Training in mould isolation, identification, handling, and evaluation of conditions leading to mycotoxin production

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Consultant U.S. Food and Drug Administration
UNDP/FAO/THA/82/004 Reduction of Post Harvest Losses In Stored Grains



In-depth training was provided to Thai personnel in the proper methods for the isolation, enumeration, handling, and disposal of food borne moulds, and in their identification. Mould species were isolated from several Thai foodstuffs including corn, beans, vegetables, flour, and from Thai soil. More than 24 species of Aspergillus, 12 species of Penicillium and at least 10 other mould genera were detected. Stressed in the identification process were the ability to recognise the macroscopic, the microscopic, and the diagnostic characteristics of each species. Several toxic mould species were found including A. flavus A. parasiticus, A. versicolor, A. nidulans, A. ochraceus, A. terreus, P. citrinum, P. islandicum, and 1:? cyclopium. A number of non-toxic but closely related species were also detected. Limited training was also provided to the employees in the areas of laboratory quality assurance and safety. Regarding Thai corn, investigations were limited since there was virtually no 1986 first crop due to drought. The following recommendations have been made:

  1. The new mycotoxin wing should be finished and become operable as soon as possible.
  2. The professional mycology staff should be given additional training abroad in a well equipped mycology laboratory.
  3. Proper methodology should be stressed. Proper laboratory quality assurance-safety measures should be drawn up and followed in all the laboratories.
  4. A mycological "clean room" should be constructed.
  5. Regarding Thai corn, it should be dried down and kept dry and clean during handling, storage and transit.



The trainees were quite familiar with the direct plating technique. However, a few refinements on their methodology were suggested and were incorporated. One such was the addition of 7.5% sodium chloride to the agar medium, in this case, potato dextrose agar. NaCI is very effective in the inhibition of fast-growing "spreader" moulds such as Mucor, Phizopus, and Trichoderma. Yet, it does not inhibit the growth of other mould species, including the mycotoxin producers. Nor does it change their macroscopic-microscopic morphology, as does some other "spreader" inhibitors such as Rose Bengal. Thus, identification is not compromised. Another suggested refinement to the direct plating technique was the addition to the medium of an antibiotic to inhibit bacterial growth. We used 40 ppm of chlortetracyline-HCL, which I brought with me from the U.S.A. It was needed and it worked. A third suggested refinement was to hold all seeds, including corn, mungbeans, and groundnuts in a freezer for 72 hours prior to direct plating in order to kill possible mites and their eggs. The trainees found that it worked.

With regard to the dilution plating technique, the trainees were not familiar with it, so I introduced it using samples of flour. However, many modifications of the standard method were necessary due to the lack of enough proper glassware, e.g., dilution bottles and pipettes. Yet, the results turned out to be quite informative to the trainees although not quantitatively accurate.



During the first 4 weeks of my consultancy, at least 24 species of Aspergillus, 12 of Penicillium, and 10 other mould genera were detected and studied by the trainees in depth. These species were isolated from Thai foodstuffs and soil and included several toxin producers, i.e., Aspergillus flavus and A. parasiticus (aflatoxins), A. versicolor and A. nidulans (sterigmatocystin) A. terreus and A. clavatus (patulin), Penicillium citrinum (penicillin acid, cyclopiazonic acid, tremorgens, etc.) and P. islandicum (luteoskyrin, etc.). Also detected were several species which are morphologically very similar to toxin producers. For instance, to the untrained eye, Aspergillus oryzee, A. tamarii, and even A. wentii, could be mistaken for A. flavus Likewise, several species of Penicillium are morphologically quite similar to - but diagnostically different from - P. citrinum, e.g., P. chrysogenum and P. oxalicum. The same is true for P. islandlcum, eg., P. variabile, P. aculeatum, P. verruculosum. I am now satisfied that the trainees can now differentiate between these toxic and non-toxic species, and, thus, will be capable of independently determining mould flora profiles of various Thai foodstuffs in the future. The possibility of a refresher course in mould identification for the trainees does bear merit, of course. After 6-12 months of even intensive work in this area, problems still can exist. Mould identification is not easy. It requires patience and determination. Often times it ends in frustration.

