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. 2025 Apr 22;27(4):e70100. doi: 10.1111/1462-2920.70100

ZfpA‐Dependent Quorum Sensing Shifts in Morphology and Secondary Metabolism in Aspergillus flavus

Benjamin Otoo 1, Dante G Calise 2, Sung Chul Park 2, Jin Woo Bok 2, Nancy P Keller 2,3,✉, Mira Syahfriena Amir Rawa 2,✉
PMCID: PMC12014256  PMID: 40262766

ABSTRACT

Development of the fungal pathogen Aspergillus flavus involves the balance of asexual spores (conidia) and overwintering hardened hyphal masses (sclerotia). This balance is achieved by an oxylipin‐based density‐dependent mechanism regulating the switch from sclerotia to conidia as population density increases in A. flavus . Here, we show the transcription factor ZfpA, required for normal oxylipin synthesis, regulates the morphology switch. ZfpA overexpression (OE::zfpA) accelerates the shift leading to increased conidial production and reduced sclerotial production under conditions normally supporting sclerotia formation. In contrast, zfpA deletion (ΔzfpA) produces more sclerotia than wild‐type control. These morphology changes are coupled with changes in tissue‐specific secondary metabolites. Specifically, the production of four sclerotial metabolites (oxyasparasone A, hydroxyaflatrem, aflavinine, and kotanin) decreases in OE::zfpA whereas the hyphal metabolite aspergillic acid is upregulated in this mutant. Chemical profiling of OE::zfpA compared to a double mutant where the aspergillic acid non‐ribosomal synthetase was deleted in the OE::zfpA background confirmed synthesis of known aspergillic acid pathway products as well as putative Val‐derived pyrazinones involved in metal chelation. These findings offer valuable insights into the quorum sensing networks connecting fungal development and tissue‐specific secondary metabolite production.

Keywords: aflatoxin, aspergillic acid, oxylipin, quorum sensing, sclerotia

ZfpA regulation on morphological development and specific secondary metabolite production in Aspergillus flavus. ZfpA overexpression increases conidiation and tissue‐specific secondary metabolite (aspergillic acid) production, while zfpA deletion increases sclerotial development and sclerotial‐related secondary metabolites.

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1. Introduction

The genus Aspergillus comprises hundreds of species of which a few are notable opportunistic pathogens. Among these pathogens, Aspergillus flavus stands out as a well‐known crossover pathogen of both crops and humans (Gauthier and Keller 2013). The fungus is the second most prevalent cause of invasive aspergillosis and a common post‐harvest pathogen where it infects seeds of various crops, contaminating them with aflatoxins, potent carcinogens (Hedayati et al. 2007; Amaike and Keller 2011). Common to all Aspergillus spp., A. flavus follows a programmed developmental process where the initial inoculum (e.g., the asexual spore or conidium) will germinate and establish a colony by alternating polar growth supported by lateral hyphal branches. With appropriate stimuli, the fungus will form hardened, pigmented hyphal masses called sclerotia that can withstand harsh environmental conditions and give rise to conidiophores and conidia when the environment is more favourable (Amaike and Keller 2011; Diener et al. 1987). The sclerotium is also the structure where sexual spores are formed when strains of opposite mating type meet (Horn et al. 2014; Wicklow et al. 1984). These programmed developmental events are critical for the ecological and pathogenic success of this fungus.

Considering the importance of A. flavus on agricultural welfare and human health, many studies have focused on elucidating genetic and environmental cues that mediate the development programme. One of the known signals mediating the ratio of asexual to sexual reproduction is endogenous oxylipins synthesised by five A. flavus oxygenases (Lox, PpoA, PpoB, PpoC, and PpoD) (Brown et al. 2009; Horowitz Brown et al. 2008). Oxylipins, a class of oxygenase‐derived unsaturated fatty acids, serve as vital signal molecules in various biological systems (Funk 2001; Niu et al. 2020). In A. flavus, each oxygenase produces one or more oxylipins from linoleic acid with different impacts on fungal development (Brown et al. 2009; Horowitz Brown et al. 2008; Niu et al. 2020). The characterisation of these oxygenases in A. flavus has revealed their significance in orchestrating a population density‐dependent morphological transition from sclerotia to conidia, where low cell density favours sclerotia production and high cell density favours asexual reproduction (Horowitz Brown et al. 2008). The deletion of A. flavus oxygenases is associated with a disruption in sclerotia and conidia development in a density‐dependent manner (Brown et al. 2009; Horowitz Brown et al. 2008), thus establishing oxylipins as quorum‐like signalling molecules. Quorum signalling in A. flavus may also be associated with density‐dependent aflatoxin production; for instance, aflatoxin levels are higher in low‐density (high sclerotial) populations (Brown et al. 2009).

