Open access
Mycology
Research Article
20 December 2024

The palmitoyl-CoA ligase Fum16 is part of a Fusarium verticillioides fumonisin subcluster involved in self-protection

ABSTRACT

Fusarium verticillioides produces the mycotoxin fumonisin B1 (FB1), which disrupts sphingolipid biosynthesis by inhibiting ceramide synthase and affects the health of plants, animals, and humans. The means by which F. verticillioides protects itself from its own mycotoxin are not completely understood. Some fumonisin (FUM) cluster genes do not contribute to the biosynthesis of the compound, but their function has remained enigmatic. Recently, we showed that FUM17, FUM18, and FUM19 encode two ceramide synthases and an ATP-binding cassette transporter, respectively, which play a role in antagonizing the toxicity mediated by FB1. In the present work, we uncovered functions of two adjacent genes, FUM15 and FUM16. Using homologous and heterologous expression systems, in F. verticillioides and Saccharomyces cerevisiae, respectively, we provide evidence that both contribute to protection against FB1. Our data indicate a potential role for the P450 monooxygenase Fum15 in the modification and detoxification of FB1 since the deletion and overexpression of the respective gene affected extracellular FB1 levels in both hosts. Furthermore, relative quantification of ceramide intermediates and an in vitro enzyme assay revealed that Fum16 is a functional palmitoyl-CoA ligase. It co-localizes together with the ceramide synthase Fum18 to the endoplasmic reticulum, where they contribute to sphingolipid biosynthesis. Thereby, FUM15-19 constitute a subcluster within the FUM biosynthetic gene cluster dedicated to the fungal self-protection against FB1.

IMPORTANCE

The study identifies a five-gene FUM subcluster (FUM15-19) in Fusarium verticillioides involved in self-protection against FB1. FUM16 (palmitoyl-CoA ligase), FUM17, and FUM18 (ceramide synthases) enzymatically supplement ceramide biosynthesis, while FUM19 (ATP-binding cassette transporter) acts as a repressor of the FUM cluster. The evolutionary conservation of FUM15 (P450 monooxygenase) in Fusarium and Aspergillus FUM clusters is discussed, and its effect on extracellular FB1 levels in both native (F. verticillioides) and heterologous (Saccharomyces cerevisiae) hosts is highlighted. These findings enhance our understanding of mycotoxin self-protection mechanisms and could inform strategies for predicting biological activity of unknown secondary metabolites, managing mycotoxin contamination, and developing resistant crop cultivars.

INTRODUCTION

Fungal secondary metabolites (SMs) are diverse organic compounds that are not essential for growth or reproduction but provide ecological advantages to the organism. These compounds help deter predators, inhibit competitors, and facilitate symbiotic relationships, enhancing the organism’s survival, reproduction, and adaptability under diverse environmental conditions (14).
Among the different SMs produced by fungi, mycotoxins are a significant group with a profound impact on the health of all living organisms, that is, microbes, plants, animals, and humans. They can contaminate food and feed, being produced by phytopathogenic fungi, which can lead to severe health issues in humans, like cancer, liver damage, and immunosuppression (5). Fungi exhibit remarkable resilience against self-intoxication by their own mycotoxins, an essential trait for their survival and ecological success. Mechanisms underlying this resistance include (i) efficient efflux pumps that secrete toxins from cells (6); (ii) enzymatic detoxification pathways (7); (iii) duplicated or resistant target enzymes (8); and (iv) subcellular separation of the toxic product (9). The biosynthetic gene clusters (BGCs) responsible for the production of mycotoxins often carry adjacent non-biosynthetic genes that are relevant for conferring self-protection (10).
The sphingolipid biosynthesis inhibitors fumonisins are a group of mycotoxins produced primarily by Fusarium and Aspergillus species (11). They are polyketide-derived aminopentol compounds with a linear C20 backbone and two tricarballylic esters. The best studied are B-series fumonisins (FBs), FB1, FB2, FB3, and FB4, which are predominantly produced by the corn-infecting fungus Fusarium verticillioides. As demonstrated in mice experiments, the most toxic FB analog is FB1, which is the final biosynthetic product accumulating in greatest abundance in F. verticillioides (approximately 80%) (1214). The fumonisin BGC (FUM) includes 16 genes (Fig. 1A) (12), expressed under the control of the Zn(II)2Cys6-type transcription factor Fum21 (15).
FIG 1
Illustration depicts fumonisin biosynthetic gene cluster inluding the subcluster FUM15–FUM19 implicated in self-protection, and their suggested roles in ceramide biosynthesis, with Fum16 as palmitoyl-CoA ligase and Fum18 as ceramide synthase in the ER.
FIG 1 The fumonisin gene cluster and its effects on ceramide biosynthesis. (A) The FUM biosynthetic gene cluster in F. verticillioides. The genes encoding FB1 biosynthetic enzymes FUM1–FUM14 are colored gray; the transcription factor (TF) FUM21 is reported in white; the non-biosynthetic genes FUM15, FUM16, and FUM18 are colored green, cyan, and orange, respectively. The table summarizes the function of FUM non-biosynthetic genes. (B) The simplified de novo synthesis of fungal ceramides. Indicated are catalyzing enzymes fatty acid CoA ligase (CoL), serine palmitoyltransferase (SPT), 3-ketoreductase (KR), C4-hydroxylase (C4H), desaturase (DES), and ceramide synthase (CerS). Ceramide intermediates analyzed in this study by HPLC-HRMS are written in bold. The FUM non-biosynthetic enzymes supplementing the pathway are included: Fum16 (cyan) and Fum18 (orange).
FB1 biosynthesis begins with the formation of the polyketide backbone by the polyketide synthase Fum1 (16) (Fig. S1). Next, the aminotransferase Fum8 catalyzes the condensation of the polyketide with l-alanine (17). Further modifications are performed by Fum6 (18), Fum13 (19), and Fum2 (20). The tricarballylic acid side chains are synthesized by Fum7 and Fum10 (21), and are attached to the backbone by Fum14 (22). FB1 biosynthesis ends with the hydroxylation at the C-5 position by Fum3 (23) (Fig. S1).
FB1 disrupts sphingolipid metabolism by inhibiting ceramide synthase (CerS), a key enzyme in ceramide biosynthesis (Fig. 1B) (24). Because FB1 is structurally similar to sphinganine, a substrate in ceramide biosynthesis, it competitively inhibits the enzyme by irreversibly binding to the active site of CerS (25). The inhibition of CerS by sphinganine analog mycotoxins triggers apoptosis, which is caused by an accumulation of sphingoid bases, rather than reduced sphingolipid biosynthesis itself (26). Targeted application of sphingolipid biosynthesis inhibitors, however, has promising potential as treatment against severe human diseases, such as cancer, schizophrenia, Alzheimer’s, multiple sclerosis, and diabetes (2729).
Ceramides constitute the backbone of complex sphingolipids. In filamentous fungi, ceramide biosynthesis begins with the condensation of l-serine with palmitoyl-CoA by a serine palmitoyltransferase to form 3-ketosphinganine, which is further reduced by a ketoreductase to sphinganine. At this point, ceramide biosynthesis branches (30). Sphinganine is converted by CerS to dihydroceramide, and subsequently to ceramide by a desaturase. Alternatively, sphinganine is converted to phytosphingosine by a C4-hydroxylase and to phytoceramide by CerS (Fig. 1B). Ceramides are vital components of cell membranes and signaling molecules involved in apoptosis, cell differentiation, and proliferation (31). This disruption thus contributes to various diseases in humans and animals (3234).
The biosynthesis of sphingolipids is compartmentalized. The first enzymatic reactions, leading up to ceramides, are catalyzed in the endoplasmic reticulum (ER), while complex sphingolipids are further synthesized in ER-derived vesicles and the Golgi (35, 36). Recently, Fum3 (the enzyme catalyzing the last reaction in FB1 biosynthesis) was shown to be localized in the cytosol in F. verticillioides (8, 37). This finding suggests a subcellular separation of the toxin FB1 from its target CerS in the ER, contributing to self-protection.
The FUM cluster in F. verticillioides entails five non-biosynthetic genes adjacent to one another (Fig. 1A), FUM15 to FUM19. Fum19 is an ATP-binding cassette (ABC) transporter that acts as a repressor of the FUM gene cluster, regulating the levels of intracellular and secreted FB1 (8). Fum17 and Fum18 are two of five CerS homologs in F. verticillioides, both co-localizing with ceramide biosynthesis in the ER. In particular, Fum18 was shown to confer resistance to FB1 self-toxicity (8). While FB1 has antifungal activity, specifically against FB nonproducers, F. verticillioides is more resistant to externally added FB1 (38).
Despite all the functional analyses of FUM cluster genes for over two decades, the function of FUM15 and FUM16 has remained unclear so far. Also, the direct effect of FUM15-FUM18 on ceramide biosynthesis has not yet been elucidated. Fum15 is a P450 monooxygenase, while Fum16 displays structural similarities with long-fatty-acid-CoA ligases (12). Our hypothesis was that they, too, may play a role in self-protection.
In this work, we uncovered the involvement of Fum15 and Fum16 in self-protection against FB1 in F. verticillioides, and additionally, in the budding yeast Saccharomyces cerevisiae as a heterologous host. Using an in vitro enzyme assay, we showed that Fum16 is a functional palmitoyl-CoA ligase, while the presence of the P450 monooxygenase Fum15 reduces extracellular levels of FB1.