Of interest, the mould species detected in the Thai foodstuffs-soils are, except for the genus Aspergillus different from what I have encountered in similar American foodstuffs-soils. This was unexpected. Of particular interest was the general lack of the genera Alternaria and Fusarium and the Penicillium species detected are not the same as those that we find regularly in America.

Listed briefly below are the diagnostic characteristics of a number of mould species Aspergillus flavus: colonies bright green often with black scierotia; conidiophores with usually 2 sets of sterigmata and with spiny stalks. Conidia globose and spiny.

Aspergillus parasiticus: same color as A. flavus but without sclerotia. Condiophore has only one set of sterigmata and stalk is spiny only at the top. Globose spiny conidia are produced.

Aspergillus oryzae: In the A. flavus group but colony color is yellow green. Just one set of sterigmata with smooth conidiophore stalks. Conidia are usually smooth and elliptical and larger than those of A. flavus.

Aspergillus tamarii: In the A. flavus group, but colonies quickly become deep chocolate brown. Conidiophores are the same as A. flavus but conidia are unique, having a double wall and color bars rather than spines.

Aspergillus wentii: Similar in color to A. tamarii, but colonies produce much floccose mycelium and conidiophores are quite long with large vesicles. Conidia are similar to A. tamarii, but with a single wall.

Aspergillus ochraceus: Colonies consistently remain bright yellow and sometimes produce purple sclerotia Conidiophores have 2 sets of sterigmata, the stalk is spiny with definite yellow pigment in the walls. Conidia are usually smooth walled and globose to slightly ellipitical.

Aspergillus versicolor Colonies are dark green but with often highly and variously colored mycelial sectors. Colony reverse is purple. Conidiophores with 2 sets of sterigmata, smooth and colorless stalk, and small vesicles. Conidia slightly roughened.

Aspergillus sydowi: In the A. versicolor group but colonies are consistently blue and produce no highly colored mycelial sectors. Reverse is red. Conidiophores similar to A. versicolor, but conidia are very definitely spiny.

Aspergillus nidulans: Colonies are dark green and produce long columns of conidia that resemble sticks or "cigarettes". Conidiophores and conidia similar to A. versicolor but stalk and vesicle are pigmented brown. The species has a sexual stage and produces cleistothecia surrounded by hulle cells plus bright red ascospores.

Aspergillus fumigates: An important species because it causes human lung infections. Colonies are blue green and produce columns of conidia similar to A. nidulans. However, conidiophores have but one set of sterigmata and conidiophore vesicles and sterigmata are pigmented blue. Colony reverse is also blue to black.

Aspergillus terreus: This species, apparently the most common in Thai soil, is like A. nidulans and A. fumigatus In producing long columns of conidia. However, colony color is tan to beige. Conidiophores are colorless and smooth, have a small vesicle and 2 sets of sterigmata.

Aspergillus glaucus group. Worldwide, perhaps the most common group of aspergilli in stored foodstuffs, the group consists of more than 12 species. However, the species are morphologically quite similar. No need to go beyond the group concept. Species colonies are blue-grey and contain numerous bright shiny yellow balls, which are the cleistothecla, or the sexual stage. Conidiophores are smooth with slight blue pigmentation and one set of sterigmata. Conidia are spiny. The colorless ascospores separate the individual species based on size, shape, and wall adornment.

Penicillium oxalicum: The most common Penicillium in American field corn, and apparently common in Thai field corn, this species should be unmistakeable. Colonies are dark green-black, reverse yellow-green. Conidia are formed in long thin chains which shine like threads of silk under illumination. When an agar plate containing P. oxalicum is tapped, the conidia fall away in crusts or clumps. The conidiophore is assymetrical. The conidia are relatively large, definitely elliptical and smooth.

Penicillium citrinum: Apparently the most common Penicillium in Thailand, it is relatively simple to identify although tremendous variation exists among isolates of this species. Colonies are blue gray, often producing yellow droplets on the surface and yellow in reverse. Conidiophores are bi-verticillate, having only metulae and flask-shaped sterigmata The metulae are not compressed to each other. Rather, there is space between them. Conidia are globose and usually smooth.

