Abstract
Sphinganine-analog mycotoxins (SAMT) are polyketide-derived natural products produced by a number of plant pathogenic fungi and are among the most economically important mycotoxins. The toxins are structurally similar to sphinganine, a key intermediate in the biosynthesis of ceramides and sphingolipids, and competitive inhibitors for ceramide synthase. The inhibition of ceramide and sphingolipid biosynthesis is associated with several fatal diseases in domestic animals and esophageal cancer and neural tube defects in humans. SAMT contains a highly reduced, acyclic polyketide carbon backbone, which is assembled by a single module polyketide synthase. The biosynthesis of SAMT involves a unique polyketide chain-releasing mechanism, in which a pyridoxal 5′-phosphate-dependent enzyme catalyzes the termination, offloading and elongation of the polyketide chain. This leads to the introduction of a new carbon–carbon bond and an amino group to the polyketide chain. The mechanism is fundamentally different from the thioesterase/cyclase-catalyzed polyketide chain releasing found in bacterial and other fungal polyketide biosynthesis. Genetic data suggest that the ketosynthase domain of the polyketide synthase and the chain-releasing enzyme are important for controlling the final product structure. In addition, several post-polyketide modifications have to take place before SAMT become mature toxins.
Similar content being viewed by others
Avoid common mistakes on your manuscript.
Introduction
Mycotoxins are fungal secondary metabolites produced by a large number of phytopathogenic and food spoilage fungi, for example, Aspergillus, Penicillium, Fusarium, and Alternaria species. These species produce some of the most important mycotoxins, including trichothecenes, aflatoxins, sterigmatocystin, fumonisins, and AAL-toxins [1, 23, 73]. Extensive research has been devoted to aflatoxins, sterigmatocystin, and trichothecenes, and their biosynthesis is relatively well understood [11, 23, 39, 53]. In recent years, significant research efforts have also been given to studying the biosynthetic mechanism for fumonisins and AAL-toxins (Fig. 1) [28, 77]. Fumonisins and AAL-toxins together are called sphinganine-analog mycotoxins (SAMT) due to their structural similarity to sphinganine (dihydrosphingosine, DHS), which is the backbone precursor of sphingolipids (Fig. 1) [9]. Fumonisins are produced by the fungus F. verticillioides (synonym F. moniliforme, teleomorph Gibberella moniliformis, synonym G. fujikuroi Mating Population A), which is a widespread pathogen of corn and contaminates maize-based food and feed worldwide [50, 56]. AAL-toxins are produced by tomato pathogen Alternaria alternata f. sp. Lycopersici [7, 8, 37]. The mechanism for SAMT to execute their toxicity is through the competitive inhibition of sphinganine N-acetyltransferase (ceramide synthase) [50, 52]. This leads to the obstruction of complex sphingolipid biosynthesis, such as the important second messenger ceramide in animal systems, and the accumulation of sphinganine. The inhibition of this enzyme leads to various diseases in animals and humans as ceramides and sphingolipids are ubiquitous constituents of eukaryotic cells and involved in crucial signal transduction of numerous cellular processes [50, 65]. It is known that fumonisins can cause leukoencephalomalacia and pulmonary edema syndrome in animals and are associated to human esophageal cancer and neural tube defects [50]. SAMT are also found to induce apoptosis [78]. In addition to their animal toxicity, AAL-toxins are known as the primary determinants of phytopathogenicity as in the case of stem-canker disease in tomatos [82]. In this review, we examine our current understanding on the biosynthesis of SAMT, with a focus on the biosynthetic mechanism for the polyketide chain.
The biosynthetic origins of sphinganine-analog mycotoxins
The biosynthetic origins of SAMT have been studied by several groups using isotope-labeled precursors (Fig. 2). For fumonisins, the 18-carbon backbone from C-3 to C-20 is derived from acetate, and C-1 and C-2, as well as the C-2 amino, are derived from alanine [5, 10]. The two methyl groups at C-12 and C-16 are derived from methionine [59]. Although multiple hydroxyls are present on the polyketide chain, only the hydroxyl at C-3 is derived from acetate; the hydroxyls at C-5, C-10, C-14, and C-15 of FB1 are derived from molecular oxygen [16]. Therefore, the initially synthesized carbon chain is a highly reduced polyketide. The origin of the two tricarballylic esters is not certain, but most likely from the citric acid cycle [5, 87]. For AAL-toxins, the biosynthetic origin is similar to that of fumonisins, except that the acetate-originated carbon chain (from C-2 to C-17) is two carbons shorter, C-1 and the amino group are derived from glycine, and only one tricarballylic ester is present (either at C-13 or C-14 hydroxyl) (Fig. 1). Biosynthetic origins, as well as the presence of methyl groups on the carbon backbone, suggest that the carbon backbone of SAMT is assembled via a polyketide biosynthetic mechanism.
Polyketides and polyketide synthases (PKS)
Polyketides are a large, structurally diverse family of metabolites [72]. They are biosynthesized through sequential decarboxylative condensations of short carboxylic acids, which are catalyzed by polyketide synthases (PKS) [19, 27, 32, 40, 67]. Throughout the biosynthesis, the growing polyketide intermediates remain covalently attached to the ACP (acyl carrier protein) domain via the 4′-phosphopantetheine cofactor [49, 76]. In bacterial modular PKSs (Type I), the mechanism by which PKS control structural variations in the products is relatively well understood. The order, number, and domain-composition of the modules dictate the length and reduction level of the polyketide carbon backbone [32, 40, 72]. Mediated by the hydrolytic function of the thioesterase (TE) domain located at the end of the modules, the product is released with a length depending on the number of elongation cycles. For example, the three bi-modular PKS, 6-deoxyerythronolide B synthases (DEBS-1, DEBS-2, and DEBS-3) that make the precursor for erythromycins in Saccharopolyspora erythraea, catalyze six cycles of decarboxylative condensation from one molecule of propionyl-CoA and six molecules of 2-methylmalonyl-CoA to synthesize the 15-carbon backbone of erythromycins [19, 26, 27, 44, 74]. Because the position of the modules dictates the course of catalytic events in the bacterial modular PKS, the pathways can be modified in semi-predictable fashion by moving, deleting, or adding modules [17, 18, 43, 45, 51, 54]. Novel polyketides with complex structures have been produced by manipulating the biosynthetic machinery.
