Genes, gene clusters, and biosynthesis of trichothecenes and fumonisins in Fusarium

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Genes, gene clusters, and biosynthesis of trichothecenes and fumonisins inFusariumNancy J. Alexander a; Robert H. Proctor a; Susan P. McCormick a

a U.S. Department of Agriculture, Agriculture Research Service, National Center for Agricultural UtilizationResearch, Mycotoxin Research Unit, Peoria, Illinois

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R E v i E w A R T i C L E

Genes, gene clusters, and biosynthesis of trichothecenes and fumonisins in Fusarium

Nancy J. Alexander, Robert H. Proctor, and Susan P. McCormick

U.S. Department of Agriculture, Agriculture Research Service, National Center for Agricultural Utilization Research, Mycotoxin Research Unit, Peoria, Illinois

Address for Correspondence: Robert H. Proctor, United States Department of Agriculture, Agriculture Research Service, National Center for Agricultural Utilization Research, Mycotoxin Research Unit, Peoria, Illinois 61604; Tel. 309-681-6380. E-mail: [email protected] mention of firm names or trade products does not imply that they are endorsed or recommended by the United States Department of Agriculture over other firms or similar products not mentioned.

(Received 10 March 2009; revised 06 April 2009; accepted 07 April 2009)

Introduction

Trichothecenes and fumonisins are families of mycotoxins produced by the filamentous fungus Fusarium and are of concern because they are toxic and can accumulate in crop plants infected with the fungus. The two toxin families differ in biogenic origin, structure, and mechanisms of toxicity. Trichothecenes are products of isoprenoid metabolism and have a tricyclic nucleus with an epoxide function. Multiple health problems in humans and animals induced by trichothecenes most likely result from the ability of the toxins to inhibit protein synthesis. Fumonisins

are products of polyketide and amino acid metabo-lism and have a linear structure with amine and tricarballylic ester functions. The multiple health problems in animals, and potentially humans, caused by fumonisins are thought to result from the ability of the toxins to disrupt sphingolipid biosynthesis.

Identification of Fusarium species can be diffi-cult, and field isolates of the fungus have frequently been misidentified. In addition, multiple species are undergoing taxonomic revisions or have under- gone revisions over the past two decades. For exam-ple, the trichothecene-producing species Fusarium graminearum was recently resolved into as many as 12

ISSN 1556-9543 print/ISSN 1556-9551 online © 2009 Informa UK LtdDOI: 10.1080/15569540903092142

AbstractTrichothecenes and fumonisins are mycotoxins produced by Fusarium, a filamentous fungus that can cause disease in barley, maize, rice, wheat, and some other crop plants. Research on the genetics and biochemistry of trichothecene and fumonisin biosynthesis has provided important insights into the genetic and biochemical pathways in Fusarium that lead to formation of these mycotoxins. In Fusarium, trichothecene biosynthetic enzymes are encoded by genes at three loci: the single-gene TRI101 locus, the two-gene TRI1-TRI16 locus, and the 12-gene core TRI cluster. In contrast, fumonisin biosynthetic enzymes identified to date are all located at one locus, the 17-gene FUM cluster. The FUM and core TRI clusters also encode proteins that regulate expression of the cluster genes and proteins that are involved in mycotoxin transport across the cell membrane. Biosynthetic pathways for both mycotoxins have been proposed based on a combination of biochemical and genetic evidence, including toxin production phenotypes of Fusarium mutants in which individual TRI or FUM genes have been inactivated. Some TRI and FUM gene mutants have also been employed to examine the role of mycotoxin production in plant pathogenesis. The studies indicate that trichothecene production can contribute to the ability of F. graminearum to cause wheat head blight, one of the most important wheat diseases in the world. Thus, studies into the genetic basis of mycotoxin production have identified a potential target to enhance resistance of wheat to a major plant disease and mycotoxin contamination problem.

Keywords: Wheat head blight; scab; corn ear rot; deoxynivalenol; T-2 toxin; fumonisin; fusarium graminearum; fusarium sporotrichioides; F. verticillioides

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Genes, gene clusters, and biosynthesis of trichothecenes and fumonisins in Fusarium 199

species (Leslie et al., 2007; O’Donnell et al., 2006; Starkey et al., 2007). Despite the complexities of Fusarium tax-onomy, it is evident that production of trichothecenes and fumonisins is limited to certain Fusarium spe-cies. Fumonisin production has been reproducibly demonstrated only in Fusarium species within the Gibberella fujikuroi species complex and rare isolates of the closely related species, F. oxysporum (Proctor et al., 2004; Rheeder et al., 2002). Trichothecene production appears to be limited to a different group of fusaria that includes the economically important species F. culmorum, F. graminearum, F. poae, F. sambucinum, and F. sporotrichioides (Desjardins, 2006; Marasas et al., 1984). Even though only a limited number of Fusarium species can produce trichothecenes and fumonisins, species in other fungal genera can also produce the toxins. Trichothecene production has been reported in certain species of Myrothecium, Stachybotrys, Trichoderma, and Trichothecium (Cole and Cox, 1981), and fumonisin production has been reported in Aspergillus niger (Frisvad et al., 2007).

This article reviews genetic and biochemical path-ways required for biosynthesis of trichothecenes and fumonisins in Fusarium. The review highlights work done at USDA-ARS-NCAUR. However, the larger body of excellent work done at other institutions in Asia, Europe, and North America is also discussed.

Trichothecenes

F. graminearum (teleomorph Gibberella zeae) has been the subject of intense study for almost two decades because it is the primary cause of wheat head blight, one of the world’s most economically important wheat diseases. It can also contaminate wheat and other crops, such as barley and maize, with trichothecenes (Windels, 2000). The Fusarium trichothecenes of greatest concern are deoxyniva-lenol (DON), acetylated DON, nivalenol (NIV), and acetylated NIV, all of which can be produced by F. graminearum, and T-2 toxin, which is produced by F. sporotrichioides (Figure 1). These and other tri-chothecenes differ in structure by the position and number of hydroxyl and ester functions. Important structural differences between trichothecenes include the presence or absence of oxygen atoms at carbon atoms 7 (C-7) and 8 (C-8). NIV and DON have a hydroxyl function at C-7 and a carbonyl (keto) function at C-8, whereas T-2 toxin lacks a C-7 oxygen and the C-8 oxygen is acylated. Another significant structural difference among trichothecenes is the presence or absence of an oxygen atom at C-4. DON and acetylated DON lack a C-4 oxygen, whereas NIV, acetylated NIV, and T-2 toxin have an oxygen at C-4.

Trichothecene biosynthesis: genes and gene cluster

Studies into the genetic basis of trichothecene produc-tion at USDA-ARS-NCAUR began in the mid-1980s using F. sporotrichioides and F. sambucinum (tele-omorph Gibberella pulicaris) as model systems. These species produce trichothecenes that lack a C-7 oxygen and that often have a C-8 hydroxyl or ester function (e.g., T-2 toxin, neosolaniol). Genetic analysis of natural variants of F. sambucinum identified a single locus, Tox1, responsible for hydroxylation of C-8 (Beremand and Desjardins, 1988; Beremand and McCormick, 1992). Genetic and biochemical analysis of UV irradiation– induced mutants of F. sporotrichioides that were blocked in T-2 toxin production identified three trichothecene biosynthetic loci: Tox1 was responsi-ble for oxygenation of C-8; Tox3 was responsible for acetylation of the oxygen atom at C-15; and Tox4 was responsible for conversion of the first pathway interme-diate, trichodiene, to an unknown product (Beremand and McCormick, 1992; McCormick et al., 1989, 1990). A more recent genetic analysis of F. graminearum identified another locus, TOX1, that affects the quantity of trichothecenes produced (Jurgenson et al., 2002).

The first trichothecene biosynthetic gene identi-fied and characterized via molecular genetic methods was the gene encoding trichodiene synthase. The gene was identified by purifying the enzyme from F. sporotrichioides, raising antibodies to the purified enzyme, and using the antibody to screen a library of F. sporotrichioides genomic DNA that was expressed in Escherichia coli (Hohn and Beremand, 1989; Hohn and Vanmiddlesworth, 1986). This effort identified a genomic clone carrying the trichodiene synthase gene, which was designated Tri5. In this review, for consistency with proposed genetic nomenclature for plant pathogenic fungi (Yoder et al., 1986), we will use TRI designations (e.g., TRI5) rather than Tri for trichothecene biosynthetic genes. In addition, we will

16 910

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Figure 1. Chemical structures of trichothecene, T-2 toxin, deoxynivalenol (DON), and nivalenol (NIV).


200 N. J. Alexander et al.

use the Tri designation (without italics) to indicate proteins encoded by TRI genes.

At about the same time that TRI5 was identified, it became evident that genes responsible for biosyn-thesis of other fungal secondary metabolites (e.g., aflatoxins and B-lactam antibiotics) were clustered (McCabe et al., 1990; Skory et al., 1992). To determine whether trichothecene biosynthetic genes were also clustered, individual cosmid clones derived from F. sporotrichioides genomic DNA were introduced, by genetic transformation, into the Tox1, Tox3, and Tox4 mutants of the fungus (Hohn et al., 1993). Two of the cosmid clones that included TRI5 restored T-2 toxin production to the Tox3 and Tox4 mutants of F. sporotrichioides, but neither clone restored produc-tion to the Tox1 mutant. These results indicated that the genes (TRI3 and TRI4) corresponding to Tox3 and Tox4 were located in the same region of DNA as TRI5. Subsequent sequence, gene disruption, and expres-sion analyses in F. sporotrichioides identified a gene cluster (Figure 2) that consisted of TRI3, TRI4, TRI5 and nine other genes, which were designated TRI6 through TRI14 (Alexander et al., 1997; Brown et al., 2002, 2004). The cluster encodes two regulatory genes and most of the biosynthetic enzymes necessary for formation of trichothecenes. The gene cluster, desig-nated here as the core TRI cluster, was subsequently identified and characterized in F. graminearum and species closely related to it (Brown et al., 2001, 2002; Ward et al., 2002). In DON-producing strains of F. graminearum, two of the cluster genes, TRI7 and TRI13, are nonfunctional as a result of multiple insertions and deletions within their coding regions, whereas in NIV-producing strains, TRI7 and TRI13 are functional (Figure 2) (Brown et al., 2002; Lee et al., 2002). This difference, combined with the find-ing that TRI13 is responsible for trichothecene C-4 hydroxylation, identified the basis for NIV versus DON production in F. gramineaerum.

