REVIEWARTICLE

Microbial Detoxification of Mycotoxins

Susan P. McCormick

Received: 24 April 2013 /Revised: 24 June 2013 /Accepted: 28 June 2013 /Published online: 12 July 2013# Springer Science+Business Media New York (outside the USA) 2013

Abstract Mycotoxins are fungal natural products that aretoxic to vertebrate animals including humans. Microbes havebeen identified that enzymatically convert aflatoxin, zeara-lenone, ochratoxin, patulin, fumonisin, deoxynivalenol, and T-2 toxin to less toxic products. Mycotoxin-degrading fungi andbacteria have been isolated from agricultural soil, infested plantmaterial, and animal digestive tracts. Biotransformation reac-tions include acetylation, glucosylation, ring cleavage, hydro-lysis, deamination, and decarboxylation. Microbial mycotoxindegrading enzymes can be used as feed additives or to decon-taminate agricultural commodities. Some detoxification geneshave been expressed in plants to limit the pre-harvest mycotox-in production and to protect crop plants from the phytotoxiceffects of mycotoxins. Toxin-deficient mutants may be usefulin assessing the role of mycotoxins in the ecology of themicroorganisms.

Keywords Mycotoxin . Fumonisin . Aflatoxin .

Trichothecene . Patulin . Ochratoxin . Microbialdegradation . Biotransformation

Introduction

Mycotoxins are a structurally disparate group of fungal naturalproducts that share the ability to cause harm to vertebrate animalor human health when they are contaminants of animal feed orfood. The termmycotoxinwas first used in the 1960s to describethe toxin associated with contaminated peanuts in animal feedand the loss of turkeys in England (Turkey X disease). Thismycotoxin was later identified as the Aspergillus flavus toxinaflatoxin B1. The term was later applied to other toxic fungal

natural products (Bennett and Klich 2003), including fumonisinB1, zearalenone, T-2 toxin, deoxynivalenol, patulin, and ochra-toxin. Regulatory agencies have set action or advisory levels formycotoxins, and food and animal feed aremonitored to keep thetoxins out of the food supply. For example, the current EuropeanUnion levels for aflatoxin B1 are 2 ppb for food and 20 ppb foranimal feed. Much of the work on mycotoxins has focused onthe biosynthetic pathways, detectionmethods, and toxicology ofthe toxins.

The functions of mycotoxins in the producing fungi or theirecological roles in interactions with other organisms or theirenvironment have not been well studied (Bu’Lock 1980).Janzen (1977) hypothesized that toxin-producing fungi com-pete with large organisms for resources such as fruit or seedsby making them unpalatable, but may also “wage antibioticwarfare” that limits microbial competition for resources.There also is evidence that mycotoxins act as chemical signalsbetween species, as anti-insectans that protect sclerotia frominsect predation, as virulence factors in plant diseases, or asinhibitors of quorum sensing (Ciegler 1983; Desjardins et al.1996; Rasmussen et al. 2005; Rohlfs et al. 2010; Wicklowet al. 1996). For example, gram-negative bacteria use a coor-dinated response to bacterial population density (quorumsensing) and form biofilms that are resistant to antimicrobialtreatments. Patulin, a mycotoxin produced by Penicillium,inhibits quorum sensing and blocks chitinase production thatthe bacteria could use to attack the fungus (Rasmussen et al.2005). Some insight into the functions of mycotoxins may begained by looking at their cellular targets. Riley and Goeger(1992) proposed some possible ecologically relevant func-tions for cyclopiazonic acid, a mycotoxin produced byAspergillus flavus, based on its activity in animal systems,This mycotoxin may alter calcium homeostasis, and maychange plasma membranes and mitochondria. However, asnoted by Utermark and Karlovsky (2007), activity in verte-brate animals may not accurately predict the ecological func-tion of the toxin in the fungus or its interactions with other

S. P. McCormick (*)Bacterial Foodborne Pathogens and Mycology Research Unit,USDA-ARS-NCAUR, Peoria, IL 61604, USAe-mail: Susan.McCormick@ars.usda.gov

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organisms. This is particularly true of the mycotoxinzearalenone, which has hormonal activity in animals but nodocumented hormonal function in the producing fungus.

