of the daily maximum temperature of 30°C increased, and/or as the number of diurnal maximum temperature cycles of 30°C increased (Gilbert et al., 1975). In maize, A. flavus infection was favoured by warm (32-38°C) rather than cool (21-26°C) temperatures under growth chamber conditions (Jones et al., 1980).

Environment in storage plays an important role in the growth of A. flavus and aflatoxin development. Ninety samples of maize grains from different regions of Brazil were inoculated with A. flavus and incubated at various controlled temperatures and RH environments for periods of 10to 30 days. The best conditions for the production of aflatoxins were 25°C and 85 and 98% RH. The lowest levels were obtained at 40°C and 61.5% RH. The lowest temperature (15°C), 61% RH and moisture content of 13% were the best conditions for storage of maize (Asevedo et al., 1993).

Wheat, sesame and faba bean substrates and their fragments containing 15% moisture, as well as the intact faba beans containing 20 or 30% moisture did not support the production of aflatoxins. However, autoclaving of the substrates stimulated the production of small amounts of aflatoxins B1 and G1. The highest level of total aflatoxins were observed from sesame seeds followed by wheat grains and faba bean seeds, and at 30% moisture. In general, dehulled seeds stimulated toxin production more than whole seeds and hulls (Ragab and Syiad, 1998).

Germination A. flavus was very rapid at >0.90 aw with an almost linear increase over time. A. flavus had very short lag times between 0.995-0.95 aw over a wide temperature range (Marin et al., 1998). When maize cv. TSZB samples re-moistened to moisture contents (mc) of 20, 25, 30 or 35% and stored with the natural microflora or sterilized before artificial inoculation with A. flavus, 20% mc sample showed a significant positive correlation between moisture content (m.c.) and fungal load. Initiation time for moulding was most rapid in 20% m.c. maize (Oyebanji and Efiuvwevwere, 1999). The minimum moisture content of sesame seeds required for aflatoxin production was 10%. There was no aflatoxin production at 15 or 40°C, and maximum production occurred at 30°C. The highest production of aflatoxin was observed after 15 days of incubation (Shahin, 1998).

Role of insects

Insects have been implicated in the infection process for cotton, groundnut and maize. The Carpel walls of green cotton bolls 12, 19, 26, 33 and 40 days were punctured after flowering to simulate damage by sucking insects or drilled to simulate the exit hole of pink bollworm (Pectinophora gossypiella) larvae. A. flavus was dusted on the wound sites and the treated bolls were harvested 4, 6, 10 and 30 days after injury-inoculation. The pattern of toxin to non-toxin seeds in locks from bolls injured by drilling and inoculated 33 days after flowering most closely resembled the pattern found in locks from naturally contaminated bolls. Results indicated that insect-vectored A. flavus entry and subsequent toxin infection are primarily of green bolls close to maturity and reinforce existing knowledge that control of insects lowers aflatoxin potential (Lee et al., 1987).

Aflatoxigenic fungi were found in or on frass from 28.6% of field-collected larvae and in 8.9% of sterilized and macerated larvae of the pyralid mite, Elasmopalpus lignosellus. More aflatoxigenic fungi tended to be found in pods from untreated plots than in plots treated with chlorpyrifos in field trials. Contamination of pods or seeds with A. flavus-type fungi was positively correlated in all four trials with scarification of pods, and this relationship has been quantified (Bowen and Mack, 1993). A. flavus-contaminated insects, such as the maize earworm (Helicoverpa zea) and the European maize borer (Ostrinia nubilalis) have been implicated as inoculum sources (Lillehoj et al., 1980; Barry et al., 1985). A. flavus-contaminated nitidulid beetles have also been recovered from piles of waste maize and in the insect traps distant from these piles (Olanya et al., 1995).

In laboratory experiments, aflatoxin B1 was quantified on maize following artificial infestation with adult weevils of Sitophilus zeamais that had each been topically treated with 100 spores of A. flavus. Findings indicated that S. zeamais facilitated the growth of A. flavus and aflatoxin production in maize by increasing the surface area susceptible to fungal infection and increasing moisture content as a result of metabolic activity. the activity of S. zeamais can have a profound effect on postharvest aflatoxin production, even though little initial inoculum is present (Beti et al., 1995). Maize kernels infested with A. flavus-contaminated weevils had significantly higher levels of aflatoxin B1 than A. flavus-inoculated maize without weevils. The presence of S. zeamais resulted in increased kernel moisture content during incubation, and grain moisture was positively correlated with aflatoxin content across treatments receiving spores. Aflatoxin B1 levels were higher in maize treated with fungus-contaminated S. zeamais compared with maize that was mechanically damaged and inoculated with spores, which in turn had more aflatoxin than undamaged maize treated with spores. Aflatoxin B1 content in maize increased with time of exposure of S. zeamais from 7 to 21 days, but decreased after 28 days of exposure. Aflatoxin levels in infested maize increased significantly with increased numbers of A. flavus-contaminated S. zeamais. S. zeamais carried spores both internally and externally; however, substantial numbers of spores were intimately associated with the exoskeleton of adult S. zeamais. These findings indicated that S. zeamais facilitated the growth of A. flavus and aflatoxin production in maize by increasing the surface area susceptible to fungal infection and increasing moisture content as a result of metabolic activity (Beti et al., 1995). In field studies, Diatraea grandiosella substantially increased aflatoxin levels when combined with A. flavus (Windham et al., 1999).

