Molecular biodiversity of mycotoxigenic fungi that threaten food safety

FOOD-06255; No of Pages 10 International Journal of Food Microbiology xxx (2013) xxx–xxx

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Molecular biodiversity of mycotoxigenic fungi that threaten food safety A. Moretti a,⁎, A. Susca a, G. Mulé a, A.F. Logrieco a, R.H. Proctor b a b

Institute of Sciences of Food Production, CNR, Bari, Italy US Department of Agriculture, ARS, NCAUR, Peoria, IL, USA

a r t i c l e Available online xxxx Keywords: Aflatoxins Aspergillus Fumonisins Fusarium Trichothecenes Biosynthetic pathway

i n f o

a b s t r a c t Fungal biodiversity is one of the most important contributors to the occurrence and severity of mycotoxin contamination of crop plants. Phenotypic and metabolic plasticity has enabled mycotoxigenic fungi to colonize a broad range of agriculturally important crops and to adapt to a range of environmental conditions. New mycotoxin-commodity combinations provide evidence for the ability of fungi to adapt to changing conditions and the emergence of genotypes that confer enhanced aggressiveness toward plants and/or altered mycotoxin production profiles. Perhaps the most important contributor to qualitative differences in mycotoxin production among fungi is variation in mycotoxin biosynthetic genes. Molecular genetic and biochemical analyses of toxigenic fungi have elucidated specific differences in biosynthetic genes that are responsible for intra- and inter-specific differences in mycotoxin production. For Aspergillus and Fusarium, the mycotoxigenic genera of greatest concern, variation in biosynthetic genes responsible for production of individual families of mycotoxins appears to be the result of evolutionary adaptation. Examples of such variation have been reported for: a) aflatoxin biosynthetic genes in Aspergillus flavus and Aspergillus parasiticus; b) trichothecene biosynthetic genes within and among Fusarium species; and c) fumonisin biosynthetic genes in Aspergillus and Fusarium species. Understanding the variation in these biosynthetic genes and the basis for variation in mycotoxin production is important for accurate assessment of the risks that fungi pose to food safety and for prevention of mycotoxin contamination of crops in the field and in storage. © 2013 Elsevier B.V. All rights reserved.

1. Introduction Mycotoxins are low molecular weight toxic compounds produced by fungi and pose a serious risk to human and animal health worldwide. These toxins occur naturally and affect approximately 25% of the global food and feed crop output (FAO, 2004). Cereals are often the most severely affected crops, and because they are a staple food for a large portion of humanity, mycotoxins are the most prevalent source of food-related health risk in field crops. Worldwide scientific and economic attention on mycotoxins also results from the significant economic losses associated with their negative impacts on human health, animal productivity, and international trade. As an example, economic losses in the US caused by contamination of wheat and barley with Fusarium mycotoxins, and the associated plant disease problems, were $2.9 billion per year (Windels, 2000), while the cost of controlling contamination of crops with aflatoxins was $1.4 billion per year (CAST, 2003). The mycotoxins of greatest concern to food and feed safety are produced primarily by three genera of filamentous fungi: Aspergillus, Fusarium and Penicillium (Marasas et al., 2008). In addition, although fungi can collectively produce hundreds of mycotoxins, only the following ⁎ Corresponding author at: Institute of Sciences of Food Production, CNR, Via Amendola 122/O, 70126 Bari, Italy. Tel.: +39 080 5929357; fax: +39 080 5929374. E-mail address: [email protected] (A. Moretti).

are of serious concern worldwide: aflatoxins, fumonisins, ochratoxins, patulin, trichothecenes, and zearalenone (Marasas et al., 2008). Some of these mycotoxins exhibit a very high degree of structural diversity, reflecting genetic diversity of species that produce them. Some mycotoxigenic species of greatest concern occur worldwide. However, other important species have more limited distributions. In some cases, mycotoxin contamination can result from the presence of multiple fungal species in a single crop. An important component of efforts to control mycotoxin contamination problems is the study of the morphological, molecular genetic, metabolic, and plant pathological diversity of mycotoxigenic fungi. Plasticity of agriculturally relevant traits contributes to the collective ability of fungi to colonize a wide variety of crop species and to adapt to a range of environmental conditions. Knowledge of environmental factors that affect the ability of fungi to grow, survive and interact with plants can affect population structures of mycotoxigenic fungi, their interactions with crop plants, and their ability to produce mycotoxins. Because climate can profoundly affect growth, distribution, and mycotoxin production in fungi, climate change has the potential to increase the risks that mycotoxigenic fungi pose to food and feed safety. The appearance of new mycotoxin-commodity combinations is of further concern and provides evidence for emergence of new fungal genotypes with higher levels of aggressiveness and altered mycotoxin production. Trans-global trade of plant products can also contribute to the spread of toxigenic fungi and has lead to increasing interest in

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molecular diversity of toxigenic fungi on a global scale (e.g. Starkey et al., 2007). Over the past two decades, analyses of fungal diversity have lead to an improved ability to identify and distinguish between species that cause mycotoxin contamination problems. These studies have also revealed some of the genetic bases for variation in mycotoxin production phenotypes (chemotypes) within and among species, including differences in the ability of fungi to produce the three mycotoxin families of greatest concern to food and feed safety: aflatoxins, fumonisins, and trichothecenes. In this paper, we review critical information on variability in fungal genes responsible for biosynthesis of these three mycotoxin families. We emphasize the importance of knowledge of biodiversity of toxigenic fungi to better understand factors that contribute to mycotoxin production, assessment of risks posed by mycotoxigenic fungi, and reduction of mycotoxin contamination in feed and food crops. 2. Biosynthetic genes and gene clusters Fungi produce numerous compounds, such as antibiotics, pigments and mycotoxins, that are termed secondary metabolites, because they are not essential for normal growth and development. Despite their diverse chemical structures and biological activities, there are multiple attributes that are common to the biosynthesis of most fungal secondary metabolites, notably biogenic origin and gene clustering. Most fungal secondary metabolites are products of multi-step biochemical pathways that often begin when a terpene synthase, polyketide synthase or nonribosomal peptide synthetase catalyzes rearrangement or condensation of a relatively simple primary metabolite(s) to form a more complex molecule, i.e. a terpene, polyketide or nonribosomal peptide (Keller et al., 2005). These molecules usually undergo a series of enzyme-catalyzed modifications to form the mycotoxin structures that are of concern to food and feed safety. Genes encoding the synthase and modifying enzymes for a given mycotoxin are typically located next to one another in a gene cluster (Keller et al., 2005). In addition to synthase and modifying enzymes, biosynthetic gene clusters can also encode metabolite transport proteins and transcription factors that regulate expression of cluster genes. Thus, the biosynthetic gene cluster is an essential genetic element for production of most mycotoxins, and the presence and absence of a cluster in fungal species are keys to whether a species produces or does not produce the corresponding mycotoxin. Most mycotoxins are families of structurally closely related molecules produced by the same biosynthetic pathway. Structural variation within a mycotoxin family can result when a pathway does not go to completion, from branches in a pathway, or from variation in function of genes within a biosynthetic cluster. This variation can result when a gene is functional in some strains but nonfunctional in others, when a gene is present in some strains but absent in others, or when the activity of an enzyme encoded by a gene varies. In all cases, however, the genetic variation results in variation in a biosynthetic pathway, and this in turn gives rise to structural variation of the corresponding mycotoxin.

A. flavus is highly variable and depends on genotype, substrate and geographic origin (Vaamonde et al., 2003; Pildain et al., 2004). In addition, strains of A. flavus produce AFB1 and AFB2 and often cyclopiazonic acid (CPA), while most strains of A. parasiticus produce AFG1 and AFG2 in addition to AFB1 and AFB2 but never produce CPA (Horn and Dorner, 1999). The aflatoxin biosynthetic gene (afl) cluster includes 25 genes (Yu et al., 2004). The gene content and organization of the cluster is highly conserved among Aspergillus species in section Flavi, which includes A. flavus and A. parasiticus. Sequence variability and deletions in various genes/regions of the afl cluster have also been used to assess variability in A. flavus (Chang et al., 2005, 2006). Moreover, differences in afl genes have been used to distinguish between aflatoxin-producing and nonproducing strains of A. flavus and A. parasiticus. Understanding such genetic differences is important because aflatoxin-nonproducing strains of A. flavus are used to control aflatoxin contamination in some crops (Cotty, 1994, 2006; Dorner and Horn, 2007). Comparative analyses of the afl cluster in multiple Aspergillus species have provided insights into the evolutionary history of the cluster and its link to species adaptation and diversification (Ehrlich et al., 2003; Carbone et al., 2007; Moore et al., 2009). In an analysis of multiple aflatoxin-nonproducing isolates of A. flavus, either part of or the entire afl cluster was deleted (Chang et al., 2005). In multiple PCR-based studies, aflatoxin nonproduction in some strains was associated with an inability to amplify selected afl genes (Criseo et al., 2008; Degola et al., 2007; Sweeney et al., 2000; Scherm et al., 2005). In one study, Gallo et al. (2012) examined a collection of aflatoxin-producing and nonproducing isolates of A. flavus for the presence of seven afl genes, two regulatory genes aflR and aflS and the structural genes aflD, aflM, aflO, aflP, and aflQ. The result was the grouping of strains into four different amplification patterns. All aflatoxin-producing isolates yielded the complete set of amplification products, whereas nonproducing isolates did not yield products for three, four or all seven genes. Although PCR-based assessments can provide evidence for afl gene deletion, they do not demonstrate deletion, because lack of an amplification product can also result from changes in primer binding sites. On the other hand, the lack of amplification of afl genes provides further evidence of the high levels of genetic variability among A. flavus isolates. Variability in the afl cluster, including deletions, recombination, inversions, translocations, and other rearrangements, could result from the proximity of the cluster to the telomere (Carbone et al., 2007). Together, analyses of variation in the afl cluster in A. flavus and A. parasiticus have illustrated the genetic complexity of these species, which includes variability in the afl cluster. This genetic variability has provided markers that can be used to monitor variation in these Aspergillus species and to evaluate the risk they pose when present on food commodities. In addition, understanding the genetic variability in the afl cluster is a key to the selection of safe and effective aflatoxin-nonproducing strains for biological control efforts aimed at limiting aflatoxin contamination in crops. 4. Trichothecenes and Fusarium