Mycotoxin-degrading bacteria and fungi have been foundin environments likely to contain mycotoxins. In some cases,selection media with a mycotoxin or structurally similar com-pounds as the sole carbon source have been used to isolatemycotoxin-degrading organisms. For example, microbes thatcould degrade the Fusarium mycotoxins fumonisin andzearalenone were isolated from Fusarium graminearuminfested wheat and corn, silage and compost (Duvick et al.1998, 2003; Duvick and Rood 1998, 2000). Other microbesthat biotransform or degrade mycotoxins have been found insoil collected from agricultural fields (Shima et al. 1997) or inthe digesta of animals (Guan et al. 2008). Bacteria or fungi inthese environments may have enzymes that protect them fromthe mycotoxins or allow them to compete with mycotoxin-producing fungi. Microbes in the digesta may help animalstolerate mycotoxins in their food. Mycotoxin-degrading bac-teria and fungi also have been found in soil contaminated withpolycyclic hydrocarbons (Hormisch et al. 2004; Teniola et al.2005). Organisms from these environments may recruit en-zymes that detoxify the pollutants for detoxification of myco-toxins that are structurally similar to the pollutants.

Levels of some mycotoxins, are diminished by physicalbinding rather than degradation. Lactic acid bacteria (e.g.,Lactobacillus sp.), gram positive bacteria that produce lacticacid as a product of carbohydrate fermentation, have beenreported to extracellularly bind zearalenone, trichothecenes,and aflatoxins (El-Nezami et al. 2002; Haskard et al. 2001;Zou et al. 2007). This type of weak non-covalent bindingmay be associated with hydrophobic pockets on the bacterialsurface.

Elimination of mycotoxins is the goal of food safety pro-grams. Current methods towards this goal include breedingfor fungal resistant varieties of plants, chemical treatmentswith ammonia or lime, competitive exclusion of toxigenicfungi, control of post-harvest seed moisture, use of fungi-cides or preservatives to prevent fungal growth, controllinginsect infestations of stored grains with insecticides, andsorting of seeds to remove those contaminated with myco-toxins. Identification of microbes that can degrade myco-toxins may lead to new or more effective way to eliminatemycotoxins from food and feed. This review will focus onthose mycotoxins for which microbial biotransformations,detoxifications, or degradations have been described.

Aflatoxins

Aflatoxin B1, aflatoxin G1 (Fig. 1) and related compoundsare difuranocoumarins produced by Aspergillus flavus and A.parasiticus and other species of Aspergillus. Aflatoxins are

associated primarily with oilseed crops including corn, pea-nuts, tree nuts, and cottonseed. They are potent carcinogensand mutagens due to their ability to form adducts with DNA.Most studies on biotransformation of aflatoxin have focusedon aflatoxin B1 which usually is produced in large quantityby A. flavus, and is the most potent naturally occurring livercarcinogen known. Ingestion of high levels of aflatoxin re-sults in liver damage including cirrhosis, bile duct prolifera-tion, edema, and death (Williams et al. 2004). In insects,consumption of aflatoxins can slow development and in-crease mortality (Zeng et al. 2006). However, some insectsare relatively tolerant of aflatoxin. For example, the navelorange worm (Amyelois transitella) avoids the adverse ef-fects of Aspergillus toxins in contaminated tree nuts and canbiotransform aflatoxin to less toxic metabolites (Niu et al.2009).

Aflatoxins are not virulence factors in plant disease. Anon-toxigenic A. flavus strain that is adapted to the agricul-tural environment has been used successfully to competitive-ly exclude aflatoxigenic A. flavus from cottonseed (Cottyand Mellon 2006; Ehrlich and Cotty 2004).

Aflatoxins are fluorescent compounds. B aflatoxins thatfluoresce blue have a lactone-cyclopentanone structure,while G aflatoxins that fluoresce green have two lactones.Ammoniation, which is an allowed method for remediationof aflatoxin-contaminated cottonseed, causes opening of thelactone ring, loss of fluorescence, and reduction in toxicity(Lee et al. 1981). Microbial biotransformations that result inloss of fluorescence can be attributed to changes to thelactone ring. However, degradation products that are morewater-soluble may escape extraction with organic solvents ordetection with methods that rely on the fluorescence of themolecule, making decomposition difficult to detect in somecases. A number of bacteria and fungi, including Rhizopussp., Dactylium dendroides (now called Hypomyces rosellus),and Cornybacterium rubrum (Mann and Rhem 1976; Coleet al. 1972) convert aflatoxin B1 to aflatoxicol (Fig. 1) byreduction of the C-3 keto on the cyclopentanone ring.Aflatoxicol was less toxic than aflatoxin B1 in a ducklingbile duct hyperplasia assay (Detroy and Hesseltine 1970).