RESULTS

Fum15 and Fum16 are co-compartmentalized with ceramide biosynthesis

We started by performing localization studies of Fum15 and Fum16 by confocal microscopy. Both proteins were C-terminally tagged with a green fluorescent protein (GFP) and expressed from the same constitutive overexpression promoter (Aspergillus nidulans PoliC). In our previous work, we showed that the CerS Fum18 localizes to the ER and ER-derived vesicles (8). We thus used our mutant producing Fum18 C-terminally tagged with red fluorescent protein (DsRed) as a recipient strain. We made sure that the fluorescent proteins did not give signals when excited with inappropriate wavelengths (Fig. S2). Fum15-GFP and Fum16-GFP both showed co-localization with Fum18-DsRed in the perinuclear ER (Fig. 2).
FIG 2
Microscopic images depict FUM18::DsRed and FUM15::GFP as well as FUM18::DsRed and FUM16::GFP co-localization in hyphal compartments. Arrows highlight overlapping signals in GFP and DsRed channels in the perinuclear ER and ER-derived vesicles.
FIG 2 Co-localization of Fum15-GFP and Fum16-GFP with Fum18-DsRed. Shown are separate channels: brightfield (BF), GFP, DsRed, and the overlay. White arrows point to the perinuclear ER. Conidia were inoculated in ICI/6 mM Gln and grown as a standing culture overnight.

Establishment of an inducible FUM21 overexpression strain

Although high enough concentrations of FB1 have been reported to have a negative impact on fungal growth, the F. verticillioides M-3125 wild type (WT) displays resistance to the toxin (8). For this reason, we designed a strain for inducible overproduction of FB1, by overexpressing the fumonisin BGC transcription factor gene FUM21. This allowed us to investigate the toxic effects on growth, ceramide biosynthesis, as well as the hypothesized contribution of FUM15 and FUM16 in self-protection. We employed the tetON promoter system, which was successfully used in other fungal species such as Aspergillus niger (39) and Fusarium fujikuroi (40), for inducible, controllable production of metabolites by the addition of the inducer molecule doxycycline hyclate (doxy). The native promoter of the transcription factor gene FUM21 was thus exchanged with the tetON sequence.
In order to choose the most suitable doxy concentration to use in further assays, we first performed a growth inhibition assay to determine the effect of doxy concentration on the WT and overproduction strain tetON::FUM21 (Fig. 3A through C). Subsequently, we relatively quantified FB1 in the supernatant and extracted mycelium of liquid cultures with and without induction (Fig. 3D). The addition of the inducer appeared to have a toxic effect by itself as even the WT growth was affected by high concentrations of doxy. A maximum of 50 µg/mL doxy was used, which impaired WT growth by 4.8% on plate and 3.3% in liquid culture. In contrast to that, tetON::FUM21 was inhibited by the addition of 50 µg/mL doxy by more than 20%, both on plate and in liquid culture (Fig. 3B and C). We hypothesized that this more severe phenotype of tetON::FUM21 was predominantly caused by an increased FB1 production. Indeed, relative quantification of FB1 via high-performance liquid chromatography coupled to high-resolution mass spectrometry (HPLC-HRMS) revealed that the addition of 50 µg/mL doxy resulted in a drastic overproduction of FB1 in tetON::FUM21, compared with the WT, both in the supernatant and mycelium (Fig. 3D). The identity of the chromatographic peak corresponding to FB1 was verified by comparison with an FB1 reference compound (Fig. S3A). Absolute quantification based on an FB1 standard curve (Fig. S3B) revealed an accumulation of 6.94 ± 0.03 µg/mL in the supernatant of the WT and of 671.74 ± 54.58 µg/mL in the supernatant of tetON::FUM21. In the absence of doxy, FB1 levels in the overproduction strain were negligible, and no expression was detected of either FUM21 or the biosynthetic gene FUM8 (Fig. S4A), suggesting limited leakiness of the promoter, so that this strain was suitable for our analysis (Fig. 3D).
FIG 3
Comparison of FvWT and tetON::FUM21 fungal strains under increasing doxycycline concentrations. Plates and bar graphs depict growth inhibition rate, dry weight reduction, and FB1 production in supernatant and mycelium under doxycycline induction.
FIG 3 Analysis of FB1 production and phenotype of F. verticillioides tetON::FUM21. (A) Growth assay performed on small complete medium plates and increasing concentration of doxycycline (doxy). The increased concentration of doxy results in the augmented expression of the TF FUM21 and consequent increase of the FUM cluster activation. (B) The average diameter of the colonies was measured after 5 days with ±SD (n = 3). The growth inhibition rate (IR) is relative to the noninduced condition for each strain. (C) Biomass measurements of lyophilized mycelia, after 5 days of growth after induction in liquid culture (ICI/6 mM Gln). Dry weight of the WT at 0 µg/mL doxy was arbitrarily set to 100%. (D) Relative FB1 production of the WT and tetON::FUM21, after 5 days of growth after induction in liquid culture (ICI/6 mM Gln). Shown are mean values with ±SD (n = 3). FB1 levels were normalized against an internal standard and related to the dry weight of the cultures. Production of the WT at 0 µg/mL doxy was arbitrarily set to 100%. Student’s t-test was used to assess statistical significance as indicated (**P < 0.01; ***P < 0.001).

Fum15 and Fum16 are involved in self-protection against FB1

In order to further investigate the function and significance of FUM15 and FUM16, single-deletion mutants in the overexpression strain were generated (tetON::FUM21fum15 and tetON::FUM21fum16). As a negative control, tetON::FUM21fum8 was also generated. The biosynthetic enzyme Fum8 is an aminotransferase that performs the second catalytic step in fumonisin biosynthesis; it catalyzes the condensation of l-alanine with the octadecanoic acid precursor. Thereby, deletion of FUM8 abolishes fumonisin production in our overexpression strain (see FB1 quantification below). Furthermore, the CerS mutant tetON::FUM21fum18 was generated as a positive control based on previously published results (8), but also to expand on the self-defense role of FUM18 as a CerS homolog. Finally, a strain was also generated, in which the whole non-biosynthetic subcluster, FUM15 to FUM19, was deleted (tetON::FUM21fum15-19).
These deletion mutants were tested both in plate assays and in liquid cultures, under uninduced and induced conditions, to evaluate their growth phenotype (Fig. 4A and B). A concentration of 50 µg/mL doxy was used in this study for FB1 overproduction in liquid cultures, while 25 µg/mL was instead used in plate assays as we observed that our strains were more susceptible to FB1 on solid medium (Fig. S5). Overproduction of FB1 resulted in irregularly shaped hyphal growth at the colony periphery of tetON::FUM21 mutants, with lack of aerial hyphae, compared with the uninduced strains (Fig. 4A). Doxy induction caused statistically significant mycelial growth inhibition after 6 days on solid medium and after 5 days in induced liquid cultures compared with the equally treated WT, with the exception of the FB1-nonproducer tetON::FUM21fum8. All three non-biosynthetic single-gene deletions (Δfum15, Δfum16, and Δfum18 deletion in the tetON::FUM21 background) caused a higher growth inhibition rate compared with tetON::FUM21 alone (Fig. 4B). Among those, deletion of FUM18 caused the most severe effect among the three, with a growth inhibition rate of 28% on solid medium and 38% in liquid culture, while deletion of FUM16 caused the least severe effect (15% and 21%, respectively) (Fig. 4B). The complete subcluster deletion of FUM15-19 did not result in an additional growth inhibition compared with tetON::FUM21fum18 (Fig. S5).
FIG 4
Colony growth of FvWT and tetON::FUM21 mutants with and without doxycycline (25 µg/mL). Growth inhibition rates on plates and in liquid culture. OD600 of S. cerevisiae strains expressing FUM15, FUM16, and FUM18 with 0 or 500 µg/mL fumonisin B1.
FIG 4 Growth phenotype analysis of the deletion mutants in F. verticillioides and of S. cerevisiae expressing FUM15-18. (A) Growth inhibition of F. verticillioides WT, tetON::FUM21 and deletion strains of the latter. The plate assay was performed on complete medium plates under uninduced and induced (50 µg/mL doxy) conditions. (B) The growth inhibition rate (IR) is relative to the noninduced condition for each strain. The average diameter of the colonies was measured after 6 days. Biomass of lyophilized mycelia was quantified after 5 days of growth after induction in liquid culture (ICI/6 mM Gln). Data are mean values ± SD (n = 3). (C) Growth curves of S. cerevisiae expressing FUM15, FUM16, or FUM18 over 20 h in SD-Ura, either in the absence or presence of 500 µg/mL FB1 (mean ± SD, n = 3). The blank was subtracted from each data point, and the curves were adjusted to an initial OD600 of 0.1. Statistical analysis was performed on the OD value at 20 h. Student’s t-test was used to assess statistical significance as indicated (*P < 0.05; **P < 0.01; ***P < 0.001).
Expression analysis of the F. verticillioides WT, tetON::FUM21 and deletion mutants thereof revealed that single deletion of FUM15, FUM16, or FUM18 did not result in a compensatory upregulation of the adjacent genes (Fig. S4A). Deletion of FUM8 in the tetON::FUM21 background, abolishing FB1 biosynthesis, failed to fully induce expression of FUM18 and FUM19, which are driven by a bidirectional promoter (Fig. 1A). We previously reported that FB1 feeding triggered FUM18 and FUM19 expression (8), and now additionally report an induction of FUM15 and FUM16 in the WT by 10 µg/mL exogenously added FB1 (Fig. S4B). We noted that the expression of FUM15 was not abolished in tetON::FUM21 in the absence of the inducer doxy, suggesting that FUM15 is not fully under the control of the cluster-specific transcription factor (Fig. S4A). This was corroborated by analysis of the Δfum21 deletion mutant (8), which showed ca. 20% of the FUM15 expression detected in the WT background (Fig. S4B). These expression data are in line with the idea that Fum15 and Fum16, in addition to Fum18, could play a protective role.
To confirm the direct involvement of Fum15 and Fum16 in self-protection against FB1, we designed a heterologous expression experiment with S. cerevisiae. S. cerevisiae strains were engineered to constitutively express FUM15 (Ptef1::FUM15), FUM16 (Ptef1::FUM16), or FUM18 (Ptef1::FUM18) (8). We incubated these strains in the absence or presence of 500 µg/mL FB1 and monitored the optical density at 600 nm (OD600) over a period of 20 h (Fig. 4C). The incubation with FB1 impaired S. cerevisiae WT growth by 47%. Expression of FUM16 and FUM18 resulted in a partial rescue as growth was impaired by only 36% and 23%, respectively. Surprisingly, S. cerevisiae expressing FUM15 displayed almost no growth inhibition compared with the control condition and could reach higher OD600 compared with the other mutants in the presence of the toxin (Fig. 4C). Expression analysis showed that all three genes were expressed in a similar range (ca. 2.5- to 3.5-fold relative to the housekeeping gene actin), excluding aberrant expression as a reason for the lower performance of FUM16 and FUM18 in the heterologous system (Fig. S4C).
Taken together, these results demonstrate that the enzymes Fum15 and Fum16 by themselves play a role in self-protection against FB1, both in homologous and heterologous hosts.