3. Aflatoxins and Aspergillus Aflatoxins are potent carcinogens that include four major structural analogues: AFB1, AFB2, AFG1 and AFG2. The International Agency for Research on Cancer (IARC) has classified AFB1 as a group 1 carcinogen in humans (IARC, 1993). In addition to hepatocellular carcinoma, aflatoxins are associated with occasional outbreaks of acute aflatoxicosis that lead to death shortly after exposure (Azziz-Baumgartner et al., 2005). Aflatoxins are produced in a diversity of agricultural commodities by several species of Aspergillus, but the two species of greatest concern are Aspergillus flavus and Aspergillus parasiticus (Frisvad et al., 2005). The ability to produce aflatoxins is highly conserved in some species but variable in others. For example, 94–97% of A. parasiticus strains that have been examined produce aflatoxins, whereas production in

The fungal genus Fusarium consists of over 90 described species and likely tens of additional undescribed but phylogenetically distinct species (Leslie and Summerell, 2006; O'Donnell et al., 2009, 2013; Geiser et al., 2013; Sarver et al., 2011). Many of these species are plant pathogens and produce a range of mycotoxins, the most agriculturally important being trichothecenes and fumonisins (Desjardins, 2006). Trichothecenes are a family of terpene-derived mycotoxins produced by multiple species of Fusarium and are among the most economically significant mycotoxins worldwide because of their potency and widespread occurrence in important grain crops such as barley, maize and wheat (CAST, 2003). These mycotoxins are toxic to plants and can contribute to pathogenesis of Fusarium on some crops (Desjardins et al., 1996; Maier et al., 2006). All trichothecenes have a tricyclic skeleton

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structure with an epoxide group, but they can be divided into two structurally distinct groups based on the absence (type A trichothecenes) and presence (type B trichothecenes) of a keto group at carbon atom 8 (C-8) of the skeleton (Fig. 1). The type A trichothecenes of greatest concern are T-2 toxin and HT-2 toxin, while the type B trichothecenes of greatest concern are deoxynivalenol (DON), nivalenol (NIV), and their acetyl-derivatives (Desjardins, 2006). 4.1. Organization of TRI loci Fusarium graminearum sensu stricto and Fusarium sporotrichioides were the two species in which the trichothecene biosynthetic gene (TRI) cluster was first characterized (Fig. 2) (Brown et al., 2001, 2002; Lee et al., 2002). In both species, the cluster consists of 12 genes that are responsible for the synthesis of the core trichothecene skeleton and several modifications to it. The cluster genes are: the terpene synthase gene TRI5; three cytochrome P450 monooxygenase genes TRI4, TRI11, and TRI13, two acyl-transferase genes TRI3 and TRI7, the esterase gene TRI8, two regulatory genes TRI6 and TRI10, and the transporter gene TRI12. Two other genes, TRI9 and TRI14 are also located in the core TRI cluster, but they lack similarity to genes with known functions and their functions in trichothecene biosynthesis are not known (Alexander et al., 2011). In F. graminearum and F. sporotrichioides, the core TRI cluster is identical with respect to gene order and orientation (Brown et al., 2002, 2004). Both species also have two smaller TRI loci (Fig. 2). The first of these consists of one gene, TRI101, encoding an acyl transferase. The second locus consists of two genes: one, TRI1, encodes a monooxygenase and the other, TRI16, encodes an acyl transferase (Kimura et al., 1998a; McCormick et al., 1999). Analysis of the F. graminearum genome sequence indicated that the three TRI loci are located on different chromosomes (Cuomo et al., 2006; Lee et al., 2008). Functional differences of the TRI1 and TRI16 enzymes in F. graminearum and F. sporotrichioides contribute to important structural differences of trichothecenes produced by the two species. In F. sporotrichioides, TRI1 is responsible for hydroxylation of the trichothecene molecule at C-8, and TRI16 is responsible for esterification of isovaleryl to the C-8 oxygen (Fig. 1) to form T-2 toxin and its acetylated derivatives, which are considered type A trichothecenes because they lack a C-8 keto (Brown et al., 2003; Meek et al., 2003; Peplow et al., 2003). In F. graminearum, the function of TRI1 differs in that it is responsible for hydroxylation of the trichothecene molecule at C-7 as well as C-8. In addition, TRI16 is nonfunctional in F. graminearum (Brown et al., 2003; McCormick et al., 2004, 2006), and as a result the C-8 oxygen does not form an isovaleryl ester. Instead, the C-8 hydroxyl undergoes

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oxidation to form the keto group characteristic of type B trichothecenes. The gene/enzyme responsible for this oxidation has not yet been identified. Analysis of TRI loci in 16 species of Fusarium (Table 1) revealed that TRI1 and TRI101 located in the core TRI cluster in four species of Fusarium (Proctor et al., 2009) are members of the F. incarnatum–Fusarium equiseti species complex (O'Donnell et al., 2009). This finding raised the question of whether the presence of TRI1 and TRI101 in the TRI cluster was the ancestral or derived state. For TRI101, it was possible to address this question, because functional or nonfunctional homologues of TRI101 are present in trichothecene-nonproducing species of Fusarium (Tokai et al., 2005; Kimura et al., 2003, 1998b) that are relatively distantly related to the clade of trichotheceneproducing species that was examined. The genomic context of TRI101 in trichothecene-nonproducing species was the same as in trichothecene-producing species in which TRI101 was located outside the TRI cluster. Based on this, Proctor et al. (2009) concluded that it was more likely that the TRI101 was located outside the cluster in the common ancestor of trichothecene-producing and nonproducing fusaria and subsequently moved into the cluster during the evolution of the F. incarnatum–F. equiseti species complex. Because there was no evidence for TRI1 homologues in trichothecenenonproducing fusaria, other evidence was used to assess whether TRI1 had moved into or out of the TRI cluster. Multiple strains of the F. incarnatum–F. equiseti species complex that were examined had an apparently functional copy of TRI16 located outside of the cluster. In phylogenetic analyses, trees based on TRI1 and TRI16 sequences were highly correlated with one another but not with a species phylogeny inferred from housekeeping genes or with the genomic context of TRI1. By contrast, trees based on TRI cluster genes and on TRI101 were highly correlated with the species phylogeny. Proctor et al. (2009) rationalized that the similar pattern of sequence variation exhibited by TRI1 and TRI16 but not by other TRI genes would have arisen more readily if TRI1 and TRI16 had originally been closely linked to one another and not to the TRI cluster. Thus, they concluded that both TRI1 and TRI101 originated outside of the TRI cluster and moved into it during divergence of the F. incarnatum–F. equiseti species complex from other lineages of trichothecene-producing fusaria. Relocation of TRI1 and TRI101 into the core TRI cluster of the F. incarnatum–F. equiseti species complex provided evidence for growth of a fungal secondary metabolite gene cluster by gene relocation rather than by gene duplication and subsequent divergence of preexisting cluster genes. Phylogenetic analysis of four F. graminearum monooxygenase genes (TRI1, TRI4, TRI11 and TRI13) involved in trichothecene biosynthesis

Fig. 1. Later steps in trichothecene biosynthesis showing the formation of the type A trichothecene T-2 toxin and type B trichothecene deoxynivalenol (DON) and how the TRI1 and TRI16-encoded enzymes contribute to the structural differences between these two mycotoxins.