Extracellular enzymes from an edible fungus, Armilleriellatabescens, converted aflatoxin B1 to a product with greatlyreduced mutagenic and toxicological activity. The degradationproduct was fluorescent, suggesting that the enzymes causedchanges to the difuran rather than the lactone portion of themolecule (Liu et al. 1998). Extracellular enzymes from thewhite rot fungus Pleurotus ostreatus (Motomura et al. 2003),also degrade aflatoxin. White rot fungi degrade lignin and alsomay have activity against polyaromatic hydrocarbons(Motomura et al. 2003). The aflatoxin-degrading abilities ofPleurotus ostreatus and another white rot fungus, Peniophorasp., were discovered in a screen for fungi with high levels ofcopper-containing polyphenol oxidases (laccases) that degrade

908 J Chem Ecol (2013) 39:907–918

the polymeric dye poly R-478 (Alberts et al. 2009). Purelaccase from another mushroom, Trametes versicolor, as wellas Aspergillus niger producing recombinant laccase fromTrametes, also degraded aflatoxin (Alberts et al. 2009).

An actinomycete bacterium, Flavobacterium aurantiacum(now called Nocardia corynebacteriodes) that could enzymat-ically degrade aflatoxin B1 was identified in a large screen of aculture collection of fungi, bacteria, yeast and algae (Ciegleret al. 1966; Smiley and Draughon 2000). Aflatoxin-degradingactivity has been found in other Actinomycetes, often thoseassociated with the ability to degrade pollutants. For example,Mycobacterium fluoranthenivorans, a soil bacterium isolatedfrom the site of a former coal gas plant, was able to utilize thepolyaromatic hydrocarbon fluoranthene as a sole carbonsource, and efficiently degraded aflatoxin B1 (Hormisch et al.2004).

Extracellular extracts of Rhodococcus erythropolis, a grampositive bacterium that can also break down polycyclic aromat-ic hydrocarbons, degraded aflatoxin to a product that was lessmutagenic and had the loss of fluorescence characteristic ofdisruption of the lactone ring (Alberts et al. 2006,2009; Teniolaet al. 2005). The mechanism for aflatoxin B1 and G1 degrada-tion in another Actinomycete,Mycobacterium smegmatis, wasrecently described. Degradation begins with reduction of theα,β-unsaturated double bond between C-2 and C-6 (Tayloret al. 2010) by a F420H2-dependent reductase, followed byspontaneous breakdown of the product (Lapalikar et al. 2012;Taylor et al. 2010).

A rich source of aflatoxin-degrading bacteria is animalfeces. Aflatoxin B1 is a furanocoumarin, and bacteria thatcan degrade other coumarins or furanocoumarins may be ableto degrade aflatoxin B1. Guan et al. (2008) isolated bacteriafrom farm soil and the feces of zoo animals, including

Bacillus, Enterobacter, Rhodococcus, Stenotrophomas, andBrachybacterium, that were able to utilize the simple phyto-chemical coumarin as a sole carbon source, and found thatmost of the isolates could also degrade aflatoxin. For example,Stenotrophomonas maltophilia, a bacterium isolated fromtapir feces, degraded over 80 % of the aflatoxin within 72 hr.Stenotrophomonas maltophilia also is able to degrade poly-cyclic aromatic hydrocarbons (Juhasz et al. 2003). No afla-toxin metabolites were identified so the mechanism for thisdegradation is not known.

Zearalenone

Zearalenone (Fig. 2) is a phenolic resorcyclic acid lactonethat can be a contaminant of cereal crops infected byFusarium species. Zearalenone has relatively low acute an-imal toxicity (Zinedine et al. 2007). Because of structuralsimilarities to the female hormone estradiol, zearalenonecauses hyperestrogenism in swine and decreased fertility incows. Structural features important for the biological activityof zearalenone include the lactone ring and the C-4 hydroxylgroup (el-Sharkawy and Abul-Hajj 1988b). Activity usuallyis assessed by assays that measure binding to an estrogenreceptor. Loss of estrogenic activity often is reported asdetoxification.

As noted by Karlovsky (2008), antibacterial or antifungalactivities that could suggest ecological functions forzearalenone may not have been considered because of thedocumented hormonal activity of zearalenone in animals.Zearalenone inhibits the growth of filamentous fungi(Utermark and Karlovsky 2007) and may help Fusariumspp. compete with other fungi for resources (Karlovsky 2008).