In addition to the core TRI cluster, two other Fusarium loci that encode trichothecene biosynthetic

enzymes have been identified. The first locus includes only one trichothecene biosynthetic gene, TRI101, and was identified by screening Fusarium cDNAs that con-fer trichothecene resistance in yeast (Kimura et al., 1998; McCormick et al., 1999). TRI101 encodes an acetyl-transferase that catalyzes esterification of acetyl to the C-3 hydroxyl of trichothecenes. This acetylation reduces the toxicity of trichothecenes to yeast and Fusarium, and therefore likely serves as a self- protection mecha-nism in trichothecene-producing species. The second locus encoding other trichothecene biosynthetic enzymes consists of two genes, TRI1 and TRI16. The F. sporotrichioides and F. graminearum orthologues of TRI1 (FsTRI1 and FgTRI1, respectively) were identified in cDNAs libraries prepared from culture conditions in which core TRI cluster genes were highly expressed (Brown et al., 2003; Meek et al., 2003; McCormick et al., 2004). TRI1 encodes a cytochrome P450 monooxygen-ase, but the deduced amino acid sequences of FsTRI1 and FgTRI1 are only 65% identical. Gene deletion and heterologous expression studies indicate that the two orthologues differ in function. The FsTRI1 enzyme catalyzes hydroxylation of trichothecenes at C-8 only, whereas the FgTRI1 enzyme catalyzes hydroxylation at C-7 and C-8 (McCormick et al., 2006b). This difference in function is consistent with the absence of a C-7 hydroxyl in trichothecenes produced by F. sporotrichioides (e.g., T-2 toxin) and presence of a C-7 hydroxyl in trichothecenes produced by F. graminearum (e.g., DON and NIV). Thus, differences in the FsTRI1 and FgTRI1 orthologues are responsible for an important structural difference in trichothecenes produced by F. sporotrichioides and F. graminearum. In addition, the FsTRI1 orthologue can functionally complement the Tox1 mutant of F. sporotrichioides (Meek et al., 2003). This indicates that TRI1 is most likely the same gene as the Tox1 identified in the early genetic analyses of F. sporotrichioides and F. sambucinum (Beremand and Desjardins, 1988; Beremand and McCormick, 1992).

TRI16 was identified in F. sporotrichioides by its location next to TRI1 and by its high level of expres-sion under conditions that promote expression of core TRI cluster genes (Brown et al., 2003; Peplow et al., 2003a). The gene encodes an acyltransferase, and gene deletion analysis indicated that it is respon-sible for esterification of an isovalerate moiety to the C-8 hydroxyl during formation of T-2 toxin. The F. graminearum TRI16 homologue is nonfunctional due to multiple insertions and deletions in its coding region. This is consistent with the absence of the isovalerate ester in trichothecenes produced by F. graminearum (Brown et al., 2003; McCormick et al., 2004).

Another gene, TRI15, was also identified by its high level of expression under conditions that promote

T-2toxin

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TRI10TRI9

TRI11TRI1

TRI16

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kb 0 4 8 12 16 20 24

Figure 2. A, Core trichothecene biosynthetic clusters and TRI1-TRI16 cluster in Fusarium that lead to formation of T-2 toxin, nivalenol (NIV), or deoxynivalenol (DON). Arrows indicate posi-tion and transcriptional orientation of genes. An X on an arrow indicates that a gene has multiple insertions and/or deletions that render it nonfunctional.



expression of core TRI cluster genes (Alexander et al., 2004). TRI15 is predicted to encode a Cys

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2-type

transcription factor and is highly expressed in an F. sporotrichioides strain with alteration to TRI10 that caused increased expression of core TRI cluster genes (Peplow et al., 2003b). TRI15 also exhibits high levels of expression in the presence of exogenous T-2 toxin. However, TRI15 mutants do not exhibit altered trichothecene production. In the F. graminearum genome sequence, TRI15 is located next to a nonri-bosomal peptide synthetase gene. Therefore, TRI15 may regulate and be located in a gene cluster involved in biosynthesis of a different secondary metabolite, i.e., nonribosomal peptide-derived metabolite. Why TRI15 is expressed so highly in the TRI10 mutant and in the presence of T-2 toxin remains to be determined.

Despite the tight linkage of genes within the core TRI cluster, F. graminearum and F. sporotrichioides have maintained three loci that encode trichothecene biosynthetic enzymes. The availability of the whole genome sequence of F. graminearum as well as genetic linkage analysis using the sexual cycle of the fungus have revealed that the core TRI cluster, TRI101, and TRI1/TRI16 are not only at different loci but also on dif-ferent chromosomes (Gale et al., 2005; Lee et al., 2008).

Coordination of expression of genes at these three loci is mediated, at least in part, by the regu-latory genes, TRI6 and TRI10, located within the core TRI cluster (Peplow et al., 2003b; Tag et al., 2001). Inactivation of either gene by gene disrup-tion reduces or eliminates expression of other TRI genes and blocks trichothecene production (Proctor et al., 1995b; Tag et al., 2001). TRI6 encodes a Cys

2-

His2-type transcription factor that binds to the DNA

sequence motif TNAGGCCT that is found in the pro-moter regions of genes in the core TRI cluster as well as TRI1, TRI16, and TRI101 (Brown et al., 2003; Hohn et al., 1999; McCormick et al., 1999). TRI10 encodes a protein without any known functional domains but does share low levels of identity (30%) with some fun-gal transcription factors (Tag et al., 2001). Thus, TRI6 and TRI10 encode proteins that activate expression of other TRI genes. Both genes can also positively affect expression of some primary metabolic genes involved in the formation of isoprenoid (e.g., farnesyl diphosphate) precursors of trichothecene biosynthe-sis (Peplow et al., 2003b).

Genetic and biochemical pathways for trichothecene biosynthesis

A trichothecene biosynthetic pathway has been pro-posed (Figure 3) based on multiple lines of evidence. Some of the earliest insight into the pathway was

obtained from precursor feeding studies, which pro-vided evidence for the origins of some of the atoms that make up trichothecenes (Desjardins et al., 1986; Zamir et al., 1990). For example, studies with isotopi-cally labeled oxygen indicate that the oxygen atoms of Fusarium trichothecenes are derived from molecular oxygen (O

2) and, therefore, their incorporation into

trichothecene molecules likely results from the activ-ity of monooxygenases or dioxygenases (Desjardins et al., 1986). Insight into the pathway has also been obtained from the diversity of trichothecenes that Fusarium can produce naturally or through manip-ulation of laboratory cultures (Desjardins et al., 1987; Hesketh et al., 1991, 1993; Zamir et al., 1991). Understanding of the pathway has also been aided by trichothecene production phenotypes of mutant strains of Fusarium in which TRI genes have been inactivated by gene disruption or deletion (Table 1) and by the ability of the mutants to convert some biosynthetic intermediates but not others (Alexander et al., 1998; McCormick et al., 1990; McCormick and Hohn, 1997). In a few cases, the roles of TRI genes in the pathway have been confirmed by their expression in nonproducing organisms (Alexander et al., 1999a; Kimura et al., 1998; McCormick et al., 2006b).

Although Figure 3 shows a step-by-step process, trichothecene biosynthesis may be a metabolic grid where the order of some reactions is not fixed (Kimura et al., 2007). Nevertheless, the first step in the pathway is the trichodiene synthase-catalyzed conversion of farnesyl diphosphate, an isoprenoid intermediate in primary metabolism, to trichodiene. This reaction has been demonstrated in vitro with purified trichodiene synthase (Hohn and Vanmiddlesworth, 1986), and it is consistent with the phenotype of F. graminearum and F. sambucinum mutants in which the trichodiene synthase gene, TRI5, has been disrupted (Hohn and Desjardins, 1992; Proctor et al., 1995a). TRI5 mutants do not produce any known trichothecenes or tri-chothecene biosynthetic intermediates.

Once formed, trichodiene undergoes a series of four oxygenation reactions: hydroxylation at C-2, epoxida-tion at C-12/C-13, hydroxylation at C-11, and hydroxy-lation at C-3 (Hesketh et al., 1991, 1993; Zamir et al., 1991, 1999). Two independent studies indicate that all four oxygenation reactions are catalyzed by the cyto-chrome P450 monooxygenase encoded by TRI4. In the first study, TRI4 was expressed in the trichothecene-nonproducing Fusarium species F. verticillioides (McCormick et al., 2006a). In the other study, TRI4 was expressed in the yeast Saccharomyces cerevisiae (Tokai et al., 2007). In both studies, the transgenic organism expressing TRI4 could carry out all four oxygenation reactions. Furthermore, F. graminearum with a functional TRI4 could metabolize the series of



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oxygenated trichodiene metabolites produced by the transgenic organisms, whereas F. graminearum with a nonfunctional TRI4 could not (Tokai et al., 2007).

The next two steps in the proposed biosynthetic pathway can occur nonenzymatically (Kimura et al., 2007; McCormick et al., 2006a). The first is formation of trichotriol by an isomerization that replaces the C-9 hydroxyl with a C-11 hydroxyl. The second step is a cyclization that uses the C-2 oxygen atom to form a pyran ring and results in the formation of isotrichode-rmol, the first biosynthetic intermediate that has the tricyclic nucleus and 12,13-epoxide structure com-mon to all trichothecenes.

From isotrichodermol, the biosynthetic path-way proceeds with esterification of acetyl to the C-3 hydroxyl. As noted earlier, this reaction is catalyzed by the TRI101-encoded acetyltransferase. Evidence for this is that TRI101 mutants of F. graminearum and F. sporotrichioides are blocked in production of C-3 acetylated trichothecenes, and yeast strains heterologously expressing TRI101 can transfer an acetyl moiety to the C-3 hydroxyl (Kimura et al., 1998; McCormick et al., 1999).

The next step in the trichothecene biosynthetic pathway is hydroxylation of the C-15 position. This reaction is catalyzed by TRI11, which encodes a cytochrome P450 monooxygenase (Alexander et al., 1998). TRI11 mutants of F. sporotrichioides accumulate trichothecenes that lack a C-15 oxygen atom (McCormick and Hohn, 1997). Following C-15

hydroxylation, the TRI3-encoded acetyltransferase catalyzes the transfer of an acetyl moiety to the C-15 oxygen. This is evident from blockage of production of C-15 acetylated trichothecenes in TRI3 mutants (McCormick et al., 1996).

The product of the TRI3 reaction, calonectrin, has been isolated from T-2 toxin, NIV and DON-producing fusaria. This suggests that the biosynthetic path-ways for all three trichothecenes are the same up to calonectrin but that the pathways can branch beyond calonectrin. In T-2 toxin and NIV-producing fusaria, trichothecene biosynthesis proceeds with C-4 hydrox-ylation, which is catalyzed by the TRI13-encoded cytochrome P450 monooxygenase (Brown et al., 2002; Lee et al., 2002). In contrast, DON-producing strains of F. graminearum lack a functional TRI13 and, there-fore, cannot hydroxylate the trichothecenes at C-4. Introduction of a functional TRI13 into a field isolate of F. graminearum that produces only trichothecenes that lack a C-4 hydroxyl (e.g. DON and acetylated DON) results in transformants that can produce C-4 hydroxylated trichothecenes (e.g. NIV) (Lee et al., 2002). In T-2 toxin and NIV-producing fusaria, the TRI7-encoded acetyltransferase esterifies the C-4 oxygen with an acetyl moiety. Inactivation of TRI7 in F. graminearum and F. sporotrichioides blocked pro-duction of C-4 acetylated trichothecenes. In addition, transformation of functional copies of TRI7 and TRI13 into F. graminearum that lacks the ability to produce C-4 hydroxylated or C-4 acetylated trichothecenes

Table 1: Function of TRI genes and phenotype of TRI mutants in Fusarium sporotrichioides and Fusarium graminearum.