Deletion of either FUM15, FUM16, or FUM18 affects the abundance of ceramide intermediates

Fum18 was already demonstrated to contribute to resistance against FB1 by functioning as a CerS on its own and thus providing supplementary CerS activity to the ceramide biosynthetic pathway (8). We asked whether Fum15 (a predicted P450 monooxygenase) or Fum16 (a predicted long-fatty-acid-CoA ligase) could also confer self-protection against the toxin similarly, supplementing the sphingolipid biosynthetic pathway. We performed an HPLC-HRMS analysis of the extracted mycelium of tetON::FUM21, as well as Δfum8, Δfum15, Δfum16, and Δfum18 deletions in the tetON::FUM21 background, grown in doxy-induced liquid cultures. We measured the abundance of 3-ketosphinganine, sphinganine, and phytosphingosine (Fig. 5A). Overproduction of FB1 by tetON::FUM21 blocked ceramide biosynthesis and caused a statistically significant increase in the precursors 3-ketosphinganine (3-fold), sphinganine (2.7-fold), and phytosphingosine (2.5-fold) compared with the induced WT (Fig. 1B). The increase was even more severe when either FUM15 or FUM18 were deleted, for all three intermediates (Fig. 5A), apparently causing a more severe inhibition of ceramide biosynthesis. These results confirmed the role of Fum18 as a CerS (8) but also provided new insight into the biosynthetic steps that are supplemented by the enzyme, which is thus acting on both of its direct substrates sphinganine and phytosphingosine (Fig. 1B). FUM16 deletion resulted in a ca. 40% decrease of sphinganine and phytosphingosine (Fig. 5A), indicating that Fum16 may have a catalytic role upstream of ceramide biosynthesis.
FIG 5
Relative abundance of 3-ketosphinganine, sphinganine, and phytosphingosine in F. verticillioides mutants under 50 µg/mL doxycycline and S. cerevisiae expressing FUM15, FUM16, and FUM18 with or without 500 µg/mL fumonisin B1.
FIG 5 Analysis of the production of ceramide intermediates. (A) Relative abundance of three ceramide intermediates, 3-ketosphinganine, sphinganine, and phytosphingosine, extracted from the mycelium of the F. verticillioides WT, tetON::FUM21 and the deletion mutants Δfum8, Δfum15, Δfum16, Δfum18 in the tetON::FUM21 background. All cultures were induced with 50 µg/mL doxy for 5 days in liquid culture (ICI/6 mM Gln), and the production of tetON::FUM21 was set to 100% (mean ± SD, n = 3). (B) Relative abundance of the abovementioned intermediates, extracted from the cell pellet of the S. cerevisiae WT (carrying the empty plasmid pYES::Ptef1), Ptef1::FUM15, FUM16, and FUM18 grown in SD-Ura. The abundance in the nontreated WT was set to 100%, and statistical analysis was performed within each condition (0 and 500 µg/mL FB1) if not specified otherwise (horizontal line between conditions). Shown are mean values ± SD (n = 3). Student’s t-test was used to assess statistical significance (**P < 0.01; ***P < 0.001).

Fum16 is directly involved in ceramide intermediate biosynthesis, while Fum15 is not

In order to understand if Fum15 and Fum16 affect ceramide intermediates directly, we analyzed 3-ketosphinganine, sphinganine, and phytosphingosine abundance in S. cerevisiae strains expressing Ptef1::FUM15, Ptef1::FUM16, and Ptef1::FUM18 in the absence and presence of 500 µg/mL FB1 (Fig. 5B). In the absence of the toxin, Fum15 was the only enzyme that did not lead to a change in ceramide intermediate abundance compared with the S. cerevisiae WT. In contrast, Fum16 activity by itself was responsible for a decrease in 3-ketosphinganine of 47%, which was the earliest tested intermediate (Fig. 1B and 5B). As expected, Fum18 caused a decrease in its direct precursors sphinganine and phytosphingosine of 54% and 18%, respectively (Fig. 5B). The ratio of sphinganine-to-phytosphingosine in Ptef1::FUM18 can be explained by sphinganine being both a substrate for dihydroceramide synthesis and a precursor of phytosphingosine, which in turn is a substrate for phytoceramide synthesis (Fig. 1B). When exposed to the toxin, both FUM15- and FUM18-expressing strains displayed a drastic decrease in all three intermediates compared with the WT (Fig. 5B), a phenotype that was opposite of the one observed for the F. verticillioides deletion mutants tetON::FUM21fum15 and tetON::FUM21fum18 (Fig. 5A).
From these results, it appears that although Fum15 has an effect on the equilibrium of ceramide biosynthesis, and counteracts CerS inhibition by FB1, it does not do so by directly supplementing the pathway. On the contrary, evidence was gained that Fum16—and Fum18, as expected—has a direct impact on ceramide biosynthesis.

Fum15 affects FB1 levels in F. verticillioides, thus alleviating self-toxicity

In the previous experiments, we observed that Fum15 performs a function that is beneficial for survival when exposed to FB1, both in F. verticillioides and S. cerevisiae (Fig. 4). We also discovered that the catalytic activity of Fum15 has an indirect effect on ceramide biosynthesis (Fig. 5). We thus raised the question whether Fum15 could possess an activity toward fumonisin itself. We analyzed the phenotype of the F. verticillioides single-deletion mutants in regard to FB1 production under inducing conditions and in comparison to the genetic background of tetON::FUM21 (Fig. 6A). As a control, we also performed the analysis on the S. cerevisiae overexpression strains treated with FB1 (Fig. 6B). FB1 production was quantified in the supernatant and mycelium of F. verticillioides liquid cultures by HPLC-HRMS. Deletion of either FUM16 or FUM18 in the tetON::FUM21 background had no impact on FB1 production (Fig. 6A). This was in accordance with previous research (8, 41). However, deletion of FUM15 resulted in a statistically significant increase of the toxin by 50% in the supernatant, compared with the background strain tetON::FUM21, but no change was detected in the mycelium (Fig. 6A). When analyzed in S. cerevisiae, the expression of FUM16 or FUM18 did not lead to a change in FB1 abundance in the supernatant. Expression of FUM15, however, led to a decrease of 60% compared with the treated S. cerevisiae WT (Fig. 6B). These results are in accordance with the change in all three quantified ceramide intermediates described above. The increase and decrease of FB1 in the used F. verticillioides and S. cerevisiae FUM15 strains, respectively, explain the stronger and weaker inhibition of ceramide biosynthesis in these organisms, and explain why they show the same trend as the CerS-encoding FUM18 deletion and overexpression strains (Fig. 5). Our data point toward the fact that the P450 monooxygenase Fum15 could chemically modify FB1 and thereby detoxify it. No new peak(s) could be detected under our cultivation conditions, neither in S. cerevisiae nor in F. verticillioides. An explanation in agreement with these observations is that the resulting metabolite could be unstable and quickly degraded.
FIG 6
Fumonisin B1 production in F. verticillioides mutants treated with 50 µg/mL doxycycline and FB1 accumulation in S. cerevisiae strains expressing FUM15, FUM16, or FUM18 under 500 µg/mL FB1 treatment.
FIG 6 Analysis of the production of FB1. (A) Relative abundance of FB1 in the supernatant and extracted mycelium of the F. verticillioides WT, tetON::FUM21, and the deletion mutants Δfum8, Δfum15, Δfum16, Δfum18 in the tetON::FUM21 background. All cultures were induced with 50 µg/mL doxy for 5 days in liquid culture (ICI/6 mM Gln), and the production of tetON::FUM21 was set to 100% (mean ± SD, n = 3). (B) Relative abundance of FB1 in the supernatant of the S. cerevisiae WT (carrying the empty plasmid pYES::Ptef1), Ptef1::FUM15, FUM16, and FUM18, grown in SD-Ura and treated with 500 µg/mL FB1. Abundance in the WT was set to 100%. Shown are mean values ± SD (n = 3). Student’s t-test was used to assess statistical significance compared with the control or as indicated (**P < 0.01; ***P < 0.001).