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Fig. 1 Aflatoxins B1, aflatoxinG1, and biotransformationproducts of aflatoxin B1

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Reported fungal transformations of zearalenone include re-duction of carbonyl ketones, reduction of double bonds, hy-droxylation, methylation, sulfation, glycosylation, and ringcleavage. Several microorganisms, including Rhizopus sp. andAlternaria alternata, reduced the C-6′ keto of zearalenone toproduce α-zearalenol and β-zearalenol, products that are notsignificantly less estrogenic than zearalenone (el-Sharkawyet al. 1991; Kamimura 1986). Aspergillus ochraceous conver-ted zearalenone to α-zearalanol and β-zearalanol (el-Sharkawyand Abul-Hajj 1988b), the former of which is more estrogenicthan zearalenone.

Modification of the C-4 hydroxyl group of zearalenonecan result in loss of estrogenic activity. For example,Cunninghamella bainieri converted zearalenone to 2,4-dimethoxyl zearalenone (el-Sharkawy and Abul-Hajj 1988b),Rhizopus arrhizus converted zearalenone to zearalenone4-sulfate (el-Sharkawy et al. 1991), and Rhizopus sp.,Thamidium elegans, andMucor bainieri converted zearalenoneto zearalenone- 4-β-D-glucoside (el-Sharkawy and Abul-Hajj1987; Kamimura 1986). None of these products was estrogenic.

Complete degradation of zearalenone has been observedwith a mixed culture of soil microorganisms collected fromthe site of a gas coal plant, but no individual isolates retained theability to break down zearalenone (Megharaj et al. 1997).Degradation of zearalenone also has been reported with speciesof Clonostachys, Rhodococcus, Norcardia, and Trichosporum(Duvick and Rood 1998; el-Sharkawy and Abul-Hajj 1988a;Molnar et al. 2004).Norcardia and Rhodococcuswere found ina screen of microbes from moldy corn that could usezearalenone as a sole carbon source (Duvick and Rood 1998).

Two fungal biotransformations involving cleavage ofzearalenone have been identified. Gliocladium roseum (nowcalled Clonostachys rosea), a mycoparasitic fungus used forbiocontrol of wilt diseases, efficiently detoxified zearalenoneby ring opening followed by spontaneous decarboxylation (el-Sharkawy and Abul-Hajj 1988a; Kakeya et al. 2002) (Fig. 2).

The lactonohydrolase for this detoxification has been purifiedand the gene (zhd101 or zes2) encoding the enzyme has beencloned (Karlovsky et al. 2004; Takahashi-Ando et al. 2002).Inactivation of the lactonase gene in Gliocladium (Utermarkand Karlovsky 2007) increased its sensitivity to zearalenone.In addition, expression of zhd101 in plants reduces zeara-lenone contamination of grain (Igawa et al. 2007) or yeast(Takahashi-Ando et al. 2004).

A second type of ring cleavage of zearalenone has beenfound in a yeast strain associated with termites, Trichosporummycotoxinivorans (Molnar et al. 2004). This yeast convertszearalenone to the cleavage product, ZOM-1 (Fig. 6) (Vekiruet al. 2010). A two step mechanism for zearalenone degrada-tion was proposed, comprising a Baeyer-Villiger type ringexpansion to give a lactone intermediate followed by cleavagewith a lactonase (Fig. 2), but neither the enzymes nor genesencoding the enzymes have been identified.

Citrinin and Ochratoxin

Citrinin and ochratoxin A (Fig. 3) are nephrotoxic myco-toxins that sometimes co-occur. Toxicity of citrinin has beenfound in poultry. Ochratoxin is associated with kidney dis-ease in dogs, rats, and swine, and also has been implicated ina human kidney disease known as Balkan endemic nephrop-athy (Castagnero et al. 2006). Ochratoxin and citrinin mayinterfere with the uptake of iron by other microbes andthereby compete for nutrients and space (Størmer andHøiby 1996).

Citrinin (Fig. 3), an antibacterial polyketide-derived my-cotoxin produced by Aspergillus, Penicillium, andMonascus(Shimizu et al. 2005), can contaminate grain and other foods.Citrinin was first identified as an antibiotic, and also dam-ages DNA. Citrinin can be introduced to foods viaMonascuspurpurea, an edible red yeast with cholesterol-lowering

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activity that is used as a food additive. The bacteriumMoraxella sp., identified in a screen of microbes that tolerateorganic solvents, converted citrinin to decarboxycitrinin(Devi et al. 2006). Decarboxylcitrinin, which is also theproduct of heat treatment of the mycotoxin, retained antibi-otic activity (Devi et al. 2006) but was not toxic to mice(Jackson and Ciegler 1978). No other microbial biotransfor-mations of citrinin have been reported.