Gene FunctionMutant Phenotypea

F. sporotrichioides F. graminearumTRI8 trichothecene-3-O-esterase 3-acetyl T-2 toxin, TAS 3,15 diADONTRI7 trichothecene-4-O-acetyltransferase HT-2 toxin NIVTRI3 trichothecene-15-O-acetyltransferase 15-decalonectrin, 3,15-didecalonectrin 15-decalonectrin, didecalonectrinTRI4 trichodiene oxygenase trichodiene trichodieneTRI6 transcription factor low levels of trichodiene NDTRI5 trichodiene synthase no trichothecenes no trichothecenesTRI10 regulatory gene no trichothecenes NDTRI9 unknown ND NDTRI11 isotrichodermin 15-oxygenase isotrichodermin NDTRI12 trichothecene efflux pump no trichothecenes NDTRI13 calonectrin 4-oxygenase 4-deoxyT-2toxin, 8-hydroxycalonectrin,

8-hydroxy-3-decalonectrinDON

TRI14 virulence factor T-2 toxin DONTRI1 C-8 or C-7,8 oxygenase 4,15-DAS calonectrin, 3-decalonectrinTRI16 C-8 acyltransferase neosolaniol TRI101 C-3 acyltransferase isotrichodermol isotrichodermola Trichothecene production profiles determined by liquid chromatography-mass spectrometry analysis of extracts and filtrates of TRI gene mutants. No trichothecenes indicates that the mutant produced less than 0.1% of the trichothecene produced by the wild-type Fusarium strain from which the mutants were derived. F. sporotrichioides wild-type produces primarily T-2 toxin and much smaller amounts of 4, 15-diacetoxyscirpenol (4,15-DAS), neosolaniol, and C-8 esters of neosolaniol. F. graminearum wild-type strains produce primarily acetylated derivatives of nivalenol (NIV) or deoxynivalenol (DON) in culture. ND indicates not determined. TAS (3,4,15-triacetoxyscirpenol).



yielded transformants that produced C-4 acetylated trichothecenes (Lee et al., 2002).

In F. sporotrichioides, trichothecene biosynthe-sis then proceeds with C-8 hydroxylation, which is catalyzed by the cytochrome P450 monooxygenase encoded by TRI1. TRI1 mutants of F. sporotrichioides are blocked in the formation of trichothecenes with an oxygen at C-8 (Brown et al., 2003; Meek et al., 2003). Further evidence for the role of the TRI1 was provided by genetic transformation of a functional FsTRI1 into F. verticillioides, which lacks tricho-thecene C-8 hydro-xylase activity and does not produce trichothecenes. The resulting transformants could hydroxylate exog-enously added trichothecenes at C-8 (McCormick et al., 2006b). In F. sporotrichioides, C-8 hydroxylation is followed by the esterification of the C-8 hydroxyl by an isovalerate moiety. This reaction is catalyzed by TRI16 as is evident by the inability of TRI16 deletion mutants to produce trichothecenes with the isovaleryl ester (Brown et al., 2003; Peplow et al., 2003a). As noted earlier, tricho-thecene biosynthesis in F. graminearum differs from that in F. sporotrichioides in that the FgTRI1 mono-oxygenase catalyzes hydroxylation of C-7 as well as C-8. This is evident from the inability of TRI1 mutants of F. graminearum to produce C-8 or C-7 oxygenated trichothecenes and from heterologous expression of FgTRI1 (McCormick et al., 2004, 2006b). When FgTRI1 was genetically transformed into F. verticillioides, the resulting transformants were able to hydroxylate exog-enously added trichothecenes at both C-7 and C-8.

The last step in trichothecene biosynthesis in both F. graminearum and F. sporotrichioides is most likely removal of the C-3 acetyl. Multiple lines of evidence indi-cate that this deacetylation reaction is catalyzed by the TRI8 protein. First, deletion of TRI8 in F. graminearum and F. sporotrichioides resulted in accumulation of trichothecenes with a C-3 acetyl (McCormick and Alexander, 2002). In the same experiments, wild-type strains of the fungus accumulated primarily tricho-thecenes with a C-3 hydroxyl rather than C-3 acetyl. Second, when C-3 acetylated trichothecenes were added to cultures of F. graminearum, wild-type strains could deacetylate the molecules, whereas TRI8 mutants could not. Finally, genetic transformation of the yeast S. cerevisiae with TRI8 resulted in transformants that were able to convert C-3-acetylated trichothecenes to C-3 hydroxylated trichothecenes. These findings are consistent with the hypothesis that C-3 acetylation catalyzed by the TRI101 protein reduces toxicity of trichothecenes relatively early in biosynthesis and facilitates additional oxygenation and acetylation of the trichothecene skeleton. Furthermore, once the bio-synthetic process is completed, Fusarium deacetylates trichothecenes to restore a higher level of toxicity prior to release of the toxins into the environment. This

hypothesis is supported by the higher level of toxicity of C-3 hydroxylated trichothecenes than C-3 acetylated trichothecenes to the green alga Chlamydomonas reinhardtii (Alexander et al., 1999b). However, C-3 acetylation of trichothecenes is not always correlated with decreased toxicity in a detached leaf assay with Arabidopsis thaliana (Desjardins et al., 2007b). Thus, the relative toxicity of C-3 hydroxylated versus C-3 acetylated trichothecenes varies among organisms.

The core TRI cluster also includes TRI9, TRI12, and TRI14. TRI9 encodes a 43-amino acid protein that does not exhibit similarity to any known protein, and its role in trichothecene production has not yet been reported (Brown et al., 2001). TRI12 encodes a major facilita-tor superfamily-type transporter that acts as a tricho-thecene efflux pump. Expression of TRI12 in yeast has revealed that the TRI12 protein, Tri12, facilitates trans-port of trichothecenes across the cell membrane and that TRI12 mutants of F. sporotrichioides are markedly reduced in trichothecene production and in ability to grow in the presence exogenous T-2 toxin (Alexander et al., 1999a). The predicted TRI14 protein does not include any known functional domains (Brown et al., 2002; Dyer et al., 2005). However, blastp analysis of the deduced amino acid sequence for TRI14 against the GenBank database revealed that it shares 30% identity over its entire length to proteins predicted from whole genome sequences of four other fungi: Aspergillus flavus, A. oryzae, Podospora anserine, and Talaromyces stipitatus. Although TRI14 deletion mutants of F. graminearum and F. sporotrichioides do not exhibit a noticeable change in trichothecene production in culture, F. graminearum TRI14 mutants do not pro-duce trichothecenes when inoculated into wheat head florets. The TRI14 mutants cause a significant amount of head blight, although less disease than caused by wild-type strains (Dyer et al., 2005). Thus, TRI14 can affect trichothecene production but only under certain conditions. Interestingly, there are TRI14 homologs that share >65% identity to the Fusarium TRI14 in the trichothecene-producing fungi Myrothecium roridum (Proctor, unpublished) and Stachybotrys chartarum (GenBank accession AAG47844).

Functional characterization of trichothecene bio-synthetic genes has provided significant insight into the genetic basis of different trichothecene production phenotypes (chemotypes) observed in Fusarium. The genetic basis for production of T-2 toxin versus DON/NIV is explained by differences in TRI1 and TRI16. Likewise, the basis for production of NIV/C-4 acetylated NIV versus DON can be explained by differ-ences in TRI7 and TRI13. Production of C-3-acetylated DON (3-ADON) versus C-15-acetylated DON (15-ADON) within populations of F. graminearum represents a third type of variation in trichothecene



chemotype. There is evidence that the 3-ADON chem-otype may result from altered expression or activity of the TRI8 protein, Tri8 (Kimura et al., 2003). This evi-dence is consistent with the C-3 deacetylase activity of Tri8. In addition, some researchers have identified genetic markers at TRI3 and TRI12 that are correlated with the 3-ADON and 15-ADON chemotypes (Starkey et al., 2007; Suga et al., 2008; Ward et al., 2008). However, a causal relationship between these mark-ers and 3-ADON versus 15-ADON production has not been demonstrated. Thus, the genetic basis for this third type of variation in trichothecene chemotype has not yet been demonstrated conclusively.

Fumonisins

Fumonisin contamination is of particular concern in maize (corn). Production of the toxins has been reproducibly demonstrated in at least eight Fusarium species, including two of the most significant fungal pathogens of maize worldwide, F. proliferatum ( teleomorph G. intermedia) and F. verticillioides (tele-omorph G. moniliformis) (Munkvold and Desjardins 1997; Proctor et al., 2004; Rheeder et al., 2002). The B-series fumonisins, fumonisin B

1, B

2, B

3, and B

4, are

generally the most abundant fumonisins in naturally contaminated maize kernels and in pure cultures of F. proliferatum and F. verticillioides. Of these four metabolites, fumonisin B

1 (FB

1) usually makes up

70% to 80% of the total fumonisin content, fumonisin B

2 (FB

2) 15% to 25% of the total content, fumonisin

B3 (FB

3) 3% to 8%, and fumonisin B

4 (FB

4) an even

lower percentage (Rheeder et al., 2002). The struc-tures of FBs were first reported in 1988 and 1989 by research groups in South Africa, New Caledonia, and France (Bezuidenhout et al., 1988; Laurent et al., 1989). The structure consists of a linear, 20-carbon backbone with an amine function at carbon atom 2 (C-2), methyl functions at C-12 and C-16, and tricarballylic esters at C-14 and C-15. FBs differ in structure by the presence or absence of hydroxyl functions at C-4, C-5, and C-10 (Figure 4). Since the first reports on the structures of FBs, over 30 fumonisin analogs have been described (Butchko et al., 2003a, 2006; Rheeder et al., 2002).

Fumonisin biosynthesis: genes and gene cluster

Studies on the genetics of fumonisin production began in the early 1990s. The first of these studies consisted of meiotic analyses of naturally occurring variants of F. verticillioides with altered fumonisin pro-duction phenotypes. The studies identified four loci: Fum1, Fum2, Fum3, and Fum4. Fum1 conferred the

ability to produce the wild-type complement of FB1,

FB2, FB

3, and FB

4, such that strains of F. verticillioides

with the variant Fum1 allele produced no fumonisins (Desjardins et al., 1992). Fum2 was responsible for hydroxylation of the fumonisin backbone at C-10, and Fum3 was responsible for hydroxylation of the back-bone at C-5 (Desjardins et al., 1996a). Fum4 affected fumonisin production quantitatively. Strains with a variant Fum4 allele produced no detectable fumoni-sins in liquid culture and only low levels in maize kernel culture (Plattner et al., 1996). Genetic linkage analysis revealed that the Fum loci were located close to one another on linkage group 1 of F. verticillioides (Desjardins et al., 1996a; Proctor et al., 1999a; Xu and Leslie, 1996). The genetic linkage of Fum genes com-bined with the growing evidence at the time for clus-tering of secondary metabolite biosynthetic genes in fungi suggested that fumonisin biosynthetic genes were located in a gene cluster.

The first fumonisin biosynthetic gene identified and characterized via molecular genetic methods was a polyketide synthase (PKS) gene (Proctor et al., 1999b). The gene was identified via a reverse transcription-PCR approach that utilized: (1) degenerate primers designed from conserved regions within ketosynthase domains of other fungal PKSs (Keller et al., 1995) and (2) F. verticillioides RNA isolated from culture condi-tions that promoted fumonisin production. Disruption of the PKS gene blocked fumonisin production in F. verticillioides and thereby confirmed its connection to fumonisin biosynthesis. When it was first identi-fied, the PKS gene was designated FUM5. However, subsequent genetic complementation and sequence analysis revealed that the molecularly defined FUM5 and the meiotically defined Fum1 were the same gene. As a result, the designation of FUM5 was changed to FUM1 (Desjardins et al., 2002; Proctor et al., 2006).