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Core TRI cluster TRI8

TRI7

TRI3

TRI4

TRI6

TRI5

TRI101 locus

TRI1-TRI16 locus

TRI101

TRI1

TRI10 TRI9

TRI11

TRI12

TRI13

TRI16

Fig. 2. Trichothecene biosynthetic loci in F. graminearum and F. sporotrichioides according to Kimura et al. (1998a) and McCormick et al. (1999). In F. graminearum, TRI7, TRI13, TRI16 are nonfunctional.

provides a further independent support for this conclusion and showed that it is unlikely that the genes encoding them evolved directly from the same ancestral TRI gene (Deng et al., 2007; Proctor et al., 2009). 4.2. TRI gene polymorphism and trichothecene structural variation As noted above, phylogenies inferred from TRI1 and TRI16 sequences from a wide range of trichothecene-producing fusaria were not congruent with the species phylogeny (Proctor et al., 2009). That is, TRI1 and TRI16 sequences in some closely related species were more distantly related to one another than to sequences from more distantly related species. TRI1 polymorphism contributes for the differences in function of TRI1 that are responsible for the different patterns of C-7 and C-8 oxygenation of the trichothecenes produced by F. graminearum and F. sporotrichioides (Fig. 1) (Meek et al., 2003; McCormick et al., 2004; McCormick et al., 2006). Phylogenetic incongruence has also been observed in the TRI cluster of members of the F. graminearum species complex, which represents a phylogenetically narrow range of species (Ward et al., 2002). Sequence variation within the F. graminearum complex was trans-specific in that sequence polymorphism was shared between some species. By contrast, of ~50 strains of Fusarium examined for TRI1 and TRI16, polymorphism existed among but not within species (Proctor et al., 2009). At least some patterns of polymorphism within the TRI cluster in the F. graminearum species complex and closely related species (e.g. Fusarium culmorum) are associated with differences in trichothecene chemotypes (Chandler et al., 2003; Ward et al., 2002). For example, polymorphism in TRI13 is responsible for variation in the presence and absence of a C-4 hydroxyl, an important structural difference that occurs within type B trichothecenes. Similar patterns of deletions within TRI13 occur in isolates of both F. graminearum and F. culmorum (Chandler et al., 2003). Strains of these fungi with a functional TRI13 produce the C-4 hydroxylated trichothecene NIV, whereas strains with a nonfunctional TRI13 (i.e. with the deletions) produce DON, which lacks a C-4 hydroxyl (Chandler et al., 2003; Lee et al., 2002). Another trichothecene chemotype difference exhibited by members of the F. graminearum species complex and closely related species is the production of 3-acetyldeoxynivalenol (3-ADON) versus 15-acetyldeoxynivalenol (15-ADON). The genetic basis for this difference resides in the TRI cluster gene TRI8. Alexander et al. (2011) identified consistent DNA sequence differences in the coding region of TRI8 in 3-ADON and 15-ADON strains. Functional analyses of the TRI8 enzyme (Tri8) from F. graminearum sensu stricto revealed that Tri8 from 3-ADON strains catalyzes deacetylation of the trichothecene biosynthetic intermediate 3,15-diacetyldeoxynivalenol at C-15 to yield 3-ADON, whereas Tri8 from 15-ADON strains catalyzes deacetylation of 3,15-diacetyldeoxynivalenol at C-3 to yield 15-ADON. Alexander et al. (2011) also showed that Tri8 in NIV-producing Fusarium strains functions like that in 15-ADON strains, and that TRI3, which encodes a trichothecene C-15 acetyltransferase, was functional in all three chemotypes. Together, data from Alexander et al. (2011) indicated that differential activity of Tri8 determines the 3-ADON and 15-ADON chemotypes in Fusarium.

Multiple studies have revealed that structural variation of trichothecene mycotoxins produced by different species of Fusarium and in some cases different strains of the same species result from DNA sequence polymorphism that causes variation in the functionality of TRI genes (e.g. Brown et al., 2001; Lee et al., 2002; Chandler et al., 2003). In some cases, the sequence differences include insertions and deletions that render genes nonfunctional, but in other cases the differences give rise to enzymes that catalyze different reactions. 5. Fumonisins and Fusarium Fumonisins are a family of polyketide-derived mycotoxins that are epidemiologically associated with esophageal cancer and neural tube defects in some human populations as well as multiple diseases in livestock animals and experimental rodents (Gelderblom et al., 1988; Marasas et al., 2004). These mycotoxins consist of a linear carbon backbone with a nitrogen-containing group, one to four hydroxyl groups, and two tricarboxylate esters at various positions along the backbone. Fumonisins exhibit considerable structural variation and can be divided into four series, designated A, B, C and P, based on variation around the nitrogen atom and differences in the length of the carbon backbone (Musser and Plattner, 1997; Sewram et al., 2005). A, B and P fumonisins have a 20-carbon backbone, whereas C fumonisins (FCs) have a 19-carbon backbone. In B fumonisins (FBs) and FCs, the nitrogen group is a free amine, in A fumonisins it is an acetylated amine, and in P fumonisins it is part of a hydroxypyridinium ring (Musser and Plattner, 1997; Sewram et al., 2005). Most known fumonisin-producing fusaria are members of the Fusarium fujikuroi species complex (FFSC) and typically produce four FB analogues (FB1, FB2, FB3 and FB4) in greatest abundance, but FB1 is usually produced at higher levels than the other three analogues (Nelson et al., 1992; Proctor et al., 2004; Rheeder et al., 2002). FB analogues differ in structure by the presence and absence of hydroxyl groups at C-5 and C-10: FB1 has hydroxyls at both C-5 and C-10; FB2 has a hydroxyl at C-5 but not C-10; FB3 has a hydroxyl at C-10 but not C-5; and FB4 lacks a hydroxyl at both C-5 and C-10. Fusarium oxysporum is closely related to but not part of the FFSC, and although it is generally regarded as a fumonisin-nonproducing species, two producing isolates of it have been identified (Seo et al., 1996; Sewram et al., 2005). In contrast to the FFSC species that have been examined, the F. oxysporum isolates produce FCs rather than FBs. The most abundant fumonisins produced by the F. oxysporum isolates are hydroxy-FC1, FC1, iso-FC1, FC2, and FC3, which like FB analogues differ in the pattern of hydroxylation along the carbon backbone (Fig. 3A). A fumonisin biosynthetic gene (FUM) cluster (Fig. 3B) has been described in F. oxysporum strain FRC O-1890 (Proctor et al., 2008) and in the FFSC species Fusarium proliferatum and Fusarium verticillioides (Proctor et al., 2003; Waalwijk et al., 2004). The cluster includes 16 genes that encode biosynthetic enzymes, transport proteins, and a transcription factor (Brown et al., 2007; Proctor et al., 2003). Functional characterization of the cluster in F. verticillioides via a variety of methods has elucidated the role of most of FUM genes in fumonisin biosynthesis

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Fig. 3. A. Analogues of B and C fumonisins produced by Fusarium and black aspergilli. All analogues have been reported in cultures of Fusarium. Asterisks (*) indicate analogues that have been reported in A. niger. B. The fumonisin biosynthetic gene (FUM) clusters in F. oxysporum, F. verticillioides and A. niger (Proctor et al., 2003, 2008). The gene in the A. niger cluster that is labeled with a question mark is not present in the Fusarium cluster and is predicted to encode a short-chain dehydrogenase.

(Butchko et al., 2003a,b, 2006; Ding et al., 2004; Yi et al., 2005; Zaleta-Rivera et al., 2006). This in combination with more limited analyses of a fumonisin-producing strain of F. oxysporum has lead to an understanding of the genetic basis for much of the structural diversity exhibited by fumonisin analogues. The cluster gene FUM8 is responsible for FB versus FC production in Fusarium (Proctor et al., 2008). FUM8 is predicted to encode an α-oxoamine synthase/class II aminotransferase, a group of enzymes that catalyze condensation of an amino acid and acyl compound, such as a fatty acid or polyketide (Christen and Mehta, 2001). A combination of precursor feeding studies (Branham and Plattner, 1993) and molecular genetic analyses of F. verticillioides and F. oxysporum (Proctor et al., 2008) indicate that during FB biosynthesis, the FUM8-encoded enzyme (Fum8) catalyzes condensation of alanine and an 18-carbon-long linear polyketide to form the 20-carbon-long FB backbone, whereas during FC biosynthesis, Fum8 catalyzes condensation of glycine and the linear polyketide to form the 19-carbon-long FC backbone. The cluster genes FUM2 and FUM3 contribute to the different patterns of hydroxylation among fumonisin analogues. FUM2 encodes a cytochrome P450 monooxygenase that catalyzes C-10 hydroxylation (Proctor et al., 2006), and FUM3 encodes a dioxygenase that catalyzes C-5 hydroxylation (Butchko et al., 2003a). Precursor feeding studies with FB2, FB3 and FB4 and biochemical analysis of multiple strains of F. verticillioides in which FUM genes were inactivated (Butchko et al., 2003a; Proctor et al., 2006; Uhlig et al., 2012) provide evidence for two branches of the fumonisin biosynthetic gene cluster; one branch leads to the formation of FB3 and FB1, and the other leads to the formation of FB4 and FB2 (Fig. 4). The genetic basis of some structural variation of fumonisin analogues has not yet been determined. Hydroxy-FC1 and iso-FC1 have a hydroxyl group at C-3 that is not present at the equivalent position (C-4) of FB1. There is a cytochrome