Fum16 is a functional palmitoyl-CoA ligase

Fum16 is predicted to be a long-fatty-acid-CoA ligase, based on the highest BLASTX score for its open-reading frame (12). In our investigation, we further pinpointed Fum16 structure and function using the web-based tool Protein Homology/analogY Recognition Engine V 2.0 (Phyre2) (42). A total of 601 residues (89% coverage) of Fum16 were modeled with a 100% confidence against several fatty-acyl-CoA ligases, spanning several domains of life: from the plant 4-coumaric-acid-CoA ligase from Populus tomentosa (Pt4CL) (43) to the bacterial fatty-acyl-AMP ligase from Mycobacterium tuberculosis (FadD23) (44). As a verification, we aligned the Phyre2 prediction with a Fum16 structure obtained via the artificial intelligence program AlphaFold2 (Fum16-AF2) (45). The alignment yielded a root mean square deviation (RMSD) of 1.840, which proved the similarity of both models (Fig. S6A and B). The alignment of Fum16-AF2 with FadD23, complexed with palmitic acid and an ATP analog, revealed an impressive residue conservation at the active site in regard to both amino acid composition and hydrogen bond interactions with the substrates (Fig. 7A). Furthermore, Fum16-AF2 also appeared to possess an N-terminal hydrophobic pocket suitable for accommodating palmitic acid or an hexadecanoyl adenylate intermediate (Fig. 7B).
FIG 7
Structure of Fum16 (AlphaFold2) bound to AMP-PNP and palmitic acid, substrate-binding pocket comparison, reaction scheme for 3-ketosphinganine formation via palmitoyl-CoA, and chromatograms depicting enzymatic activity of Fum16 in combination with SPT.
FIG 7 In vitro activity assay of Fum16 for fatty-acid-CoA ligase activity. (A and B) Stereo representations of two FadD23 complexes (orange), aligned with the active site of the Fum16 structure predicted by AlphaFold2 (cyan). (A) Hydrogen bonds (black dotted lines) in the active site of the AMP-PNP-FadD23 complex. AMP-PNP and palmitic acid are shown in gray. (B) The hydrophobic AMP-C16-FadD23 complex. (C) Graphical representation of the reaction catalyzed by Fum16, namely the ligation of CoA with palmitic acid. (D) Shown are the total ion chromatograms of the HPLC-HRMS measurements of the assay. When Fum16 was present in the reaction, a peak corresponding to 3-ketosphinganine (1) appeared (cyan chromatogram). The negative control contained only the SPT enzyme from S. paucimobilis (white chromatogram), while the positive control contained SPT with the bacterial CoA ligase FadD (orange chromatogram).
Based on our predictions, we next aimed to prove Fum16’s activity in ceramide intermediate biosynthesis, which was expected to be the ligation of coenzyme A (CoA) with palmitic acid to yield palmitoyl-CoA. We thus established an in vitro assay. Fum16 was rendered soluble for bacterial expression and purification by truncating its predicted N-terminal transmembrane domain and N-terminally fusing it to a maltose-binding protein (Fig. S6C through F). Palmitic acid was supplied in the reaction and Fum16 was indeed able to synthesize palmitoyl-CoA, which in turn could be condensed with serine by a serine palmitoyl transferase (SPT, from Sphingomonas paucimobilis) to yield the final product 3-ketosphinganine. The consumption of palmitic acid and the production of 3-ketosphinganine were detected by HPLC-HRMS (Fig. 7C and D). A purified fatty-acid-CoA ligase from Escherichia coli (FadD) was used as a positive control. The negative control only included SPT with the substrates (Fig. 7D).
Thus, we demonstrated the function of Fum16 as a palmitoyl-CoA ligase. This finding is in accordance with what was described above for the ceramide intermediate analysis (Fig. 5) and unequivocally provides proof that Fum16 contributes to self-protection against FB1 as functional palmitoyl-CoA ligase (Fig. 1B).

DISCUSSION

Mycotoxin-producing organisms evolved several strategies to overcome self-toxicity, like the duplication of target enzymes, efficient toxin secretion mechanisms, enzymatic detoxification, and pathway compartmentalization (69). In our previous work, we identified the function of three genes in the FUM cluster of F. verticillioides that are located in a subcluster relevant for self-protection, FUM17, FUM18, and FUM19 (8). In the present study, we expanded on this knowledge by postulating the role of two additional genes, FUM15 (P450 monooxygenase) and FUM16 (palmitoyl-CoA ligase), and discovered that they, too, carry out self-protective functions (Fig. 1B). We are therefore presenting evidence for the existence of a five-gene subcluster FUM15-19 specifically dedicated to self-protection against the sphingolipid biosynthesis inhibitor FB1 in F. verticillioides.
The production of fungal SMs is tightly controlled not only on a temporal level but also spatially (46). In our previous work, we showed that the CerS homologs Fum17 and Fum18 are compartmentalized and co-localize with ceramide biosynthesis in the ER, while the final biosynthetic step to FB1 by Fum3 is performed in the cytosol (8). Here, we demonstrated an equivalent localization for Fum15 and Fum16 in the ER. While certain P450 monooxygenases have been observed in various cellular compartments, they are typically located in the ER (47) where they associate with NADPH-cytochrome P450 reductases for electron supply (48). In contrast to that, long-fatty-acid-CoA ligases are predominantly found in peroxisomes (49). The localization of Fum16 to the ER thus offered an initial indication of its role in ceramide biosynthesis.
In this work, the involvement of FUM15 and FUM16 in protection against FB1 was demonstrated both in the native host F. verticillioides and in the heterologous host S. cerevisiae. Deletions of either FUM15, FUM16, or FUM18 in an FB1-overproducing strain of F. verticillioides resulted in growth inhibition, both on solid medium and in liquid culture. On the opposite, constitutive heterologous expression of these genes in S. cerevisiae conferred protection against FB1. FUM18 appears to be the non-biosynthetic gene conferring the highest degree of protection in the F. verticillioides experimental setup. This was confirmed by observing no difference in the growth inhibition phenotype both when deleting FUM18 by itself and in combination with the other non-biosynthetic genes FUM15-19. In contrast, in the S. cerevisiae system, it was FUM15 expression that resulted in the highest survival, with a growth phenotype almost comparable to the untreated S. cerevisiae WT. While all three genes were expressed to similar strength from the overexpression promoter, we cannot exclude differences in the resulting protein levels as we did not perform codon optimization. Alternatively, it might be due to variations in enzyme activities between the different host organisms.
Subsequently, we observed that the deletion of either FUM15, FUM16, or FUM18 affects ceramide homeostasis in F. verticillioides. In S. cerevisiae, in the absence of FB1, the expression of FUM15 did not alter the levels of ceramide intermediates, which suggests that it does not have a direct enzymatic function in ceramide biosynthesis. Conversely, FUM16 expression in untreated S. cerevisiae cultures was sufficient to influence ceramide intermediate biosynthesis. There was no effect of FUM16 expression on FB1 levels in S. cerevisiae, which was consistent with findings for the FUM16 deletion mutant of F. verticillioides (this study; 41), and supported the hypothesis that it plays a direct role in supplementing the ceramide pathway. This could be unequivocally verified by an in vitro enzymatic assay, where we demonstrated that Fum16 is a functional palmitoyl-CoA ligase (Fig. 1B). Recently, we described a similar case of enzymatic supplementation of sphingolipid biosynthesis serving as a self-protection mechanism in Aspergillus. We explored the interactions between sphingolipid metabolism and sphingofungin biosynthesis in Aspergillus fumigatus, with the latter being a sphingolipid biosynthesis inhibitor acting on SPT (37). We demonstrated that the biosynthetic enzyme SphA (aminotransferase) plays a dual role in both sphingofungin and sphingolipid biosynthesis as it performs SPT activity in the presence of the serine and palmitoyl-CoA precursors (50).
From an evolutionary perspective, FUM15 seems to play an important role. The fumonisin BGC exhibits significant variation among black aspergilli (51), which are reported to primarily produce the FB analogs FB2 and FB4 (precursor of FB2), but not FB1 (52). Among the non-biosynthetic genes identified for self-protection in Fusarium (FUM15-19), only FUM15 is present in the FUM cluster of both fumonisin-producing and nonproducing strains of A. niger (53). Furthermore, in nonproducing isolates of Aspergillus luchuensis, Aspergillus brasiliensis, and Aspergillus tubingensis, the partial FUM cluster exclusively retained FUM1 and FUM15 (51). Our expression data revealed that FUM15 was not under the tight control of the transcription factor Fum21, similar to what was reported for A. niger previously (54).
Our data do not provide evidence for a direct role of Fum15 in ceramide biosynthesis. Instead, we suggest that it chemically modifies and thereby detoxifies FB1 due to the fact that FUM15 deletion in F. verticillioides resulted in increased extracellular FB1 levels, while the opposite was observed for the heterologous expression of FUM15 in FB1-supplemented S. cerevisiae. P450 monooxygenases have been largely reported to be involved in the detoxification of mycotoxins through biotransformation and chemical modification (55). In F. fujikuroi, P450 monooxygenase(s) were shown to be involved in the detoxification of fusaric acid (56) and gibepyrone A (40). Other examples are the hydroxylation and epoxidation of aflatoxin B1 (7, 55), as well as the hydroxylation of deoxynivalenol (7). It is apparent that other cluster-independent mechanisms cooperate to confer protection against self-toxicity by FB1 in F. verticillioides since the deletion of the complete subcluster FUM15-19 still displayed viability. These are likely cluster-independent ABC transporters compensating for the loss of FUM19 (8). Additionally, a gene directly adjacent to the FUM cluster, FvZBD1, is postulated to act as a genetic repressor of fumonisin production (57).
Based on the results of this study, we conclude that the non-biosynthetic genes, FUM15-19, in the F. verticillioides FUM cluster compose a subcluster of contiguous genes, which are able to confer self-resistance against the toxic effects of FB1. FUM16, FUM17, and FUM18 do so by supplementing ceramide biosynthesis as palmitoyl-CoA ligase and CerSs, respectively (this study; 8)). FUM19 encodes an ABC transporter that acts as a repressor of the FUM gene cluster (8), while FUM15 is possibly involved in chemical detoxification of FB1. Addressing mycotoxin contamination is crucial for maintaining a safe and sustainable food supply, highlighting the importance of continued research and monitoring (1). In particular, future identification of these self-protection subclusters could uncover mycotoxins with currently unknown biological functions and potentially assist in developing plant cultivars with enhanced resistance.