Despite this understanding of oxylipin mediated quorum signalling in A. flavus and the identification of oxylipin generating oxygenases in this species, regulatory networks governing ppo expression and downstream density dependent development are largely unknown. Recently, significant progress on this front came with the finding in the human pathogen Aspergillus fumigatus that a C2H2 zinc finger transcription factor, ZfpA, regulates PpoA activity and synthesis of the PpoA oxylipin product 5,8‐diHODE (Niu et al. 2020; Calise et al. 2024). Both 5,8‐diHODE and overexpression of zfpA induce hyperbranching and cell wall chitin deposition in this species, leading to resistance to the echinocandin class of antifungals (Calise et al. 2024). This work also supported that a similar resistance mechanism existed in A. flavus .

We hypothesised that the identification of ZfpA, the first transcription factor found to regulate a specific Aspergillus ppo gene and its corresponding oxylipin, would allow for a new approach to understand mechanisms and consequences of quorum sensing in A. flavus. Here, we find that ZfpA accelerates the switch of sclerotial to conidial development where deletion of zfpA (ΔzfpA) increases sclerotial production and overexpression of zfpA (OE::zfpA) increases conidial development in a pattern that shifts but does not eliminate the density‐dependent influence on these processes. These morphological changes were correlated to corresponding ‘switches’ in production of specific secondary metabolites (SMs). Depending on the population densities, the OE::zfpA strain showed a 5–60 fold‐change decrease in production of several sclerotial associated metabolites and a 5–15 fold‐change increase in aspergillic acid, a hyphal metabolite (Calvo and Cary 2015; TePaske et al. 1992; Cary et al. 2018). Further, through deletion of asaC, encoding the non‐ribosomal peptide synthetase (NRPS) required for aspergillic acid synthesis (Lebar et al. 2018), we were able to obtain putative aspergillic acid related products including Val‐derived products and demonstrate the metal chelating properties of these mycelial secondary metabolites.

2. Results

2.1. Expression of ppoA and the 5,8‐diHODE Branching Response Are Mediated by the Transcription Factor ZfpA

A previous study on Aspergillus developmental signals identified an endogenously produced chemical cue, the diol oxylipin 5,8‐diHODE, which induced secondary branching in both A. fumigatus and A. flavus (Niu et al. 2020). The enzyme PpoA, which produces 5,8‐diHODE, is conserved in Aspergilli (Brown et al. 2009; Niu et al. 2020; Tsitsigiannis et al. 2004). Thus, we were interested to assess the dynamics of A. flavus 5,8‐diHODE production and if this metabolite impacted other developmental processes of the fungus. Examination of the parental wild‐type (WT) strain NRRL3357 at a concentration of 106 spores/mL in liquid shake and incubated for 3 days showed that 5,8‐diHODE was abundantly present in fungal tissue but was barely present in the liquid culture supernatant (Figure 1A), following a similar pattern reported for A. fumigatus (Niu et al. 2020).

FIGURE 1.

FIGURE 1

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ppoA expression, 5,8‐diHODE production and lateral branching are regulated by ZfpA in A. flavus . (A) Quantification of 5,8‐diHODE from liquid shake glucose minimal medium (GMM) in A. flavus . p‐values were calculated using one‐way ANOVA with Tukey's multiple comparisons test (B) Concentration dependent increase in branching in A. flavus when treated with 5,8‐diHODE. p‐values were calculated using one‐way ANOVA with Tukey's multiple comparisons test. (C) Northern blot result shows the downregulation of ppoA expression in ΔzfpA compared to WT and OE::zfpA mutants. OE::zfpA mutant shows an upregulation of ppoA transcripts compared to the WT control. Transcript signals of ppoA and gpdA were quantified using ImageJ and signal ratios were normalised by the mean of the WT samples. (D) The bar graph shows 5,8‐diHODE production after 7 days of growth at 104 spore density. The intensity of 5,8‐diHODE increased in the OE::zfpA mutant and decreased in the ΔzfpA mutant, comparable to the WT control. p‐values were calculated using one‐way ANOVA with Tukey's multiple comparisons test (E) Examination of branching in WT, ΔzfpA, and OE::zfpA strains grown in a 96‐well plate containing liquid GMM 21‐h post‐incubation at 30°C. p‐values were calculated using one‐way ANOVA with Tukey's multiple comparisons test. (F) The graph illustrates zfpA expression is needed for 5,8‐diHODE response. OE::zfpA mutant and WT control significantly increased branching in response to 0.1 μg/mL 5,8‐diHODE treatment. p‐values were calculated using two‐way ANOVA with Tukey's multiple comparisons test.