Because genetic linkage analysis indicated that fumonisin biosynthetic genes (FUM) were likely

OH

2 4 6 8 10 12 14 16 18

R2

R1NH2

Fumonisin B1 H OH OHIsofumonisin B1 OH H OHFumonisin B2 H OH HFumonisin B3 H H OHFumonisin B4 H H H

R1 R2 R3

R3

O

OO OH

OH

OH

OH

O

OO

O

O

Figure 4. Chemical structure of B-series fumonisins.



clustered, FUM1 was used as a hybridization probe to isolate overlapping cosmid clones in order to identify genes adjacent to it. Sequence analysis of the clones and subsequent expression analysis revealed the presence of 14 genes immediately downstream of FUM1 with patterns of expression that were similar to each other and correlated with fumonisin produc-tion. These 14 putative fumonisin biosynthetic genes were designated FUM6 through FUM19 (Figure 5) (Proctor et al., 2003; Seo et al., 2001). A subsequent expressed sequence tag (EST) database prepared from F. verticillioides revealed the presence of two addi-tional genes (FUM20 and FUM21) in the FUM cluster (Brown et al., 2005, 2007). As a result, the FUM cluster is now considered to include 17 genes (Figure 5). Since the initial descriptions of the cluster, comple-mentation and sequence analyses have indicated that two of the cluster genes, FUM9 and FUM12, were the same as the meitoically defined loci Fum3 and Fum2, respectively (Butchko et al., 2003b; Desjardins et al., 1996a; Proctor et al., 2006). For consistency with rules of genetic nomenclature, the designation for FUM9 was changed to FUM3, and the designation for FUM12 was changed to FUM2. Since its first description in F. verticillioides, the presence of the FUM gene cluster has also been confirmed in F. proliferatum (Waalwijk et al., 2004) and F. oxysporum (Proctor et al., 2008).

The general biochemical functions of the FUM genes were predicted based on similarities of their deduced amino acid sequences to proteins with known functions (Table 2). However, the roles of the genes in fumonisin biosynthesis have also been exam-ined by functional analyses. Each FUM gene, except FUM20, has been inactivated in F. verticillioides by gene deletion or disruption, and the resulting effects on fumonisin production have been determined (Table 2). The fumonisin analogs produced by some of the FUM gene mutants were metabolites that had not been identified previously and have provided insight into FUM gene function and the biochemistry of fumonisin biosynthesis (Butchko et al., 2003a, 2003b, 2006; Zaleta-Rivera et al., 2006). A fumonisin

biosynthetic pathway (Figure 6) has been proposed based on these insights as well as on other molecular genetic, biochemical and analytical chemical studies.

Genetic and biochemical pathways for fumonisin biosynthesis

The proposed fumonisin biosynthetic pathway begins when the FUM1 protein (Fum1p) catalyzes the con-densation of nine acetate and two methyl units to form a linear, 18-carbon-long polyketide. The polyketide should be identical or similar in structure to 10,14-dimethyl octadecanoic acid. However, it is possible that the polyketide does not exist as a free acid but instead remains covalently attached to the phosphopanteth-einyl cofactor of the PKS (Gerber et al., 2009). The role of FUM1 in synthesis of the fumonisin polyketide is supported by the fact that the FUM1 protein, Fum1p, is a PKS. In addition, precursor feeding studies indi-cate that C-3 through C-20 of the fumonisin backbone are derived from acetate and are, therefore, likely to be a product of polyketide or fatty acid metabolism. Furthermore, fumonisin production can be blocked by disruption of FUM1 and by some alterations to the ketosynthase and methylation-coding regions of the gene (Proctor et al., 1999b; Yu et al., 2005; Zhu et al., 2007). Despite the substantial evidence for the role of Fum1p in the first step committed to fumonisin bio-synthesis, the putative 10,14-dimethyl octadecanoic acid-like polyketide has not yet been isolated and subjected to structural analysis. Therefore, additional studies are required to elucidate the structure of the product of the Fum1p-catalyzed reaction.

In the second step of the pathway, the FUM8 pro-tein, Fum8p, catalyzes the condensation of the linear polyketide and alanine to yield a linear molecule that is 20 carbons long and has an amine at C-2, a carbo-nyl at C-3, and methyl functions at C-12 and C-16 (Du et al., 2008; Proctor et al., 2008; Seo et al., 2001). This conclusion is consistent with precursor feeding stud-ies that demonstrate that C-1, C-2 and the amine of FB1

are derived from alanine (Branham and Plattner, 1993). The role of Fum8p in this step is consistent with its similarity to oxoamine synthases, a class of enzymes that catalyze condensation of amino acids and acyl compounds, and with the fact that FUM8 dis-ruption mutants are blocked in production of known fumonisin analogs. A study of a strain of F. oxysporum that produces C-series fumonisins (FCs) has provided further evidence for the function of Fum8p (Seo et al., 1996). FCs and FBs have the same structures except that FCs lack the terminal methyl group that is at C-1 in FBs. This difference in structure suggests that FCs result from condensation of the polyketide with

FUM

21

FUM

1

FUM

6FU

M7

FUM

8FU

M3

FUM

10FU

M11

FUM

2FU

M20

FUM

13FU

M14

FUM

15FU

M16

FUM

17FU

M18

FUM

19

kb10 20 30 40 50

Figure 5. The fumonisin biosynthetic gene cluster in F. verticillioides. Arrows indicate position and transcriptional orientation of genes. The order and orientation of genes in the F. proliferatum and F. oxysporum clusters are the same, although, distances between genes can vary (Proctor et al., 2008; Waalwijk et al., 2004).



glycine rather than with alanine, because alanine and glycine differ only by a methyl group (Sewram et al., 2005). Complementation of the F. verticillioides FUM8 disruption mutant with a wild-type copy of F. verticillioides FUM8 restored FB production, whereas complementation of the mutant with the F. oxysporum FUM8 restored production but resulted in produc-tion of predominantly FCs rather than FBs (Proctor et al., 2008). This indicates that FUM8 orthologues in different Fusarium species control whether FBs or FCs are produced and is consistent with the hypothesis that Fum8p catalyzes the polyketide-amino acid conden-sation. Du and coworkers have found evidence that Fum8p also facilitates release of the polyketide from the PKS (Du et al., 2008; Gerber et al., 2009). Despite the considerable evidence for the function of Fum8p, the product resulting from the Fum8p-catalyzed reaction has not yet been isolated or characterized chemically. Therefore, additional studies are required to confirm the structure of the product of the Fum8p-catalyzed reaction.

The third step in the proposed fumonisin biosyn-thetic pathway is catalyzed by the FUM6 protein, Fum6p, and consists of the hydroxylation of the polyketide-amino acid condensation product at

C-14 and C-15 (Figure 6) (Bojja et al., 2004; Proctor et al., 2008; Seo et al., 2001). Several indirect lines of evidence are consistent with the role of Fum6p in this reaction. First, FUM6 deletion mutants of F. verticillioides do not produce detectable fumonisin-like compounds, which suggests that Fum6p catalyzes a reaction that occurs early in fumonisin biosynthesis (Bojja et al., 2004; Seo et al., 2001). Second, the pre-dicted amino acid sequence of Fum6p is a hybrid protein consisting of a cytochrome P450 monooxy-genase and a NADPH-dependent reductase. This is consistent with precursor feeding studies indicating that the oxygen atoms at C-14 and C-15 are derived from molecular oxygen (Caldas et al., 1998). Finally, all other FUM gene-encoded enzymes that are pre-dicted to utilize molecular oxygen catalyze other reactions (i.e., Fum2p and Fum3p) or do not affect fumonisin production (Fum15p). This leaves Fum6p as the only likely candidate for a FUM cluster-encoded C-14/C-15 hydroxylase. The predicted product of the Fum6p reaction is 3-keto hydrolyzed FB

4. The produc-

tion of this metabolite by F. verticillioides has not yet been rigorously confirmed. However, two structur-ally similar metabolites, 3-keto FB

4 and hydrolyzed

FB4, have been isolated from cultures of FUM13 and

Table 2: Function of FUM genes and fumonisin production phenotypes of FUM mutants in F. verticillioides.Gene Predicted Function a Mutant Phenotype b

FUM21 Cys-6 transcription factor no fumonisinsFUM1 polyketide synthase no fumonisinsFUM6 cytochrome P450 monooxygenase and reductase no fumonisinsFUM7 alcohol dehydrogenase tetradehydro-FBFUM8 -oxoamine synthase no fumonisins

FUM3 dioxygenase FB2 & FB

4

FUM10 acyl-CoA synthetase/acyl- protein synthetase hydrolyzed FB3 & hydrolyzed FB

4

FUM11 tricarboxyllic acid transporter FB1, FB

2, FB

3, & FB

4

Half-hydrolyzed FB1, FB

2, FB

3, & FB

4

Keto half-hydrolyzed FB1, FB

2, FB

3, FB

4

FUM2 cytochrome P450 monooxygenase FB2 & FB

4

FUM20 unknown NDFUM13 short-chain dehydrogenase/reductase 3-keto FB

3 & 3-keto FB

4

FUM14 nonribosomal peptide synthase (peptidyl and condensation domains)

hydrolyzed FB3 & hydrolyzed FB

4

FUM15 cytochrome P450 monooxygenase no effectFUM16 acyl-CoA synthetase/acyl- protein synthetase no effectFUM17 ceramide synthase no effectFUM18 ceramide synthase no effectFUM19 ABC transporter increased ratio FB

1:FB

3a The predicted function is based on similarity of deduced amino acid sequence to proteins with known functions based on blastx and blastp analysis against the GenBank database at the National Center for Biotechnology Information.b Fumonisin production profiles determined by liquid chromatography-mass spectrometry analysis of extracts and filtrates of FUM gene mutants. No fumonisins indicates that the mutant produced less than 0.1% of the fumonisin produced by the wild-type F. verticillioides strain from which the mutants were derived. Tetradehydro-fumonisins have a carbon-carbon double bond in each of the tricarballylic ester functions. Hydrolyzed fumonisins have hydroxyl functions rather than tricarballylic ester functions at C-14 and C-15. In half-hydrolyzed fumonisins, one tricarballylic ester function is replaced by a hydroxyl function, and in keto half-hydrolyzed fumonisins one tricarballylic ester function is replaced by a carbonyl (keto) function. 3-keto fumonisins have a carbonyl (keto) function at C-3 rather than a hydroxyl function. ND indicates not determined.



FUM14 mutants, respectively (Butchko et al., 2003a, 2006). Thus, it should be possible to obtain the pre-dicted Fum6p reaction product by generating a strain of F. verticillioides in which both FUM13 and FUM14 have been deleted.