Fungal polyketides and fungal PKS
So far, most of the biosynthetic investigations have been carried out on polyketides that are isolated from soil bacteria, especially in the order of Actinomycetes. Although filamentous fungi are known to produce numerous polyketide natural products, our understanding of the biosynthetic mechanism for these products is still at the infant stage. Many fungal polyketides are biologically active, economically important natural products [20]. These polyketides have significant impacts on human health because many of the fungal producers are unavoidable species in our daily lives. For example, aflatoxins produced by the peanut pathogens Aspergillus flavus and A. parasiticus are very potent carcinogens [53, 57], fumonisins produced by the corn pathogen F. verticillioides cause several fatal diseases in livestock and are associated with human esophageal cancer and neural tube defects [50, 65], fusarin C produced by F. verticillioides is a potent mycotoxin [70], zearalenones produced by the wheat pathogens F. graminearum and F. roseum are potent estrogens [60], melanins produced by the human pathogen A. fumigatus are virulence factors for this fungus to cause pneumonia and invasive disseminated diseases [75, 80], T-toxins produced by the corn pathogen Cochliobolus heterostrophus [84] and AAL-toxins by the tomato pathogen Alternaria alternata f. sp. Lycopersici [37] are the pathogenicity factors essential for the fungi to infect plants, equisetin produced by the corn/wheat pathogen F. heterosporum is an HIV-1 integrase inhibitor [69], squalestatin produced by Phoma sp. is a potent inhibitor for mammalian squalene synthase [21], tenellin produced by the insect pathogen Beauveria bassiana is toxic to mammalian erythrocytes and probably important for this fungus to cause white muscardine disease in the domestic silk worm [29, 83], 6-methylsalicylic acid produced by several pathogenic Penicillium sp. is a precursor of the mycotoxin patulin [24, 71], and lovastatin/compactin produced by A. terreus is an important cholesterol-lowering drug [42, 68].
Like bacterial type I PKS, fungal PKS are modular enzymes, but typically contain only a single set of domains [72] (Fig. 3). Therefore, the single module must be iteratively used during the polyketide chain elongation. Fungal PKS are divided into three groups based on the reduction level of their products, non-reducing PKS (NR-PKS), partially reducing PKS (PR-PKS), and highly reducing PKS (HR-PKS) [4, 48, 58]. NR-PKSs synthesize aromatic polycyclic compounds, such as tetrahydroxy naphthalene synthase (THNS) in Colletotrichum lagenarium [33] and Wangiella dermatitidis [30]. Their domain architecture has recently been redefined, which includes several unique domains [22] (Fig. 3a). The domains consist of SAT (starter unit ACP transacylase), KS (β-ketoacylsynthase), AT (acyl transferase), PT (product template), ACP (acyl carrier protein), and TE/CLC (thiolesterase/Claisen-like cyclase). KS-AT-ACP are the “minimal PKS” found in all PKS systems and required for chain extension. The studies of aflatoxin PKS (norsolorinic acid synthase) showed that SAT controls chain initiation and PT involves chain length control [20] (Townsend, personal communication). TE/CLC controls cyclization and chain releasing and probably also chain-length determination [35, 79].
Fungal partially reducing PKS synthesize cyclic compounds like 6-methylsalicylic acid (6-MSA), which has an aromatic ring but results from a partially reduced intermediate (Fig. 3b). The domain architecture of PR-PKS differs considerably from NR-PKS [2, 34]. 6-MSAS consists of KS-AT-DH-Core-KR-ACP. It does not have SAT, PT, or TE/CLC, but has β-keto processing domains, KR (β-ketoacyl reductase), DH (dehydratase), which are selectively used during the biosynthesis to produce “partially” reduced intermediates. The so-called Core domain is probably involved in the subunit–subunit interaction, as 6-MSAS is a homotetramer [55].
Fungal highly reducing PKS synthesize non-aromatic compounds like fumonisins, AAL-toxins, and lovastatin (Fig. 3C). The domain architecture resembles a module of bacterial Type-I PKSs, typically consisting of KS-AT-DH-MT-ER-KR-ACP. Again, it does not have SAT, PT, or TE/CLC, but contains a complete set of β-keto processing domains, KR, DH, ER (enoylreductase), as well as a methyltransferase (MT) that is found only in HR-PKS among the three groups [20]. The mechanism for HR-PKS programming has been a mystery. It is not clear what controls the number of chain elongation and the reduction level of the carbon chain, as HR-PKS with identical domain architecture can synthesize products with huge variations in chain-lengths and reduction level [20]. In addition, the PKS does not have an obvious way of terminating and offloading the linear polyketide chain.