We also confirmed the ability of 5,8‐diHODE to induce lateral branching in A. flavus . We treated A. flavus WT spores with different concentrations of purified 5,8‐diHODE. After 21 h post incubation, a dose‐dependent hyperbranching response to 5,8‐diHODE was observed, with the highest branching at a concentration of 10 μg/mL (Figure 1B), the highest concentration tested. This finding extended an initial report that 5,8‐diHODE induced branching in A. flavus (Calise et al. 2024).

Expression of ppoA and subsequent 5,8‐diHODE biosynthesis are positively regulated by the transcription factor ZfpA in A. fumigatus (Calise et al. 2024). To determine if ZfpA regulated ppoA expression in A. flavus and subsequent 5,8‐diHODE synthesis, the ΔzfpA and OE::zfpA mutants (Calise et al. 2024) were compared with WT. After determining that all strains germinated at equal rates (Figure S1), the three strains were grown for 24 h to assess ppoA expression and 7 days to quantify 5,8‐diHODE production. ZfpA positively regulates both ppoA gene expression (Figure 1C) and 5,8‐diHODE synthesis (Figure 1D). Assessment of the mutants compared to WT showed that the OE::zfpA strain exhibited a significant hyperbranching pattern whereas the deletion strain was similar to that of WT (Figure 1E). Unlike the A. fumigatus OE::zfpA strain, the OE::zfpA strain of A. flavus was still responsive to 0.1 μg/mL of 5,8‐diHODE and exhibited an even higher hyperbranching phenotype when thus treated (Figure 1E,F). In contrast, the ΔzfpA mutant remained unresponsive to 5,8‐diHODE treatment (Figure 1E,F), similar to the lack of response in the A. fumigatus deletion mutant (Niu et al. 2020). Considering the collective findings, this data show that ZfpA plays a conserved and pivotal role in the 5,8‐diHODE response cascade, exerting regulatory control over hyphal branching in A. flavus .

2.2. ZfpA Regulates Density Dependent Morphological Transitions During A. flavus Development

Previous studies have highlighted the involvement of oxylipins in density‐dependent quorum‐like spore and sclerotia development in A. flavus (Brown et al. 2009; Horowitz Brown et al. 2008). At spore densities below 103/plate, the fungus predominantly generates sclerotia with very little conidia production. Conversely, at spore densities of 106/plate and above, the fungus primarily produces conidia with minimal or no sclerotia formation. Intermediate spore densities represent a state of equilibrium between sclerotia and conidia production. Given the evidence implicating ZfpA in mediating 5,8‐diHODE oxylipin signalling, our interest was piqued regarding the potential impact of ZfpA on density‐dependent morphological transitions in A. flavus . To investigate this, the WT, ΔzfpA and OE::zfpA mutants were cultivated at three initial spore densities: 102 spores/plate (low), 104 spores/plate (intermediate), and 106 spores/plate (high). After seven days of incubation, spore enumeration and sclerotia collection and weighing were performed for all treatments. Notably, the OE::zfpA strain produced more conidia at low density compared to the ΔzfpA and WT control strains, although the difference was not statistically significant (Figure 2A,B). At the intermediate density level, the overexpression mutant displayed significantly higher conidial production, contrasting with the ΔzfpA and WT control (Figure 2B). However, at high spore density, there was no difference in spore production among the strains (Figure 2B). The results for sclerotia formation after 7 days revealed a reversal of the conidiation findings. At low and intermediate densities, sclerotia mass was significantly higher for the ΔzfpA and WT control compared to the OE::zfpA strain (Figure 2C,D). The OE::zfpA strain also showed a different pattern of sclerotial formation at the low density where sclerotia were clumped unlike the typical dispersed formation in both WT and ΔzfpA strains (Figure 2E,F). No sclerotia were produced at high spore density for any strain following the established density‐dependent development programme. These findings indicate that ZfpA is a key protein in mediating the density transition of sclerotial/conidiation development in A. flavus .

FIGURE 2.