P450 monooxygenase gene (CPM1) adjacent to the known FUM cluster in F. oxysporum that could be responsible for the C-3 hydroxylation (Fig. 3), but whether it plays a role in fumonisin biosynthesis has not been determined (Proctor et al., 2008). In addition, the genetic basis, if any, for the formation of the acetylated amine of A fumonisins and the hydroxypyridinium ring of P fumonisins has not been determined. Comparisons of the FUM clusters in F. verticillioides, F. oxysporum, and F. proliferatum indicate that the order and orientations of genes within the three species are the same (Proctor et al., 2003, 2008; Waalwijk et al., 2004). However, other observations suggest that other attributes of the cluster are more complex than the uniformity of gene organization would indicate. For example, the sequences flanking the clusters differ in the three species examined, indicating that the cluster is in a different genomic context in each species, which in turn suggests that the cluster has undergone translocation during the evolutionary history of fumonisin-producing fusaria. In addition, phylogenetic analyses of FUM1, the fumonisin polyketide synthase gene, and FUM8 in seven fumonisin-producing fusaria yielded trees (Proctor et al., 2004) with topologies that differed from those generated with housekeeping genes in another study (O'Donnell et al., 1998). For example, the six FFSC species employed in the FUM1/FUM8 gene analysis are all more closely related to one another than any is to F. oxysporum (O'Donnell et al., 1998). However, the FUM1 and FUM8 sequences of the FFSC species Fusarium nygamai and F. verticillioides were more closely related to those of the fumonisin-producing isolate of F. oxysporum than to other FFSC species included in the analysis (Proctor et al., 2004). Thus, the analysis provided evidence that the phylogenetic relationships of FUM genes do not necessarily mirror the phylogenetic relationships of the species in which they occur. Similar phylogenetic discord observed for the aflatoxin biosynthetic gene (afl) cluster in Aspergillus and the TRI cluster in the F. graminearum species complex

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Fig. 4. Two branches of the fumonisin biosynthetic pathway. One branch leads to the formation of FB3 and FB1 and occurs in Fusarium but not in A. niger. The other branch leads to the formation of FB4 and FB2 and occurs in both Fusarium and A. niger (Butchko et al., 2003a; Proctor et al., 2006; Uhlig et al., 2012).

has been attributed to balancing selection.1 Studies are currently underway to determine whether this or other evolutionary processes have contributed to the apparent phylogenetic discord observed between FUM1 and FUM8 and the species in which they occur (Proctor et al., 2013). The FFSC consists of over 50 phylogenetically distinct species, including some that are pathogens and endophytes of many economically important plants (Kvas et al., 2009; O'Donnell et al., 1998). Nucleotide-based phylogenetic analyses have resolved the complex into three major lineages that are designated as the African, American, and Asian clades (O'Donnell et al., 1998, 2000; Summerell et al., 2010). Fumonisin production has been reported in at least one species in each clade (Nelson et al., 1992; Rheeder et al., 2002). However, multiple FFSC and almost all F. oxysporum strains do not produce fumonisins and do not have FUM genes (Glenn et al., 2008; Mirete et al., 2004; Proctor et al., 2004; Stepien et al., 2010; Van Hove et al., 2011). In the FFSC, the presence of FUM genes and fumonisin production is discontinuous, in that their occurrence is not correlated with phylogenetic relationships of species (Fig. 5) (O'Donnell et al., 2000; Proctor et al., 2004; Rheeder et al., 2002). For example, F. proliferatum (Asian clade) and F. verticillioides (African clade) are relatively distantly related, but both have the FUM cluster and both can produce fumonisins (Proctor et al., 2003; Waalwijk et al., 2004). By contrast, Fusarium musae, the closest known relative of F. verticillioides, has only a small remnant of the FUM cluster and, therefore, does not produce fumonisins (Glenn et al., 2008; Mirete et al., 2004; Van Hove et al., 2011). Furthermore, the remnant of the FUM cluster in F. musae is in the same genomic context as the intact cluster in F. verticillioides (Fig. 6). This indicates that the cluster was present in the common ancestor of F. musae and F. verticillioides, but that as F. musae diverged the cluster was deleted. This finding indicates that the FUM cluster could have been present in the common ancestor of the FFSC, and that the current discontinuous distribution of the cluster is the result of differential cluster loss and retention during the evolutionary history of FFSC species.

distribution and occur on a large variety of substrates, including soil, cotton textiles, and meat products, and vegetable, cereal, fruit and nut crops (Raper and Fennell, 1965; Pitt and Hocking, 2007, 2009). Some species are also used extensively for industrial production of enzymes and organic acids, including enzymes and acids used for food products (Raper and Fennell, 1965; Varga et al., 2000). The black aspergilli include multiple species that are morphologically similar or undistinguishable. However, DNA sequence analysis of housekeeping genes (e.g. the calmodulin-encoding gene) serves as a useful tool to identify and distinguish between phylogenetically distinct species. Several species in section Nigri can produce mycotoxins that are of significant concern. Multiple species are reported to produce ochratoxins (Abarca et al., 2003, 2004; Wicklow et al., 1996; Varga et al., 2000; Cabanes et al., 2002; Sage et al., 2004; Samson et al., 2004), and at least two, Aspergillus niger and Aspergillus awamori, are reported to produce

6. Fumonisins and black aspergilli Aspergillus species in section Nigri (Gams et al., 1985) are commonly known as the black aspergilli. These fungi have a worldwide 1 Balancing selection selective processes by which multiple alleles are actively maintained in a population at frequencies above that of gene mutation.

Fig. 5. Distribution of fumonisin production and FUM genes among members of the Asian, African and American clades of the Fusarium fujikuroi species complex and F. oxysporum (Proctor et al., 2004; Van hove et al., 2011). A “+” indicates that fumonisin production or FUM genes have been detected in the species indicated, whereas a “–” indicates that production or the genes have not been detected.

Please cite this article as: Moretti, A., et al., Molecular biodiversity of mycotoxigenic fungi that threaten food safety, International Journal of Food Microbiology (2013), http://dx.doi.org/10.1016/j.ijfoodmicro.2013.06.033

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Fig. 6. Region of the F. verticillioides genome with the fumonisin biosynthetic gene (FUM) cluster and the same region in F. musae that has only a remnant of the FUM cluster (Glenn et al., 2008; Van Hove et al., 2011).

fumonisins (Frisvad et al., 2007; Mansson et al., 2010; Noonim et al., 2009; Palumbo and O'Keeffe, 2013). Analyses of A. niger revealed that it can produce FB2, FB4 and FB6, but not FB1 or FB3. In a survey of black aspergilli isolated from raisins, 77% of A. niger isolates produced FB2 and FB4, but none of these strains produced ochratoxins (Medina et al., 2005; Noonim et al., 2009). These results have prompted multiple investigations into the genetic bases for differences in the abilities of black aspergilli to produce mycotoxins and an evaluation of the distribution of ochratoxin and fumonisin production in these fungi. In a PCR and Southern blot-based survey of 32 isolates of A. niger from grapes from the Mediterranean basin, the FUM/fum cluster gene fum8 (genetic nomenclature for Aspergillus employs lower case letters) was detected in 11 isolates, only nine of which produced FB2 under the conditions examined (Susca et al., 2010). A similar PCR-based survey of 197 isolates of A. niger recovered from California raisins also detected fumonisin production and the fum cluster genes fum1 and fum19 in 66 and 96%, respectively, of the isolates (Palumbo et al., 2011). The two latter studies suggest that fum genes are present in some fumonisin-nonproducing isolates of A. niger but not others. Further insight into the presence and absence of fum genes was provided when A. niger was resolved, by DNA sequence-based analyses, into two morphologically indistinguishable species, A. niger sensu stricto and A. awamori (Perrone et al., 2011). In a

Table 1 Fusarium species analyzed for trichothecene production ability, TRI101, TRI1 gene occurrence and location and orientation of TRI101 flanking regions. Species

TRICHa producer

TRI101/PHO5 orientation

TRI101/URA7 orientation

TRI1 location

F. F. F. F. F. F. F. F. F. F. F. F. F. F. F. F. F. F. F. F.

graminearum boothi crockwellense culmorum sambucinum venenatum kyushuense poae chlamydosporum sporotrichioides armeniacum longipes equiseti scirpi semitrectum camptoceras verticillioides fujikuroi oxysporum solani

Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Not Not Not Not

FGb-FSc like FG-FS like FG-FS like FG-FS like FG-FS like FG-FS like FG-FS like FG-FS like FG-FS like FG-FS like FG-FS like FG-FS like FEd clade FE clade FE clade FE clade TRI101 remnant TRI101 remnant TRI101 remnant Unique

FG-FS like FG-FS like FG-FS like FG-FS like FG-FS like Unkown Unkown FG-FS like FG-FS like FG-FS like FG-FS like Unkown FE clade FE clade FE clade FE clade TRI101 remnant TRI101 remnant TRI101 remnant Unique

FG-like FG-like FG-like FG-like FG-like FG-like FG-like FG-like FS-like FS-like FS-like Unique FE clade FE clade FE clade FE clade TRI1 absent TRI1 absent TRI1 absent TRI1 absent

a

Trichothecenes. F. graminearum. F. sporotrichioides. F. equiseti.