The significant interaction between Mussidia nigrivenella infestation and A. flavus inoculation indicated that higher concentrations of aflatoxin B1 were found when the fungus was associated with borers than with the fungus alone. M. nigrivenella was the major field pest connected with A. flavus infection and subsequent aflatoxin production in preharvest maize in Benin (Setamou et al., 1998). Maize samples from Benin had no aflatoxins when free of insect damage by M. nigrivenella. In maize, of 70% of cobs damaged by insects, 30.3% were aflatoxin-positive, with a mean aflatoxin contamination of 77.8 p.p.b. Grain moisture increased with the level of damage (Hell et al., 2000).

Resistance to pink bollworm could result in reduced aflatoxin contamination when bollworm pressure coincides with conditions conducive to A. flavus infection. However, Bt cultivars are not resistant to aflatoxin increases occurring after boll opening and large quantities of aflatoxin can form during this period (Cotty et al., 1997). A. flavus, was isolated from dead Vespula vulgaris larvae collected from the field in New Zealand. Disease symptoms developed in some larvae within 24 h after inoculation with A. flavus and killed 40% of larvae (Glare et al., 1996).

Variation

Large variations in strains of A. flavus have been reported for morphological traits, virulence patterns, aflatoxin production potential and genetic diversity.

The A. flavus group consists of two morphological groups, S and L, on the basis of the production of small or large sclerotia, respectively. Phylogenetic analysis based on gene sequences, morphological data and aflatoxin production indicated that A. flavus groups I and II represent a deep divergence within this species. Most group I strains produced some B aflatoxins, but none produced G aflatoxins. Of the six group II strains, four produced both B and G aflatoxins. Group II isolates were all of the S phenotype; group I strains included both S and L phenotypes (Geiser et al., 2000). In general, S strain isolates produce greater quantities of aflatoxins than L strain isolates. Boll age at inoculation influenced the formation of sclerotia. Frequent sclerotia formation during boll infection may favour success of the S strain in cotton fields and increase the toxicity of A. flavus-infected cottonseed. Atoxigenic A. flavus L strain isolate AF36 reduced the formation of sclerotia and aflatoxin when coinoculated with S strain isolates. The use of atoxigenic L strain isolates to prevent contamination through competitive exclusion may be particularly effective where S strain isolates are common (Garber and Cotty, 1997). In the southern USA, A. flavus incidence increased with temperature and decreased with latitude. The aflatoxin-producing potential of A. flavus differed between areas and was correlated with the incidence of the S strain. L strain isolates produced only 33% as much aflatoxin B1 as S strain isolates. No S strain isolate produced both aflatoxin B1 and aflatoxin G1. Correlations suggested that L strain toxigenicity varied geographically. Differences among communities may reflect geographic isolation and/or adaptation, and may cause different vulnerabilities to aflatoxin contamination among crops planted in diverse locations (Cotty, 1997).

The diversity of a naturally occurring population of A. flavus and its ability to contaminate grain with aflatoxin and produce bright greenish yellow fluorescent (BGYF) kernels suggests that substantial variation exists among A. flavus genotypes in their ability to produce aflatoxin in the germ and endosperm of infected BGYF kernels. Naturally occurring A. flavus population may include a majority of strains that produced no aflatoxin but exhibited BGYF and were thus aflatoxin 'false positives' when maize grain was examined with UV light at 365 nm (Wicklow, 1999).

The spatial and temporal patterns of A. flavus strain incidence were compared with patterns of A. flavus propagule density in the soils of Yuma County, Arizona. Strain S isolates were found in all sampled fields but the percentage of strain S isolates ranged from 4 to 93%. For both variables, the largest component of variance occurred among fields within areas at a spatial scale of 1 to 5 km. Temporal patterns were similar for both variables (Orum et al., 1997).

Isolates of A. flavus can be differentiated on the basis of the production of the polygalacturonase P2c. In general, the group that produces P2c causes more damage and spreads to a greater extent in cotton bolls indicating that P2c contributes to the invasion and spread of A. flavus during infection of cotton bolls (Shieh et al., 1997). Two A. flavus isolates, AF12 (low virulence and lacking pectinase P2C) and AF13 (high virulence and producing pectinase P2C) produced amylases, proteases and xylanases, whereas cellulases were not detected in either isolate. AF13 produced more amylase than AF12, and this difference was supported by amylase isoform differences between isolates. These variations in other hydrolytic activities (besides pectinases) may contribute to virulence differences in cotton bolls between AF12 and AF13 (Brown et al., 2001).

Of 147 A. flavus isolates recovered from dried fruit slices, 23.14% of the isolates were aflatoxinogenic, producing aflatoxins B1 and B2 in varying amounts. The biochemical composition of dry fruit slices of quince, along with climatic conditions are favourable for aflatoxin production by toxigenic A. flavus strains (Sharma et al., 1999). A non-toxigenic strain (4351) had greater growth potential than a toxigenic strain (2754) over a wide temperature range, suggesting a mechanism for possible competitive advantage in the soil. Optimum temperature for growth and aflatoxin production in the toxigenic strain was substantially different at 25 and 35°C, respectively (Schearer et al., 1999).