Although Figure 6 depicts C-3 carbonyl reduction, C-10 hydroxylation, and C-14/C-15 esterification to

be the fourth, fifth, and sixth steps, respectively, in fumonisin biosynthesis, production profiles of sev-eral F. verticillioides mutants indicate that each of the reactions can occur in the absence of the others. Thus, the positions of the three reactions in the fumonisin biosynthetic pathway are likely not fixed. C-3 carbonyl (keto) reduction is catalyzed by Fum13p. Three lines of

HO

O

O

O

O OH

OH

OH

OH

OH

OH OH

OH

OH

OH

OH

OH

OH

OH

OH

OH

OH

OH

OH

OH

OH

OH

NH2

NH2

NH2

NH2

OH

OH

OH

O

OH

OH

O

OO

OO

HO

HO

OH

OH

OOO

HO

Mito

chod

rial l

umen

Cyt

opla

sm

NH2

NH2

NH2

Alinine

Acetate

Fum1p

Fum8p

Fum6p

Fum11p

Fum7p

Fum13p

Fum2p

Fum10pFum14p

Fum3pFumonisin B3

Fumonisin B1

O

O

O O O

OO

O

OOO

O

O O O

O

O

23 5

10 12 14 1615

HO

Figure 6. Proposed biosynthetic pathway for fumonisin B1 showing functions of FUM gene/proteins. Fumonisin B

2 results when C-10 of

the fumonisin backbone is not hydroxlated. Fumonisin B4 results when C-5 and C-10 are not hydroxylated.



evidence support this role for Fum13p. First, FUM13 deletion mutants of F. verticillioides produce fumoni-sin analogs with a carbonyl group at C-3 (Butchko et al., 2003a). Second, the predicted Fum13p sequence is most similar to short-chain dehydrogenases/reductases, a group of enzymes that includes carbonyl reductases (Butchko et al., 2003a). Finally, purified Fum13p can catalyze reduction of the fumonisin C-3 carbonyl to a hydroxyl in vitro (Yi et al., 2005).

Fumonisin C-10 hydroxylation is most likely catalyzed by Fum2p. The evidence for this is that FUM2 deletion mutants and natural variants pro-duce fumonisins that lack the C-10 hydroxyl (Proctor et al., 2006). In addition, the predicted amino acid sequence of Fum2p is most similar to cytochrome P450 monooxygenases, which, as noted above, are enzymes that often catalyze hydroxylation reactions.

Esterification of the tricarballylic moieties to the hydroxyls at C-14 and C-15 of fumonisins is cata-lyzed by Fum14p. The most direct evidence for this is that purified Fum14p catalyzes the reaction in vitro (Zaleta-Rivera et al., 2006). In addition, FUM14 mutants produce fumonisin analogs that lack the tricarballylic esters, i.e., hydrolyzed FB

3 (HFB

3) and

FB4 (HFB

4) (Butchko et al., 2006; Zaleta-Rivera et al.,

2006). The predicted amino acid sequence of Fum14p has two functional domains, a condensation domain and a peptidyl carrier protein domain. Both domains occur in nonribosomal peptide synthetases (NRPSs), and in NRPSs condensation domains catalyze amide bond formation. The Fum14p-catalyzed esterification of the fumonisin backbone indicates that amide and ester bond formation can involve similar enzymatic mechanisms.

Although Fum14p catalyzes C-14/C-15 esterification, gene deletion analysis indicates that Fum7p, Fum10p, and Fum11p also contribute to formation of the tricar-ballylic esters (Butchko et al., 2006). FUM10 deletion mutants of F. verticillioides have the same phenotype as FUM14 mutants; they produce only fumonisins that lack the tricarballylic esters. Fum10p is predicted to be an acyl-CoA synthetase/acyl-protein synthetase, a group of enzymes that catalyze formation of thioesters of carboxylic acids to coenzyme A (CoA) or proteins, respectively. Based on these data, Zaleta-Rivera et al. (2006) proposed that Fum10p first catalyzes activation of a tricarballylic precursor with adenosine mono-phosphate and then catalyzes formation of a tricarbal-lylic thioester to the peptidyl carrier protein domain of Fum14p. The thioester is then used as a substrate in the formation of the C-14 and C-15 tricarballylic esters, a reaction that is catalyzed by the Fum14p condensation domain. FUM7 deletion mutants produce fumonisin analogs with a single carbon-carbon double bond in

each of the tricarballylic esters (e.g., tetradehydro-FB1

and -FB3) (Butchko et al., 2006). Fum7p is most similar

to iron-containing dehydrogenases, a class of enzymes that sometimes catalyze reduction of carbon-carbon double bonds. Based on these data, the proposed role for Fum7p is reduction of a carbon-carbon double bond in a precursor (e.g., aconitic acid) of the tricarbal-lylic ester. FUM11 mutants produce a complex mixture of fumonisin analogs that lack one of the tricarballylic esters (i.e., half hydrolyzed and keto-half hydrolyzed fumonisins) in addition to reduced levels of FB

1, FB

2,

FB3, and FB

4 (Butchko et al., 2006). Fum11p is similar

to proteins embedded in the inner mitochrondrial membrane that transport tricarboxylic acid intermedi-ates from the TCA/Krebs cycle out of the mitochondrial lumen. Based on these data, Fum11p was proposed to transport tricarboxylic acid(s) out of the inner mito-chondrial lumen in order to make them available for fumonisin biosynthesis in the cytoplasm and on the ER membrane (Figure 6). Production of reduced lev-els of FB

1, FB

2, FB

3 and FB

4 by FUM11 mutants also

suggests that other tricarboxylic acid transports in F. verticillioides can partially compensate for absence of Fum11p.

The final step in the proposed fumonisin biosyn-thetic pathway is the Fum3p-catalyzed hydroxylation of the fumonisin backbone at C-5. Fum3p is predicted to encode a dioxygenase, and its role in fumonisin biosynthesis is evident from enzyme assays in which the purified protein catalyzed fumonisin C-5 hydroxylation (Ding et al., 2004). In addition, FUM3 mutants of F. verticillioides produce only fumonisins that lack the C-5 hydroxyl (Butchko et al., 2006). The phenotypes of FUM10, FUM13, and FUM14 mutants provide evidence that C-5 hydroxylation is indeed the final step in fumonisin biosynthesis. FUM10 and FUM14 mutants produce analogs of hydrolyzed fumonisins that lack the C-5 hydroxyl but not analogs with the C-5 hydroxyl. Likewise, FUM13 mutants produce only 3-keto fumonisin analogs that lack the C-5 hydroxyl. Thus, it appears that the C-3 carbonyl must be reduced and the C-14 and C-15 tricarballylic esters must be present before the C-5 hydroxylation of the fumonisin backbone can occur.

The FUM gene functions described above account for all the major enzymatic activities predicted to be necessary for formation of the predominant fumonisins produced by F. verticillioides (i.e., FB

1, FB

2,

FB3, or FB

4). The other FUM cluster genes (FUM15,

FUM16, FUM17, FUM18, FUM19, and FUM21) that have been examined for function either have indirect roles in fumonisin biosynthesis or are not essential for production of FB

1, FB

2, FB

3, or FB

4. Based on its

amino acid sequence, Fum21p is predicted to be a Zn(II)2Cys6 transcription factor, a class of proteins



that bind to promoter regions of other genes and acti-vate their transcription (Brown et al., 2007). FUM21 deletion mutants of F. verticillioides produce little or no fumonisins and exhibit markedly reduced FUM gene expression. Based on these data, Fum21p is most likely a transcription factor that regulates expression of genes in the FUM cluster, a function analogous to the TRI6 protein in trichothecene biosynthesis. FUM19 is also indirectly involved in fumonisin bio-synthesis. Fum19p is predicted to be an ABC trans-porter, a class of proteins that transport metabolites across the plasma membrane. Therefore, Fum19p likely pumps fumonisins out of the cells in which they are produced and into the surrounding environment, a function analogous to the trichothecene efflux pump encoded by TRI12. However, TRI12 mutants are markedly reduced in trichothecene production, whereas FUM19 mutants exhibit only a slight change in fumonisin production, a small increase in the ratio of FB

1 to FB

3 (Proctor et al., 2003). The fact that

fumonisins are produced by FUM19 deletion mutants and excreted into the surrounding environment suggests that one or more other transport proteins partially compensate for the absence of Fum19p in the mutants.

Mutants of F. verticillioides in which both FUM17 and FUM18 have been deleted do not exhibit detect-able changes in production of FB

1, FB

2, FB

3, or FB

4

(Proctor et al., 2003). This indicates that the genes are not essential for production of the predominant F. verticillioides fumonisins. Both Fum17p and Fum18p are similar in sequence to a class of proteins first clas-sified as longevity assurance factors but more recently determined to be components of ceramide synthase (sphinganine N-acyltransferase). Ceramide synthase catalyzes acylation of sphinganine with a long-chain fatty acid during de novo sphingolipid biosynthesis (Merrill et al., 1997). In eukaryotic cells, fumonisins inhibit ceramide synthase and thereby disrupt sphin-golipid metabolism (Riley et al., 1996). In addition, some tomato varieties have a ceramide synthase gene, Asc-1, that confers insensitivity to fumonisins and AAL toxin, a group of fumonisin-like metabolites pro-duced by the tomato pathogen Alternaria alternata (Brandwagt et al., 2000). Tomato plants with asc-1, a defective allele of Asc-1, are sensitive to fumonisins and AAL toxin. Based on these findings, Asc-1 likely encodes a ceramide synthetase that is resistant to the effects of fumonisins and AAL toxin. Thus, the presence of FUM17 and FUM18 in the FUM cluster suggests that they encode fumonisin-resistant ceramide synthases that serve as a fumonisin self-protection mechanism in F. verticillioides. This hypothesis is seemingly not consistent with the observations that FUM17/FUM18 double mutants do not exhibit increased sensitivity

to fumonisins on agar media (Proctor and Plattner, unpublished). However, the F. verticillioides genome includes three other ceramide synthase genes, one or more of which may be able to compensate for the absence of Fum17p and Fum18p in FUM17/FUM18 mutants. Thus, further studies are required to examine the potential role of ceramide synthases in fumonisin self-protection in F. verticillioides.

FUM15 and FUM16 deletion mutants do not exhibit altered production of FB

1, FB

2, FB

3, and FB

4 (Butchko

et al., 2006). Thus, these genes are not essential for production of the predominant fumonisins produced by F. verticillioides. FUM15 is predicted to encode a cytochrome P450 monooxygenase and, therefore, may function as a hydroxylase. F. verticillioides can produce low levels of iso-FB

1, which is identical to FB

1 except

that it has a hydroxyl function at C-4 rather than C-5 (MacKenzie et al., 1998). In addition, the predominant fumonisins produced by F. oxysporum strain O-1890 have a hydroxyl at the equivalent position (C-3) of the C fumonisin backbone. Thus, a possible function for Fum15p is hydroxylation of the C-4 and C-3 position of B and C fumonisins, respectively. When F. verticillioides FUM15 mutants were examined for fumonisin produc-tion, they were not analyzed for production of fumoni-sins with a C-4 hydroxyl, and attempts to disrupt FUM15 in F. oxysporum O-1890 have so far not been successful (unpublished data). FUM16 is predicted to encode an acyl-CoA synthetase/acyl-protein synthetase. Initially, the proposed role of Fum16p was activation of the polyketide product of Fum1p with CoA to facilitate condensation of the polyketide with alanine (Proctor et al., 2003). This proposed function was analogous to CoA activation of palmitate prior to condensation with serine to form 3-ketosphinganine during de novo sphingolipid biosynthesis. However, in mechanistic studies, Gerber et al. (2009) have demonstrated that condensation of an 18-carbon polyketide/fatty acid and alanine can occur in the presence of Fum8p and absence of an acyl-CoA synthetase/acyl-protein syn-thetase. Thus, the functions of Fum15p and Fum16p remain enigmas.