b c d

subsequent study using a multiplex PCR assay for eight fum genes, all eight fum genes were detected in fumonisin-nonproducing isolates of A. niger but only two (fum1 and fum19) were detected in nonproducing isolates of A. awamori (Palumbo and O'Keeffe, 2013). These results suggest partial deletion of or marked sequence variation in the fum cluster of nonproducing isolates of A. awamori but not in A. niger. However, gene expression analysis revealed reduced fum gene expression in fumonisin-nonproducing isolates of A. niger relative to producing isolates (Palumbo and O'Keeffe, 2013). Analyses of black aspergilli have revealed a diversity of fum gene-related genotypes within A. niger and A. awamori, that the presence of fum genes based solely on PCR amplification does not reliably predict the ability of some isolates to produce FB2, and suggest that in some strains the loss of FB2 production could be a result of structural or regulatory mutations that alter gene expression or function. To address these issues, additional studies are underway to determine the extent of the deletion of the fum cluster in fumonisin-nonproducing isolates of A. awamori as well as the nucleotide sequence of the entire fum cluster in nonproducing isolates of A. niger and to assess abiotic factors that affect FB2 production in producing isolates (Susca et al., unpublished). In addition, a re-examination of large collections of black aspergilli that were formally considered to consist only of A. niger is warranted to discern the frequency of A. niger and A. awamori isolates in these collections, the relative frequencies of fumonisin-producing and nonproducing isolates, and the extent to which the ecology of these two species overlap (Perrone et al., 2011; Varga et al., 2010). The ecological role of FB2 production in A. niger and A. awamori should also be examined to gain insight into why these two species exist as mixed populations of fumonisin-producing and nonproducing individuals. It may also be possible to exploit fumonisin-nonproducing strains of black aspergilli as biocontrol agents for reduction of fumonisin contamination of vineyard crops, in a manner similar to the use of aflatoxinnonproducing isolates of A. flavus to control aflatoxin contamination in cotton, peanuts and corn (Abbas et al., 2006; Cotty, 2006; Dorner and Horn, 2007). Moreover, development of non-toxigenic strains of A. niger for industrial applications may become important, given that, according to a recent study, 83% of A. niger strains used for production of citric acid also produce fumonisins, and that toxigenic strains optimized for industrial uses via random mutagenesis usually retained the ability to produce mycotoxins (Frisvad et al., 2011). Finally, clarification of the genetic potential for and frequency of fumonisin production among strains of A. niger has increased concern about this fungus with respect to food safety, because of its widespread use in food production and its frequent occurrence on food crops. 7. Conclusions Functional analyses and multi-species comparisons of mycotoxin biosynthetic genes are providing important insights into the genetic

Please cite this article as: Moretti, A., et al., Molecular biodiversity of mycotoxigenic fungi that threaten food safety, International Journal of Food Microbiology (2013), http://dx.doi.org/10.1016/j.ijfoodmicro.2013.06.033

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bases for variation in production of aflatoxins, fumonisins and trichothecenes, three of the mycotoxins of greatest concern to food and feed safety worldwide. The studies indicate that the variation in mycotoxin production is often the result in three types of genetic variation: i) presence versus absence of one or more biosynthetic genes; ii) presence of functional versus nonfunctional genes; and iii) polymorphic sequences in enzyme-encoding genes that give rise to homologous enzymes with different activities or substrate specificities. The distribution of such sequence variation has had two opposing effects on variation in mycotoxin production. In some cases, sequence variation is distributed such that closely related species and even individual species exhibit dissimilar mycotoxin chemotypes, whereas in other cases, more distantly related species have similar or the same chemotypes. Balancing selection has contributed to the current distribution of some variation within the afl cluster in Aspergillus and TRI cluster in the F. graminearum species complex (Carbone et al., 2007; Ward et al., 2002). In addition, deletion of biosynthetic genes appears to have contributed to the distribution of fumonisin-production and nonproduction chemotypes in the F. fujikuroi species complex and in black aspergilli (Glenn et al., 2008; Van Hove et al., 2011; Susca et al., 2010) and of aflatoxin production in A. flavus (Chang et al., 2005). It is not clear whether other evolutionary processes (e.g. horizontal transfer, gene duplication, and interspecies hybridization) have also contributed to mycotoxin chemotype variation in Aspergillus and Fusarium. Horizontal transfer of the sterigmatocystin (an aflatoxin analogue) biosynthetic gene cluster from Aspergillus to the fungus Podospora (Slot and Rokas, 2011) and of the bikaverin (a secondary metabolite pigment) from Fusarium to the fungus Botrytis (Campbell et al., 2012) has been reported. It has yet to be determined whether horizontal transfer of mycotoxin biosynthetic gene clusters can occur between Aspergillus and Fusarium. Additional analyses are warranted to evaluate the extent to which horizontal transfer and other evolutionary processes have contributed to variation in mycotoxin chemotypes in Aspergillus and Fusarium. The results of such analyses will contribute to the understanding of how and why fungi differ in ability to produce mycotoxins and aid in the assessment of the risk to food and feed safety that exists between and within fungal species. Such information can also contribute to the development of strategies to control these fungi and the mycotoxin contamination problems they cause.

References Abarca, M.L., Accensi, F., Bragulat, M.R., Castella, G., Cabanes, F.J., 2003. Aspergillus carbonarius as the main source of ochratoxin A contamination in dried wine fruits from the Spanish market. Journal of Food Protection 66, 504–506. Abarca, M.L., Accensi, F., Cano, J., Cabanes, F.J., 2004. Taxonomy and significance of black aspergilli. Antonie Van Leeuwenhoek 86, 33–49. Abbas, H.K., Zablotowicz, R.M., Bruns, H.A., Abel, C.A., 2006. Biocontrol of aflatoxin in corn by inoculation with non-aflatoxigenic Aspergillus flavus isolates. Biocontrol Science and Technology 16, 437–449. Alexander, N.J., McCormick, P., Waalwijk, C., van der Lee, T., Proctor, R.H., 2011. The genetic basis for 3-ADON and 15-ADON trichothecene chemotypes in Fusarium. Fungal Genetics and Biology 48, 485–495. Azziz-Baumgartner, E., Lindblade, K., Gieseker, K., Rogers, H.S., Kieszak, S., Njapau, H., Schleicher, R., McCoy, L.F., Misore, A., DeCock, K., Rubin, C., Slutsker, L., 2005. Case–control study of an acute aflatoxicosis outbreak, Kenya. Environmental Health Perspectives 113, 1779–1783. Branham, B.E., Plattner, R.D., 1993. Alanine is a precursor in the biosynthesis of fumonisin B1 by Fusarium moniliforme. Mycopathologia 124, 99–104. Brown, D.W., McCormick, S.P., Alexander, N.J., Proctor, R.H., Desjardins, A.E., 2001. A genetic and biochemical approach to study trichothecene diversity in Fusarium sporotrichioides and Fusarium graminearum. Fungal Genetics and Biology 32, 121–133. Brown, D.W., McCormick, S.P., Alexander, N.J., Proctor, R.H., Desjardins, A.E., 2002. Inactivation of a cytochrome P-450 is a determinant of trichothecene diversity in Fusarium species. Fungal Genetics and Biology 36, 224–233. Brown, D.W., Proctor, R.H., Dyer, R.B., Plattner, R.D., 2003. Characterization of a Fusarium 2-gene cluster involved in trichothecene C-8 modification. Journal of Agricultural and Food Chemistry 51, 7936–7944. Brown, D.W., Dyer, J.M., McCormick, S.P., Kendra, D.F., Plattner, R.D., 2004. Functional demarcation of the Fusarium core trichothecene gene cluster. Fungal Genetics and Biology 41, 454–462.