A. flavus is thought to be cosmopolitan and clonal because it has uniform asexual morphology. The assumptions of clonality and conspecificity were investigated in a sample of 31 Australian isolates by assaying restriction site polymorphisms from 11 protein encoding genes, and DNA sequences from five of those genes. A. flavus isolates were grouped into two reproductively isolated classes (I and II). The lack of concordance among gene genealogies among isolates in group I was consistent with a history of recombination (Geiser et al. 1998). The genotypic diversity (DNA fingerprinting) of 269 A. flavus strains, from grain sampled at harvest, field soil, maize insects and air-spora, from a maize field in Illinois, USA, showed that 98% of the A. flavus genotypes produced sclerotia and 53% produced aflatoxin. DNA fingerprints revealed two matches involving subpopulations from grain and soil, one match for grain and maize insects and no matches for grain and air-spora (Wicklow et al., 1998).

Of 60 isolates of A. flavus and A. parasiticus from a maize field in Illinois, 33 (55%) distinct DNA fingerprint groups were identified (each group sharing <80% pAF28 band similarity), including 50 distinct genotypes (83%) with <100% pAF28 band similarity. The 83% genotypic diversity of the A. parasiticus population was equivalent to the 81% genotypic diversity 31 A. flavus isolates (McAlpin, 1998). The genetic relationship between 20 toxigenic and non-toxigenic isolates of A. flavus and A. parasiticus was examined using RAPD and Neighbour Joining analysis. A total of 24 RAPD amplifications using a combination of 17 primers discriminated 20 isolates into two distinct groups with both toxigenic and non-toxigenic isolates. There was no association between RAPD genotype and the ability to produce toxin. Five non-toxigenic isolates of A. parasiticus were separated into two groups, in which the isolates were similar but not identical. These groups of non-toxigenic isolates occurred on branches in which toxigenic isolates also occurred suggesting that either multiple losses of toxigenicity have occurred, or that recombination has reassorted this phenotype into a variety of different genetic backgrounds (Tran et al., 1999).

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Bacillus thuringiensis kurstaki Pathogen
Hypocrea rufa Mycoparasite
Trichoderma hamatum Antagonist
Trichoderma harzianum Antagonist
Trichoderma koningii Antagonist
Trichoderma longibrachiatum Antagonist
Trichoderma pseudokoningii Antagonist

Seedborne Aspects

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Incidence

Under conditions favourable for infection and aflatoxin development, very high incidences of seed infection by A. flavus can be expected in maize, groundnuts and cotton (Handoo and Aulakh, 1979). A. flavus has also been recorded as seedborne on barley (Ramakrishna et al., 1993), rice (Almeida et al., 1991), sorghum (Abou-Zeid, 1995), sunflower (Suryanarayanan and Suryanarayanan, 1990), Vigna mungo (Ahmad, 1993), cowpea (Mishra et al., 1990), wheat (Sinha and Sinha, 1991), onion (Gupta et al., 1984), pigeon pea (Shukla and Bhargava, 1978), cedar (Mittal, 1983), chickpea (Shukla and Bhargava, 1978), coriander (Prasad, 1979), soyabean (Tsay, 1990; Krishnamurty and Raveesha, 1996), mangral (Srivastava and Chandra, 1983) and castor bean (Kanwar and Khanna, 1979).

In field and greenhouse studies of the process of infection of maize kernels, inoculation of external maize silks that were yellow-brown resulted in more extensive colonization of the silks and a greater number of infected kernels than inoculation of brown silks. The pattern of tissue colonization was very similar in silk-inoculated and non-inoculated ears; growth generally proceeded from the ear tip towards the base, colonizing the silks first, then the glumes and (by the late milk stage) the kernel surfaces, but rarely penetrating the cob pith. Silk senescence and the subsequent growth of A. flavus down the silks was rapid in environment chambers at 30/34°C; the fungus reached the base of some ears in 4 days. In the field, equivalent progress took 4-13 days. The mycelium of A. flavus spread quickly from the silks onto the kernel surfaces, forming a clustered distribution. The percentage of kernels colonized within an ear half was correlated with the extent of contamination of the associated silks by A. flavus, but showed no relationship to the location or extent of visible insect damage. Internal infection of kernels did not appear until the early dent stage (Marah and Payne, 1984).

Colonization of silks and kernels of preharvest maize was examined by SEM. Silks of three physiological ages (green unpollinated, yellow-brown and brown) were examined at 4, 8 and 24 h after inoculation. The few conidia that germinated on unpollinated silks failed to colonize. Conidia on yellow-brown silks germinated in 4-8 h and colonized the silks extensively, especially near pollen grains, where thick hyphal mats produced numerous conidiophores. Indirect and direct penetration of silk was observed. Conidia germinated on brown silks but hyphal growth was sparse. SEM observation of split kernels from ears inoculated with A. flavus (inoculum applied to kernels) showed early hyphal growth localized in the tip cap (Marsh and Payne, 1984).

Evidence in support of soil invasion of groundnuts by A. flavus, as opposed to aerial invasion of flowers, includes the following observations: there is a higher percentage invasion of kernels rather than flower or aerial pegs by either wild-type A. flavus or mutants; significant invasion by an A. parasiticus colour mutant occurred only in groundnuts from soil supplemented with the mutant, whereas adjacent plants in close proximity but in untreated soil were only invaded by wild-type A. flavus or A. parasiticus; aflatoxin data from drought-stressed, visibly undamaged kernels showed that samples from soil not supplemented with a mutant strain contained a preponderance of aflatoxin Bs (from wild-type A. flavus), whereas adjacent samples from mutant-supplemented soil contained a preponderance of aflatoxin Bs plus Gs (from wild-type and mutant A. parasiticus); preliminary data from two air samplings showed an absence of propagules of A. flavus or A. parasiticus in air around the experimental facility (Cole et al., 1986). However, infection of flowers does occur, as indicated by recovery of A. flavus from field-collected flowers (Griffin and Garren, 1976a).