Penicillium islandicum: Another common Thai species, it is bi-verticillate, but unlike P. citrinum, the metulae are closely compressed but bear flask shaped sterigmata Conidia are elliptical and smooth. Colonies are orange with green areas of sporulation and produce abundent orange liquid droplets. Reverse is orangebrown.

Pencicillium variabile: This species is dark green with abundant areas of yellow mycelium, especially along colony margins. Its conidiophore differs from that of P. islandicum only in that the sterigmata are long and tapered, not flask shaped.

Penicillium verruculosum: Another biverticillate species, it is quite similar to P. variabile except that it does not produce abundant yellow mycelium and the conidia are globose and spiny.

Penicillium funiculosum Another bi-verticallate species (most Penicillium species in Thailand appear to be bi-verticillate), the colonies are gray and produce definite ropes or funicles of sponulating mycelium on the surface No other Penicillium species does this. Reverse is usually purple. Conidiophores are identical with those of P. variabile.

Penicillium purpurogenum: Another bi-verticillate species, it produces dark green colonies with abundant red droplets and a bright red reverse with the pigment diffusing into the surrounding agar. Its conidiophore is similar to that of P. verruculosum with the spiny globose conidia.

Penicillium cyclopium: This species was only detected once here, but I have drilled on this species since it is the most important Pencillium in storage in America. Colonies are bright blue, granular and usually produce maroon surface droplets and reverse. The colony produces a strong mouldy odor. Conidiophores as assymetrical with a usually spiny stalk. Conidia are globose and slightly roughened.



This area was also stressed. Trainees were taught how to make proper microscope mounts and how to dispose of the slides after use. Also taught were proper ways of inoculating plates and of observing the plates on a daily basis. Dish and lids should never be removed unless one is certain that the mould is harmless. Regarding maintenance, the trainees are routinely keeping the cultures on agar slants. However, with time, many mould species change morphologically when maintained this way and suddenly become unidentifiable. I am recommending that a freeze-dryer be purchased for purposes of proper culture maintenance. Such an appliance is relatively inexpensive and once a culture has been freeze-dried it remains viable for years with little or no morphological change. Another alternative method of maintenance is preservation of spores in sterile soil. In soil, cultures remain relatively stable. However, depending on the species, periods of viability may be relatively short. Another maintenance method would be to immerse the mould propaguies in liquid nitrogen. This is perhaps the most acceptable method, but is also the most costly. Regarding the proper disposal of mould. the only acceptable way is by auto-craving. But I pointed out that if the mold or molds were mycotoxin producers, in addition to growing they undoubtedly were also producing mycotoxins. So after autoclaving, decontamination, with e.g., NaOCI is in order.

Laboratory Safely-Quality Assurance

Although not a "term of reference' upon arrival at the laboratory I noticed a number of safety-quality assurance violations and attempted to point them out to the laboratory leaders. I obtained copies of the FDA, Bureau of Foods, Safety Regulations and of the FDA Division of Microbiology Quality Assurance Regulations (see attachment B). I discussed these documents with the laboratory leaders. They agreed that many violations existed but were in hopes that when the new wing for mycotoxin work became operable, that most of the violations could be corrected. The new wing is finished but has not been occupied due to the lack of air conditioning needed where volatile solvents are kept, laboratory hoods, benches, etc., and equipment, and a needed electronic transformer. I was then violations with the Director General of the Department of Agriculture. He decided that the new Department of Agriculture. He decided that the new wing should become operable as soon as possible. Within 5 days, the air conditioning units arrived and have been installed. We are still waiting the benches, equipment, and transformer.

A primary reason for the violations is the overcrowding that exists in the present facility. The added new space would sharply alleviate the crowded conditions. Yet, additional items will need to be purchased such as fire extinguishers, flame-proof solvent cabinets, clamps for compressed gas cylinders, fire blankets, etc. Also, additional electrical outlets are needed in the present facility. Too many extension cords are used. And special areas for eating lunch should be designated and employees should be urged to wear lab coats and hardtoe shoes when doing bench work.