Sphinganine-analog mycotoxins as a model system to study the biosynthesis of fungal highly reduced polyketides
The study of the biosynthetic mechanism for fungal highly reduced polyketides has been challenging due to the difficulties in detecting the intermediates with a linear, non-aromatic structure and in manipulating the multi-domain PKS genes in the filamentous fungi. Our laboratory has been studying the biosynthesis of SAMT and using it as a model system for the biosynthesis of fungal highly reduced polyketides [28, 88, 89]. This system has several advantages. Fumonisins and AAL-toxins are structurally similar, and the main difference is the length of polyketide carbon chain. Fumonisins have a dimethylated 18-carbon chain, while AAL-toxins have a dimethylated 16-carbon chain. The PKS for fumosinins (Fum1p, 2507 residues, encoded by FUM1) [62] and AAL-toxins (Alt1p, 2521 residues, encoded by ALT1) (personal communication with Dr. Motoichiro Kodama at Tottori University, Japan) have an identical domain architecture, KS-AT-DH-MT-ER-KR-ACP, and are highly homologous to each other (72%/60% similarity/identity for the entire amino acid sequence). However, Fum1p synthesizes an 18-carbon chain, while Alt1p makes a 16-carbon chain. Thus, a simple domain swapping experiment could reveal interesting insights into the mechanism for the chain-length control. Most intriguingly, Fum1p and Alt1p do not contain a thioesterase (TE) domain for releasing bacterial reduced polyketides [40] or a Claisen-like cyclase (CLC) domain for releasing fungal non-reduced, aromatic polyketides [35]. Furthermore, since the products are acyclic compounds, the polyketide-releasing through simultaneous cyclization, as seen in partially reducing PKS (Fig. 3b), cannot be involved. In addition, in fumonisin biosynthesis the linear 18-carbon chain is offloaded by condensing with the α-carbon of l-alanine, whereas in AAL-toxins, the linear 16-carbon is offloaded by condensing with the α-carbon of glycine (Fig. 3c). Therefore, the study of SAMT biosynthesis could also potentially reveal novel mechanisms for polyketide structure determination.
In order to study the biosynthetic mechanism, we first developed a genetic system that enables specific and functional manipulations of the PKS domains in F. verticillioides [86]. In bacteria, the genetic manipulation of PKS domains has proven a powerful way to elucidate the domain structure and function of PKS [40, 72]. In our system, we used a two-stage screening strategy, including both positive and negative screenings, to manipulate the domains of SAMT PKS. Since our goal is not only to specifically change the domains, but also to maintain the activity of the chimeric PKS, the one-stage protocol developed previously [62] to screen resistant colonies will not be appropriate for our purpose. In the first stage screening, hygromycin-resistant colonies were obtained by transforming fungal protoplasts with a properly constructed vector. Any of the single crossovers would result in a disrupted PKS gene, which should be nonfunctional. PCR was used to quickly distinguish putative homologous recombinants from those mutants resulted from random insertions. The putative homologous recombinants were further confirmed by Southern hybridization. In the second stage, the mutants were grown on a liquid medium without hygromycin for 4–5 generations to encourage the second crossover. The individual single colonies were then replica plated on two plates, one with hygromycin and one without. Colonies that did not grow on the plate with hygromycin but grew on the plate without hygromycin were selected as putative domain replacement mutants, which were confirmed by PCR and Southern analysis.
Genetic manipulations of the PKS for sphinganine-analog mycotoxin biosynthesis
Using this genetic system, we replaced the KS domain of fumonisin FUM1 in F. verticillioides with the KS domain of T-toxin PKS1 from C. heterostrophus. KS is the catalytic domain responsible for the carbon–carbon formation during polyketide chain assembly. A replacement of this domain will most likely lead to structural variations in products. T-toxins are a family of mycotoxins with carbon chain lengths varying from 35 to 45, and the C41 analog is most abundant (50%) [47]. Fum1p and PKS1 share a sequence similarity of 44.1% and identity of 34.2% for the entire amino acid sequence [84]. When the KS domain of FUM1 was replaced with the KS domain of PKS1, the F. verticillioides strain produced fumonisins [88]. The result shows that the heterologous KS is able to functionally interact with the six other domains of Fum1p. It also suggests that KS domain alone may not be sufficient to control the product’s structure. This represents the first successful domain swapping in fungal HR-PKS. To further test if the whole fumonisin PKS could be functionally replaced by a PKS that has a similar domain architecture, we replaced entire FUM1 with PKS1 [88]. This F. verticillioides strain did not produce any fumonisin or new metabolites. Previous heterologous expression and in vitro assays of 6-methylsalicylic acid synthase [3, 41, 64] and squalestatin tetraketide synthase [21] have shown that each of the single module fungal PKS contains the necessary information for synthesizing a distinct polyketide product. The results obtained from fumonisin biosynthesis suggest that the intrinsic interactions between the intact PKS and downstream enzymes in the biosynthetic pathway may play a key role in the control of the biosynthesis of fungal reduced polyketides.
To obtain more insights, we replaced the KS domain of FUM1 with the KS domain of LovF, which encodes lovastatin diketide synthase (LDKS) responsible for the 2-methylbutyryl side chain of lovastatin in Aspergillus terreus [42]. Since Fum1p, T-toxin PKS1 and LDKS synthesize very different polyketide chains, the outcome of this experiment is expected to reveal interesting new information on KS domain’s role in product structure determination. The KS-replaced F. verticillioides strains produced four new compounds with a yield comparable to fumonisins in the wild type. HRMS and NMR data showed that these compounds are not linear, acyclic polyketides found in SAMT, T-toxins, or lovastatin, but are rather aromatic metabolites, dihydroisocoumarins [89]. The result is surprising because, as discussed earlier, the enzyme architecture for fungal aromatic (non-reduced) polyketides is very different from that for fungal highly reduced (non-aromatic) polyketides. A possible way for the dihydroisocoumarins to arise is through inter-molecular ring formation between two short (C8 and C10), linear polyketide intermediates [89]. A series of possibly spontaneous reactions including aldol condensation, Claisen condensation, aromatization, and lactonization between the intermediates would account for the formation of the aromatic products. These results show that fungal chimeric PKS with different KS domain makes different products, suggesting that the KS domain does have an effect on the product structure. It is possible that the substrate-binding pocket of the KS domain of T-toxin PKS1 is large enough to allow the chimeric PKS to synthesize a C18 chain required for fumonisin production. This is reasonable considering that PKS1 is involved in the synthesis of polyketides (T-toxins) larger than fumonisins. Thus, T-toxin KS within the context of Fum1p is able to produce a proper polyketide chain that can be processed by the downstream enzymes in the pathway, which eventually leads to fumonisin production in the fungus. On the other hand, the substrate-binding pocket of the KS domain of LDKS could be smaller than that of Fum1p or T-toxin PKS1, since LDKS synthesizes a short carbon chain (C4). Thus, the chimeric Fum1p with the KS from LDKS could only make “short” chains that cannot be processed by the downstream enzymes and thus “stuck” on the PKS. One possible way for the release of these stuck short chains is through intermolecular ring formation, leading to the aromatic compounds. Therefore, both the KS domain and downstream processing enzymes are important for the production of a distinct group of products during fungal highly reduced polyketide biosynthesis [89].