MATERIALS AND METHODS

General molecular methods

For amplification of desired DNA fragments from template DNA, the Phusion Flash High-Fidelity PCR Master Mix (Thermo Fisher Scientific, Dreieich, Germany) was used. All primers used for PCR amplification, diagnostic PCR, and Southern blots are listed in Table S1. For DNA enzymatic digestions, restriction enzymes by New England Biolabs (Frankfurt am Main, Germany) were used. DNA extraction from agarose gel was performed using the GeneJET Gel Extraction Kit (Thermo Fisher Scientific, Dreieich, Germany).
Vector assembly was achieved using either S. cerevisiae transformation-associated recombination (TAR) (58) or Gibson assembly (59) and checked with restriction digestion and sequencing. Plasmid DNA purification was performed using the NucleoSpin Plasmid Mini Kit (Macherey-Nagel, Düren, Germany).

Plasmid construction

Promoter exchange of F. verticillioides FUM21 with the inducible tetON promoter (Fig. S7) started out by amplifying upstream and (intragenic) downstream sequences (relative to the start codon) with the primer pairs fum21_5F/nat1_fum21_5R and TETon_FUM21_3F/TetON_FUM21_3R2, respectively. tetON (39) was amplified from pYES2-ptrA-TetON (37) with nat1_tetON_F2/gpda_for_vv. natR was amplified from pZPnat1 (GenBank accession no. AY631958.1) with hph_trpC_F/nat1_R. The obtained fragments were cloned into the BamHI/HindIII-digested shuttle vector pYES2 (Life Technologies, Darmstadt, Germany) using Gibson assembly.
Deletion vectors harbored ~1 kb upstream and downstream flanks of the gene of interest, as well as a deletion cassette. Flanks were amplified using primer pairs 5F/5R and 3F/3R (Table S1), as well as F. verticillioides genomic DNA. FUM8, FUM15, FUM16, FUM18, and FUM15-19 were deleted by exchange with the hygromycin B resistance gene under the control of A. nidulans PgpdA (Fig. S7). For that purpose, hphR was amplified from pUC-hph (60) with gpda_for_vv/Hph_Rev_VV2. The complete vectors were assembled under the use of a linearized (HindIII/XbaI) pYES2, applying TAR cloning.
Microscopy vectors were obtained by fusing the genes of interest to the constitutive A. nidulans promoter PoliC and C-terminal fusion to GFP (Fig. S8). To this end, the insert was cloned into NcoI-digested pNDH-OGG (61), applying TAR cloning.
S. cerevisiae expression vectors (Fig. S9) were constructed by amplifying the inserts from F. verticillioides cDNA. The inserts were then cloned into the amplified (primer pair TEF_Rv/pYes2_cyc1T_Fw) backbone pYES2::Ptef1 (62) using Gibson assembly.
Finally, the bacterial overexpression vector was obtained by amplifying an N-terminally truncated version of FUM16 from F. verticillioides cDNA. The insert was then cloned into the BamHI/HindIII-digested vector pMALC2HTEV (Addgene no. 75286) using Gibson assembly.

F. verticillioides transformation and analysis of transformants

Protoplast transformation of F. verticillioides was carried out as described elsewhere (63). Here, 20–30 µg of the circular vector was transformed to achieve overexpression of GFP-tagged FUM15 and FUM16, while PCR-amplified fragments were transformed for inducible expression of FUM21, and for gene deletions. Transformants of tetON::FUM21 were maintained on complete medium (CM) plates (64) containing 200 µg/mL nourseothricin (Jena Bioscience, Jena, Germany). This strain was then used as background for the single-gene deletions, and the transformants were grown on CM plates containing both hygromycin B (200 µg/mL, InvivoGen Europe, Toulouse, France) and nourseothricin (200 µg/mL). Homologous recombination of the flanks and absence of untransformed nuclei were tested by diagnostic PCR, while Southern blot experiments excluded additional ectopic integration events. Thus, 2–10 independent transformants were verified for tetON::FUM21, tetON::FUM21fum8, tetON::FUM21fum15, tetON::FUM21fum16, tetON::FUM21fum18, and tetON::FUM21fum15-19 mutants (Fig. S7). The microscopy strain FUM18::DsRed (8) was maintained on CM plates containing 200 µg/mL nourseothricin and was used as background for the transformation of FUM15::GFP and FUM16::GFP. The transformants were grown on CM plates containing both hygromycin B (200 µg/mL) and nourseothricin (200 µg/mL). The ectopic integration of the plasmids was verified by diagnostic PCR (Fig. S8).

E. coli media and growth conditions

For cloning, E. coli DH5α cells were incubated on Luria-Bertani (LB) plates supplemented with 60 µg/mL carbenicillin (Carl Roth, Karlsruhe, Germany) and incubated at 37°C overnight. In order to isolate plasmids, single colonies were inoculated in LB medium with carbenicillin and incubated at 37°C and 180 rpm overnight. For protein production, E. coli BL21 (DE3) competent cells were transformed with the appropriate plasmid, grown overnight (37°C, 180 rpm) in LB medium with kanamycin (25 µg/mL, Carl Roth, Karlsruhe, Germany).

S. cerevisiae media and growth conditions

S. cerevisiae BY4741 (Euroscarf, Oberursel, Germany) was used as a background strain and is thus referred to as ScWT. General maintenance of the generated S. cerevisiae mutants was performed on solidified synthetic defined medium lacking uracil (SD-Ura). S. cerevisiae cells for plate assays were pre-cultured in liquid SD-Ura and shaken overnight in 100 mL flasks with baffles at 30°C and 180 rpm. All cultivations for growth curves, survival assays, as well as the analysis of ceramide intermediates were carried out in SD-Ura at 30°C. For gene expression analysis, 20 mL SD-Ura in 100 mL flasks with baffles was inoculated with a dense overnight culture to an OD600 of 0.2 and grown out to an OD600 of 1 before harvesting and freeze-drying.

F. verticillioides media and growth conditions

F. verticillioides M-3125 (65) was used as parental FvWT strain for the analysis. General maintenance of fungal strains was performed on solidified CM, with or without appropriate selection (200 µg/mL nourseothricin, 200 µg/mL hygromycin B). For the cultivation of strains in liquid culture, 100 mL of Darken pre-culture (66) in 300 mL Erlenmeyer flasks was inoculated with a mycelial plug and shaken for 3 days at 180 rpm and 28°C. For the main culture, 500 µL of the pre-culture was transferred to 100 mL of synthetic ICI medium (Imperial Chemical Industries, Ltd., London, UK) (67) supplemented with 6 mM glutamine (Gln) and shaken under the same conditions for 2 days. Gene expression of the FUM cluster under the tetON promoter was induced via the addition of 50 µg/mL doxycycline hyclate (Applichem, Darmstadt, Germany), and the cultures were shaken under the same conditions for an additional 5 days for FB1 and ceramide intermediate analyses.

F. verticillioides growth assay on plates

F. verticillioides strains were first incubated on CM plates with an appropriate resistance marker at 30°C for 7 days from which spores were harvested and counted using a Cell Counter (Beckmann Coulter, Krefeld, Germany). 10 µL containing 104 spores was spotted on the middle of fresh CM plates with and without supplementation of doxy (10–50 µg/mL). Each analyzed strain was spotted in triplicates. Plates were incubated for 3–6 days at 30°C. The colony diameters on plates containing doxy were adjusted by considering the growth of the strains in the absence of the inducer. The inhibition rate was then calculated with a formula: inhibition rate (IR) = (CT)/C × 100 (68), where C (control) represents the average growth of the strain without doxy and T (treated) represents the growth of the respective strain and replicate with doxy. For gene expression analysis, strains were grown for 3 days on CM plates with or without 25 µg/mL doxy. Cells were covered with a layer of cellophane to enable harvesting and subsequent freeze-drying of the mycelium.

Gene expression analysis via quantitative PCR

Fungal cultures were prepared as described above. Induction of 2-day-old ICI/6 mM Gln cultures with 10 µg/mL FB1 for 2 h was done previously, and expression analysis was essentially performed as previously described (8). Reactions were run on an Agilent Mx3000P qPCR System with the respective Agilent plastics (Santa Clara, CA, USA). Expression of the genes of interest and of the constitutively expressed reference genes (FVEG_07930/FvACT and YFL039C/ScACT1 [69] encoding actin) was determined in triplicates with the primers listed in Table S1. Expression relative to actin was calculated using the ∆Ct method (70).