Application of genetic and biochemical information

Knowledge of the genetic and biochemical pathways for biosynthesis of trichothecenes and fumonisins has been used to study enzyme structure and func-tion (Rynkiewicz et al., 2001), evolution of mycotoxins (Ward et al., 2002, 2008), and regulation of mycotoxin production (Flaherty and Woloshuk, 2004; Shim and Woloshuk, 2001). Variation in TRI genes has also been used to develop genetic markers for prediction



of trichothecene chemotypes of F. graminearum (Chandler et al., 2003; Desjardins, 2008; Lee et al., 2001). One area that has received a great deal of attention is the study of the role of trichothecenes and fumonisins in host–pathogen interactions. The identification of genes (TRI5 and FUM1) responsible for the first committed steps in trichothecene and fumonisin biosynthesis has facilitated the generation of Fusarium mutants that are blocked in production of the mycotoxins as well as their biosynthetic interme-diates. Studies with FUM1 mutants of F. verticillioides indicate that fumonisin production does not contrib-ute significantly to the ability of the fungus to cause maize ear rot in the field (Desjardins et al., 2002), but can contribute to maize seedling blight under some conditions (Glenn et al., 2008) and to nonsympto-matic colonization of seedlings (Desjardins et al., 1995, 2007a).

Studies with TRI5 mutants of F. graminearum and F. sambucinum indicate that trichothecene production can contribute to the ability of these species to cause disease on some hosts but not on others (Desjardins et al., 1993, 1996b; Harris et al., 1999; Maier et al., 2006). A highly significant finding from these studies is that trichothecene production in F. graminearum contributes to its ability to cause wheat head blight. Thus, improving trichothecene resistance in wheat has the potential to enhance resistance to both head blight and trichothecene contamination. Three proteins with potential to enhance crop resistance have been identified: (1) the Fusarium TRI101 protein catalyzes trichothecene C-3 acetylation and serves as a trichothecene self-protection mechanism in the fungus (Kimura et al., 1998; McCormick et al., 1999); (2) an Arabidopsis thaliana glucosyl transferase (DOGT1) catalyzes trichothecene C-3 glycosylation and enhances resistance of the plant to trichothecenes (Poppenberger et al., 2003); and (3) the ribosomal protein L3 (Rpl3), which is the target of trichothecenes, can be modified by amino acid substitutions to confer trichothecene resistance (Harris and Gleddie, 2001; Mitterbauer et al., 2004). When expressed in some plants, TRI101, DOGT1, and modified forms of RPL3 can enhance trichothecene resistance. However, expression of TRI101 in wheat and barley has not been effective at providing high levels of resistance to head blight or trichothecene contamination caused by F. graminearum (Manoharan et al., 2006; Okubara et al., 2002). It is possible that high levels of resistance were not achieved because C-3 acetylation is revers-ible and/or may not markedly reduce trichothecene toxicity to wheat. In addition, the experiments with transgenic wheat and barley employed the F. sporotrichioides TRI101 (FsTRI101), but structure and kinetic studies indicate that the FsTRI101 protein has

a 70-fold lower affinity for DON than the F. gramine-arum TRI101 protein (Garvey et al., 2008). There are also limitations with Rpl3, because amino acid sub-stitutions that increase its trichothecene resistance can negatively alter ribosome assembly (Mitterbauer et al., 2004). Thus, additional genes/proteins that confer resistance to trichothecenes may be necessary to enhance crop resistance to Fusarium head blight and trichothecene contamination. An enzyme that fundamentally and irreversibly alters trichothecene structure (e.g., an epoxide hydrolase) could be highly effective for this purpose (Desjardins et al., 2007b). Because the levels of trichothecene and fumonisin contamination in grain crops are generally corre-lated with the amount of disease in the affected crops (Desjardins et al., 1998; Paul et al., 2006; Robertson-Hoyt et al., 2007), other factors that enhance resist-ance to Fusarium should also contribute to reduction of these and other Fusarium mycotoxins in crops. The recent availability of whole genome sequences for F. verticillioides and F. graminearum should greatly facilitate efforts to identify such factors.

Acknowledgments

Declaration of interest: The authors report no con-flicts of interest.

References

Alexander NJ, Hohn TM, McCormick SP. (1998). The TRI11 gene of Fusarium sporotrichioides encodes a cytochrome P450 monooxygenase required for C-15 hydroxylation in trichothecene biosynthesis. Appl Environ Microbiol. 64: 221–225.

Alexander NJ, Hohn TM, McCormick SP. (1999a). TRI12, a tri-chothecene efflux pump form Fusarium sporotrichioides: gene isolation and expression in yeast. Mol Gen Genet. 261: 977–984.

Alexander NJ, McCormick SP, Larson TM, Jurgenson JE. (2004). Expression of Tri15 in Fusarium sporotrichioides. Curr Genet. 45: 157–162.

Alexander NJ, McCormick SP, Ziegenhorn, SL (1999b). Phytotoxicity of selected trichothecenes using Chlamydomonas reinhardtii as a model system. Nat Toxins. 7: 265–269.

Alexander NJ, Proctor RH, McCormick SP, Plattner RD. (1997). Genetic and molecular aspects of the biosynthesis of trichothecenes by Fusarium. Cereal Res Commun. 25: 315–320.

Beremand MN, Desjardins AE. (1988). Trichothecene biosynthe-sis in Gibberella pulicaris: inheritance of C-8 hydroxylation. J Ind Microbiol. 3: 167–174.

Beremand MN, McCormick SP. (1992). Biosynthesis and regu-lation of trichothecene production by Fusarium species. In: Bhatnagar D, Lillehoj EB, Arora DK, eds. Handbook of Applied Mycology: Mycotoxins in Ecological Systems, Vol 5. New York, NY: Marcel Dekker, Inc, pp. 359–384.



Bezuidenhout SC, Gelderblom WCA, Gorst-Allman CP, Horak RM, Marasas WFO, Spiteller G, Bleggaar R. (1988). Structure elucidation of the fumonisins, mycotoxins from Fusarium moniliforme. J Chem Soc Chem Commun. 1988: 743–745.

Bojja RS, Cerny RL, Proctor RH, Du L. (2004). Determining the biosynthetic sequence in the early steps of the fumoni-sin pathway by use of three gene-disruption mutants of Fusarium verticillioides. J Agric Food Chem. 52: 2855–2860.

Brandwagt BF, Mesbah LA, Takken FLW, Laurent PL, Kneppers TJA, Hille J, Nijkamp HJJ. (2000). A longevity assurance gene homolog of tomato mediates resistance to Alternaria alternata f. sp. lycopersici toxins and fumonisin B1. Proc Natl Acad Sci U S A. 97: 4961–4966.

Branham BE, Plattner RD. (1993). Alanine is a precursor in the biosynthesis of fumonisin B

1 by Fusarium moniliforme.

Mycopathologia. 124: 99–104.Brown DW, Butchko RAE, Busman M, Proctor RH. (2007). The

Fusarium verticillioides FUM gene cluster encodes a Zn (II)2Cys6 protein that affects FUM gene expression and fumonisin production. Eukaryot Cell. 6: 1210–1218.

Brown DW, Cheung F, Proctor RH, Butchko RAE, Zheng L, Lee Y, Utterback T, Smith S, Feldblyum T, Glenn AE, Plattner RD, Kendra DF, Town CD, Whitelaw CA. (2005). Analysis of 87,000 expressed sequence tags reveals alternatively spliced introns in multiple genes of the fumonisin gene cluster. Fungal Genet Biol. 42: 848–861.

Brown DW, Dyer JM, McCormick SP, Kendra DF, Plattner RD. (2004). Functional demarcation of the Fusarium core tri-chothecene gene cluster. Fungal Genet Biol. 41: 462.

Brown DW, McCormick SP, Alexander NJ, Proctor RH, Desjardins AE. (2001). A genetic and biochemical approach to study trichothecene diversity in Fusarium sporotrichioides and Fusarium graminearum. Fungal Genet Biol. 32: 121–133.

Brown DW, McCormick SP, Alexander NJ, Proctor RH, Desjardins AE. (2002). Inactivation of a cytochrome P-450 is a determinant of trichothecene diversity in Fusarium species. Fungal Genet Biol. 36: 224–233.

Brown DW, Proctor RH, Dyer RB, Plattner RD. (2003). Characterization of a Fusarium 2-gene cluster involved in trichothecene C-8 modification. J Agric Food Chem. 51: 7936–7944.

Butchko RAE, Plattner RD, Proctor RH. (2003a). FUM13 encodes a short chain dehydrogenase/reductase required for C-3 carb-onyl reduction during fumonisin biosynthesis in Gibberella moniliformis. J Agric Food Chem. 51: 3000–3006.

Butchko RAE, Plattner RD, Proctor RH. (2003b). FUM9 is required for C-5 hydroxylation of fumonisins and complements the meiotically defined Fum3 locus in Gibberella moniliformis. Appl Environ Microbiol. 69: 6935–6937.

Butchko RAE, Plattner RD, Proctor RH. (2006). Deletion analysis of FUM genes involved in tricarballylic ester formation during fumonisin biosynthesis. J Agric Food Chem. 54: 9398–9404.

Caldas ED, Sadilkova K, Ward BL, Jones AD, Winter CK, Gilchrist DG. (1998). Biosynthetic studies of fumonisin B

1 and AAL

toxins. J Agric Food Chem. 46: 4734–4743.Chandler EA, Simpson DR, Thomsett MA, 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. Physiol Mol Plant Pathol. 62: 355–367.

Cole JR, Cox RJ. (1981). Handbook of Toxic Fungal Metabolites. New York, NY: Academic Press.

Desjardins AE (2006). Fusarium Mycotoxins Chemistry, Genetics and Biology. St. Paul, MN: APS Press.

Desjardins AE. (2008). Natural product chemistry meets genet-ics: when is a genotype a chemotype? J Agric Food Chem. 56: 7587–7592.

Desjardins AE, Busman M, Muhitch MJ, Proctor RH. (2007a). Complementary host-pathogen genetic analyses of the role of fumonisins in the Zea mays-Gibberella moniliformis inter-action. Physiol Mol Plant Pathol. 70: 149–160.

Desjardins AE, Hohn TM, McCormick SP. (1993). Trichothecene biosynthesis in Fusarium species: chemistry, genetics, and significance. Microbiol Rev. 57: 595–604.

Desjardins, AE, McCormick, SP, and Appell, M. (2007b). Structure-activity relationships of trichothecene toxins in an Arabidopsis thaliana leaf assay. J Agric Food Chem. 55: 6487–6492.

Desjardins AE, Munkvold GP, Plattner RD, Proctor RH. (2002). FUM1–a gene required for fumonisin biosynthesis but not for maize ear rot and ear infection by Gibberella moniliformis in field tests. Mol Plant-Microbe Interact. 15: 1157–1164.

Desjardins AE, Plattner RD, Beremand MN. (1987). Ancymidol blocks trichothecene biosynthesis and leads to accumulation of trichodiene in Fusarium sporotrichioides and Gibberella pulicaris. Appl Environ Microbiol. 53: 1860–1865.

Desjardins AE, Plattner RD, Lu M, Claflin LE. (1998). Distribution of fumonisins in maize ears infected with strains of Fusarium moniliforme that differ in fumonisin production. Plant Dis. 82: 953–958.

Desjardins AE, Plattner RD, Nelson PE, Nelsen TC, Leslie JF. (1995). Genetic analysis of fumonisin production and viru-lence of Gibberella fujikuroi mating population A (Fusarium moniliforme) on maize (Zea mays) seedlings. Appl Environ Microbiol. 61: 79–86.