In greenhouse and field studies, cotton flowers were inoculated with A. flavus at the involucral nectaries. Bolls developing from early-season flowers had significantly higher percentages of A. flavus-infected seed than bolls from flowers formed later in the season. Seeds from bolls inoculated 2 weeks after anthesis had the same infection levels as those from flowers inoculated at anthesis. These results indicate that early-season flowers are predisposed to A. flavus infection and that the degree of susceptibility at anthesis is retained through early boll development (Klich, 1990).

Cottonseeds having fluorescent fibres were harvested from fields in Arizona, USA, and examined utilizing light microscopy and TEM. The occurrence of fluorescent fibres indicated that seeds had been infected by A. flavus during development. The presence of A. flavus was verified by plating portions of seeds with fluorescent fibres. Hyphae, conidial heads and conidia were identified readily in differentially-stained cotyledon tissue processed for light microscopy. Utilization of TEM permitted observations on lignified seed coats and cotyledons of mature cottonseeds. Hyphae were located throughout the cotyledon and in the non-lignified layers of the seed coat. The identification of hyphae in cross sections of vessel elements within the seed coat provided ultrastructural evidence supporting the hypothesis that A. flavus may enter seeds via the vascular tissue. Controls for the microscopy studies included observations on cottonseeds with no visual signs of infection and on laboratory growth cultures of A. flavus. The observations demonstrated that the hyphae localized within fluorescent seeds had features characteristic of A. flavus and that fungal-like structures do not occur within uninfected seeds (Huizar et al., 1990).

Dry seed examination of soyabean seeds grown in Karnataka, India, indicated the frequent occurrence of orange-yellow discoloured seeds that were highly distorted, shrivelled and smaller than normal seeds. A. flavus was consistently isolated from these seeds. The fungus was isolated from seed coats, cotyledons and the hypocotyl-radicle axis. Histopathological investigations revealed hyphae and conidia of A. flavus in the internal tissues of the discoloured seeds (Krishnamurty and Raveesha, 1996).

Effect on Seed Quality

Severely infected seeds will be discoloured or rotten. Seed germination can be reduced. Seeds may also be contaminated with aflatoxin (Christensen and Meronuck, 1986). Total protein and starch contents decreased, whereas amino acid and reducing sugars increased in groundnut seeds infected with A. fumigatus, A. flavus and A. terreus (Bindu and Kumar, 2003).

Seed Transmission

There is no evidence for transmission of A. flavus from naturally infected seeds. However, it has been demonstrated for artificially inoculated maize and groundnut seeds.

Caryopses of maize, wet-heat treated to reduce inherent infection, were inoculated with conidiospores of A. flavus var. columnaris and stored for 1 month, during which the fungus became established within all the seeds. The fungus was isolated from roots, stems and leaves of plants produced from the infected seeds at all stages during growth, and also from male and female inflorescences, despite competition from Fusarium species. A significant proportion of the seeds produced by these plants was also infected by A. flavus var. columnaris, which was never isolated from control plants grown from uninoculated seeds, nor were the propagules either air- or soil-borne (Mycock et al., 1992).

Further evidence for systemic transmission of the pathogen was evident when A. flavus was isolated from the internal tissues of emerging shoots grown from inoculated seeds after 48 and 72 h , but not from the 24-h samples or the controls. Light and electron microscopy showed that A. flavus var. columnaris was able to invade the internal tissues of the emerging shoot via wounds in the surface (caused by physical injury during germination in the sand or vermiculite), through the stomata, and also by penetrating the apparently intact cuticle. Although the mycelium was concentrated in the space between the coleoptile and the primary leaves, both tissue types were infected. Seeds that had been infected in the same manner were allowed to germinate on sterile filter paper before being planted in sterile soil. Uninoculated seeds were used as controls. A. flavus var. columnaris was isolated from all tissues of the experimental seedlings over the 6-week test period, but never from control material (Mycock et al., 1992).

When groundnut seeds were soaked in a spore and mycelium suspension of A. flavus or sown in inoculated soil, pre- and post-emergence symptoms developed. The latter included stunted growth, development of lanceolate, pale green leaves and strong inhibition of root growth. Injury of kernels before inoculation affected post-emergence loss only slightly but greatly increased pre-emergence rot, particularly in the seed inoculations. Post-emergence symptoms were dependent on temperature. At high temperature (28-29°C) plants developed only slight, temporary symptoms (El Khadem, 1975).

Seed Treatments

Seeds of M-13 groundnuts were dressed with fungicides (carbendazim, copper oxychloride, quintozene, mancozeb and carboxin) and incubated at 28±1°C for 6 days. All fungicide treatments except copper oxychloride and quintozene significantly reduced infection by A. flavus. Acid delinting of seeds did not reduce fungal infection and decreased germination percentage (Dharmalingam and Sundararaj, 1973).