The production of dihydroisocoumarins in F. verticillioides represents the first successful genetic manipulation of an iterative polyketide synthase gene to produce new, biologically active metabolites in a filamentous fungus. The dihydroisocoumarins are known to have anti-malarial, antifungal, and anti-tuberculosis activities [46] and had never been isolated from a Fusarium species. The new F. verticillioides strains not only stop producing mycotoxins but also produce a group of active metabolites. The fungal strains could have potential applications in agricultural biotechnology.
Polyketide chain-releasing in sphinganine-analog mycotoxin biosynthesis
The genetic manipulation of the KS domain of FUM1 has revealed interesting results for SAMT biosynthesis. Both the genetic data and biosynthetic origins of SAMT suggest that the enzymes for terminating-offloading the SAMT polyketide chains are important in determining the structure of the final products. As shown in Fig. 3, the terminating–offloading of SAMT polyketide is accomplished by a new carbon–carbon bond formation, between C-3 and C-2 in fumonisins and C-2 and C-1 in AAL toxins. Thus, a distinct chain-releasing mechanism is expected for SAMT biosynthesis. Within the FUM gene cluster, FUM8 is predicted to encode a homolog (Fum8p, 839 residues) of l-serine palmitoyltransferase, which is a member of the 2-oxoamino synthase family [66]. Similarly, in the ALT gene cluster, ALT4 is predicted to encode a homolog (Alt4p, 871 residues) of the same family of enzymes (personal communication with Dr. Motoichiro Kodama at Tottori University, Japan). The amino acid sequence of Fum8p and Alt4p shares a similarity/identify of 50%/38% for the entire sequence.
2-Oxoamino synthases are a small family of enzymes involved in several very important primary biosynthetic pathways, including l-serine palmitoyltransferase (SPT) in ceramide-sphingolipid biosynthesis [38], 5-aminolevulinate synthase (ALAS) in heme biosynthesis [31], and 8-amino-7-oxononanoate synthase (AONS) in biotin biosynthesis [81]. The enzymes are known to use pyridoxal 5′-phosphate as cofactor to catalyze the acyltransfer reactions [38]. The new carbon–carbon bond formation during the terminating–offloading of SAMT polyketide is catalytically analogous to the reactions catalyzed by the 2-oxoamino synthases. Thus, Fum8p and Alt4p are most likely the enzymes responsible for the polyketide chain releasing in the biosynthesis of fumonisins and AAL-toxins, respectively. While ALAS and AONS are homodimers, SPT is a heterodimer of two separate subunits, Lcb1p and Lcb2p. Lcb2p contains the active site lysine residue to form a Schiff’s base with pyridoxal 5′-phosphate [36]. Fum8p and Alt4p are natural fusions of Lcb2p and Lcb1p, with the N-terminal half being homologous to Lcb2p and the C-terminal half being homologous to Lcb1p.
Based on the general mechanism known for the 2-oxoamine synthases, we propose a pyridoxal 5′-phosphate-dependent releasing of polyketide acyl chain that is covalently attached to the 4′-phosphopantetheine cofactor of the ACP domain of PKS (Fum1p) (Fig. 4a). The reaction mechanism follows the sequence of deprotonation–condensation–decarboxylation–reprotonation. The PLP-enzyme uses a carbon nucleophile (carbanion derived from the α-carbon of l-alanine by deprotonation) to attack the carbonyl of polyketide acyl-S-ACP, which results in the release of the PKS (ACP-SH) and the formation of a new carbon–carbon bond in the PLP-bound aldimine intermediate (Fig. 4a). It is likely that the substrate specificity of the chain-releasing enzymes is important for controlling the size of the polyketide chain in the final products. This chain-releasing mechanism is fundamentally distinct from the TE/CLC-mediated polyketide chain releasing (Fig. 4b), which typically uses a nitrogen or oxygen nucleophile to attack the carbonyl and involves a catalytic triad (e.g. Ser-Asp-His) at the active site of TE/CLC [12, 35]. The subsequent decarboxylation, reprotonation, and transimination of this aldimine intermediate result in the regeneration of the PLP-enzyme and free 3-keto intermediate for fumonisins (Fig. 4c). The net results are (1) the termination and release of the polyketide chain from PKS, (2) the elongation of the polyketide acyl chain, and (3) the incorporation of the amino group.