Brown, D.W., Butchko, R.A.E., Busman, M., Proctor, R.H., 2007. The Fusarium verticillioides FUM gene cluster encodes a Zn(II)2Cys6 protein that affects FUM gene expression and fumonisin production. Eukaryotic Cell 6, 1210–1218. Butchko, R.A.E., Plattner, R.D., Proctor, R.H., 2003a. FUM9 is required for C-5 hydroxylation of fumonisins and complements the meitotically defined Fum3 locus in Gibberella moniliformis. Applied and Environmental Microbiology 69, 6935–6937. Butchko, R.A.E., Plattner, R.D., Proctor, R.H., 2003b. FUM13 encodes a short chain dehydrogenase/reductase required for C-3 carbonyl reduction during fumonisin biosynthesis in Gibberella moniliformis. Journal of Agricultural and Food Chemistry 51, 3000–3006. Butchko, R.A.E., Plattner, R.D., Proctor, R.H., 2006. Deletion analysis of FUM genes involved in tricarballylic ester formation during fumonisin biosynthesis. Journal of Agricultural and Food Chemistry 54, 9398–9404. Cabanes, F.J., Accensi, F., Bragulat, M.R., Abarca, M.L., Castella, G., Minguez, S., Pons, A., 2002. What is the source of ochratoxin A in wine? International Journal of Food Microbiology 79, 213–215. Campbell, M.A., Rokas, A., Slot, J.C., 2012. Horizontal transfer and death of fungal secondary metabolic gene cluster. Genome Biology and Evolution 4, 289–293. Carbone, I., Ramirez-Prado, J.H., Jakobek, J.L., Horn, B.W., 2007. Gene duplication, modularity and adaptation in the evolution of the aflatoxin gene cluster. BMC Evolutionary Biology 7, 111 (http://www.biomedcentral.com/1471-2148/7/111). CAST (Council for Agricultural Science, Technology), 2003. Mycotoxins. Risks in plants, animals and human systems. Task Force Report,139 January, 2003 (Ames, Iowa, U.S.A.). Chandler, E.A., Simpson, D.R., Thomsett, M.A., Nicholson, P., 2003. Development of PCR assays to Tri7 and Tri13 trichothecene biosynthetic genes, and characterisation of chemotypes of Fusarium graminearum, Fusarium culmorum and Fusarium cerealis. Physiology and Molecular Plant Pathology 62, 355–367. Chang, P.K., Horn, B.W., Dorner, J.W., 2005. Sequence breakpoints in the aflatoxin biosynthesis gene cluster and flanking regions in nonaflatoxigenic Aspergillus flavus isolates. Fungal Genetics and Biology 42, 914–923. Chang, P.K., Ehrlich, K.C., Hua, S.S.T., 2006. Cladal relatedness among Aspergillus oryzae isolates and Aspergillus flavus S and L morphotype isolates. International Journal of Food Microbiology 108, 172–177. Christen, P., Mehta, P.K., 2001. From cofactor to enzymes: the molecular evolution of pyridoxal-5′-phosphate-dependent enzymes. Chemical Record 1, 436–447. Cotty, P.J., 1994. Influence of field application of an atoxigenic strain of Aspergillus flavus on the populations of A. flavus infecting cotton bolls and on the aflatoxin content of cottonseed. Phytopathology 84, 1270–1277. Cotty, P.J., 2006. Biocompetitive exclusion of toxigenic fungi. In: Barug, D., Bhatnagar, D., van Egmond, H.P., van der Kamp, J.W., van Osenbruggen, W.A., Visconti, A. (Eds.), The Mycotoxin Factbook. Wageningen Academic Publishers, The Netherlands (400 pp.). Criseo, G., Racco, C., Romeo, O., 2008. High genetic variability in non-aflatoxigenic Aspergillus flavus strains by using Quadruplex PCR-based assay. International Journal of Food Microbiology 125, 341–343. Cuomo, C.A., Güldener, U., Xu, J.R., Trail, F., Turgeon, B.G., Pietro, A.D., Walton, J.D., Ma, L.J., Rep, M., Adam, G., Antoniw, J., Balswin, T., Calvo, S., Chang, Y.L., DeCaprio, D., Gale, L.R., Desjardins, A.E., 2006. Fusarium Mycotoxins Chemistry, Genetics and Biology. APS Press, St. Paul 1–260. Degola, F., Berni, E., Dall'Asta, C., Spotti, E., Marchelli, R., Ferrero, I., Restivo, F.M., 2007. A multiplex RT-PCR approach to detect aflatoxigenic strains of Aspergillus flavus. Journal of Applied Microbiology 103, 409–417. Deng, J., Carbone, I., Dean, R.A., 2007. The evolutionary history of cytochrome P450 genes in four filamentous ascomycetes. BMC Evolutionary Biology 7, 30. Desjardins, A.E., 2006. Fusarium Mycotoxins Chemistry, Genetics and Biology. APS Press, St. Paul. Desjardins, A.E., Proctor, R.H., Bai, G., McCormick, S.P., Shaner, G., Buechley, G., Hohn, T.M., 1996. Reduced virulence of trichothecene-nonproducing mutants of Gibberella zeae in wheat field tests. Molecular Plant-Microbe Interactions 9, 775–781. Ding, Y., Bojja, R.S., Du, L., 2004. Fum3p, a 2-ketoglutarate-dependent dioxygenase required for C-5 hydroxylation of fumonisins in Fusarium verticillioides. Applied and Environmental Microbiology 70, 1931–1934. Dorner, J.W., Horn, B.W., 2007. Separate and combined applications of non toxigenic Aspergillus flavus and A. parasiticus for biocontrol of aflatoxin in peanuts. Mycopathologia 163, 215–223. Ehrlich, K.C., Montalbano, B.G., Cotty, P.J., 2003. Sequence comparison of aflR from different Aspergillus species provides evidence for variability in regulation of aflatoxin production. Fungal Genetics and Biology 38, 63–74. FAO (Food and Agriculture Organization), 2004. Worldwide Regulations for Mycotoxins in Foods and Feeds in 2003. FAO Food and Nutrition Paper 81 (Rome, Italy). Frisvad, J.C., Skouboe, P., Samson, R.A., 2005. Taxonomic comparison of three different groups of aflatoxin producers and a new efficient producer of aflatoxin B1, sterigmatocystin and 3-O-methylsterigmatocystin, Aspergillus rambellii sp. nov. Systematic and Applied Microbiology 28, 442–453. Frisvad, J.C., Smedsgaard, J., Samson, R.A., Larsen, T.O., Trane, U., 2007. Fumonisin B2 production by Aspergillus niger. International Journal of Food Microbiology 55, 9727–9732. Frisvad, J.C., Larsen, T.O., Thrane, U., Meijer, M., Varga, J., Samson, R.A., 2011. Fumonisin and ochratoxin production in industrial Aspergillus niger strains. PLoS One 6 (8), e23496. http://dx.doi.org/10.1371/journal.pone.0023496. Gallo, A., Stea, G., Battilani, P., Logrieco, A.F., Perrone, G., 2012. Molecular characterization of an Aspergillus flavus population isolated from maize during the first outbreak of aflatoxin contamination in Italy. Phytopathologia Mediterranea 51, 198–206. Gams, W., Christensen, M., Onions, A.H.S., Pitt, J.I., Samson, R.A., 1985. Infrageneric taxa of Aspergillus. In: Samson, R.A., Pitt, J.I. (Eds.), Advances in Aspergillus Systematics. Plenum Press, New York, pp. 55–64.

Please cite this article as: Moretti, A., et al., Molecular biodiversity of mycotoxigenic fungi that threaten food safety, International Journal of Food Microbiology (2013), http://dx.doi.org/10.1016/j.ijfoodmicro.2013.06.033