FIGURE 2

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ZfpA impacts conidiation and sclerotia production at low and intermediate spore densities. (A) The images illustrate density‐dependent conidiation of ZfpA mutants after 7 days of growth. An overlay of low, intermediate, and high spore densities of mutants was cultivated on GMM containing 2% sorbitol, then incubated at 30°C. Four replicates of each mutant were grown for reproducibility. (B) The graph displays quantitative spore counts for ZfpA mutants. Plugs were taken from four replicates plates of each mutant after 7 days for conidia counts. p‐values were calculated using two‐way ANOVA with Tukey's multiple comparisons test. (C) The images show density‐dependent sclerotia production of ZfpA mutants after 7 days of growth. The black structures observed are sclerotia. (D) Graph illustrates gravimetric measurement of sclerotia collected from four replicates plates of each mutant after 7 days of growth. p‐values were calculated using two‐way ANOVA with Tukey's multiple comparisons test. (E) The images show sclerotia formation for WT plate. Large round black structures represent sclerotia. Cultures were grown in the dark using GMM plates supplemented with sorbitol to induce sclerotia. (F) The images show clustered sclerotia formation for OE::zfpA mutant plate. Cultures were grown in the dark using GMM plates supplemented with sorbitol to induce sclerotia.

2.3. ZfpA Modulates the Chemical Profile of A. flavus

A characteristic of many fungal natural products is the spatial production of specific metabolites in distinct developmental tissues (Lim and Keller 2014). Clear examples include the preferential synthesis of a subset of SMs in A. fumigatus asexual spores (Blachowicz et al. 2020), bikaverin synthesis in chlamydospores of Fusarium spp. (Spraker et al. 2018), fusarubin synthesis of Fusarium fujikuroi perithecia (Studt et al. 2012) and, pertinent to A. flavus, the numerous metabolites produced in sclerotia including aflavarins, aflavazoles, and cyclopiazonic acids (Calvo and Cary 2015; TePaske et al. 1992; Cary et al. 2018). Driven by these previous findings, we hypothesised that the morphological development associated with density dependence or mediated by ZfpA would regulate the SM profile in A. flavus , particularly sclerotial SMs.

To thoroughly examine the density dependent and ZfpA effects on A. flavus chemical composition, the WT, ΔzfpA and OE::zfpA mutants were cultivated at three initial spore densities similar to the morphological development analysis (Figure 2A,C). After 7 days of incubation, the strains were extracted with organic solvent and subjected to UHPLC–MS/MS and metabolomic analysis. Principal component analysis (PCA) showed that the WT control and ΔzfpA strain were grouped together, denoting comparable chemical profiles between these two strains at the low and intermediate density levels, with some separation only at the high‐density level (Figure 3A). On the other hand, the OE::zfpA mutant displayed a distinct chemical profile at all three population densities, especially at 106 spore density, where it singularly clustered. Volcano plots (p‐values > 0.05, abs(log2fold‐change) > 1, intensity threshold: 5 × 105) of the 106 spore density cultures visually illustrate UHPLC–MS/MS chemical profile differences between WT and the two mutants, respectively (Figure 3B,C). The OE::zfpA strain exhibited 15.10% upregulated features and 25.95% downregulated features compared to WT (Figure 3B) whereas the ΔzfpA strain showed 7.90% chemical feature upregulation and 11.20% downregulation compared to WT (Figure 3C). This result is consistent with the PCA result in which metabolites are differentially synthesised in ZfpA mutants, especially at high spore density cultures.

FIGURE 3.

FIGURE 3

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Interactive principal component analysis and metabolomic analyses of A. flavus WT and ZfpA mutants across three spore densities. (A) Scores plot of the WT, ΔzfpA, and OE::zfpA strains of A. flavus at three different spore densities generated by XCMS v. 3. 7. 1. (B) Volcano plot shows upregulated and downregulated chemical features in the OE::zfpA mutant compared to WT control of 106 spore density cultures. (C) Volcano plot shows upregulated and downregulated chemical features in the ΔzfpA mutant compared to WT control of 106 spore density cultures. The plots were constructed based on Log2 (fold‐change) of intensity with p‐values > 0.05 generated by XCMS at minimum intensity: 5 × 105. The green and red colours indicate statistically significant features with Log2(fold‐change) > 1 and < −1 on x–axis and –Log10(p‐value) > 1.30103 on y–axis.

2.4. Interdependence of ZfpA and Density on Production of Sclerotial and Hyphal Metabolites

Although nearly 100 biosynthetic gene clusters (BGCs) have thus far been identified across 100 s of isolates of the Aspergillus flavus pan‐genome (Calvo and Cary 2015; Cary et al. 2018; Drott et al. 2021; Gangurde et al. 2024; Uka et al. 2019) only approximately a dozen end products (and over 50 known derivatives) have been linked to specific BGCs (Table S1). Among the characterised SMs, a subset is found primarily in sclerotia (cyclopiazonic acids, aflatrems, aflavinines, aflavarin, asparasones, and kotanin) (Calvo and Cary 2015; Cary et al. 2018) The well‐known carcinogen aflatoxin has been found in all A. flavus tissues, dependent on strain (Wicklow and Shotwell 1983). We were interested in investigating the production of known metabolites mediated by density dependence and ZfpA, following the logic that some of the metabolites would be primarily extracted from sclerotial growth and some primarily from conidial/hyphal growth.