Confocal microscopy

Microscopy experiments were performed on an Axio Observer Spinning Disc Confocal Microscope (Carl Zeiss, Jena, Germany) using 63×/1.2 oil or 100×/1.4 oil objectives with a numerical aperture (NA) value of 0.55. Laser lines of 488 nm and 561 nm were used for fluorophore excitation. Analysis was done with the ZEN 2.6 software, and in postprocessing, microscopy images were adjusted for brightness using the ImageJ software (https://imagej.nih.gov [71, 72]). Fungal hyphae were grown out from 104 conidia in 300 µL of ICI/6 mM Gln as adherent cultures in 8-well dishes (ibidi, Gräfelfing, Germany) at 30°C for 16 h.

Fum16 expression and purification

In order to render Fum16p soluble for bacterial expression, purification, and in vitro activity assay, its predicted N-terminal transmembrane domain had to be truncated; this corresponds to amino acids 1–88. Fum16p was expressed in E. coli BL21 (DE3) as a fusion construct N-terminally tagged with MBP under the control of an inducible T7 promoter (Fig. S6). A pre-culture was incubated overnight in LB medium containing 25 µg/mL kanamycin (37°C, 180 rpm). Then, 50 mL of Terrific Broth (TB) medium (73) was inoculated with the pre-culture to an OD600 of 0.1. The main culture was incubated (37°C, 180 rpm) until an OD600 of 0.5 was reached. Next, the temperature was lowered to 18°C, expression was induced after 1 h with 1 mM isopropyl β-d-1-thiogalactopyranoside (Carl Roth, Karlsruhe, Germany), and the culture was incubated overnight. The cell pellet was harvested by centrifugation (4,000 × g, 4°C, 20 min) and stored at −20°C until further processing.
Bacterial cells were thawed, resuspended in buffer A (20 mM Tris, 0.2 M NaCl, pH 8.0) containing 0.5 mM 4-(2-aminoethyl)benzenesulfonyl fluoride (AEBSF) and EDTA-free protease inhibitor cocktail (Roche, Grenzach-Wyhlen, Germany), and lysed using sonication (Sonopuls 2070, Bandelin, cycle 6, 75% intensity, 2 × 2 min) on ice. The protein preparations were centrifuged (16,000 × g, 4°C, 15 min), and the supernatants were applied to an MBP-Trap HP 1 mL column connected to an Aekta FPLC system (both GE Healthcare, Münich, Germany). After washing with 25 mM imidazole, the proteins were eluted with 500 mM imidazole using buffer B (20 mM Tris, 0.2 M NaCl, 10 mM maltose, pH 8.0). Protein-containing fractions were analyzed by Coomassie-stained SDS-PAGE, pooled, and concentrated via an Amicon filter (10 kDa cutoff) to 500 µL. This was applied to a Superdex 200 Increase 10/300 (Cytiva, Freiburg im Breisgau, Germany), and the protein was further purified by size-exclusion chromatography. Again, protein-containing fractions were analyzed (Fig. S6), pooled, flash-frozen in liquid nitrogen, and stored at −80°C. Protein concentrations were determined using the Bradford assay.

Fum16 in vitro activity assay

The reaction was performed in a 100 µL volume containing 100 mM HEPES (pH 7.5), 0.5 mM TCEP, 2.5 mM MgCl2, 0.5 mM ATP, 0.5 mM CoA, 5 mM serine, 0.5 mM palmitic acid, 20 µg/mL FadD (purified in references 37, 74), 100 µg/mL SPT (purified in reference 74), and 100 µg/mL Fum16p. The reaction was incubated overnight at 37°C and extracted with 50% (vol/vol) methanol and filtered through a 0.2 µm PTFE filter (Chromafil, Macherey-Nagel, Düren, Germany). The filtrate was then subjected to HPLC-HRMS analysis. The negative control did not contain any SPT enzyme.

S. cerevisiae cell viability assay

FB1 growth inhibition of S. cerevisiae strains expressing F. verticillioides genes FUM8, FUM15, FUM16, and FUM18 was evaluated by monitoring the OD600 every 15 min for up to 24 h. The experiments were conducted in an Infinite M200 plate reader (Tecan, Crailsheim, Germany) with either sterile black 96-well plates (BRAND plates; VWR, Darmstadt, Germany), or sterile black 24-well plates (ibidi, Gräfelfing, Germany). Incubation was performed with SD-Ura at 30°C. Each well contained either 500 µg/mL FB1 (Cayman Chemicals, Ann Arbor, MI) or water as a control, and 0.1 OD600 cells diluted in SD-Ura.

FB1 measurement for F. verticillioides liquid cultures

The supernatant was separated from mycelium through Miracloth and further clarified via centrifugation. Then, 100 µL of the clarified supernatant was mixed with 100% (vol/vol) MeOH and 1% (vol/vol) naringenin as an internal standard (1 mg/mL in methanol; Sigma-Aldrich, Steinheim, Germany) to give 1 mL (thereby diluted 1:10) and filtered through a 0.2 µm PTFE filter.
Filtered mycelia were washed with water, lyophilized, and weighed. Approximately 100 mg of dry mycelium was powdered using liquid nitrogen and resuspended in 1 mL MeOH:CHCl3 (1:1, vol/vol). The mycelium suspension was vigorously shaken (1,400 rpm) at 40°C overnight. Cell debris were pelleted (16,000 × g, 4°C, 15 min), and the supernatants were transferred to a fresh tube. The pellets were resuspended again in 1 mL MeOH:CHCl3 (1:1, vol/vol) and shaken for 1 h. The process was repeated two more times. The combined supernatants were collected, evaporated using SpeedVac, and resuspended overnight in 100% (vol/vol) MeOH. The samples were then filtered with a 0.2 µm PTFE filter prior to HPLC-HRMS analysis.

FB1 measurement for S. cerevisiae liquid cultures

The whole content from each well of the cell viability assay plate (see above) was centrifuged (10,000 × g, 4°C, 5 min) and 100 µL supernatant was mixed with 100% (vol/vol) MeOH to give 1 mL and filtered through a 0.2 µm PTFE filter (thereby diluted 1:10).

Extraction of ceramide intermediates from F. verticillioides and S. cerevisiae

For F. verticillioides, the same extraction method was used as described above for FB1 extraction from mycelium. For S. cerevisiae cells, the obtained pellet (10,000 × g, 4°C, 5 min) was washed twice with 900 µL water and resuspended in 1 mL MeOH:CHCl3 (1:1, vol/vol). The suspension was mixed with 0.5 mm glass beads, and the cells were lysed using a SpeedMill Plus (Analytic Jena, Jena, Germany, two cycles of 1 min). Cell debris were pelleted (10,000 × g, 4°C, 10 min), and the supernatant was collected. The pellet was resuspended again in 1 mL MeOH:CHCl3 (1:1, vol/vol) and lysed once more. The process was repeated once more. The combined supernatants were collected, evaporated using SpeedVac, and resuspended overnight in 100% (vol/vol) MeOH. The samples were then filtered with a 0.2 µm PTFE filter for HPLC-HRMS analysis.

HPLC-HRMS analysis

HPLC-HRMS analysis was performed using an LC-MS system consisting of a Q-Exactive Plus Hybrid Quadrupole Orbitrap mass spectrometer using electrospray ionization and a Dionex UltiMate 3000 UHPLC system (Thermo Fisher Scientific, Dreieich, Germany). Sample separation via HPLC was performed with a Kinetex C18 column (2.1 × 150 mm, 2.5 µm, 100 Å, Phenomenex) at a flowrate of 0.3 mL/min and column oven at 40°C. For analysis of FB1 and ceramide intermediates from both F. verticillioides and S. cerevisiae, an injection volume of 3 µL was set. The following elution gradient was used for FB1 detection (solvent A, H2O plus 0.1% [vol/vol] HCOOH; solvent B, acetonitrile plus 0.1% [vol/vol] HCOOH): 5% B for 0.5 min, 5 to 97% B in 11.5 min, and 97% B for 3 min. The following gradient was used for ceramide intermediates detection: 5% B for 0.5 min, 5 to 97% B in 54.5 min, and 97% B for 3 min. Raw LC-MS data were analyzed using XCalibur (Thermo Fisher Scientific, Dreieich, Germany) with a mass resolution of 10 ppm (Fig. S3 and S10). FB1 peak areas were normalized against the internal standard to account for variability between runs. FB1 and ceramide intermediate levels were related to the dry weight of the F. verticillioides cultures, which were performed in biological triplicate. The standard curve was prepared by injecting 3 µL of the following FB1 concentrations prepared in ethanol in triplicate: 0.1 µg/mL, 0.5 µg/mL, 1 µg/mL, 10 µg/mL, and 100 µg/mL.

ACKNOWLEDGMENTS

We thank Daniela Hildebrandt and Ahsan Ullah (Leibniz-HKI) for excellent technical assistance.
This work was supported by the German Research Foundation (DFG grant 453246485 to V.V. and S.J.) and by the Free State of Thuringia and the European Social Fund (project SphinX, 2017FGR0073) to V.V. Work in S.J.’s laboratory is supported by the Free State of Thuringia and the European Social Fund Plus (project FusInfect, 2022FGR0007).
The authors declare no competing interests. The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
Conceptualization, V.V. and S.J.; methodology, F.G., K.J., Y.H., S.H., S.J., and V.V.; investigation, F.G., K.J., Y.H., S.H., and S.J.; writing – original draft, F.G.; writing – editing, S.J.; writing – review, all authors; funding acquisition, V.V. and S.J.; supervision, V.V. and S.J.