A. Moretti et al. / International Journal of Food Microbiology xxx (2013) xxx–xxx Geiser, D.M., Aoki, T., Bacon, C.W., Baker, S.E., Bhattacharyya, M.K., Brandt, M.E., Brown, D.W., Burgess, L.W., Chulze, S., Coleman, J.J., Correll, J.C., Covert, S.F., Crous, P.W., Cuomo, C.A., De Hoog, G.S., Di Pietro, A., Elmer, W.H., Epstein, L., Frandsen, R.J.N., Freeman, S., Gagkaeva, T., Glenn, A.E., Gordon, T.R., Gregory, N.F., Hammond-Kosack, K.E., Hanson, L.E., del Mar Jiménez-Gasco, M., Kang, S., Kistler, H.C., Kuldau, G.A., Leslie, J.F., Logrieco, A., Lu, G., Lysøe, E., Ma, L.-J., McCormick, S.P., Migheli, Q., Moretti, A., Munaut, F., O'Donnell, K., Pfenning, L., Ploetz, R.C., Proctor, R.H., Rehner, S.A., Robert, V.A.R.G., Rooney, A.P., bin Salleh, B., Scandia, M.M., Scauflaire, J., Short, D.P.G., Steenkamp, E., Suga, H., Summerell, B.A., Sutton, D.A., Thrane, U., Trail, F., Van Diepeningen, A., VanEtten, H.D., Viljoen, A., Waalwijk, C., Ward, T.J., Wingfield, M.J., Xu, J.-R., Yang, X.-B., Yli-Mattila, T., Zhang, N., 2013. One fungus, one name: defining the genus Fusarium in a scientifically robust way that preserves longstanding use. Phytopathology 103, 400–408. Gelderblom, W.C.A., Jazwinski, S.M., Marasas, W.F.O., Thiel, P.G., Vleggaar, R., Kriek, N.P.J., 1988. Fumonisins — novel mycotoxins with cancer-promoting activity produced by Fusarium moniliforme. Applied and Environmental Microbiology 54, 1806–1811. Glenn, A.E., Zitomer, N.C., Zimeri, A.M., Williams, L.D., Riley, R.T., Proctor, R.H., 2008. Transformation-mediated complementation of a FUM gene cluster deletion in Fusarium verticillioides restores both fumonisin production and pathogenicity on maize seedlings. Molecular Plant-Microbe Interactions 87–97. Horn, B.W., Dorner, J.W., 1999. Regional differences in production of aflatoxin B1 and cyclopiazonic acid by soil isolates of Aspergillus flavus along a transect within the United States. Applied and Environmental Microbiology 65, 1444–1449. IARC (International Agency for Cancer Research), 1993. Some naturally occurring substances: food items and constituents, heterocyclic aromatic amines and mycotoxins. IARC Monographs on the Evaluation of Carcinogenic Risks to Humans 56, 1–599. Keller, N.P., Turner, G., Bennett, J.W., 2005. Fungal secondary metabolism — from biochemistry to genomics. Nature Reviews Microbiology 3, 937–947. Kimura, M., Kaneko, I., Komiyama, M., Takatsuki, A., Koshino, H., Yoneyama, K., Yamaguchi, I., 1998a. Trichothecene 3-O-acetyltransfease protects both the producing organism and transformed yeast from related mycotoxins. Journal of Biological Chemistry 273, 1654–1661. Kimura, M., Matsumoto, G., Shingu, Y., Yoneyama, K., Yamaguchi, I., 1998b. The mystery of the trichothecene 3-O-acetyltransferase gene analysis of the region around Tri101 and characterization of its homologue from Fusarium sporotrichioides. FEBS Letters 435, 163–168. Kimura, M., Tokai, T., O'Donnell, K., Ward, T.J., Fujimura, M., Hamamoto, H., Shibata, T., Yamaguchi, I., 2003. The trichothecene biosynthesis gene cluster of Fusarium graminearum F15 contains a limited number of essential pathway genes and expressed non-essential genes. FEBS Letters 359, 105–110. Kvas, M., Marasas, W.F.O., Wingfield, B.D., Wingfield, M.J., Steenkamp, E.T., 2009. Diversity and evolution of Fusarium species in the Gibberella fujikuroi complex. Fungal Diversity 34, 1–21. Lee, T., Han, Y.K., Kim, K.H., Yun, S.H., Lee, Y.W., 2002. Tri13 and Tri7 determine deoxynivalenol- and nivalenol-producing chemotypes of Gibberella zeae. Applied and Environmental Microbiology 68, 2148–2154. Lee, J., Jurgenson, J.E., Leslie, J.F., Bowden, R.L., 2008. Alignment of genetic and physical maps of Gibberella zeae. Applied and Environmental Microbiology 74, 2349–2359. Leslie, J.F., Summerell, B.A., 2006. The Fusarium Laboratory Manual. Blackwell Publishing, Ames (1-388 pp.). Maier, F.J., Miedaner, T., Hadeler, B., Felk, A., Salomon, S., Lemmens, M., Kassner, H., Schäfer, W., 2006. Involvement of trichothecenes in fusarioses of wheat, barley and maize evaluated by gene disruption of the trichodiene synthase (Tri5) gene in three field isolates of different chemotype and virulence. Molecular Plant Pathology 7, 449–461. Mansson, M., Klejnstrup, M.L., Phipps, R.K., Nielsen, K.F., Frisvad, J.C., Gotfredsen, C.H., Larsen, T.O., 2010. Isolation and NMR characterization of fumonisin B2 and a new fumonisin B6 from Aspergillus niger. Journal of Agricultural and Food Chemistry 58, 949–953. Marasas, W.F.O., Riley, R.T., Hendricks, K., Stevens, V.L., Sadler, T.W., Gelineau-van Waes, J., Missmer, Cabrera, J., Torres, O., Gelderblom, W.C.A., Allegood, J., Martínez, C., Maddox, J., Miller, J.D., Starr, L., Sullards, M.C., Roman, A.V., Voss, K.A., Wang, E., Merrill, A.H., 2004. Fumonisins disrupt shingolipid metabolism, folate transport and neural tube development in embryo culture and in vivo: a potential risk factor for human neural tube defects among populations consuming fumonisin-contaminated maize. Journal of Nutrition 134, 711–716. Marasas, W.F.O., Gelderblom, W.C.A., Vismer, H.F., 2008. Mycotoxins: a global problem. In: Leslie, J.F., Bandyopadhayay, R., Visconti, A. (Eds.), Mycotoxins. CABI, Oxfordshire, UK, pp. 29–40. McCormick, S.P., Alexander, N.J., Trapp, S.C., Hohn, T.M., 1999. Disruption of TRI101, the gene encoding trichothecene 3-O-acetyltransferase, from Fusarium sporotrichioides. Applied and Environmental Microbiology 65, 5252–5256. McCormick, S.P., Harris, L.J., Alexander, N.J., Ouellet, T., Saparno, A., Allard, S., Desjardins, A.E., 2004. Tri1 in Fusarium graminearum encodes a P450 oxygenase. Applied and Environmental Microbiology 70, 2044–2051. McCormick, S.P., Alexander, N.J., Proctor, R.H., 2006. Heterologous expression of two trichothecene P450 genes in Fusarium verticillioides. Canadian Journal of Microbiology 52, 220–226. Medina, A., Mateo, R., Lopez-Ocana, L., Valle-Algarra, F.M., 2005. Study of Spanish grape microbiota and ochratoxin A production by isolates of Aspergillus tubingensis and other members of Aspergillus section nigri. Applied and Environmental Microbiology 71, 4696–4702. Meek, I.B., Peplow, A.W., Ake, C., Phillips, T.D., Beremand, M.N., 2003. Tri1 encodes the cytochrome P450 monooxygenase for C-8 hydroxylation during trichothecene biosynthesis in Fusarium sporotrichioides and resides upstream of another new Tri gene. Applied and Environmental Microbiology 69, 1607–1613.

9

Mirete, S., Vásquez, C., Mulè, G., Jurado, M., Gonzáles-Jaén, M.T., 2004. Differentiation of Fusarium verticillioides from banana fruits by IGS and EF-1 sequence analyses. European Journal of Plant Pathology 110, 515–523. Moore, G.G., Singh, R., Horn, B.W., Carbone, I., 2009. Recombination and lineage-specific gene loss in the aflatoxin gene cluster of Aspergillus flavus. Molecular Ecology 18, 4870–4887. Musser, S.M., Plattner, R.D., 1997. Fumonisin composition in cultures of Fusarium moniliforme, Fusarium proliferatum and Fusarium nygami. Journal of Agricultural and Food Chemistry 45, 1169–1173. Nelson, P.E., Plattner, R.D., Shackelford, D.D., Desjardins, A.E., 1992. Fumonisin B1 production by Fusarium species other than F. moniliforme in Section Liseola and by some related species. Applied and Environmental Microbiology 58, 984–989. Noonim, P., Wahakarnchanakul, W., Nielsen, K.F., Frisvad, K.F., Samson, R.A., 2009. Fumonisin B2 production by Aspergillus niger in Thai coffee beans. Food Additives and Contaminants. Part A 26, 94–100. O'Donnell, K., Cigelnik, E., Nirenberg, H.I., 1998. Molecular systematics and phylogeography of the Gibberella fujikuroi species complex. Mycologia 90, 465–493. O'Donnell, K., Nirenberg, H.I., Aoki, T., Cigelnik, E., 2000. A multigene phylogeny of the Gibberella fujikuroi species complex: detection of additional phylogenetically distinct species. Mycoscience 41, 61–78. O'Donnell, K., Sutton, D.A., Rinaldi, M.G., Gueidan, C., Crous, P.W., Geiser, D.M., 2009. Novel multilocus sequence typing scheme reveals high genetic diversity of human pathogenic members of the Fusarium incarnatum–F. equiseti and F. chlamydosporum species complexes within the United States. Journal of Clinical Microbiology 47, 3851–3861. O'Donnell, K., Rooney, A.P., Proctor, R.H., Brown, D.W., McCormick, S.P., Ward, T.J., Frandsen, R.J.N., Lysøe, E., Rehner, S.A., Aoki, T., Robert, V.A., Crous, P.W., Groenewald, J.Z., Kang, S., Geiser, D.M., 2013. Phylogenetic analyses of RPB1 and RPB2 strongly support a middle Cretaceous origin for a clade comprising all agriculturally and medically important fusaria. Fungal Genetics and Biology 52, 20–31. Palumbo, J.D., O'Keeffe, T.L., 2013. Multiplex PCR analysis of fumonisin biosynthetic genes in fumonisin-nonproducing Aspergillus niger and A. awamori strains. Mycologia 105 (2), 277–284 (March/April). Palumbo, J.D., O'Keeffe, T.L., Vasquez, S.J., Mahoney, N.E., 2011. Isolation and identification of ochratoxin A-producing Aspergillus section Nigri strains from California raisins. Letters in Applied Microbiology 52, 330–336. Peplow, A.W., Meek, I.B., Wiles, M.C., Phillips, T.D., Beremand, M.N., 2003. Tri16 is required for esterification of position C-8 during trichothecene mycotoxin production by Fusarium sporotrichioides. Applied and Environmental Microbiology 69, 5935–5940. Perrone, G., Stea, G., Epifani, F., Varga, J., Frisvad, J.C., Samson, R.A., 2011. Aspergillus niger contains the cryptic phylogenetic species A. awamori. Fungal Biology 115, 1138–1150. Pildain, M.B., Vaamonde, G., Cabral, D., 2004. Analysis of population structure of Aspergillus flavus from peanut based on vegetative compatibility, geographic origin, mycotoxin and sclerotia production. International Journal of Food Microbiology 93, 31–40. Pitt, J.I., Hocking, A.D., 2007. Fungi and Food Spoilage, 2nd edn. Blackie, London. Pitt, J.I., Hocking, A.D., 2009. Fungi and Food Spoilage, 3rd edn. Springer, London. Proctor, R.H., Brown, D.W., Plattner, R.D., Desjardins, A.E., 2003. Co-expression of 15 contiguous genes delineates a fumonisin biosynthetic gene cluster in Gibberella moniliformis. Fungal Genetics and Biology 38, 237–249. Proctor, R.H., Plattner, R.D., Brown, D.W., Seo, J.A., Lee, Y.W., 2004. Discontinuous distribution of fumonisin biosynthetic genes in the Gibberella fujikuroi species complex. Mycological Research 108, 815–822. Proctor, R.H., Plattner, R.D., Desjardins, A.E., Busman, M., Butchko, R.A.E., 2006. Fumonisin production in the maize pathogen Fusarium verticillioides: genetic basis of naturally occurring chemical variation. Journal of Agriculture and Food Chemistry 54, 2424–2430. Proctor, R.H., Busman, M., Seo, J.A., Lee, Y.W., Plattner, R.D., 2008. A fumonisin biosynthetic gene cluster in Fusarium oxysporum strain O-1890 and the genetic basis for B versus C fumonisin production. Fungal Genetics and Biology 45, 1016–1026. Proctor, R.H., McCormick, S.P., Alexander, N.J., Desjardins, A.E., 2009. Evidence that a secondary metabolic biosynthetic gene cluster has grown by gene relocation during evolution of the filamentous fungus Fusarium. Molecular Microbiology 74, 1128–1142. Proctor, R.H., Van Hove, F., Susca, A., Stea, G., Busman, M., van der Lee, T., Waalwijk, C., Moertti, A., Ward, T.J., 2013. Evidence for birth-and-death evolution and horizontal transfer of the fumonisin mycotoxin biosynthetic gene cluster in Fusarium. Proceedings of the Twenty-Seventh Fungal Genetics Conferences. Pacific Grove California (Abstract). Raper, K.B., Fennell, D.I., 1965. The genus Aspergillus. Williams & Wilkins Co., Baltimore. Rheeder, J.P., Marasas, W.F.O., Vismer, H.F., 2002. Production of fumonisin analogs by Fusarium species. Applied and Environmental Microbiology 68, 2101–2105. Sage, L., Garon, D., Seigle-Murandi, F., 2004. Fungal microflora and ochratoxin A risk in French wineyards. Journal of Agricultural and Food Chemistry 52, 5764–5768. Samson, R.A., Hoekstra, E.S., Frisvad, J.C., 2004. Introduction to Food and Airborne Fungi, 7th edn. Centraalbureau voor Schimmelcultures, Utrecht. Sarver, B.A., Ward, T.J., Gale, L.R., Broz, K.L., Kistler, H.C., Aoki, T., Nicholson, P., Carter, J., O'Donnell, K., 2011. Novel Fusarium head blight pathogens from Nepal and Louisiana revealed by multilocus genealogical concordance. Fungal Genetics and Biology 48, 1077–1152. Scherm, B., Palomba, M., Serra, D., Marcello, A., Migheli, Q., 2005. Detection of transcripts of the aflatoxin genes aflD, aflO, and aflP by reverse transcription-polymerase chain reaction allows differentiation of aflatoxin-producing and non-producing isolates of Aspergillus flavus and Aspergillus parasiticus. International Journal of Food Microbiology 98, 201–210. Seo, J.A., Kim, J.C., Lee, Y.W., 1996. Isolation and characterization of two new type C fumonisins produced by Fusarium oxysporum. Journal of Natural Products 59, 1003–1005.