The effects of 11 plant essential oils for maize (cv. Pozolero) kernel protection against A. flavus were studied. Principal constituents of eight essential oils were tested for ability to protect maize kernels. Essential oils of Cinnamomum zeylanicum (cinnamon), Mentha piperita (peppermint), Ocimum basilicum (basil), Origanum vulgare (origanum), Teloxys ambrosioides (the flavoring herb epazote), Syzygium aromaticum (clove) and Thymus vulgaris (thyme) caused a total inhibition of fungal development on maize kernels. Thymol and o-methoxycinnamaldehyde significantly reduced maize grain contamination. The optimal dosage for protection of maize varied from 3 to 8%. Combinations of C. zeylanicum with the remaining oils gave efficient control. A residual effect of C. zeylanicum was detected after 4 weeks of kernel treatment. No phytotoxic effect on germination and maize growth was detected with any of these oils (Montes-Belmont and Carvajal, 1998).

Seed Health Tests

Culture plate (Sauer et al., 1982)

1. Seeds are surface sterilized in 5.25% NaOCl for 1 min., then rinsed in sterile water.
2. Incubate seeds on malt agar with 4% NaCl and 200 p.p.m. tergitol for 3-7 days at 5°C.

Blotter (Handoo and Aulakh, 1979)

1. Seeds are surface sterilized in 2% NaOCl for 10 min.
2. Incubate seeds on moist blotter at 20°C for 2 h and transfer to a freezer at -20°C for 24 h, before incubation at 20°C for 7 days under cycles of 12 h UV light followed by 12 h darkness.

Plant Trade

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Plant parts liable to carry the pest in trade/transportPest stagesBorne internallyBorne externallyVisibility of pest or symptoms
Flowers/Inflorescences/Cones/Calyx fungi/hyphae; fungi/spores Yes Yes Pest or symptoms usually visible to the naked eye
Fruits (inc. pods) fungi/hyphae; fungi/spores Yes Yes Pest or symptoms usually visible to the naked eye
Growing medium accompanying plants fungi/spores Yes Pest or symptoms usually invisible
True seeds (inc. grain) fungi/hyphae; fungi/spores Yes Yes Pest or symptoms usually invisible
Plant parts not known to carry the pest in trade/transport
Bark
Leaves
Roots
Seedlings/Micropropagated plants
Stems (above ground)/Shoots/Trunks/Branches
Wood

Impact

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A. flavus is a member of a group of fungi known as storage fungi which cause deterioration of grain or seeds of all plant species stored at seed moisture contents in the range 13-20% (Christensen and Meronuck, 1986). It also produces aflatoxin in infected seeds that can cause death or other symptoms of toxicity when ingested by animals or humans (Diener et al., 1987). Aflatoxin contamination is of major economic importance in cotton, maize, groundnuts and tree nuts due to invasion of seeds in the field, but it may also develop in seeds and grains of other crop species when they are improperly stored (Diener et al., 1987).

Surveys throughout the world emphasize the enormous problem presented by aflatoxin to human health and to the economics of crop production. The incidence of aflatoxin-contaminated samples in major commodities in the USA in 1976 was estimated at 35% for groundnuts, 41% for cottonseed from the far west, 8% for cottonseed from the south-east, 44% for maize from the south-east, and 2.5% for maize from the mid-west (Stoloff, 1976). A total of 455 food samples representing 22 different food types were collected from several localities in Alexandria, Egypt; aflatoxins B1 and M1 were detected in five samples (1.1%). Of 206 fungal isolates obtained, 32 (15.5%) produced aflatoxins. A. flavus was the predominant isolate; A. parasiticus was also isolated from a few food samples. The highest incidence of aflatoxin contamination occurred in groundnuts (three of 40 samples, 7.5%). Aflatoxin B1 levels varied from 98 to 1056 p.p.b. in groundnut samples and were 28 p.p.b. in rice (El-Gohary, 1995).

In South Africa during the period 1982-83, just under 800 samples of agricultural commodities, comprising cereals, compound feeds, hay and silage, were examined. Aflatoxin B1 occurred in over 27% of all samples and in groundnut meal was 1500 ng/g. The most prevalent fungi were A. flavus and A. parasiticus, which occurred in over 22% of all samples (Dutton and Westlake, 1985).

In Malawi, export losses due to aflatoxins ranged from K 0.16 million in 1988/89 to K 1.58 million in 1985/86 with a mean of K 0.943 million. The export losses in groundnut as a percentage of the trade balance ranged from 0.01% in 1988/89 to 1.77% in 1981/82 (Babu et al., 1994).

In the USA, seven truck-loads of maize were tested for mycotoxin contamination. Aflatoxin was identified in all seven at concentrations of 3-501 ng/g (aflatoxin B1 + B2) (Lee and Hagler, 1991). During 1977 and 1980, losses to individuals, firms and public expenditures due to aflatoxin contamination of maize in the south-eastern USA amounted to approximately US$ 200 million and US$ 238 million, respectively (Nichols, 1983). A decline in maize production in Georgia, USA, followed, which was attributed to chronic insect damage and associated aflatoxin contamination in preharvest grain (McMillian et al., 1991). In the mid-west of the USA, where hot, dry conditions favourable for aflatoxin contamination of maize occur less frequently than in the south-east, epidemics occurred in 1977, 1983 and 1988, but in non-drought years, aflatoxin causes minimal economic loss (Hurburgh et al., 1991).

During the cultivation of oyster mushroom variety Aruppukotai-1 (Pleurotus eous), pre-pasteurized straw was used for bed preparation and spawned at the rate of 5% (wet weight basis). These bags caused yield reductions of 17.2-77.3% (Anandh et al., 1999).