The most serious quality assurance problem in the laboratory area where the mould identification training is being conducted is severe insect and mite infestation. I brought 11 standard Aspergillus Penicillium cultures with me. We plated them out. But within 72 hours, 6 of the cultures were overrun with bacteria and other mould species due to infestation with ants. We were able to construct a temporary structure that has, so far, prevented additional infestation, stackable shelves whose bottom legs are immersed in dishes of glycerine, thus trapping insects and mites. But this is only temporary. What is urgently needed once space becomes available is the construction of a mycological "clean room". That is, a room that is physically isolated from the rest of the laboratory facility, that is sealed to prevent insect-mite infestation, and that is used only for culturing, identifying, and storing mould species. The room should be sprayed weekly with insecticide and should be restricted to selected personnel. Plans are under way for the construction of such a room.

Moulds Detected In Thai Corn

Although corn samples were in limited supply, we had access to a few samples. Table 3 lists our results from 5 samples of "excellent quality" shelled corn that had been stored for 8 months at 10C. When placed in storage, this corn was of high quality with very low levels of aflatoxin. Initial mould floras had not been determine for this corn. Table 3 shows, however, that the corn had become extremely mouldy, 100% before surface disinfection and 87% after. In addition, A. flavus was the predominant species. This was unexpected since A. flavus grows poorly below 12C But I was informed that the 10C room sometimes loses power and warms up, thus, resulting in water condensation on the corn. In addition, the corn was infested with mites which are capable of mechanically transmitting mould propagules, from seed to seed and from plate to plate.

Table 4 lists results of the few samples of freshly harvested, wet season, corn that we were able to obtain. Of interest, although A. flavus was present, its levels were lower Rather, the A. glaucus group seemed to predominate. But, mites were also detected in these "fresh" samples. Thus, the mould flora can be expected to change.




Mould proliferation and subsequent metabolise (mycotoxin) production is dependent upon 3 necessary and inter-related factors, namely: 1. the physical presence of the generating organism; 2. a substrate suitable for growth; 3. an environment suitable for growth. All 3 factors must be met in order for growth and toxin production to occur. These 3 factors are discussed below and include some comments, based upon my 6-week stay in Thailand.

Physical Presence of the Generating Organism

As tables 1 and 2 show, many toxic mould species are present in Thailand. Although their sources are numerous, the chief source of the organisms would appear to be the soil, the air, former crop organic material, and contaminated storage facilities. If present, these moulds can be spread by the air, by insects, mites, higher animals, and even facility personnel. For instance, upon one field trip here, I observed that a number of animals were present in the areas of the drying corn, including dogs, cats, and chickens. I also noted dried organic material, possibly from an earlier crop, lying nearby. In my laboratory studies, large populations of mites were detected along with lower levels of insects. And on my field trip I watched personnel indiscriminantly walking over the drying corn with shoes on. All the above are potential sources of the spread of the mould. Granted, there is no way to eliminate the presence of these toxic mould species in Thailand, but ways do exist to lessen their spread and contamination potential. For instance, where corn is being dried there should be no freewalking animals around nor any indiscriminatingly walking humans. Nor should there be any residues of earlier crops. And after drying, just prior to placing in storage, the corn should be fumigated to eliminate mites and insects.

Regarding storage facilities, although I did not visit any here since most were empty due to this season's crop failure in the field, they can be an Important source of mould presence if not kept clean. Thus, these facilities should routinely be cleaned up between storage loads, including fumigation. In addition, vehicles used to transport these foodcrops should also be kept clean. There is nothing to be done about the presence of toxic moulds in Thailand, but things can be done to alleviate their spread.