Known SMs were suggested based on comparisons of their MS/MS fragmentation patterns with published data (Uka et al. 2019) except aflatoxin B1, which was assessed by a standard purchased commercially (Figures S2 and S3). The height intensity of each putative SM was quantified, and bar graphs of intensity were plotted in Log10 scale (Figure 4). Overall, the production levels of four known sclerotial metabolites—aflavanine, hydroxyaflatrem, oxyasparasone A, and kotanin—decreased with increasing population densities in WT (Figure 4A). We found that ZfpA negatively regulates the production of these sclerotia‐related metabolites (Figure 4B). Specifically, the OE::zfpA strain compared to WT showed a 5 to 20‐fold decrease in the production of these compounds at the low spore density and a nearly 30 to 35‐fold decrease at the intermediate spore density (Figure 4B). At the high spore density, the production of aflavinine and kotanin was undetectable in OE::zfpA (Figure 4B). These findings correlated with sclerotial formation in the three strains and support the spatial linkage of these metabolites to sclerotial formation.

FIGURE 4.

FIGURE 4

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Production of secondary metabolites in A. flavus mediated by density dependence and ZfpA. (A) Scatter plot of production of six SMs mediated by population density dependence in WT in Log10 height intensity. p‐values were calculated using two‐way ANOVA with Tukey's multiple comparisons test. (B) Bar charts of production of six SMs and their molecular structures in WT, ΔzfpA, and OE::zfpA strains presented in Log10 height intensity at three spore densities. p‐values were calculated using two‐way ANOVA with Tukey's multiple comparisons test.

Aside from the noted regulation of sclerotial metabolite synthesis by density and/or ZfpA, most known metabolites were either not detected or not significantly affected by either parameter (e.g., α‐cycloziapionic acid, and ditryptophenaline) (Figure S4). OE::zfpA positively regulated leporin B production at intermediate intensity but not at high spore density (Figure S4). On the other hand, aspergillic acid production was favourably increased at high spore density in WT (Figure 4A) and positively regulated by ZfpA with a 5 to 15‐fold increase (Figure 4B). Aspergillic acid production was largely unaffected in the ΔzfpA strain, suggesting ZfpA is a positive regulator of this metabolite.

Aflatoxin biosynthesis largely followed the same trend as sclerotial metabolite production. Many studies have established a linkage between sclerotia production and A. flavus aflatoxin synthesis, where high sclerotia development correlated with high aflatoxin levels (Zhao et al. 2017; Kale et al. 2008; Duran et al. 2007) although one study supports an inverse under certain conditions (Cho et al. 2022), and these differences might be isolate specific (Wicklow and Shotwell 1983). Overall, UHPLC–MS/MS analysis showed that the production of aflatoxin B1 followed the previously noted pattern of more aflatoxin synthesis in lower spore densities for all three strains, although differences in their synthesis in the mutants led to an unclear regulation pattern by ZfpA (Figure 4).

2.5. ZfpA Upregulates Pathway Analogues of Aspergillic Acid Synthesis

Our metabolomic data also showed that the OE::zfpA strain upregulated other products, especially at 106 spore density (Figure 5A). Given the positive regulation of aspergillic acid by both ZfpA and density dependence, especially in OE::zfpA, we hypothesised that these upregulated peaks could be precursors or alternate end metabolites from the aspergillic acid pathway.

FIGURE 5.

FIGURE 5

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Metabolomic analyses of aspergillic acid related products and metal chelation assessment in WT and A. flavus mutants. (A) Comparison of total ion chromatograms in positive ion mode between WT and mutants at 106 spore density. The OE::zfpA strain shows 11 upregulated features (B) Extracted ion chromatograms (m/z 225.1607) of the OE::zfpA strain at three spore densities. (C) Biosynthetic gene cluster (BGC) and pathway of asa producing aspergillic acid adapted from Lebar et al. (2018). (D) Molecular structures of analogues or derivatives of aspergillic acid. (E) Colony formation of WT and mutant strains grown on GMM with and without Fe supplementation. The OE::zfpA strain produced an orange colony due to ferriaspergillin production. (F) Chrome azurol S (CAS) assay assessing the metal chelation (distance of clearance) in WT and mutant strains. p‐values were calculated using ordinary one‐way ANOVA with Tukey's multiple comparisons test.