The maximum loss in dry weight of seeds of cluster bean (Cyamopsis tetragonoloba) was recorded as 27.51% due to A. flavus after 180 days of storage. There was a gradual decrease in protein content (from 35.75% to 12.25%) after 180 days of storage (Dwivedi, 1996).

Diagnosis

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Contaminated seeds may be detected by visual examination, by a blue-green-yellow fluorescence under UV light or by ELISA (Candlish et al., 1987).

BGY-F reaction products prepared from three preparations: kojic acid (KA) + peroxidase (soyabean peroxide or horseradish type VI and type II) + H2O2, or detached fresh cotton locules + KA + H2O2, or attached field cotton locules that were treated with a spore suspension of aflatoxigenic A. flavus, all resulted in identical chromatographic characteristics, and all exhibited a molecular weight of 282. Further characterization of the BGY-F reaction product with 1H-and 13C-NMR spectroscopic analysis revealed that it was a dehydrogenator dimer of 2 KA, linked through the C-6 positions (Zeringue et al., 1999).

Transformants of A. flavus containing the Aequorea victoria gfp gene fused to a viral promoter or the promoter region and 483 bp of the coding region of A. flavus aflR expressed green fluorescence detectable without a microscope or filters. Expression of green fluorescent protein fluorescence was correlated with resistance to aflatoxin accumulation in five maize genotypes inoculated with these transformants (Du WangLei et al., 1999).

A rapid identification method was developed for aflatoxin-producing strains of A. flavus and A. parasiticus using ammonia vapour. The colony reverse of aflatoxin (AF)-producing strains of A. flavus and A. parasiticus turned pink when their cultures were exposed to ammonia vapour. The colour change was visible for colonies grown on media suitable for AF production such as potato dextrose, coconut and yeast extract sucrose agars after 2 days incubation at 25°C. The colour change occurred immediately after the colony was exposed to ammonia vapour (Saito and Machida, 1999).

Detection and Inspection

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Seeds are discoloured and rotten. A greenish-yellow mould may be evident.

BGY-F reaction products prepared from three preparations: kojic acid (KA) + peroxidase (soyabean peroxide or horseradish type VI and type II) + H2O2, or detached fresh cotton locules + KA + H2O2, or attached field cotton locules treated with a spore suspension of aflatoxigenic A. flavus, all resulted in identical chromatographic characteristics and exhibited a molecular weight of 282. Further characterization of the BGY-F reaction product with 1H-and 13C-NMR spectroscopic analysis revealed that it was a dehydrogenator dimer of 2 KA, linked through the C-6 positions (Zeringue et al., 1999).

Transformants of A. flavus containing the Aequorea victoria gfp gene fused to a viral promoter or the promoter region and 483 bp of the coding region of A. flavus aflR expressed green fluorescence detectable without a microscope or filters. Expression of green fluorescent protein fluorescence was correlated with resistance to aflatoxin accumulation in five maize genotypes inoculated with these transformants (Du et al., 1999).

A rapid identification method has been developed for aflatoxin-producing strains of A. flavus and A. parasiticus using ammonia vapour. The colony reverse of aflatoxin (AF)-producing strains of A. flavus and A. parasiticus turned pink when their cultures were exposed to ammonia vapour. The colour change was visible for colonies grown on media suitable for AF production such as potato dextrose, coconut and yeast extract sucrose agars after 2 d incubation at 25°C and occurred immediately after the colony came into contact with the vapor (Saito and Machida, 1999).

Prevention and Control

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Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.

Cultural Control and Sanitary Methods

Aflatoxin accumulation due to A. flavus infection in cottonseed grown in Arizona is influenced by the timing of irrigation (Russell et al., 1976). Reduced populations of A. flavus and other fungi in the soil, rhizosphere and geocarposphere of groundnuts occurred when the previous crop was vegetables or rice as opposed to groundnuts (Subrahmanyam and Rao, 1974). The use of green rye manure in the rotation tended to increase populations of A. flavus in groundnut production fields (Griffin and Garren, 1976b). In silk-inoculated maize, aflatoxin contamination was less each year in plots that were either irrigated or subsoiled. Although several factors may contribute to high levels of aflatoxin in the field, water stress appears to be a major factor because both subsoiling and irrigation reduced aflatoxin contamination (Payne et al., 1986). Tillage and rotation practices had little impact on soilborne A. flavus populations. A. flavus was recovered at lower frequencies from maize residues from a soyabean-maize rotation than from a plots grown with continuous maize (McGee et al., 1996).

Fermentation of groundnut press cake by black mould (Rhizopus oligosporus) to produce black 'oncom' reduced the total aflatoxin content to 13.43% of the original content, while fermentation by red mould (Neurospora sitophila) to produce red 'oncom' reduced the total aflatoxin content to 41.07% of the original content. Processing of groundnuts into groundnut butter reduced the total aflatoxin content to 20.58% of the original content. Oil extracted from aflatoxin-contaminated groundnuts by wet rendering, hydraulic pressing and solvent extraction methods reduced the total aflatoxin levels at by 78, 47 and 75%, respectively (Fardiaz, 1991). Intact pods and kernels normally contain less aflatoxin than damaged and shrivelled pods and kernels. Sorting out intact and undamaged pods and kernels can help to reduce aflatoxin contamination in food and feed.