Ochratoxin A (OTA) (Fig. 3) is a chloro-isocoumarin pro-duced by Aspergillus ochraceus, A. ostianus, and Penicilliumverricosum. This mycotoxin can contaminate many agriculturalcommodities including cereals, coffee, and grapes (Kuiper-Goodman and Scott 1989). Ochratoxin may be synthesized tomaintain chlorine levels (Schmidt-Heydt et al. 2012).

Yeast, including Saccharomyces,Rhodotorula,Cryptococcus,and Trichosporon mycotoxinivorans, can hydrolyze the amidebond of OTA to produce nontoxic products phenylalanine andochratoxinα (Fig. 3) (Abrunhosa et al. 2010;Molnar et al. 2004;Schatzmayr et al. 2003). It is interesting that Trichosporonmycotoxinivorans did not attack the lactone of ochratoxin sincelactone cleavage is the mode of detoxification of zearalenone bythis yeast (Vekiru et al. 2010). A bacterium, Phenylobacteriumimmobile, which can use phenylalanine as a sole carbon source,converted ochratoxin A to ochratoxin α and three other metab-olites resulting from the breakdown of the phenyl moiety ofphenylalanine (Wegst and Lingens 1983).

Fumonisins

Fumonisin B1 (Fig. 4) and related compounds are polyketide-derived mycotoxins produced by Fusarium verticillioides and

F. proliferatum. These fungi cause corn ear rot and contami-nated corn or corn products are the most common sources offumonisin in the diet. The mycotoxin has two tricarballylateside groups attached to a 22 carbon aminopentol (Gelderblomet al. 1988). Fumonisins are structurally similar to the sphin-golipid bases sphinganine and sphingasine and they disruptsphingolipid metabolism (Merrill et al. 1993). Toxic effectsassociated with exposure to fumonisins include equineleukoencephalomalacia (Ross et al. 1992), porcine edema(Ross et al. 1992), and esophageal (Myburg et al. 2002) andliver cancer (Gelderblom et al. 2001). Structure-function stud-ies have shown that both the tricarballylate groups and the freeamino group are important for animal toxicity (Abbas et al.1993).

Other organisms, including plants, bacteria and fungi, havesphingolipids that may be fumonsin targets. Disruption ofsphingolipidmetabolismmay also account for the phytotoxicityof fumonisins and related compounds (Abbas et al. 1994).Fumonisins also have been reported to inhibit the growth offungi including Alternaria alternata, Botrytis cinerea, andPenicillium expansum (Keyser and Vismer 1999).

Bacteria and fungi that can degrade fumonisins have beenisolated from soil and from field-grown corn kernels(Benedetti et al. 2006; Duvick et al. 2003; see also Falardeauet al. 2013). These microbes were able to use fumonisin as asole carbon source. The first step of fumonisin detoxification,in both bacteria and fungi (Blackwell et al. 1999; Duvick et al.2003; Heinl et al. 2010), is the removal of tricarballylategroups at C-14 and C-15 with carboxylesterases, convertingFumonisin B1 (FB1) to hydrolyzed fumonisin B1 (HFB1)(Fig. 4). The carboxylesterase from the fumonisin-degradingbacterium Sphingopyxis sp. has been commercialized as a feed

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additive (FUMzyme®) by Biomin. HFB1 also is producedwhen fumonisin-containing corn is treated with lime(nixtamalization) (Palencia et al. 2003).

In bacteria, hydrolysis of the two side chains is followedby deamination catalyzed by an aminotransferase to produce2-keto HFB1 (Fig. 4) (Hartinger et al. 2011; Heinl et al.2010, 2011). In Sphingopyxis, the carboxylesterase gene(fumD) and the aminotransferase gene (fumI) are within acatabolic gene cluster that also contains transcriptional reg-ulatory genes, transporters, and other genes that may beinvolved in fumonisin degradation (Hartinger et al. 2011;Heinl et al. 2010, 2011).

Black yeasts isolated from field grown corn kernels canuse fumonisin as a sole carbon source (Duvick 2001; Duvicket al. 1998, 2003). In the black yeast Exophiala spinifera,hydrolysis of the tricarballylic side chains is followed bydeamination by an amine oxidase to produce 2-keto HFB(Blackwell et al. 1999; Duvick 2001; Duvick et al. 1998;2003). 2-keto HFB1 may be converted to hemiketal deriva-tives by cyclization of the hydroxyl at C-5 and the C-2carbon (Blackwell et al. 1999; Hartinger et al. 2011).