Polyketide chain modifications in sphinganine-analog mycotoxin biosynthesis
On the PLP-dependent polyketide chain releasing, a 3-keto compound is produced as an intermediate. This 3-keto intermediate is stereospecifically reduced by the product of FUM13 [14, 85]. We have expressed FUM13 in E. coli and purified the 46-kDa Fum13p. When 3-keto FB3 was incubated with Fum13p in the presence of NADPH, it was converted to FB3. This demonstrates that Fum13p is an NADPH-dependent ketoreductase required for the reduction of 3-keto to 3-hydroxy in fumonisins (Fig. 5). The further modifications of the SAMT polyketide chain include several hydroxylations and esterifications [6, 13–15, 25, 28, 61, 63, 77, 85, 87]. The hydroxylation at carbon C-14, C-15, and C-10 are probably catalyzed by the P450 monooxygenases (Fum6p and Fum12p) encoded by FUM6 and FUM12 [6, 63]. However, the nature and specificity of the enzymes have not been demonstrated in vitro. The esterifications of the hydroxylated intermediate are catalyzed by a nonribosomal peptide synthetase complex (Fum10p-Fum14p-Fum7p, note that the putative tricarboxylate transporter Fum11p is likely to involve in vivo) [15, 87]. We have expressed FUM14 in E. coli. This gene was predicted to encode the peptidyl carrier protein (PCP) and condensation (C) domain of a nonribosomal peptide synthetase [87]. Fum14p was shown to convert HFB3 and HFB4, which are biosynthetic precursors of fumonisins lacking the tricarballylic esters, to the tricarballylic esters-containing fumonisins, FB3 and FB4, respectively, when incubated with tricarballylic thioester of N-acetylcysteamine. Interestingly, the nonribosomal peptide synthetase catalyzes a C–O bond (ester) formation, instead of the typical C–N bond (amide) formation. The final step of biosynthesis is the hydroxylation at C-5, which is catalyzed by the product of FUM3 gene [13, 25]. We have expressed FUM3 in Saccharomyces cerevisiae and purified the 33-kDa Fum3p. The enzyme was able to convert FB3 to FB1 in the presence of α-ketoglutarate, Fe2+, ascorbic acid, and catalase. This demonstrates that Fum3p is a α-ketoglutarate-dependent dioxygenases required for C-5 hydroxylation of fumonisins (Fig. 5).
Conclusion
As an important group of mycotoxins, SAMT represent an unusual type of fungal highly reduced polyketides. Like other fungal polyketides, the biosynthesis of SAMT involves a single module PKS. The architecture of fungal HR-PKS is distinct from that of NR-PKS and PR-PKS. The study of SAMT biosynthesis is starting to reveal a unique chain-releasing mechanism that is not found in any other polyketides. Most notably, this mechanism allows the formation of a new carbon–carbon bond and introduction of a new functional group during the polyketide chain-releasing step, which is an unprecedented way of creating structural diversity of polyketides. Filamentous fungi produce numerous polyketide metabolites. Many of the metabolites have important biological activities (such as virulence factors, carcinogens, and therapeutics), fascinating molecular architectures (such as aflatoxins and T-toxins), and enormous commercial value (such as lovastatin). These polyketides play a significant role in human and animal health because of their presence in many indispensable fungal species that affect our everyday lives. The understanding of the molecular mechanism for SAMT biosynthesis will contribute to the development of new strategies toward mycotoxin reduction and elimination in food industry and agriculture.
References
Abbas HK, Duke SO, Shier WT, Riley RT, Kraus GA (1996) The chemistry and biological activities of the natural products AAL-toxin and the fumonisins. Adv Exp Med Biol 391:293–308
Beck J, Ripka S, Siegner A, Schiltz E, Schweizer E (1990) The multifunctional 6-methylsalicylic acid synthase gene of Penicillium patulum. Its gene structure relative to that of other polyketide synthases. Eur J Biochem 192:487–498
Bedford DJ, Schweizer E, Hopwood DA, Khosla C (1995) Expression of a functional fungal polyketide synthase in the bacterium Streptomyces coelicolor A3(2). J Bacteriol 177:4544–4548
Bingle LE, Simpson TJ, Lazarus CM (1999) Ketosynthase domain probes identify two subclasses of fungal polyketide synthase genes. Fungal Genet Biol 26:209–223
Blackwell BA, Edwards OE, Fruchier A, ApSimon JW, Miller JD (1996) NMR structural studies of fumonisin B1 and related compounds from Fusarium moniliforme. Adv Exp Med Biol 392:75–91
Bojja RS, Cerny RL, Proctor RH, Du L (2004) Determining the biosynthetic sequence in the early steps of the fumonisin pathway by use of three gene-disruption mutants of Fusarium verticillioides. J Agric Food Chem 52:2855–2860
Bottini AT, Bowen JR, Gilchrist DG (1981) Phytotoxins. II. A characterization of a phytotoxic fraction from Alternaria alternata f. sp. lycopersici. Tetrahedron Lett 22:2723–2726
Bottini AT, Gilchrist DG (1981) Phytotoxins. I. A 1-aminodimethylheptadecapentol from Alternaria alternata f. sp. lycopersici. Tetrahedron Lett 22:2719–2722