Please cite this article as: Moretti, A., et al., Molecular biodiversity of mycotoxigenic fungi that threaten food safety, International Journal of Food Microbiology (2013), http://dx.doi.org/10.1016/j.ijfoodmicro.2013.06.033

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Sewram, V., Mshicileli, N., Shepard, G.S., Vismer, H.F., Rheeder, J.P., Lee, Y.W., Leslie, J.F., Marasas, W.F.O., 2005. Production of fumonisin B and C analogues by several Fusarium species. Journal of Agricultural and Food Chemistry 53, 4861–4866. Slot, J.C., Rokas, A., 2011. Horizontal transfer of a large and highly toxic secondary metabolic gene cluster between fungi. Current Biology 21, 134–139. Starkey, D.E., Ward, T.J., Aoki, T., Gale, L.R., Kistler, H.C., Geiser, D.M., Suga, H., Tóth, B., Varga, J., O'Donnell, K., 2007. Global molecular surveillance reveals novel Fusarium head blight species and trichothecene toxin diversity. Fungal Genetics and Biology 44, 1191–1204. Stepien, L., Koczyk, C., Waskiewicz, A., 2010. FUM cluster divergence in fumonisinsproducing Fusarium species. Fungal Biology 115, 112–123. Summerell, B.A., Laurence, M.H., Liew, E.C.Y., Leslie, J.F., 2010. Biogeography and phylogeography of Fusarium: a review. Fungal Diversity 44, 3–13. Susca, A., Proctor, R.H., Mule, G., Stea, G., Ritieni, A. Logrieco A., Moretti, A., 2010. Correlation of mycotoxin fumonisin B2 production and presence of the fumonisin biosynthetic gene fum8 in Aspergillus niger from grape. Journal of Agricultural and Food Chemistry 58, 9266–9272. Sweeney, M.J., Pàmies, P., Dobson, A.D.W., 2000. The use of reverse transcriptionpolymerase chain reaction (RTPCR) for monitoring aflatoxin production in Aspergillus parasiticus 439. International Journal of Food Microbiology 56, 97–103. Tokai, T., Fujimura, M., Inoue, H., Aoki, T., Ohta, K., Shibata, T., Yamaguchi, I., Kimura, M., 2005. Concordant evolution of trichothecene 3-O-acetyltransferase and rDNA species phylogeny of trichothecene-producing and nonproducing fusaria and other ascomycetous fungi. Microbiology 151, 509–519. Uhlig, S., Busman, M., Shane, D.S., Rønning, H., Rise, F., Proctor, R.H., 2012. Identification of early fumonisin biosynthetic intermediates by inactivation of the FUM6 gene in Fusarium verticillioides. Journal of Agricultural and Food Chemistry 60, 10293–10301. Vaamonde, G., Patriarca, A., Pinto, V.F., Comerio, R., Degrossi, C., 2003. Variability of aflatoxin and cyclopiazonic acid production by Aspergillus section Flavi from different substrates in Argentina. International Journal of Food Microbiology 88, 79–84.

Van Hove, F., Waalwijk, C., Logrieco, A., Munaut, F., Moretti, A., 2011. Gibberela musae (Fusarium musae) sp. nov.: a new species from banana is sister to F. verticillioides. Mycologia 103, 570–585. Varga, J., Kevei, F., Hamari, Z., Tóth, B., Téren, J., Croft, J.H., Kozakiewicz, Z., 2000. Genotypic and phenotypic variability among black aspergilli. In: Samson, R.A., Pitt, J.I. (Eds.), Integration of Modern Taxonomic Methods for Penicillium and Aspergillus Classification. Harwood Academic Publishers, Amsterdam, pp. 397–411. Varga, J., Kocsubé, S., Suri, S., Szigeti, G., Szekeres, A., Varga, M., Tóth, B., Bartók, T., 2010. Fumonisin contamination and fumonisin producing black Aspergilli in dried vine fruits of different origin. International Journal of Food Microbiology 143, 143–149. Waalwijk, C., van der Lee, T., de Vries, I., Hesselink, T., Arts, J., Kema, G.H.J., 2004. Synteny in toxigenic Fusarium species: the fumonisin gene cluster and the mating type region as examples. European Journal of Plant Pathology 110, 533–544. Ward, T.J., Bielawski, J.P., Kistler, H.C., Sullivan, E., O'Donnell, K., 2002. Ancestral polymorphism and adaptive evolution in the trichothecene mycotoxin gene cluster of phytopathogenic Fusarium. Proceedings of National Academy of Sciences USA 99, 9278–9283. Wicklow, D.T., Dowd, P.F., Alftafta, A.A., Gloer, J.B., 1996. Ochratoxin A: an antiinsectan metabolite from the sclerotia of Aspergillus carbonarius NRRL 369. Canadian Journal of Microbiology 42, 1100–1103. Windels, C.E., 2000. Economic and social impacts of Fusarium head blight: changing farms and rural communities in the Northern great plains. Phytopathology 90, 17–21. Yi, H., Bojja, R., Fu, J., Du, L., 2005. Direct evidence for the function of FUM13 in 3ketoreduction of mycotoxin fumonisins in Fusarium verticillioides. Journal of Agricultural and Food Chemistry 53, 5456–5460. Yu, J., Chang, P.K., Ehrlich, K.C., Cary, J.W., Bhatnagar, D., Cleveland, T.E., Payne, G.A., Linz, J.E., Woloshuk, C.P., Bennett, J.W., 2004. Clustered pathway genes in aflatoxin biosynthesis. Applied and Environmental Microbiology 70, 1253–1262. Zaleta-Rivera, K., Xu, C., Yu, F., Butchko, R.A.E., Proctor, R.H., Hidalgo-Lara, M.E., Raza, A., Dussault, P.H., Du, L., 2006. A bidomain nonribosomal peptide synthetase encoded by FUM14 catalyzes the formation of tricarballylic esters in the biosynthesis of fumonisins. Biochemistry 45, 2561–2569.

Please cite this article as: Moretti, A., et al., Molecular biodiversity of mycotoxigenic fungi that threaten food safety, International Journal of Food Microbiology (2013), http://dx.doi.org/10.1016/j.ijfoodmicro.2013.06.033