Patulin

Patulin (Fig. 5) is a polyketide lactone produced by speciesof Penicillium expansum, an important postharvest pathogenof fruits, and other species of Penicillium, Aspergillus, andByssochylamys. Patulin is a mycotoxin usually associatedwith contaminated fruit or fruit juices leading to food safetyconcerns for apple juice (González-Osnaya et al. 2007).Acute exposure to patulin can cause nausea and gastritis

(Castoria et al. 2011), while chronic exposure can causeneurotoxic, immunotoxic, teratogenic, and carcinogenic ef-fects (Moake et al. 2005). Patulin was first isolated as part ofan antibiotic screening program (Puel et al. 2010). It formscovalent adducts with thiol-containing compounds, such ascysteine or glutathione, and can disable enzymes that containsulfhydryl groups in their active sites (Castoria et al. 2011;Fliege and Metzler 2000), but may also disrupt membraneintegrity, protein synthesis, RNA synthesis, and other cellu-lar functions. Both the hemiacetal and the lactone ring areimportant for toxicity (Wallen et al. 1980).

Approaches used to limit patulin contamination in foodsand juices include culling visibly damaged fruit, cutting outthe damaged portions of the fruit, and the use of fungicides.Alternatives to fungicides are biocontrol microbes that in-hibit the fungus and thereby production of the patulin.

Although patulin is a potent antibiotic that may have a role inmicrobial competition, some yeast and bacteria can degradepatulin. Saccharomyces cerevisiae converts patulin to (E)-ascladiol by opening the pyran ring (Fig. 5) (Moss and Long2002). Fermentation of fruit juice to wine or vinegar is aneffective way to remove patulin (Stinson et al. 1978). Thegram-negative bacterium Gluconobacter oxydans, isolatedfrom rotted apple puree containing high levels of patulin,converts patulin to (E)-ascladiol (Ricelli et al. 2007). (E)-Ascladiol is the immediate biosynthetic precursor of patulin(Sekiguchi et al. 1983) and is 25 % as toxic as patulin. Likepatulin, it can form adducts with thiol groups (Coelho et al.2008; Stinson et al. 1978). It has been isolated from Aspergillusclavatus (Suzuki et al. 1971).

Some yeast that inhibit the growth of patulin-producingfungi also have been found to efficiently degrade patulin,

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including Pichia ohmeri, isolated from ant hills, Pichiamembranifaciens, and Sporobolomyces roseus (Coelho et al.2006; Levy et al. 2002). However, no degradation productshave been identified. The mechanism for patulin detoxifica-tion was identified in the biocontrol yeast, Rhodosporidumkratochvilovae (Castoria et al. 2011), which converts patulinto desoxypatulinic acid (Fig. 5). This compound has also beenisolated from Penicillium patulum (Scott et al. 1972). It lacksboth hemiacetal and lactone functions and is unable to reactwith thiol groups. Desoxypatulinic acid was much less toxicthan patulin in an assay with human lymphocytes (Castoriaet al. 2011).

Trichothecenes

Trichothecenes are a group of sesquiterpenoid myco-toxins produced by Fusarium sp. and other members ofthe Hypocreales, including Trichoderma, Trichothecium,Stachybotrys, Myrothecium, and Spicellum. Trichothecenesinhibit eukaryotic protein synthesis by preventing peptidebond formation. Some trichothecenes are virulence factors inplant diseases, such as deoxynivalenol in Fusarium head scabof cereals (Desjardins et al. 1996). Trichothecenes such asdeoxynivalenol, T-2 toxin, and diacetoxyscirpenol can becontaminants of agricultural products including corn, wheat,and potato. Exposure to trichothecenes can cause emesis, feedrefusal, hemorrhaging, and dermatitis in vertebrate animals(Bennett and Klich 2003).

Fusarium trichothecenes are characterized by an oxygenfunction at C-3, and are grouped based on the oxygen func-tion at C-8. Type A trichothecenes (e.g., T-2 toxin; Fig. 6)have a hydroxyl or acyl group at C-8, while type B tricho-thecenes (e.g. deoxynivalenol; Fig. 7) have a C-8 keto func-tion (McCormick et al. 2011). Structure-activity studies in-dicate that the epoxide and the C-3 hydroxyl group areimportant for toxicity (Kimura et al. 1998a; Zhou et al.2008), but differences in hydroxylation or acetylation at theC-4, C-15, C-7 and C-8 also contribute to the relative toxicityof individual trichothecenes (Desjardins et al. 2007).

Microbial biotransformations of trichothecenes include acet-ylation (Alexander et al. 2002; Kimura et al. 1998a),deacetylation (Beeton and Bull 1989; Ueno et al. 1983), oxy-genation (Baldwin et al. 1986), de-epoxidation (Islam et al.