Desjardins AE, Plattner RD, Proctor RH (1996a). Linkage among genes responsible for fumonisin biosynthesis in Gibberella fujikuroi mating population A. Appl Environ Microbiol. 62: 2571–2576.

Desjardins AE, Plattner RD, Shackelford DD, Leslie JF, Nelson PE. (1992). Heritability of fumonisin B

1 production in Gibberella

fujikuroi mating population A. Appl Environ Microbiol. 58: 2799–2805.

Desjardins AE, Plattner RD, Vanmiddlesworth F. (1986). Trichothecene biosynthesis in Fusarium sporotrichio-ides: origin of the oxygen atoms of T-2 toxin. Appl Environ Microbiol. 51: 493–497.

Desjardins AE, Proctor RH, Bai G, McCormick SP, Shaner G, Buechley G, Hohn TM. (1996b). Reduced virulence of tri-chothecene-nonproducing mutants of Gibberella zeae in wheat field tests. Mol Plant Microbe Interact. 9: 775–781.

Ding Y, Bojja RS, Du L. (2004). Fum3p, a 2-ketoglutarate- dependent dioxygenase required for C-5 hydroxylation of fumonisins in Fusarium verticillioides. Appl Environ Microbiol. 70: 1931–1934.

Du L, Zhu X, Gerber R, Huffman J, Lou L, Jorgenson J, Yu F, Zaleta-Rivera K, Wang Q. (2008). Biosynthesis of sphinganine-analog mycotoxins. J Ind Microbiol Biotechnol. 35: 455–464.

Dyer RB, Plattner RD, Kendra DF, Brown DW. (2005). Fusarium graminearum TRI14 is required for high virulence and DON production on wheat but not for DON synthesis in vitro. J Agric Food Chem. 53: 9281–9287.

Flaherty JE, Woloshuk CP. (2004). Regulation of fumonisin biosyn-thesis in Fusarium verticillioides by a zinc binuclear cluster-type gene, ZFR1. Appl Environ Microbiol. 70: 2653–2659.

Frisvad JC, Smedsgaard J, Samson RA, Larsen TO, Thrane U. (2007). Fumonisin B

2 production by Aspergillus niger. J Agric

Food Chem. 55: 9727–9732.Gale LR, Bryant JD, Calvo S, Giese H, Katan T, O’Donnell K,

Suga H, Taga M, Usgaard TR, Ward TJ, Kistler HC. (2005).



Chromosome complement of the fungal plant pathogen Fusarium graminearum based on genetic and physical map-ping and cytological observations. Genetics. 171: 985–1001.

Garvey GS, McCormick SP, Rayment, I. (2008). Structural and functional characterization of the TRI101 trichotehecene 3-O-acetyltransferase from Fusarium sporotrichioides and Fusarium graminearum: kinetic insights to combating Fusarium head blight. J Biol Chem. 283: 1660–1669.

Gerber R, Lou L, Du L. (2009). A PLP-dependent polyketide chain releasing mechanism in the biosynthesis of mycotoxin fumonisins in Fusarium verticillioides. J Am Chem Soc. 131: 3148–3149.

Glenn AE, Zitomer NC, Zimeri AM, Williams LD, Riley RT, Proctor RH. (2008). Transformation-mediated complementa-tion of a FUM gene cluster deletion in Fusarium verticilioides restores both fumonisin production and pathogenicity on maize seedlings. Mol Plant-Microbe Interact. 21: 87–97.

Harris LJ, Gleddie SC. (2001). A modified Rpl3 gene from rice confers tolerance of the Fusarium graminearum mycotoxin deoxynivalenol to transgenic tobacco. Physiol Mol Plant Pathol. 58: 1–9.

Harris LJ, Desjardins AE, Plattner RD, Nicholson P, Butler G, Young JC, Weston G, Proctor RH, Hohn TM. (1999). Possible role of trichothecene mycotoxins in virulence of Fusarium graminearum on maize. Plant Dis. 83: 954–960.

Hesketh AR, Gledhill L, Bycroft BW, Dewick PM, Gilbert J. (1993). Potential inhibitors of trichothecene biosynthesis in Fusarium culmorum. Phytochemistry. 32: 93–104.

Hesketh AR, Gledhill L, Marsh DC, Bycroft BW, Dewick PM, Gilbert J. (1991). Biosynthesis of trichothecene mycotoxins: identification of isotrichodiol as a post-trichothecene inter-mediate. Phytochemistry. 30: 2243.

Hohn TM, Beremand, P. (1989). Isolation and nucleotide sequence of a sesquiterpene cyclase gene from the trichothecene- producing fungus Fusarium sporotrichioides. Gene. 79: 131–138.

Hohn TM, Desjardins AE. (1992). Isolation and gene disruption of the Tox5 gene encoding trichodiene synthase in Gibberella pulicaris. Mol Plant Microbe Interact. 5: 249–256.

Hohn TM, Krishna R, Proctor RH. (1999). Characterization of a transcriptional activator controlling trichothecene toxin biosynthesis. Fungal Genet Biol. 26: 224–235.

Hohn TM, McCormick SP, Desjardins AE. (1993). Evidence for a gene cluster involving trichothecene-pathway biosynthetic genes in Fusarium sporotrichioides. Curr Genet. 24: 291–295.

Hohn TM, Vanmiddlesworth F. (1986). Purification and character-ization of the sesquiterpene cyclase trichodiene synthetase from Fusarium sporotrichioides. Arch Biochem Biosphys. 251: 756–761.

Jurgenson JE, Bowden RL, Zeller KA, Leslie JF, Alexander NA, Plattner RD. (2002). A genetic map of Gibberella zeae (Fusarium graminearum). Genetics. 160: 1431–1460.

Keller NP, Brown DW, Butchko RA, Fernandes M, Kelkar H, Nesbitt C, Segner S, Bhatnagar D, Cleveland TE, Adams TH. (1995). A conserved polyketide mycotoxin gene cluster in Aspergillus nidulans. In Eklund M, Richard JL, Mise K, eds. Molecular Approaches to Food Safety Issues Involving Toxic Microorganisms. Alaken Inc, Fort Collins, pp. 263–277.

Kimura M, Kaneko I, Komiyama M, Takatsuki A, Koshino H, Yoneyama K, Yamaguchi I. (1998). Trichothecene 3-O-acetyltransfease protects both the producing organism and transformed yeast from related mycotoxins. J Biol Chem. 273: 1654–1661.

Kimura M, Tokai T, O’Donnell K, Ward TJ, Fujimura M, Hamamoto H, Shibata T, Yamaguchi I. (2003). The trichothecene biosynthesis

gene cluster of Fusarium graminearum F15 contains a lim-ited number of essential pathway genes and expressed non- essential genes. FEBS Lett. 359: 105–110.

Kimura M, Tokai T, Takahashi-Ando N, Ohsato S, Fujimura M. (2007). Molecular and genetic studies of Fusarium tri-chothecene biosynthesis pathways, genes, and evolution. Biosci Biotechnol Biochem. 71: 2105–2123.

Laurent D, Platzer N, Kohler F, Sauviat MP, Pellegrin G. (1989). Macrofusine et micromoniline: duex nouvelles mycotoxines isolées de maïs infesté par Fusarium moniliforme. Microbiol Alim Nutr. 7: 9–16.

Lee J, Jurgenson JE, Leslie JF, Bowden RL. (2008). Alignment of genetic and physical maps of Gibberella zeae. Appl Environ Microbiol. 74: 2349–2359.

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. Appl Environ Microbiol. 68: 2148–2154.

Lee T, Oh D-W, Kim H-S, Lee J, Kim Y-H, Yun S-H, Lee Y-W. (2001). Identification of deoxynivalenol- and nivalenol-producing chemotypes of Gibberella zeae by using PCR. Appl Environ Microbiol. 67: 2966–2972.

Leslie JF, Anderson LL, Bowden RL, Lee Y-W. (2007). Inter- and intra-specific genetic variation in Fusarium. Int J Food Microbiol. 119: 25–32.

MacKenzie SE, Savard ME, Blackwell BA, Miller JD, ApSimon JW. (1998). Isolation of a new fumonisin from Fusarium monili-forme grown in liquid culture. J Nat Prod. 61: 367–369.

Maier FJ, Miedaner T, Hadeler B, Felk A, Salomon S, Lemmens M, Kassner H, Schäfer W. (2006). Involvement of trichothecnes 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. Molec Plant Pathol. 7: 449–461.

Manoharan M, Dahleen LS, Hohn TM, Neate SM, Yu X, Alexander NJ, McCormick SP, Bregitzer P, Schwarz PB, Horsley RD. (2006). Expression of 3-OH trichothecene acetyl-transferase in barley (Hordeum vulgare L ) and effects on deoxynivalenol. Plant Sci. 71: 699–796.

Marasas WFO, Nelson PE, Toussoun TA. (1984). Toxigenic Fusarium Species: Identity and Mycotoxicology. University Park, PN: The Pennsylvania State University Press.

McCabe AP, Riach MBR, Unkles SE, Kinghorn JR. (1990). The Aspergillus nidulans npeA locus consists of three contiguous genes required for penicillin biosynthesis. EMBO J. 9: 279–287.

McCormick SP Alexander NA. (2002). Fusarium TRI8 encodes a trichothecene C-3 esterase. Appl Environ Microbiol. 68: 2959–2964.

McCormick SP, Alexander NA, Proctor RH. (2006a). Fusarium Tri4 encodes a multifunctional oxygenase required for trichothecene biosynthesis. Can J Microbiol. 52: 636–642.

McCormick SP, Alexander NJ, Proctor RH. (2006b). Heterologous expression of two trichothecene P450 genes in Fusarium verticillioides. Can J Microbiol. 52: 220–226.

McCormick SP, Alexander NJ, Trapp SC, Hohn TM. (1999). Disruption of TRI101, the gene encoding trichothecene 3-O-acetyltransferase, from Fusarium sporotrichioides. Appl Environ Microbiol. 65: 5252–5256.

McCormick SP, Harris LJ, Alexander NJ, Ouellet T, Saparno A, Allard S, Desjardins AE. (2004). Tri1 in Fusarium graminearum encodes a P450 oxygenase. Appl Environ Microbiol. 70: 2044–2051.

McCormick SP, Hohn TM. (1997). Accumulation of trichothecenes in liquid cultures of a Fusarium sporotrichioides mutant



lacking a functional trichothecene C-15 hydroxylase. Appl Environ Microbiol. 63: 1685–1688.

McCormick SP, Hohn TM, Desjardins AE. (1996). Isolation and char-acterization of Tri3, a gene encoding 15-O- acetyltransferase from Fusarium sporotrichioides. Appl Environ Microbiol. 62: 353–359.

McCormick SP, Taylor SL, Plattner RD, Beremand MN. (1989). New modified trichothecenes accumulated in solid culture by mutant strains of Fusarium sporotrichioides. Appl Environ Microbiol. 55: 2195–2199.

McCormick SP, Taylor SL, Plattner RD, Beremand MN. (1990). Bioconversion of possible T-2 toxin precursors by a mutant strain of Fusarium sporotrichioides NRRL 3299. Appl Environ Microbiol. 56: 702–706.

Meek IB, Peplow AW, Ake C, Phillips TD, Beremand MN. (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. Appl Environ Microbiol. 69: 1607–1613.