Brandwagt BF, Mesbah LA, Takken FL, Laurent PL, Kneppers TJ, Hille J, Nijkamp HJ (2000) A longevity assurance gene homolog of tomato mediates resistance to Alternaria alternata f. sp. lycopersici toxins and fumonisin B1. Proc Natl Acad Sci USA 97:4961–4966
Branham BE, Plattner RD (1993) Alanine is a precursor in the biosynthesis of fumonisin B1 by Fusarium moniliforme. Mycopathologia 124:99–104
Brown DW, Yu JH, Kelkar HS, Fernandes M, Nesbitt TC, Keller NP, Adams TH, Leonard TJ (1996) Twenty-five coregulated transcripts define a sterigmatocystin gene cluster in Aspergillus nidulans. Proc Natl Acad Sci USA 93:1418–1422
Bruner SD, Weber T, Kohli RM, Schwarzer D, Marahiel MA, Walsh CT, Stubbs MT (2002) Structural basis for the cyclization of the lipopeptide antibiotic surfactin by the thioesterase domain SrfTE. Structure 10:301–310
Butchko RA, Plattner RD, Proctor RH (2003) FUM9 is required for C-5 hydroxylation of fumonisins and complements the meitotically defined Fum3 locus in Gibberella moniliformis. Appl Environ Microbiol 69:6935–6937
Butchko RA, Plattner RD, Proctor RH (2003) FUM13 encodes a short chain dehydrogenase/reductase required for C-3 carbonyl reduction during fumonisin biosynthesis in Gibberella moniliformis. J Agric Food Chem 51:3000–3006
Butchko RA, 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 B1 and AAL toxins. J Agric Food Chem 46:4734–4743
Cane DE, Walsh CT, Khosla C (1998) Harnessing the biosynthetic code: combinations, permutations, and mutations. Science 282:63–68
Cheng Q, Xiang L, Izumikawa M, Meluzzi D, Moore BS (2007) Enzymatic total synthesis of enterocin polyketides. Nat Chem Biol 3:557–558
Cortes J, Haydock SF, Roberts GA, Bevitt DJ, Leadlay PF (1990) An unusually large multifunctional polypeptide in the erythromycin-producing polyketide synthase of Saccharopolyspora erythraea. Nature 348:176–178
Cox RJ (2007) Polyketides, proteins, genes in fungi programmed nano-machines begin to reveal their secrets. Org Biomol Chem 5:2010–2026
Cox RJ, Glod F, Hurley D, Lazarus CM, Nicholson TP, Rudd BA, Simpson TJ, Wilkinson B, Zhang Y (2004) Rapid cloning and expression of a fungal polyketide synthase gene involved in squalestatin biosynthesis Chem Commun (Camb) 20:2260–2261
Crawford JM, Dancy BC, Hill EA, Udwary DW, Townsend CA (2006) Identification of a starter unit acyl-carrier protein transacylase domain in an iterative type I polyketide synthase. Proc Natl Acad Sci USA 103:16728–16733
Desjardins AE, Proctor RH (2007) Molecular biology of Fusarium mycotoxins. Int J Food Microbiol 119(1-2):47–50 (Epub ahead of print, PMID: 17707105)
Dimroth P, Ringelmann E, Lynen F (1976) 6-Methylsalicylic acid synthetase from Penicillium patulum. Some catalytic properties of the enzyme and its relation to fatty acid synthetase. Eur J Biochem 68:591–596
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
Donadio S, Katz L (1992) Organization of the enzymatic domains in the multifunctional polyketide synthase involved in erythromycin formation in Saccharopolyspora erythraea. Gene 111:51–60
Donadio S, Staver MJ, McAlpine JB, Swanson SJ, Katz L (1991) Modular organization of genes required for complex polyketide biosynthesis. Science 252:675–679
Du L, Yu F, Zhu X, Zaleta-Rivera K, Bojja RS, Ding Y, Yi H, Wang Q (2007) Biochemical and molecular analysis of the biosynthesis of fumonisins. In: Baerson SR (ed) Polyketides: biosynthesis, biological activities and genetic engineering, American Chemical Society, Washington, DC, pp 81–96
Eley KL, Halo LM, Song Z, Powles H, Cox RJ, Bailey AM, Lazarus CM, Simpson TJ (2007) Biosynthesis of the 2-pyridone tenellin in the insect pathogenic fungus Beauveria bassiana. Chembiochem 8:289–297
Feng B, Wang X, Hauser M, Kaufmann S, Jentsch S, Haase G, Becker JM, Szaniszlo PJ (2001) Molecular cloning and characterization of WdPKS1, a gene involved in dihydroxynaphthalene melanin biosynthesis and virulence in Wangiella (Exophiala) dermatitidis. Infect Immun 69:1781–1794
Ferreira GC, Vajapey U, Hafez O, Hunter GA, Barber MJ (1995) Aminolevulinate synthase: lysine 313 is not essential for binding the pyridoxal phosphate cofactor but is essential for catalysis. Protein Sci 4:1001–1006
Fischbach MA, Walsh CT (2006) Assembly-line enzymology for polyketide and nonribosomal Peptide antibiotics: logic, machinery, and mechanisms. Chem Rev 106:3468–3496
Fujii I, Mori Y, Watanabe A, Kubo Y, Tsuji G, Ebizuka Y (1999) Heterologous expression and product identification of Colletotrichum lagenarium polyketide synthase encoded by the PKS1 gene involved in melanin biosynthesis. Biosci Biotechnol Biochem 63:1445–1452
Fujii I, Ono Y, Tada H, Gomi K, Ebizuka Y, Sankawa U (1996) Cloning of the polyketide synthase gene atX from Aspergillus terreus and its identification as the 6-methylsalicylic acid synthase gene by heterologous expression. Mol Gen Genet 253:1–10