The synthesis of aspergillic acid, a hydroxamic acid‐containing pyrazinone, is initiated by the NRPS AsaC that incorporates leucine (Leu) and isoleucine (Ile) to produce Leu‐Ile pyrazinone‐structured compounds (Figure 5C) (Lebar et al. 2022). We observed 11 highly upregulated features, including aspergillic acid from the total ion chromatogram (TIC) under the positive ionisation mode of the OE::zfpA strain grown at 106 spore density (Figure 5A). The precursor ions of these features ranged from m/z 195.1494 to 241.1458 at retention times between 5.40 and 7.88 min (Table 1). Their molecular formulae were proposed using SIRIUS v. 5.7.3 software (Figures S5–S15) based on [M + H]+ ions, within the error range of 5 ppm. Peak 2 was predicted to be aspergillic acid due to its mass fragmentation pattern matching with the previous reports (Uka et al. 2019; Saldan et al. 2018). Peaks 2, 10, and 11 (Figure 5A) shared an identical precursor ion of m/z 225.1607 (Δ = −1.8 ppm), suggesting that aspergillic acid isomers such as deoxyhydroxyaspergillic acid and neoaspergillic acid were possibly produced. These isomers differ by the position of the hydroxyl group or carbon atom arrangement on the butyl groups derived from either Leu or Ile (Lebar et al. 2022). Notably, the intensities of peaks 10 and 11 were increased in OE::zfpA and higher than aspergillic acid as spore density increased (Figure 5B), suggesting that aspergillic acid and derivatives are differentially regulated.

TABLE 1.

Chemical information of 11 upregulated peaks observed in the OE::zfpA strain.

Ion peak R.T. (min) Observed m/z Theoretical m/z Mass error (ppm) Tentative M.W. (Da) Tentative M.F.
1 5.40 241.1556 241.1552 −1.6 240 C12H20N2O3
2 5.63 225.1607 225.1603 −1.8 224 C12H20N2O2
3 5.78 241.1556 241.1552 −1.6 240 C12H20N2O3
4 5.97 207.1504 207.1497 −3.2 206 C12H18N2O
5 6.18 195.1501 195.1497 −1.9 194 C11H18N2O
6 6.46 195.1501 195.1497 −1.9 194 C11H18N2O
7 6.73 211.1451 211.1446 −2.2 210 C11H18N2O2
8 6.86 209.1658 209.1654 −2.0 208 C12H20N2O
9 7.19 211.1442 211.1446 2.1 210 C11H18N2O2
10 7.52 225.1607 225.1603 −1.8 224 C12H20N2O2
11 7.88 225.1607 225.1603 −1.8 224 C12H20N2O2
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Abbreviations: M.F. = molecular formula; M.W. = molecular weight; R.T. = retention time.

Peaks 1, 3, 4, and 8 with precursor ions of m/z 241.1548, 207.1494, and 209.1650 (Figure 5A, Table 1) were predicted to be aspergillic acid derivatives, including hydroxyaspergillic acid, deoxyaspergillic acid, flavacol, and dehydrodeoxyaspergillic acid (Lebar et al. 2018, 2022). Peaks 7 and 9 (m/z 211.1443) suggest a CH2 loss (14 Da) from aspergillic acid (Figure 5A, Table 1). Previous studies identified pyrazinone products with m/z 211.1443 in Aspergillus oryzae A21 (Ueno et al. 1977) or via Val‐feeding in A. flavus (MacDonald 1970), attributed to valine incorporation replacing one isobutyl group with an isopropyl group. Peaks 5 and 6 (m/z 195.1149) suggest Val‐derived products with a 30 Da mass reduction (loss of COH2) relative to aspergillic acid (Figure 5A, Table 1), similar to deoxymutaspergillic acid (Shaala et al. 2016) and leuvalin (Zhu et al. 2024; Zimmermann and Fischbach 2010).

We posited that aspergillic acid and the other 10 upregulated features are produced through the same biosynthetic pathway. To address this hypothesis, we deleted the synthetase gene (asaC) in WT and OE::zfpA backgrounds (TJW368.1 ΔasaC and TJW369.3 OE::zfpA ΔasaC respectively). Metabolomic data comparison between five strains (WT, ΔzfpA, OE::zfpA, OE::zfpA ΔasaC, and ΔasaC) showed aspergillic acid and the 10 features were absent in the asaC deletion strains (Figure 5A), confirming they are generated from the AsaC pathway. Ferriaspergillin (m/z 726.3782, Figure S16), likely contributing to the orange colour (Figure 5E) in the OE::zfpA strain, was observed at 7.52 min but was absent in both asaC deletion strains.