2012; King et al. 1984; Swanson et al. 1987a), epimerization(Ikunaga et al. 2011), and glucosylation (McCormick et al.2012). Unlike other mycotoxins, there have been no reportsthat suggest ring cleavage or complete mineralization oftrichothecenes.

Because trichothecenes are toxic to eukaryotes, producingorganisms have self-protection mechanisms. In Fusarium, thebiosynthesis of trichothecenes is via a series of C-3 acetylatedintermediates (McCormick et al. 2011) with acetylation con-trolled by an acetyltransferase gene (Tri101) (Kimura et al.1998b; McCormick et al. 1999). C-3 acetylation of trichothe-cenes decreases their toxicity (Alexander et al. 1999).Trichothecene C-3 acetyltransferase activity is and has beenfound in ascomycetes including Aspergillus, Saccharomyces,Blastobotrys, and Trichomonascus, which do not producetrichothecenes (Alexander et al. 2002; Khatibi et al. 2011;Kimura et al. 1998b; McCormick et al. 2012; Tokai et al.2005), but may be exposed to them. It is not known what rolethese acetyltransferases may play in the interactions betweentrichothecene-producing Fusarium sp. and other ascomycetesin the soil or in plants. The Fusarium trichothecene acetyl-transferase gene has been engineered into plants to improveplant resistance to F. graminearum (Alexander 2008; Ohsatoet al. 2007). There is a need to identify other trichothecenedetoxification genes that can be expressed in cereals to increaseresistance to Fusarium head blight.

Yeast in the Trichomonascus clade were found to detoxifyT-2 toxin in three ways: 1) C-3 acetylation; 2) hydrolysis ofthe C-8 isovaleryl group to neosolaniol; or 3) C-3 gluco-sylation (McCormick et al. 2012). All three biotransformationproducts (Fig. 6) are significantly less toxic than T-2 toxin.The three yeast species (Kurtzman 2007; Middelhoven andKurtzman 2003) that glucosylated T-2 toxin were isolatedfrom moss, rotted log, and decaying fungal matter, but thenatural substrate for the glucosyltransferases in not known. T-2 toxin 3-glucoside is unusual in having an alpha linked sugar.Most glycosylation of xenobiotics is via a beta linkage, suchas zearalenone-4-β-glycoside (el-Sharkawy and Abul-Hajj1987; Kamimura 1986). Both T-2 toxin 3-glucoside, andzearalenone-4-glucoside have been found as plant metabolitesof the mycotoxins (Lattanzio et al. 2011; Schneweis et al.2002). Mycotoxin glycosides in plants have been calledmasked mycotoxins because they may escape detection withcurrent analytical methods and may be hydrolyzed during

Patulin

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OO

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OH

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OH

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(E)-ascladiol Desoxypatulinic acid

Fig. 5 Patulin and itsbiotransformation products, (E)-ascladiol and desoxypatulinicacid

J Chem Ecol (2013) 39:907–918 913

digestion in animals or humans (Berthiller et al. 2013). It ispossible that the report of complete microbial degradation oftrichothecenes by Mucor racemosus (Bocarov-Stamcic et al.2011) may be due to the formation of masked mycotoxins.

There are two types of bacterial degradation of the tricho-thecene deoxynivalenol (DON): 1) oxidation to 3-keto DON,along with isomerization to 3-epi DON (3-β-hydroxy), byaerobic bacteria; and 2) deepoxidation to de-epoxy DON(DOM-1), generally by rumen or intestinal anaerobic bacte-ria. In addition, the soil fungus Aspergillus tubingensis (Heet al. 2008) converted DON to a product with a molecularweight 18 kDa, which is larger than that of DON, but theproduct was not further characterized.

Shima et al. (1997) isolated a soil microbe that convertedthe C-3 hydroxyl group to a C-3 keto function with a loss oftoxicity. Both 3-keto DON and epi-DON (Fig. 6) have beenreported as products of DON detoxification by strains of thebacteriaNocardioides andDevosia (Ikunaga et al. 2011; Satoet al. 2012; Shima et al. 1997).

Anaerobic microbial degradation of trichothecenes has beenfound in animal systems. The toxic effects of trichothecenes inanimals may be less impacted by rumen or intestinal bacteria thatcan detoxify the compounds. Deepoxy deoxynivalenol (DOM-1)(Fig. 6) has been found inmilk, urine, and feces of lactating cowsthat had been fed deoxynivalenol contaminated corn (Côté et al.