Fujii I, Watanabe A, Sankawa U, Ebizuka Y (2001) Identification of Claisen cyclase domain in fungal polyketide synthase WA, a naphthopyrone synthase of Aspergillus nidulans. Chem Biol 8:189–197
Gable K, Han G, Monaghan E, Bacikova D, Natarajan M, Williams R, Dunn TM (2002) Mutations in the yeast LCB1 and LCB2 genes, including those corresponding to the hereditary sensory neuropathy type I mutations, dominantly inactivate serine palmitoyltransferase. J Biol Chem 277:10194–10200
Gilchrist DG, Grogan RG (1976) Production and nature of a host-specific toxin from Alternaria alternata f. sp. lycopersici. Phytopathology 66:165–171
Hanada K (2003) Serine palmitoyltransferase, a key enzyme of sphingolipid metabolism. Biochim Biophys Acta 1632:16–30
Hoffmeister D, Keller NP (2007) Natural products of filamentous fungi: enzymes, genes, and their regulation. Nat Prod Rep 24:393–416
Hopwood DA (1997) Genetic contributions to understanding polyketide synthases. Chem Rev 97:2465–2498
Kealey JT, Liu L, Santi DV, Betlach MC, Barr PJ (1998) Production of a polyketide natural product in nonpolyketide-producing prokaryotic and eukaryotic hosts. Proc Natl Acad Sci USA 95:505–509
Kennedy J, Auclair K, Kendrew SG, Park C, Vederas JC, Hutchinson CR (1999) Modulation of polyketide synthase activity by accessory proteins during lovastatin biosynthesis. Science 284:1368–1372
Khosla C, Gokhale RS, Jacobsen JR, Cane DE (1999) Tolerance and specificity of polyketide synthases. Annu Rev Biochem 68:219–253
Khosla C, Tang Y, Chen AY, Schnarr NA, Cane DE (2007) Structure and mechanism of the 6-deoxyerythronolide B synthase. Annu Rev Biochem 76:195–221
Kittendorf JD, Sherman DH (2006) Developing tools for engineering hybrid polyketide synthetic pathways. Curr Opin Biotechnol 17:597–605
Kongsaeree P, Prabpai S, Sriubolmas N, Vongvein C, Wiyakrutta S (2003) Antimalarial dihydroisocoumarins produced by Geotrichum sp., an endophytic fungus of Crassocephalum crepidioides. J Nat Prod 66:709–711
Kono Y, Daly JM (1979) Characterization of the host-specific pathotoxin produced by Helminthosporium maydis race T affecting corn with Texas male sterile cytoplasm. Bioorg Chem 8:391–397
Kroken S, Glass NL, Taylor JW, Yoder OC, Turgeon BG (2003) Phylogenomic analysis of type I polyketide synthase genes in pathogenic and saprobic ascomycetes. Proc Natl Acad Sci USA 100:15670–15675
Lambalot RH, Gehring AM, Flugel RS, Zuber P, LaCelle M, Marahiel MA, Reid R, Khosla C, Walsh CT (1996) A new enzyme superfamily—the phosphopantetheinyl transferases. Chem Biol 3:923–936
Marasas WF, Riley RT, Hendricks KA, Stevens VL, Sadler TW, Gelineau-van Waes J, Missmer SA, Cabrera J, Torres O, Gelderblom WC, Allegood J, Martinez C, Maddox J, Miller JD, Starr L, Sullards MC, Roman AV, Voss KA, Wang E, Merrill AH Jr (2004) Fumonisins disrupt sphingolipid 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. J Nutr 134:711–716
Menzella HG, Reeves CD (2007) Combinatorial biosynthesis for drug development. Curr Opin Microbiol 10:238–245
Merrill AH Jr, van Echten G, Wang E, Sandhoff K (1993) Fumonisin B1 inhibits sphingosine (sphinganine) N-acyltransferase and de novo sphingolipid biosynthesis in cultured neurons in situ. J Biol Chem 268:27299–27306
Minto RE, Townsend CA (1997) Enzymology and molecular biology of aflatoxin biosynthesis. Chem Rev 97:2537–2556
Moore BS, Hertweck C (2002) Biosynthesis and attachment of novel bacterial polyketide synthase starter units. Nat Prod Rep 19:70–99
Moriguchi T, Ebizuka Y, Fujii I (2006) Analysis of subunit interactions in the iterative type I polyketide synthase ATX from Aspergillus terreus. Chembiochem 7:1869–1874
Nelson PE, Desjardins AE, Plattner RD (1993) Fumonisins, mycotoxins produced by Fusarium species: biology, chemistry, and significance. Annu Rev Phytopathol 31:233–252
Nesbitt BF, O’Kelly J, Sargeant K, Sheridan A (1962) Aspergillus flavus and turkey X disease. Toxic metabolites of Aspergillus flavus. Nature 195:1062–1063
Nicholson TP, Rudd BA, Dawson M, Lazarus CM, Simpson TJ, Cox RJ (2001) Design and utility of oligonucleotide gene probes for fungal polyketide synthases. Chem Biol 8:157–178
Plattner RD, Shackelford DD (1992) Biosynthesis of labeled fumonisins in liquid cultures of Fusarium moniliforme. Mycopathologia 117:17–22
Price WD, Lovell RA, McChesney DG (1993) Naturally occurring toxins in feedstuffs: Center for Veterinary Medicine Perspective. J Anim Sci 71:2556–2562
Proctor RH, Brown DW, Plattner RD, Desjardins AE (2003) Co-expression of 15 contiguous genes delineates a fumonisin biosynthetic gene cluster in Gibberella moniliformis. Fungal Genet Biol 38:237–249
Proctor RH, Desjardins AE, Plattner RD, Hohn TM (1999) A polyketide synthase gene required for biosynthesis of fumonisin mycotoxins in Gibberella fujikuroi mating population A. Fungal Genet Biol 27:100–112
Proctor RH, Plattner RD, Desjardins AE, Busman M, Butchko RA (2006) Fumonisin production in the maize pathogen Fusarium verticillioides: genetic basis of naturally occurring chemical variation. J Agric Food Chem 54:2424–2430