Lastly, we assessed Fe and Cu chelation ability of the strains via the chrome azurol S (CAS) assay based on observations in previous studies (Lebar et al. 2022; Zhu et al. 2011). The OE::zfpA demonstrated both Fe and Cu chelation (Figures 5F and S17). Interestingly, the metabolomic analysis also showed that leporin A production increased in the OE::zfpa ΔasaC and ΔasaC strains compared to the other three strains (Figure S18). As leporins are known to chelate Fe (and presumably Cu) (Cary et al. 2018, 2015), this finding might explain why both asaC deletion strains have increased metal chelation over the WT and ΔzfpA strain.

3. Discussion

Oxylipins are a broad class of signalling molecules known to direct developmental processes in an expansive range of eukaryotes and prokaryotes (Gessler et al. 2017; Pohl and Kock 2014) as well as direct pathogenic and symbiotic interactions across Kingdoms (Erb‐Downward and Huffnagle 2006; Beccaccioli et al. 2022). In filamentous fungi, several oxylipins are involved in directing differentiation processes including spore germination, hyphal branching, and balancing asexual versus sexual development (Niu et al. 2020; Calise et al. 2024; Tsitsigiannis et al. 2004). Studies of A. flavus have shown that oxylipins and their producing oxygenases drive a quorum‐like sensing programme that promotes sexual (sclerotia) development at low population densities and asexual (conidia) production at high population densities (Brown et al. 2009; Horowitz Brown et al. 2008). Here, we present the involvement of the oxylipin responsive zinc finger protein ZfpA in facilitating this programme in A. flavus .

ZfpA was identified in A. fumigatus as the transcription factor mediating the hyperbranching oxylipin signal 5,8‐diHODE generated by the oxygenase PpoA (Niu et al. 2020; Calise et al. 2024). ZfpA orchestrates chitin deposition, hyperbranching, septation, virulence, and resistance to echinocandins in A. fumigatus (Niu et al. 2020; Calise et al. 2024; Schoen et al. 2023). ZfpA activity is largely dependent on 5,8‐diHODE signalling in WT A. fumigatus , but an OE::zfpA strain supersedes the oxylipin signal and presents a locked‐in hyperbranching phenotype in A. fumigatus. Here, we find a similar function for ZfpA in A. flavus where OE::zfpA leads to a hyperbranching phenotype (Figure 1E). We also confirm the production of the PpoA oxylipin 5,8‐diHODE during A. flavus development, which elicits the ZfpA‐mediated hyperbranching response, and find the OE::zfpA strain produces high concentrations of 5,8‐diHODE (Figure 1D,F).

Because oxylipin production is tied in with a density‐dependent quorum sensing programme in A. flavus , we examined the hypothesis that loss of or overexpression of zfpA could impact the density‐dependent morphological transition from sclerotia to conidia in this species. We found that ZfpA overexpression promotes early conidia development at lower spore population densities than either the WT or zfpA deletion mutant (Figure 2). We speculate that the increase in 5,8‐diHODE production in the overexpression mutant may play a signalling activity that promotes conidiation and/or represses sclerotial synthesis, a hypothesis that may explain some of the observations made in previous oxygenase deletion studies in A. flavus (Brown et al. 2009; Horowitz Brown et al. 2008).

An outstanding contribution of this work is that not only morphology, but also secondary metabolite synthesis is regulated by population density and ZfpA activity. One characteristic of fungal SMs is the localization and synthesis of specific metabolites in distinct tissues in the fungus, including asexual spores, conidiophores, sexual spores, fruiting bodies, hyphae and other specialised structures such as appressoria and/or chlamydospores (Lim and Keller 2014; Blachowicz et al. 2020; Spraker et al. 2018; Studt et al. 2012). A. flavus is well known for SMs that predominately are synthesised in sclerotia and are thought to protect the sclerotia from insect predators or abiotic stressors (Calvo and Cary 2015; TePaske et al. 1992; Cary et al. 2018). Asparasone A and its known analogues give dark brown colour pigmentation to sclerotia and provide protection against UV and heat stress (Calvo and Cary 2015; Cary et al. 2014). Inactivation of the asparasone A polyketide synthase pks27 causes sclerotia to turn greyish yellow and decreases sclerotia viability (Cary et al. 2014). In this study, we measured putative oxyasparasone A and found its production was increased when sclerotia formation was favourable (Figure 4). Likewise, we also found increased production of sclerotial SMs such as kotanin, hydroxyaflatrem (an aflatrem derivative), and aflavinine (Figure 4). Aflatrem and aflavinine are both associated with sclerotia and have been shown to be protective against insect predation due to their toxic and anti‐insectan activities (Gloer et al. 1988; Gallagher and Wilson 1979). Like asparasone, kotanin is also a polyketide, and its role in A. flavus may also be associated with anti‐insectan properties (Buechi et al. 1971). Our current study provides evidence of the co‐regulation of sclerotial development with kotanin production, possibly suggesting a protective role of this metabolite in A. flavus ecology as well.