Merrill AH, Schmelz E-M, Dillehay DL, Spiegel S, Shayman JA, Schroeder JJ, Riley RT, Voss KA, Wang E. (1997). Sphingolipids–the enigmatic lipid class: biochemistry, physiology, and pathophysiology. Toxicol Appl Pharmacol. 142: 208–225.

Mitterbauer R, Poppenberger B, Raditschnig A, Lucyshyn D, Lemmens M, Glössl J, Adam G. (2004). Toxin-dependent utilization of engineered ribosomal protein L3 limits tri-chothecene resistance in transgenic plants. Plant Biotech J. 2: 329–340.

Munkvold GP, Desjardins AE. (1997). Fumonisins in maize: can we reduce their occurrence? Plant Dis. 81: 556–565.

O’Donnell K, Ward TJ, Geiser DM, Kistler HC, Aoki T. (2006). Genealogical concordance between the mating type locus and seven other nuclear genes supports formal recognition of nine phylogenetically distinct species within the Fusarium graminearum clade. Fungal Genet Biol. 41: 600–623.

Okubara PA, Blechl AE, McCormick SP, Alexander NJ, Dill-Macky R, Hohn TM. (2002). Engineering deoxynivalenol metabolism in wheat through the expression of a fungal trichothecene acetyltransferase gene. Theor Appl Genet. 106: 74–83.

Paul PA, Lipps PE, Madden LV. (2006). Meta-analysis of regression coefficients for the relationship between Fusarium head blight and deoxynivalenol content of wheat. Phytopathology. 96: 951–961.

Peplow AW, Meek IB, Wiles MC, Phillips TD, Beremand MN. (2003a). Tri16 is required for esterification of position C-8 during trichothecene mycotoxin production by Fusarium sporotrichioides. Appl Environ Microbiol. 69: 5935–5940.

Peplow AW, Tag AG, Garifullina GF, Beremand MN. (2003b). Identification of new genes positively regulated by TRI10 and a regulatory network for trichothecene mycotoxin pro-duction. Appl Environ Microbiol. 69: 2731–2736.

Plattner RD, Desjardins AE, Leslie JF, Nelson PE. (1996). Identification and characterization of strains of Gibberella fujikuroi mating population A with rare fumonisin produc-tion phenotypes. Mycologia. 88: 416–424.

Poppenberger B, Berthiller F, Lucyshyn D, Sieberer T, Schuhmacher R, Krska R, Kuchler K, Glössl J, Luschnig C, Adam G. (2003). Detoxification of the Fusarium myco-toxin deoxynivalenol by a UDP-glucosyltransferase from Arabidopsis thaliana. J Biol Chem. 278: 47905–47914.

Proctor RH, Brown DW, Plattner RD, Desjardins AE. (2003). Co-expression of 15 contiguous genes delineates a fumoni-sin biosynthetic gene cluster in Gibberella moniliformis. Fungal Genet Biol. 38: 237–249.

Proctor RH, Busman M, Seo J-A, Lee Y-W, Plattner RD. (2008). A fumonisin biosynthetic gene cluster in Fusarium oxysporum

strain O-1890 and the genetic basis for B versus C fumonisin production. Fungal Genet Biol. 45: 1016–1026.

Proctor RH, Desjardins AE, Plattner RD. (1999a). Biosynthetic and genetic relationships of B-series fumonisins produced by Gibberella fujikuroi mating population A. Nat Toxins. 7: 251–258.

Proctor RH, Desjardins AE, Plattner RD, Hohn TM. (1999b). A polyketide synthase gene required for biosynthesis of fumonisin mycotoxins in Gibberella fujikuroi mating popu-lation A. Fungal Genet Biol. 27: 100–112.

Proctor RH, Hohn TM, McCormick SP. (1995a). Reduced virulence of Gibberella zeae caused by disruption of a trichothecene toxin biosynthectic gene. Mol Plant Microbe Interact. 8: 593–601.

Proctor RH, Hohn TM, McCormick SP, Desjardins AE. (1995b). Tri6 encodes an unusual zinc finger protein involved in regulation of trichothecene biosynthesis in Fusarium sporo-trichioides. Appl Environ Microbiol. 61: 1923–1930.

Proctor RH, Plattner RD, Brown DW, Seo J-A, Lee Y-W. (2004). Discontinuous distribution of fumonisin biosynthetic genes in the Gibberella fujikuroi species complex. Mycol Res. 108: 815–822.

Proctor RH, Plattner RD, Desjardins AE, Busman M, Butchko RAE. (2006). Fumonisin production in the maize pathogen Fusarium verticillioides: genetic basis of naturally occurring chemical variation. J Agric Food Chem. 54: 2424–2430.

Rheeder JP, Marasas WFO, Vismer HF. (2002). Production of fumonisin analogs by Fusarium species. Appl Environ Microbiol. 68: 2101–2105.

Riley RT, Wang E, Schroeder JJ, Smith ER, Plattner RD, Abbas H, Yoo H-S, Merrill AF. (1996). Evidence for disruption of sphin-golipid metabolism as a contributing factor in the toxicity and carcinogenicity of fumonisins. Nat Toxins. 4: 3–15.

Robertson-Hoyt LA, Betrán J, Payne GA, White DG, Isakeit T, Maragos CM, Molnár TL, Holland JB. (2007). Relationships among resistances to Fusarium and Aspergillus ear rots and contamination by fumoinisin and aflatoxin in maize. Phytopathology. 97: 311–317.

Rynkiewicz MJ, Cane DE, Christianson DW. (2001). Structure of trichodiene synthase from Fusarium sporotrichioides provides mechanistic inferences on the terpene cyclization cascade. Proc Natl Acad Sci U S A. 98: 13543–13548.

Seo J-A, Kim J-C, Lee Y-W. (1996). Isolation and characteriza-tion of two new type C fumonisins produced by Fusarium oxysporum. J Nat Prod. 59: 1003–1005.

Seo J-A, Proctor RH, Plattner RD. (2001). Characterization of four clustered and coregulated genes associated with fumonisin biosynthesis in Fusarium verticillioides. Fungal Genet Biol. 34: 155–165.

Sewram V, Mshicileli N, Shepard GS, Vismer HF, Rheeder JP, Lee Y-W, Leslie JF, Marasas WFO. (2005). Production of fumonisin B and C analogues by several Fusarium species. J Agric Food Chem. 53: 4861–4866.

Shim W-B, Woloshuk CP. (2001). Regulation of fumonisin B1 biosynthesis and conidiation in Fusarium verticillioides by a cyclin-like (C-type) gene, FCC1. Appl Environ Microbiol. 67: 1607–1612.

Skory CD, Chang P-K, Cary J, Linz JE. (1992). Isolation and charac-terization of a gene from Aspergillus parasiticus associated with the conversion of versicolorin A to sterigmatocystin in afla-toxin biosynthesis. Appl Environ Microbiol. 58: 3527–3537.

Starkey DE, Ward TJ, Aoki T, Gale LR, Kistler HC, Geiser DM, Suga H, Tóth B, Varga J, O’Donnell K. (2007). Global molecular surveillance reveals novel Fusarium head blight species and trichothecene toxin diversity. Fungal Genet Biol. 44: 1191–1204.



Suga H, Karugia GW, Ward T, Gale LR, Tomimura K, Nakajima T, Miyasaka A, Koizumi S, Kageyama K, Hyakumachi M. (2008). Molecular characterization of the Fusarium graminearum species complex in Japan. Phytopathology. 98: 159–166.

Tag AG, Garifullina GF, Peplow AW, Ake C, Phillips TD, Hohn TM, Beremand MN. (2001). A novel regulatory gene, Tri10, con-trols trichothecene toxin production and gene expression. Appl Environ Microbiol. 67: 5294–5302.

Tokai T, Koshino H, Takahashi-Ando N, Sato M, Fujimura M, Kimura M. (2007). Fusarium Tri4 encodes a key multifunc-tional cytochrome P450 monooxygenase for four consecutive oxygenation steps in trichothecene biosynthesis. Biochem Biophys Res Commun. 353: 412–417.

Waalwijk C, van der Lee T, de Vries I, Hesselink T, Arts J, Kema GHJ. (2004). Synteny in toxigenic Fusarium species: the fumoni-sin gene cluster and the mating type region as examples. Eur J Plant Pathol. 110: 533–544.

Ward TJ, Bielawski JP, Kistler HC, Sullivan E, O’Donnell K. (2002). Ancestral polymorphism and adaptive evolution in the tri-chothecene mycotoxin gene cluster of phytopathogenic Fusarium. Proc Natl Acad Sci U S A. 99: 9278–9283.

Ward TJ, Clear RM, Rooney AP, O’Donnell D, Gaba, D, Patrick S, Starkey DE, Gilbert J, Geiser DM, Nowicki TW. (2008). An adaptive evolutionary shift in Fusarium head blight patho-gen populations is driving the rapid spead of more toxigenic Fusarium graminearum in North America. Fungal Genet Biol. 45: 473–484.

Windels C. (2000). Economic and social impacts of Fusarium head blight: changing farms and rural communities in the northern Great Plains. Phytopathology. 90: 17–21.

Xu J-R, Leslie JF. (1996). A genetic map of Gibberella fujikuroi mating population A (Fusarium moniliforme). Genetics. 143: 175–189.

Yi H, Bojja RS, Fu J, Du L. (2005). Direct evidence for the function of FUM13 in 3-ketoreduction of mycotoxin fumonisins in Fusarium verticillioides. J Agric Food Chem. 53: 5456–5460.

Yoder OC, Valent, B Chumley F. (1986). Genetic nomenclature and practice for plant pathogenic fungi. Phytopathology. 76: 383–385.

Yu F, Zhu X, Du L. (2005). Developing a genetic system for functional manipulations of FUM1, a polyketide syn-thase gene for the biosynthesis of fumonisins in Fusarium verticillioides. FEMS Microbiol Lett. 248: 257–264.

Zaleta-Rivera K, Xu C, Yu F, Butchko RAE, Proctor RH, Hidalgo-Lara ME, Raza A, Dussault PH, Du L. (2006). A bi-domain non-ribosomal peptide synthetase encoded by FUM14 catalyzes the formation of tricarballylic esters in the biosynthesis of fumonisins. Biochemistry. 45: 2561–2569.

Zamir LO, Devor KA, Morin N, Sauriol F. (1991). Biosynthesis of trichothecenes: oxygenation steps post-trichodiene. J Chem Soc Chem Commun. 1991: 1033–1034.

Zamir LO, Devor KA, Nikolakakis A, Sauriol F. (1990). Biosynthesis of Fusarium culmorum trichothecenes: the roles of isotri-chodermin and 12,13-epoxytrichothec-9-ene. J Biol Chem. 265: 6713–6725.

Zamir LO, Nikolakakis A, Huang L, St-Pierre P, Sauriol F, Sparace S, Mamer O. (1999). Biosynthesis of 3-acetyldeoxynivalenol and sambucinol: identification of the two oxygenation steps after trichodiene. J Biol Chem. 274: 12269–12277.

Zhu X, Yu F, Li X-C, Du L. (2007). Production of dihydroisocour-marins in Fusarium verticillioides by swapping ketosynthase domain of the fungal iterative polyketide synthase Fum1p with that of lovastatin diketide synthase. J Am Chem Soc. 129: 36–37.


Genes, gene clusters, and biosynthesis of trichothecenes and fumonisins in Fusarium

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