1986). Deoxynivalenol is 55 times more toxic than DOM-1(Eriksen et al. 2004). Other deepoxy trichothecenes were pre-pared by fermentation with bacteria from rumenal fluid(Swanson et al. 1987a, b). A Eubacterium with deepoxidaseactivity was isolated from rumen (Fuchs et al. 2002). Bacteriafrom the intestines of bullhead catfish and chicken also canconvert deoxynivalenol and other trichothecenes to their deepoxyanalogs (Guan et al. 2009; Young et al. 2007; Yu et al. 2010;).Although rumen and intestinal bacteria are usually anaerobic,some can live in aerobic conditions. For example, a Bacillusstrain isolated from fish intestines was successfully used todetoxify the deoxynivalenol in moldy corn that was then fed topigs (Li et al. 2011). Islam et al. (2012) recently reported thatmixed populations of enterobacteria, including Serratia,Clostridium, Citrobacter, and Enterococcus, found in soil fromagricultural fields, could convert deoxynivalenol to DOM-1under aerobic conditions. There is a great interest in cloningdeepoxidase genes from these microorganisms in order to engi-neer trichothecene detoxification into cereal plants.

Applications and Future Directions

Microbial detoxification is now an important componentof strategies to eliminate mycotoxins from food and feed.

O

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OH

OAcOAcO

C

O

23

4157

8

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OAc

OAcOAcO

C

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OH

OAcHOAcO

O

OO

OAcOAcO

C

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HO

OH

OH

OH

T-2 toxin 3-acetyl T-2 toxin

T-2 toxin 3-glucoside Neosolaniol

Fig. 6 T-2 toxin and itsbiotransformation products,3-acetyl T-2 toxin, neosolaniol,and T-2 toxin 3-glucoside

O

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OOAc

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OOH

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deoxynivalenol 3-acetyl-deoxynivalenol

DOM-1 3-keto DON 3-Epi-DON

Fig. 7 Deoxynivalenol (DON)and its biotransformationproducts 3-acetyldeoxynivalenol, 3-ketodeoxynivalenol, 3-epi-deoxynivalenol anddeepoxydeoxynivalenol(DOM-1)

914 J Chem Ecol (2013) 39:907–918

Bacteria and fungi that can biotransform mycotoxins to lesstoxic products serve as a source of enzymes that can be used todecontaminate agricultural commodities or used as feed addi-tives. In addition, genes that control the detoxifications ofmycotoxins, such as the trichothecene acetyltransferase(TRI101) (Alexander 2008) and the zearalenone lactonohy-drolase (zhd101) (Igawa et al. 2007), have been expressed inplants to limit pre-harvest contamination of crops. In cases inwhich the mycotoxin is also a virulence factor in a plant diseasesuch as Fusarium head blight, introduction of resistance to thetoxin can increase resistance to the disease.

Molecular approaches may offer insight into the interac-tions of mycotoxin-producing fungi and other organisms in-cluding mycotoxin-degrading microbes. Biosynthetic genesfor several mycotoxins have been identified (Gaffoor andTrail 2006; Hohn and Beremand 1989; O’Callaghan et al.2003; Proctor et al. 2003; Sanzani et al. 2012; Shimizu et al.2005;White et al. 2006; Yu et al. 2004), andmutants producedby disruption or deletion of toxin genes have been used toassess the role of mycotoxins in plant disease. For example,trichothecenes and patulin are both pathogenicity factors inplant disease (Desjardins et al. 1996; Sanzani et al. 2012). Incontrast, fumonisins are not required for corn infection or cornear rot (Desjardins et al. 2002). Molecular tools are nowavailable to control the production of mycotoxins and in somecases, their degradation. Toxin-deficient mutants could beused to assess other ecological functions of the toxins.Similarly, disruption of genes that control biotransformationsof mycotoxins could be used to assess how changes in theability to detoxify a mycotoxin contributes to the overallfitness of the organisms. For example, Gliocladium mutantsthat lack the zearalenone lactonase gene have increased sen-sitivity to the toxin (Utermark and Karlovsky 2007).

Finally, a better understanding of the ecology of mycotoxin-producing organisms, in agricultural fields, on plant surfaces, orwithin animal digestive systems, may in turn lead to new sourcesof mycotoxin degrading enzymes and genes. Mycotoxins haveheretofore received little attention from chemical ecologists, andthe field is ripe for development.

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Microbial Detoxification of MycotoxinsAbstractIntroductionAflatoxinsZearalenoneCitrinin and OchratoxinFumonisinsPatulinTrichothecenesApplications and Future DirectionsReferences