SUPPLEMENTAL MATERIAL

Supplemental material - mbio.02681-24-s0001.pdf
Supplemental figures and table.
ASM does not own the copyrights to Supplemental Material that may be linked to, or accessed through, an article. The authors have granted ASM a non-exclusive, world-wide license to publish the Supplemental Material files. Please contact the corresponding author directly for reuse.

REFERENCES

1.
Keller NP, Turner G, Bennett JW. 2005. Fungal secondary metabolism - from biochemistry to genomics. Nat Rev Microbiol 3:937–947.
2.
Netzker T, Fischer J, Weber J, Mattern DJ, König CC, Valiante V, Schroeckh V, Brakhage AA. 2015. Microbial communication leading to the activation of silent fungal secondary metabolite gene clusters. Front Microbiol 6:299.
3.
Venkatesh N, Keller NP. 2019. Mycotoxins in conversation with bacteria and fungi. Front Microbiol 10:403.
4.
Li GH, Zhang KQ. 2014. Nematode-toxic fungi and their nematicidal metabolites. In Zhang KQ, Hyde K (ed), Nematode-trapping fungi. Fungal diversity research series. Vol. 23. Springer, Dordrecht.
5.
Bennett JW, Klich M. 2003. Mycotoxins. Clin Microbiol Rev 16:497–516.
6.
Del Sorbo G, Schoonbeek H, De Waard MA. 2000. Fungal transporters involved in efflux of natural toxic compounds and fungicides. Fungal Genet Biol 30:1–15.
7.
Liu L, Xie M, Wei D. 2022. Biological detoxification of mycotoxins: current status and future advances. Int J Mol Sci 23:1064.
8.
Janevska S, Ferling I, Jojić K, Rautschek J, Hoefgen S, Proctor RH, Hillmann F, Valiante V. 2020. Self-protection against the sphingolipid biosynthesis inhibitor fumonisin B1 is conferred by a FUM cluster-encoded ceramide synthase. MBio 11:e00455-20.
9.
Roze LV, Chanda A, Linz JE. 2011. Compartmentalization and molecular traffic in secondary metabolism: a new understanding of established cellular processes. Fungal Genet Biol 48:35–48.
10.
Nützmann HW, Scazzocchio C, Osbourn A. 2018. Metabolic gene clusters in eukaryotes. Annu Rev Genet 52:159–183.
11.
Khaldi N, Wolfe KH. 2011. Evolutionary origins of the fumonisin secondary metabolite gene cluster in Fusarium verticillioides and Aspergillus niger. Int J Evol Biol 2011:423821.
12.
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.
13.
Howard PC, Couch LH, Patton RE, Eppley RM, Doerge DR, Churchwell MI, Marques MM, Okerberg CV. 2002. Comparison of the toxicity of several fumonisin derivatives in a 28-day feeding study with female B6C3F(1) mice. Toxicol Appl Pharmacol 185:153–165.
14.
Merrill AH, Wang E, Vales TR, Smith ER, Schroeder JJ, Menaldino DS, Alexander C, Crane HM, Xia J, Liotta DC, Meredith FI, Riley RT. 1996. Fumonisin toxicity and sphingolipid biosynthesis. Adv Exp Med Biol 392:297–306.
15.
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.
16.
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.
17.
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.
18.
Uhlig S, Busman M, Shane DS, Rønning H, Rise F, Proctor R. 2012. Identification of early fumonisin biosynthetic intermediates by inactivation of the FUM6 gene in Fusarium verticillioides. J Agric Food Chem 60:10293–10301.
19.
Butchko RAE, 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.
20.
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.
21.
Li Y, Lou L, Cerny RL, Butchko RAE, Proctor RH, Shen Y, Du L. 2013. Tricarballylic ester formation during biosynthesis of fumonisin mycotoxins in Fusarium verticillioides . Mycology: An International Journal on Fungal Biology 4:179–186.
22.
Zaleta-Rivera K, Xu C, Yu F, Butchko RAE, 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. Biochemistry 45:2561–2569.
23.
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.
24.
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.
25.
Wang E, Norred WP, Bacon CW, Riley RT, Merrill AH Jr. 1991. Inhibition of sphingolipid biosynthesis by fumonisins. Implications for diseases associated with Fusarium moniliforme. J Biol Chem 266:14486–14490.
26.
Merrill AH, Sullards MC, Wang E, Voss KA, Riley RT. 2001. Sphingolipid metabolism: roles in signal transduction and disruption by fumonisins. Environ Health Perspect 109 Suppl 2:283–289.
27.
Park JW, Park WJ, Futerman AH. 2014. Ceramide synthases as potential targets for therapeutic intervention in human diseases. Biochim Biophys Acta 1841:671–681.
28.
Narayan S, Head SR, Gilmartin TJ, Dean B, Thomas EA. 2009. Evidence for disruption of sphingolipid metabolism in schizophrenia. J of Neuroscience Research 87:278–288.
29.
He X, Huang Y, Li B, Gong CX, Schuchman EH. 2010. Deregulation of sphingolipid metabolism in Alzheimer’s disease. Neurobiol Aging 31:398–408.
30.
Fernandes CM, Goldman GH, Del Poeta M. 2018. Biological roles played by sphingolipids in dimorphic and filamentous fungi. MBio 9:e00642-18.
31.
Hannun YA, Obeid LM. 2008. Principles of bioactive lipid signalling: lessons from sphingolipids. Nat Rev Mol Cell Biol 9:139–150.
32.
Mencarelli C, Martinez-Martinez P. 2013. Ceramide function in the brain: when a slight tilt is enough. Cell Mol Life Sci 70:181–203.
33.
Coant N, Sakamoto W, Mao C, Hannun YA. 2017. Ceramidases, roles in sphingolipid metabolism and in health and disease. Adv Biol Regul 63:122–131.
34.
Ogretmen B. 2018. Sphingolipid metabolism in cancer signalling and therapy. Nat Rev Cancer 18:33–50.
35.
Tidhar R, Futerman AH. 2013. The complexity of sphingolipid biosynthesis in the endoplasmic reticulum. Biochim Biophys Acta 1833:2511–2518.
36.
Körner C, Fröhlich F. 2022. Compartmentation and functions of sphingolipids. Curr Opin Cell Biol 74:104–111.
37.
Bissell AU, Rautschek J, Hoefgen S, Raguž L, Mattern DJ, Saeed N, Janevska S, Jojić K, Huang Y, Kufs JE, Herboeck B, Guo H, Hillmann F, Beemelmanns C, Valiante V. 2022. Biosynthesis of the sphingolipid inhibitors sphingofungins in filamentous fungi requires aminomalonate as a metabolic precursor. ACS Chem Biol 17:386–394.
38.
Keyser Z, Vismer HF, Klaasen JA, Snljman· PW, Marasas· WFO. 1999. The antifungal effect of fumonisin B1 on Fusarium and other fungal species. Res Lett S Afr J Sci 95:455–458.
39.
Meyer V, Wanka F, van Gent J, Arentshorst M, van den Hondel CAMJJ, Ram AFJ. 2011. Fungal gene expression on demand: an inducible, tunable, and metabolism-independent expression system for Aspergillus niger. Appl Environ Microbiol 77:2975–2983.
40.
Janevska S, Arndt B, Baumann L, Apken LH, Mauriz Marques LM, Humpf HU, Tudzynski B. 2017. Establishment of the inducible Tet-on system for the activation of the silent trichosetin gene cluster in Fusarium fujikuroi. Toxins (Basel) 9:126.
41.
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.
42.
Kelley LA, Mezulis S, Yates CM, Wass MN, Sternberg MJE. 2015. The Phyre2 web portal for protein modeling, prediction and analysis. Nat Protoc 10:845–858.
43.
Hu Y, Gai Y, Yin L, Wang X, Feng C, Feng L, Li D, Jiang X-N, Wang D-C. 2010. Crystal structures of a Populus tomentosa 4-coumarate:CoA ligase shed light on its enzymatic mechanisms. Plant Cell 22:3093–3104.
44.
Yan M, Cao L, Zhao L, Zhou W, Liu X, Zhang W, Rao Z. 2023. The key roles of Mycobacterium tuberculosis FadD23 C-terminal domain in catalytic mechanisms. Front Microbiol 14:1090534.
45.
Jumper J, Evans R, Pritzel A, Green T, Figurnov M, Ronneberger O, Tunyasuvunakool K, Bates R, Žídek A, Potapenko A, et al. 2021. Highly accurate protein structure prediction with AlphaFold. Nature New Biol 596:583–589.
46.
Kistler HC, Broz K. 2015. Cellular compartmentalization of secondary metabolism. Front Microbiol 6:68.
47.
Neve EPA, Ingelman-Sundberg M. 2008. Intracellular transport and localization of microsomal cytochrome P450. Anal Bioanal Chem 392:1075–1084.
48.
Malonek S, Rojas MC, Hedden P, Gaskin P, Hopkins P, Tudzynski B. 2004. The NADPH-cytochrome P450 reductase gene from Gibberella fujikuroi is essential for gibberellin biosynthesis. J Biol Chem 279:25075–25084.
49.
Lim FY, Keller NP. 2014. Spatial and temporal control of fungal natural product synthesis. Nat Prod Rep 31:1277–1286.
50.
Jojić K, Gherlone F, Cseresnyés Z, Bissell AU, Hoefgen S, Hoffmann S, Huang Y, Janevska S, Figge MT, Valiante V. 2024. The spatial organization of sphingofungin biosynthesis in Aspergillus fumigatus and its cross-interaction with sphingolipid metabolism. MBio 15:e0019524.
51.
Susca A, Proctor RH, Butchko RAE, Haidukowski M, Stea G, Logrieco A, Moretti A. 2014. Variation in the fumonisin biosynthetic gene cluster in fumonisin-producing and nonproducing black aspergilli. Fungal Genet Biol 73:39–52.
52.
Shimizu K, Nakagawa H, Hashimoto R, Hagiwara D, Onji Y, Asano K, Kawamoto S, Takahashi H, Yokoyama K. 2015. The α-oxoamine synthase gene fum8 is involved in fumonisin B2 biosynthesis in Aspergillus niger. Mycoscience 56:301–308.
53.
Pel HJ, de Winde JH, Archer DB, Dyer PS, Hofmann G, Schaap PJ, Turner G, de Vries RP, Albang R, Albermann K, et al. 2007. Genome sequencing and analysis of the versatile cell factory Aspergillus niger CBS 513.88. Nat Biotechnol 25:221–231.
54.
Aerts D, Hauer EE, Ohm RA, Arentshorst M, Teertstra WR, Phippen C, Ram AFJ, Frisvad JC, Wösten HAB. 2018. The FlbA-regulated predicted transcription factor Fum21 of Aspergillus niger is involved in fumonisin production. Antonie Van Leeuwenhoek 111:311–322.
55.
Li P, Su R, Yin R, Lai D, Wang M, Liu Y, Zhou L. 2020. Detoxification of mycotoxins through biotransformation. Toxins (Basel) 12:121.
56.
Studt L, Janevska S, Niehaus EM, Burkhardt I, Arndt B, Sieber CMK, Humpf HU, Dickschat JS, Tudzynski B. 2016. Two separate key enzymes and two pathway-specific transcription factors are involved in fusaric acid biosynthesis in Fusarium fujikuroi. Environ Microbiol 18:936–956.
57.
Gao M, Glenn AE, Gu X, Mitchell TR, Satterlee T, Duke MV, Scheffler BE, Gold SE. 2020. Pyrrocidine, a molecular off switch for fumonisin biosynthesis. PLoS Pathog 16:e1008595.
58.
Kouprina N, Larionov V. 2008. Selective isolation of genomic loci from complex genomes by transformation-associated recombination cloning in the yeast Saccharomyces cerevisiae. Nat Protoc 3:371–377.
59.
Gibson DG, Young L, Chuang RY, Venter JC, Hutchison CA, Smith HO. 2009. Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods 6:343–345.
60.
Liebmann B, Müller M, Braun A, Brakhage AA. 2004. The cyclic AMP-dependent protein kinase A network regulates development and virulence in Aspergillus fumigatus. Infect Immun 72:5193–5203.
61.
Schumacher J. 2012. Tools for Botrytis cinerea: new expression vectors make the gray mold fungus more accessible to cell biology approaches. Fungal Genet Biol 49:483–497.
62.
Hoefgen S, Lin J, Fricke J, Stroe MC, Mattern DJ, Kufs JE, Hortschansky P, Brakhage AA, Hoffmeister D, Valiante V. 2018. Facile assembly and fluorescence-based screening method for heterologous expression of biosynthetic pathways in fungi. Metab Eng 48:44–51.
63.
Tudzynski B, Homann V, Feng B, Marzluf GA. 1999. Isolation, characterization and disruption of the areA nitrogen regulatory gene of Gibberella fujikuroi. Mol Gen Genet 261:106–114.
64.
Pontecorvo G, Roper JA, Chemmons LM, Macdonald KD, Bufton AWJ. 1953. The genetics of Aspergillus nidulans. Adv Genet 5:141–238.
65.
Leslie JF, Doe FJ, Plattner RD, Shackelford DD, Jonz J. 1992. Fumonisin B1 production and vegetative compatibility of strains from Gibberella fujikuroi mating population “A” (Fusarium moniliforme). Mycopathologia 117:37–45.
66.
DARKEN MA, JENSEN AL, SHU P. 1959. Production of gibberellic acid by fermentation. Appl Microbiol 7:301–303.
67.
Geissman TA, Verbiscar AJ, Phinney BO, Cragg G. 1966. Studies on the biosynthesis of gibberellins from (−)-kaurenoic acid in cultures of Gibberella fujikuroi. Phytochemistry 5:933–947.
68.
Wonglom P, Ito S, Sunpapao A. 2020. Volatile organic compounds emitted from endophytic fungus Trichoderma asperellum T1 mediate antifungal activity, defense response and promote plant growth in lettuce (Lactuca sativa). Fungal Ecol 43:100867.
69.
De-Souza EA, Pimentel FSA, Machado CM, Martins LS, da-Silva WS, Montero-Lomelí M, Masuda CA. 2014. The unfolded protein response has a protective role in yeast models of classic galactosemia. Dis Model Mech 7:55–61.
70.
Livak KJ, Schmittgen TD. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods 25:402–408.
71.
Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez JY, White DJ, Hartenstein V, Eliceiri K, Tomancak P, Cardona A. 2012. Fiji: an open-source platform for biological-image analysis. Nat Methods 9:676–682.
72.
Rueden CT, Schindelin J, Hiner MC, DeZonia BE, Walter AE, Arena ET, Eliceiri KW. 2017. ImageJ2: ImageJ for the next generation of scientific image data. BMC Bioinformatics 18:529.
73.
Elbing KL, Brent R. 2019. Recipes and tools for culture of Escherichia coli. Curr Protoc Mol Biol 125:e83.
74.
Hoefgen S, Bissell AU, Huang Y, Gherlone F, Raguž L, Beemelmanns C, Valiante V. 2022. Desaturation of the sphingofungin polyketide tail results in increased serine palmitoyltransferase inhibition. Microbiol Spectr 10:e0133122.