Substrate Suitable for Growth

Although most of the toxic moulds I detected in Thailand could conceivably grow on any Thai foodstuff, I doubt that this really is the case. We know that A. flavus thrives on corn and groundnuts. But it was rare on fresh vegetable crops. However, A. versicolor was the principal toxic species detected on black beans. Thus, it should be within the interests of my trainees to begin determining mould floras of Thai foodstuffs other than corn and groundnuts. For instance, A. versicolor may indeed be the chief toxic mould in black beans. Thus, the possibility of sterigmatocystin. Determining mould floras of individual substrates is indeed important when considering the overall picture of mycotoxin contamination.

Environment Suitable for Growth

The most effective means to control and prevent aflatoxin contamination of corn and groundnuts, and the mycotoxin contamination of any foodstuff, is the ability to control the environment so as to prevent mould growth. Several environmental parameters are involved, including:

1. Atmosphere All mold species are obligate aerobes, and cannot grow in the absence of free oxygen. Although probably not economically feasible, one sure way to prevent mycotoxin contamination of corn, groundout, and small grains is to store them under anaerobic conditions, e.g. CO2 or nitrogen. For instance, this could be done in large airtight silos. High moisture would be no problem. The moulds just would not grow But for the farmer and the middleman, this type of environment control is understandably unrealistic.

2. Temperature In my experience, virtually all of the mycotoxin producing species I detected in Thailand grow poorly, it at all, at 10C. Thus, another environmental way to control or prevent mycotoxin production is a storage temperature of 10C or less. However, low temperature facilities are virtually nonexistent. Thus, temperature control is not the answer.

3. Moisture Without question, moisture control is the best and most economical means to control the environment to prevent mould growth and mycotoxin production. To reiterate the comments of previous consultants, Mr. Nesheim, Mr. Ware, and Dr. Smalley, corn must be dried down to 14.5% moisture, wet weight basis, to avoid aflatoxin contamination. The same for groundnuts. Certain of the small grains require even lower moisture contents, cat 13%. The problem in Thailand is that these desired moistures are not achieved, or at least, not quickly enough. Regarding corn, the farmer often harvests the crop when moisture content to too high, e.g., 25-35% or more, wet weight basis. Upon harvest, the farmers may attempt to sundry the corn on plastic sheets or concrete slabs-which may be effective with the dry season second harvest or they may deliver it directly to the buyer at its original field moisture, if for no other reason than need of the money. If the farmer could be convinced to delay harvesting of the corn, that is, if the ears were allowed to remain on the stalks in the field for an additional 1, 2, or even 3 weeks (especially the first, wet season, crop), the moisture content of the corn would dry down naturally, maybe to as low as 18-22%. And rain would not significantly effect this natural drying in the field. Sundrying then could effectively bring the corn moisture close to the needed 14.5%. However, even after acceptable drying, the corn must be handled-by the farmer, the buyer, the exporter - in a manner that will not allow the moisture to return. It should never be exposed to free water and manner of transportation of the corn should be such as to minimize actual water condensation. For instance, corn transportation in trucks, railcars, and even water going vessels may be subject to moisture build-up due to condensation if transport time is lengthy. Hot day time temperatures followed by cooler nightime temperatures invariably will cause the corn to "sweat", thus, free water available for mould growth. And once a mould has begun to grow, it does not dissipate the water. It just transfers it. Thus, proper handling and shipment of the corn, even after acceptable drying, is essential in order to avoid moisture build-up, mould growth and toxin formation. To the farmer, to the buyer, to the exporter, to the importer, transport must be proper and rapid. And as referred to in the section, "Physical Presence of the Generating organism", transportation vehicles should be clean. No leftover mould or bacteria, no excrete, no unusual amounts of soil or organic debris.