Richardson MT, Pohl NL, Kealey JT, Khosla C (1999) Tolerance and specificity of recombinant 6-methylsalicyclic acid synthase. Metab Eng 1:180–187
Riley RT, Wang E, Schroeder JJ, Smith ER, Plattner RD, Abbas H, Yoo HS, Merrill AH Jr (1996) Evidence for disruption of sphingolipid metabolism as a contributing factor in the toxicity and carcinogenicity of fumonisins. Nat Toxins 4:3–15
Seo JA, 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
Shen B (2000) Biosynthesis of aromatic polyketides. Top Curr Chem 209:1–51
Shiao MS, Don HS (1987) Biosynthesis of mevinolin, a hypocholesterolemic fungal metabolite, in Aspergillus terreus. Proc Natl Sci Counc Repub China B 11:223–231
Sims JW, Fillmore JP, Warner DD, Schmidt EW (2005) Equisetin biosynthesis in Fusarium heterosporum. Chem Commun (Camb) 2:186–188
Song Z, Cox RJ, Lazarus CM, Simpson TT (2004) Fusarin C biosynthesis in Fusarium moniliforme and Fusarium venenatum. Chembiochem 5:1196–1203
Spencer JB, Jordan PM (1992) Investigation of the mechanism and steric course of the reaction catalyzed by 6-methylsalicylic acid synthase from Penicillium patulum using (R)-[1-13C;2-2H]- and (S)-[1-13C;2-2H]malonates. Biochem 31:9107–9116
Staunton J, Weissman KJ (2001) Polyketide biosynthesis: a millennium review. Nat Prod Rep 18:380–416
Sweeney MJ, Dobson AD (1999) Molecular biology of mycotoxin biosynthesis. FEMS Microbiol Lett 175:149–163
Tang Y, Kim CY, Mathews II, Cane DE, Khosla C (2006) The 2.7-Angstrom crystal structure of a 194-kDa homodimeric fragment of the 6-deoxyerythronolide B synthase. Proc Natl Acad Sci USA 103:11124–11129
Tsai HF, Fujii I, Watanabe A, Wheeler MH, Chang YC, Yasuoka Y, Ebizuka Y, Kwon-Chung KJ (2001) Pentaketide melanin biosynthesis in Aspergillus fumigatus requires chain-length shortening of a heptaketide precursor. J Biol Chem 276:29292–29298
Walsh CT, Gehring AM, Weinreb PH, Quadri LE, Flugel RS (1997) Post-translational modification of polyketide and nonribosomal peptide synthases. Curr Opin Chem Biol 1:309–315
Wang Q, Wang J, Yu F, Zhu X, Zaleta-Rivera K, Du L (2006) Mycotoxin fumonisins: health impacts and biosynthetic mechanism. Prog Nat Sci 16:7–15
Wang W, Jones C, Ciacci-Zanella J, Holt T, Gilchrist DG, Dickman MB (1996) Fumonisins and Alternaria alternata lycopersici toxins: sphinganine analog mycotoxins induce apoptosis in monkey kidney cells. Proc Natl Acad Sci USA 93:3461–3465
Watanabe A, Ebizuka Y (2004) Unprecedented mechanism of chain length determination in fungal aromatic polyketide synthases. Chem Biol 11:1101–1106
Watanabe A, Fujii I, Tsai H, Chang YC, Kwon-Chung KJ, Ebizuka Y (2000) Aspergillus fumigatus alb1 encodes naphthopyrone synthase when expressed in Aspergillus oryzae. FEMS Microbiol Lett 192:39–44
Webster SP, Alexeev D, Campopiano DJ, Watt RM, Alexeeva M, Sawyer L, Baxter RL (2000) Mechanism of 8-amino-7-oxononanoate synthase: spectroscopic, kinetic, and crystallographic studies. Biochem 39:516–528
Winter CK, Gilchrist DG, Dickman MB, Jones C (1996) Chemistry and biological activity of AAL toxins. Adv Exp Med Biol 392:307–316
Wright JL, Vining LC, McInnes AG, Smith DG, Walter JA (1977) Use of 13C in biosynthetic studies. The labelling pattern in tenellin enriched from isotope-labelled acetate, methionine, and phenylalanine. Can J Biochem 55:678–685
Yang G, Rose MS, Turgeon BG, Yoder OC (1996) A polyketide synthase is required for fungal virulence and production of the polyketide T-toxin. Plant Cell 8:2139–2150
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
Yu F, Zhu X, Du L (2005) Developing a genetic system for functional manipulations of FUM1, a polyketide synthase gene for the biosynthesis of fumonisins in Fusarium verticillioides. FEMS Microbiol Lett 248:257–264
Zaleta-Rivera K, Xu C, Yu F, Butchko RA, Proctor RH, Hidalgo-Lara ME, Raza A, Dussault PH, Du L (2006) A bidomain nonribosomal peptide synthetase encoded by FUM14 catalyzes the formation of tricarballylic esters in the biosynthesis of fumonisins. Biochem 45:2561–2569
Zhu X, Yu F, Bojja RS, Zaleta-Rivera K, Du L (2006) Functional replacement of the ketosynthase domain of FUM1 for the biosynthesis of fumonisins, a group of fungal reduced polyketides. J Ind Microbiol Biotechnol 33:859–868
Zhu X, Yu F, Li XC, Du L (2007) Production of dihydroisocoumarins 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
Acknowledgments
This work was supported in part by NSF (MCB-0614916) and NSF China (No. 30428023). The research was performed in facilities renovated with support from NIH (RR015468-01).
Author information
Authors and Affiliations
Corresponding author
Rights and permissions
About this article
Cite this article
Du, L., Zhu, X., Gerber, R. et al. Biosynthesis of sphinganine-analog mycotoxins. J Ind Microbiol Biotechnol 35, 455–464 (2008). https://doi.org/10.1007/s10295-008-0316-y
Received:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1007/s10295-008-0316-y