Information & Contributors

Information

Published In

cover image mBio
mBio
Volume 16Number 25 February 2025
eLocator: e02681-24
Editor: James W. Kronstad, The University of British Columbia, Vancouver, British Columbia, Canada
PubMed: 39704544

History

Received: 30 August 2024
Accepted: 2 December 2024
Published online: 20 December 2024

Keywords

  1. Fusarium
  2. fumonisin B1
  3. FUM cluster
  4. self-protection
  5. ceramide biosynthesis
  6. palmitoyl-CoA ligase

Contributors

Authors

Fabio Gherlone
(Epi-)Genetic Regulation of Fungal Virulence, Leibniz Institute for Natural Product Research and Infection Biology-Hans Knöll Institute (Leibniz-HKI), Jena, Germany
Biobricks of Microbial Natural Product Syntheses, Leibniz Institute for Natural Product Research and Infection Biology-Hans Knöll Institute (Leibniz-HKI), Jena, Germany
Author Contributions: Investigation, Methodology, and Writing – original draft.
Katarina Jojić
Biobricks of Microbial Natural Product Syntheses, Leibniz Institute for Natural Product Research and Infection Biology-Hans Knöll Institute (Leibniz-HKI), Jena, Germany
Author Contributions: Investigation, Methodology, and Writing – review and editing.
Ying Huang
Biobricks of Microbial Natural Product Syntheses, Leibniz Institute for Natural Product Research and Infection Biology-Hans Knöll Institute (Leibniz-HKI), Jena, Germany
Author Contributions: Investigation, Methodology, and Writing – review and editing.
Sandra Hoefgen
Biobricks of Microbial Natural Product Syntheses, Leibniz Institute for Natural Product Research and Infection Biology-Hans Knöll Institute (Leibniz-HKI), Jena, Germany
Author Contributions: Investigation, Methodology, and Writing – review and editing.
Present address: Jena Bioscience GmbH, Jena, Germany
Vito Valiante
Biobricks of Microbial Natural Product Syntheses, Leibniz Institute for Natural Product Research and Infection Biology-Hans Knöll Institute (Leibniz-HKI), Jena, Germany
Author Contributions: Conceptualization, Funding acquisition, Methodology, Supervision, and Writing – review and editing.
(Epi-)Genetic Regulation of Fungal Virulence, Leibniz Institute for Natural Product Research and Infection Biology-Hans Knöll Institute (Leibniz-HKI), Jena, Germany
Author Contributions: Conceptualization, Funding acquisition, Investigation, Methodology, Supervision, and Writing – review and editing.

Editor

James W. Kronstad
Editor
The University of British Columbia, Vancouver, British Columbia, Canada

Notes

Fabio Gherlone and Katarina Jojić contributed equally to this article. Author order was determined alphabetically.
The authors declare no conflict of interest.

Metrics & Citations

Metrics

Note:

  • For recently published articles, the TOTAL download count will appear as zero until a new month starts.
  • There is a 3- to 4-day delay in article usage, so article usage will not appear immediately after publication.
  • Citation counts come from the Crossref Cited by service.

Citations

If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. For an editable text file, please select Medlars format which will download as a .txt file. Simply select your manager software from the list below and click Download.

View Options

Figures

Tables

Media

Share

Share

Share the article link

Share with email

Email a colleague

Share on social media

American Society for Microbiology ("ASM") is committed to maintaining your confidence and trust with respect to the information we collect from you on websites owned and operated by ASM ("ASM Web Sites") and other sources. This Privacy Policy sets forth the information we collect about you, how we use this information and the choices you have about how we use such information.
FIND OUT MORE about the privacy policy