  1. Barnett, H. L.1972. Illustrated Manual of Imperfect Fungi. Burgess Publishing Co., Minneapolis.
  2. Eillis, M. B.1971. Dematiaceous Hyphomycetes. Commonwealth Mycological Institute. Kew, Surrey, England.
  3. Gilman, J. C. 1959. A Manual of the Soil Fungi. Iowa State University Press, Ames, Iowa.
  4. Mislivec, P. B. 1978. The Genus Penicillium. In T. Wyllie and H. Morehouse (ED). Mycotoxic Fungi, Mycotoxins, and Mycotoxicoses. Marcel Dekker, NY.
  5. Nelson, P., T.A. Toussoun, and W. Marasas. 1984. Illustrated Manual of the Genus Fusarium. Pennsylvania State University Press, State College, Pennsylvania.
  6. Raper, K. and D. Fennell. 1965. The Genus Aspergillus. Williams and Wilkins Press. Baltimore, MD.
  7. Raper, K., and C. Thom. 1949. A Manual of the Penicillia. Williams and Wilkins Press, Baltimore, MD.
  8. U. S. Food and Rrug Administration. 1984. Bacteriological Analytical Manual. Association of Official Analytical Chemists, Arlington, VA.

Table 1. Mould species Detected In Various Thai Foodstuffs (maize, groundauts, mungbeans, fresh vegetables)

The Genus Aspergillus The Genus Penicillium

A. aculeatus P. aculeatum
A. amstelodami P. chrysogenum
A. cadidus P. citrinum
A. chevalier) P. cyclopium
A. flavus P. funiculosum
A. flavus var. co/umnaris P. implicatum
A. fumigatus P. islandicum
A. niger P. oxalicum
A. ochraceus P. purpurogenum
A. oryzae P. variabile
A. parasiticus P. verruculosum
A. repens  
A. restrictus  
A. sydowi  
A. tamarii  
A. terreus  
A. versicolor  
A. wentii  
Other Mould Genera:  
Alternaria a/temata  
Cephalosporium spp.  
Cladosporium spp.  
Curvularia spp.  
Fusarium spp.  
Paecilomyces varioti  
Pestalotia spp.  
Rhizactonia spp.  
Rhizopus nigricans  
Syncephalastrum spp.  
Trichoderma viride  


Table 2. Mould Species Detected in 16 Soil Samples Collected from Thai Maize Farms

Mould Species Number of Samples Positive
Aspergillus aculeatus 5
A. candidus 8
A. clavatus 1
A. flavus 12
A. flavipes 7
A. fumigatus 7
A. glaucus group 3
A. nidulans 13
A. niger 15
A. ochraceus 1
A. tamarii 2
A. terreus 14
A. terrcus var. africanus 2
A. ustus 3
A. versicolor 1
Chaetomium spp. 1
Cladosporium herbarum 1
Mucor spp. 2
Penicillium citrinum 14
P. funiculosum 1
P. Iuteum 1
P. variabile 3
P. verruculosum 2
Rhizopus nigricans 4
Syncephalastrum spp. 2
Trichoderma viride 2


Table 3. Mould Flora of Thai Corn Kernels after 8 Month's Storage at 10C

(% Mouldiness = 100%)
Surface Disinfected
(% Mouldiness = 87%)
Species Detected Species Detected(a)
Aspergillus flavus: 100% Aspergillus flavus: 52%
A. glaucus group: 3% A. glaucus group: 19%
A. niger: 15% A. niger: 8%
Penicillium citrinum: 9% A. tamarii: 2%
  A. wentii: 3%
  Cephalosporium spp.: 3%
  Mucor spp.: 3%
  Penicillium citrinum: 10%
  Phizopus nigficans: 1%

(a) The following species were detected at a rate of less than 1%: Aspergillus aculeatus; A. candidus; A. fumigates; A. nidulans; A. restrictus; A. terreus; Penicillium chrysogenum.

Table 4. Mould Flora of Freshly Harvested 1986 Thai Corn Kernels

(% Mouldiness = 100%)
Surface Disinfected
(% Mouldiness = 88%)
Species Detected Species Detected
Aspergillus flavus: 88% Aspergillus flavus: 29%
A. glaucus group: 39% A. glaucus group: 55%
A. niger 24% A. niger 8%
A. fumigates: 2% A. fumigates: 1%
A. wentii: 2% A. wentii: 5%
  A. tamarii: 2%
  A. aculeatus: 1%
  A. terreus: 1%
  Penicillium citrinum: 5%
  P. oxalicum: 1%
  Rhizopus nigricans: 3%
  Cephalosporium spp.: 1%
  Syncephalastrum spp.: 1%

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