Development of mechanical/electronic methods for sorting pods and seeds to reduce aflatoxin content has been suggested (Bockelee-Morvan and Gillier, 1974). The most practical method for salvaging aflatoxin-contaminated maize is by ammoniation. A described procedure is introduction of ammonia that reduces aflatoxin in maize from more than 1000 p.p.b. to less than 10 p.p.b. (Anderson, 1983). Various other techniques used to ammoniate aflatoxin-contaminated maize in the field have been summarized by Hammond et al. (1991).

The application of starter N rates and the interaction of starter with N timing and N rates significantly affected aflatoxin levels. Rates of 50-250 lb N/acre were 34-43% lower in aflatoxin contamination than plots receiving no N. The application of 10 lb N/acre starter reduced aflatoxin levels by 20% compared to the no-starter control (Tubajika et al., 1999).

Regression analysis indicated significant relationships between gin date and aflatoxin content in 1995 and 1996. Aflatoxin content increased with later ginning in 1995 and 1996 in western Arizona, USA. Overall, 89 and 79% of seed lots exceeded 20 p.p.b. aflatoxin in 1995 and 1996, respectively. Means separation confirmed later ginned crops had a significantly greater aflatoxin content. Transgenic Bt and non-Bt cottonseed were similarly contaminated. The mean aflatoxin content of Bt cottonseed was 413 p.p.b. and that of non-Bt cottonseed was 598 p.p.b. These observations suggest that, in Arizona, losses from aflatoxin contamination of cottonseed can be reduced by early harvest (Bock and Cotty, 1999).

Aflatoxin contamination was greater during the spring when high temperatures occurred during maize reproduction and maturation. The crop management system consisting of early planting, a well-adapted hybrid (H-422), 55 000 plants/ha, adequate irrigation and ear insect control by insecticides gave high yield and low aflatoxin levels (Rodriguez del Bosque, 1996).

Management of mid-season and end-season drought reduces aflatoxin contamination. Drought stress increased A. flavus infection in seeds of all five genotypes. TMV 2, Ec 76446 (292) and NcAc 17090 had higher levels of A. flavus seed infection than J 11 or PI 337394 F. Aflatoxin contamination was higher in seed from drought-stressed crops than from those receiving full irrigation (Saleha Nahdi, 1996).

Host-Plant Resistance

In the early 1970s, more than 1200 genotypes of Arachis hypogaea and ten other Arachis species were evaluated for seed resistance to strains NRRL A13794 and 2999 of A. flavus. Two Argentine types (PI337394 and PI337409) were resistant to both strains, and the commercial varieties Wilco 1, Florunner and Argentine were moderately resistant. The two resistant genotypes showed a consistently low level of seed infection through four generations of evaluation (Mixon and Rogers, 1973). The Spanish groundnut (Arachis hypogaea ssp. fastigiata var. vulgaris) germplasms ICGV88145 (PI585006) and ICGV89104 (PI585007), derived from single crosses involving PI337409 and FESR12 for ICGV88145 and J11 and U4-7-5 for ICGV89104, were released in 1993 for their high levels of resistance to natural seed infection by A. flavus. Natural seed infection averaged 0.7% in ICGV88145 and 1.0% in ICGV89104, compared with 1.3% in the best resistant control; seed colonization by A. flavus under artificial inoculation conditions averaged 22.2, 24.0 and 15.6% in these lines, respectively. Average pod yields of ICGV88145 and ICGV89104 in trials in India averaged 2.17 and 2.20 t/ha, respectively, 22 and 18% more than J11. Both lines matured in 110-120 days in the rainy season at Patancheru (Rao et al., 1995).

A maize population has been developed from a single ear on a maize hybrid grown in a field in Georgia, USA, in 1980. This population is resistant to aflatoxin development (McMillian et al., 1991). The population (MAS:gk) expected to be resistant on the basis of the healthy appearance of kernels selected from the original ear had significantly less contamination than its counterpart (MAS:pw,nf) in three out of four field and two out of three laboratory experiments. Over 5 years, the contamination of MAS:pw,nf exceeded that of MAS:gk in all seven trials (P < 0.01). The variation in kernel phenotype, on which the initial selections were based, disappeared with time and populations were produced which did not differ in insect resistance or other plant characters (Widstrom et al., 1987).

In Illinois, USA, the inheritance of resistance to ear rot of maize caused by A. flavus was studied in progeny derived from crosses between resistant (LB31, L317, CI2, N6, 75-R001, B37Ht2, OH513, Tex6 and H103) and susceptible (B73 and/or Mo17) inbreds following inoculation. During 1992-93, the parental, F1, F2, F3 and both backcross generations of 11 crosses were tested. The number of generations evaluated for each cross was dependent on the year. Parental and F1 generations, together with five F2 and three F3 populations, were evaluated for aflatoxin content in both 1992 and 1993. Generation mean analysis indicated that additive and dominance gene action were of primary importance in resistance to Aspergillus ear rot. Dominance genetic effects estimates ranged from 0 to 87.3% of the variation between generation means. Inbreds Tex6, LB31, CI2 and OH513 consistently had the highest levels of resistance. Frequency distribution of aflatoxin content of ears on F2 plants and ears on F3 families (lines) of the Mo17 x Tex6 and B73 x LB31 populations were highly skewed towards the resistant parent, indicating gene dominance. The F2 and F3 generations indicated various levels of transgressive segregation for resistance to A. flavus and to aflatoxin production (Campbell and White, 1995). Breeding lines have been released from Mississippi as sources of resistance to grain infection by A. flavus (Scott and Zummo, 1990, 1992).

Sources of resistance to aflatoxin development are being sought using a grain-screening laboratory assay. A total of 31 maize inbreds and the highly resistant GT-MAS:gk maize population were screened for resistance to aflatoxin production by A. flavus. Intact grains from each line were evaluated in three trials. Significant differences among genotypes for resistance to aflatoxin production were found. Certain genotypes, previously shown to be resistant in field trials, demonstrated resistance in the grain-screening assay. Of the genotypes, 22 had intact-grain resistance that was comparable to GT-MAS:gk.

A separate experiment was carried out to visualize fungal colonization of internal tissue in susceptible and resistant maize grains and to further elucidate the relationship between fungal colonization and aflatoxin production. Five genotypes, screened in the inbred evaluation, were inoculated with an A. flavus aflatoxin-producing strain containing the Escherichia coli beta-D-glucuronidase (GUS) reporter gene linked to an A. flavus beta-tubulin gene promoter. Histochemical staining on non-wounded and wounded seeds detected differences in GUS expression among genotypes, and there was a relationship between GUS expression and the amount of aflatoxin detected in grains. Minimum GUS expression was related to low aflatoxin production in wounded seeds of two inbreds previously identified in field trials as having moderate-to-high levels of resistance to aflatoxin production. It is suggested that resistance to aflatoxin production is directly related to resistance to fungal colonization in certain genotypes (Brown et al., 1995).

Investigation of five inoculation techniques showed that maize genotypes can be screened in the field for resistance to grain infection by A. flavus using the pinbar (side needle through the husks) and needle in the silk channel inoculation techniques (Zummo and Scott, 1989). Susceptibility of five maize hybrids to inoculum of A. flavus released from waste maize infested with A. flavus gave the same results in screening lines for resistance as did a pinbar inoculation device. This natural method additionally showed that kernel resistance was reflected in reduced silk (Olanya et al., 1995).

A method was developed to study the induction and regulation of aflatoxin biosynthesis by examining the expression of one aflatoxin pathway gene, ver1. The promoter region of ver1 was fused to the beta-D-glucuronidase (GUS) gene (uidA) from E. coli to form the reporter construct GAP13. A. flavus 656-2 was transformed with this construct. Aflatoxin production, GUS activity and transcript accumulation were determined in transformants after shifting the cultures from a non-conducive medium to a medium conducive to aflatoxin biosynthesis. Transformants harbouring GAP13 displayed GUS expression only when aflatoxin was detected in culture. The transcription of the uidA gene driven by the ver1 promoter followed the same profile as for the ver1 genes. The results indicated that the GAP13 construct may be useful as a genetic tool to study the induction of aflatoxin in situ and to identify substances that affect the expression of genes involved in aflatoxin biosynthesis. The utility of this construct to detect inducers of aflatoxin biosynthesis in maize kernels was tested in a bioassay. A heat-stable inducer of aflatoxin with a molecular size of less than 10 kDa was detected in extracts from maize kernels colonized by A. flavus (Flaherty et al., 1995).

Chemical Control

The effect of propionic acid was investigated on mycoflora of rice, sorghum and groundnut during storage at 90% RH. Incidence of A. flavus was greatly reduced in all three seed types (Patkar et al., 1995). During a survey of grain from 146 Swedish farms using acid preservation, aflatoxins were found in grain samples from 41 and 35% of farms using 70 and 85% mixtures of formic acid, respectively; aflatoxins were found in 5% of samples from farms using propionic acid. In laboratory studies, fungal growth and toxin production in moist barley inoculated with aflatoxinogenic A. flavus or A. parasiticus strains were favoured by the presence of formic acid, while propionic acid suppressed both fungal growth and aflatoxin formation (Holmberg and Kaspersson, 1987).

The use of chemicals as a preventive method to control aflatoxin development was investigated in the wet season in Thailand during farm-level maize-cob storage prior to drying and shelling. Dipping and spraying methods using a range of propionic acid concentrations controlled A. flavus and very low aflatoxin levels were found. Chemical residues remained detectable after 2 months' storage, indicating their persistence for control of mould growth (Ilangantileke et al., 1989).

Biological Control

Several experiments were employed to test the role of competition in the ability of an atoxigenic strain of A. flavus to inhibit the aflatoxin contamination of developing cotton bolls. In initial tests, mutants which could not utilize nitrate were used to follow seed infection by toxigenic and atoxigenic strains of A. flavus in coinoculated bolls. Competitive exclusion contributed to the effect of the atoxigenic strain on contamination, but results suggested a second mechanism may also have been in effect. Aflatoxin contamination by the toxigenic strain was similarly inhibited by an atoxigenic strain in vivo and in liquid fermentation, and the atoxigenic strain was equally effective when applied at spore concentrations either equal to or half those of the toxigenic strain. The atoxigenic strain reduced aflatoxin production in vitro when mycelial balls of the two strains were mixed after a 48 h fermentation period, suggesting that close intertwining of mycelia was not required and that aflatoxin biosynthesis could be interrupted even after initiation. The atoxigenic strain did not degrade aflatoxins in vitro, and both culture filtrates and mycelial extracts of the atoxigenic strain simulated aflatoxin production by the toxigenic strain. It is suggested that the atoxigenic strain may interfere with the contamination process both by physically excluding the toxigenic strain during infection and by competing for nutrients required for aflatoxin biosynthesis (Cotty and Bayman, 1993).

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