Aflatoxin B1 production has been associated with low density populations where it was assumed that increased numbers of sclerotia increase toxin production (Brown et al. 2009) although some studies have not found a correlation between sclerotia formation and aflatoxin synthesis (Cho et al. 2022; Bennett et al. 1979). We found production at all densities but at much lower levels at the high densities, particularly in the OE::zfpA strain (Figure 4). However, interpretation of how ZfpA may regulate aflatoxin was not clear, as despite producing less sclerotia in low densities, aflatoxin synthesis was high in the overexpression mutant in low densities, which we speculate may be associated with stress response pathway activation, which has been associated with ZfpA regulation (Niu et al. 2020; Schoen et al. 2023).

In contrast to the sclerotial pigments, the OE::zfpA strain greatly increased production of putative aspergillic acid and related pathway metabolites, and this production was correlated with increased asexual spore production. The hydroxamic acid group in aspergillic acid and its hydroxy‐analogue permits Fe binding and the formation of ferriaspergillin, with some experimentation suggesting they function as siderophores (Haas 2014). Other SMs in Aspergillus species containing a hydroxamic acid moiety are known siderophores, such as triacetylfusarinine C produced by A. fumigatus and A. nidulans (Aguiar et al. 2022; Haas et al. 2003). Fungi utilise siderophores for iron acquisition, transport, and storage, essential for fungal development and providing microbial defence and virulence in plant and animal hosts, and success in environmental competition with other microbes (Haas 2014; Patil et al. 2021; Johnson 2008). We found that both Fe and Cu chelation was increased in the OE::zfpA strain (Figure 5F) and venture that high aspergillic acid content in spores could increase their ecological fitness, especially in regards to competition with other microbes in scarce metal environments. It is not clear if aspergillic acid is important for virulence, as although two studies have shown an increase in gene expression level of asaC during maize kernel infection by A. flavus , no aspergillic acid or its analogues were detected in infected live maize kernels (Lebar et al. 2018; Dolezal et al. 2013).

Regardless of function, the high production of aspergillic acid and its related products enabled us to suggest its derivatives included likely Val‐derived analogues from the same pathway (Figure 5A, Table 1). Lebar et al. (2022) suggested that AsaC in different Aspergillus species could selectively activate different amino acids to generate species‐specific pyrazinone products. A. flavus CA14 (section Flavi) is capable of producing neoaspergillic acid and its precursor flavacol when the A. flavus asaC gene was replaced with Aspergillus sclerotiorum NRRL 415 (section Circumdati) asaC (Lebar et al. 2022). The Circumdati AsaC orthologs exhibit an average pairwise amino acid sequence identity of 71.6% in comparison to A. flavus . To further illustrate the potential flexibility of AsaC amino acid preference, another study showed that various pyrazinone products related to aspergillic acid, including Val‐derived products, were biosynthesized in A. flavus PRL 932 when the growth media were supplemented with specific amino acids (MacDonald 1970). These findings are in line with our data showing the potential of AsaC synthesising varying aspergillic acid related pyrazinone products.

In conclusion, our study elucidates a pivotal role of ZfpA in orchestrating morphological and chemical transitions in A. flavus . We demonstrated that ZfpA influences secondary branching, reaffirming the conserved role of ZfpA in governing branching response in Aspergilli. Moreover, our results confirmed that the PpoA oxylipin 5,8‐diHODE is produced during A. flavus development and triggers a ZfpA‐mediated hyperbranching response in a dose‐dependent manner. Further investigation revealed the involvement of ZfpA in coordinating density‐dependent morphological transitions. Additionally, we have established that SM production is impacted by ZfpA expression in a density‐dependent manner, linking specific metabolites to sclerotial development and asexual spore production. Overall, our study sheds light on the multifaceted role of ZfpA in A. flavus biology, providing valuable insights into the fungal quorum networks linking fungal development to tissue‐specific SM pathways.

4. Materials and Methods

4.1. Fungal Strains and Growth Conditions

The A. flavus strains used and generated in this study are shown in Table S2. Aspergillus flavus conidial stocks were maintained at −80°C in glycerol suspension until being streaked on media plates. All strains employed for physiological examinations were prototrophic and cultured at 30°C on glucose minimal medium (GMM) (Shimizu and Keller 2001) unless specified otherwise. The cultures were cultivated in the absence of light to allow normal sclerotial formation.

4.2. Strain Construction

To make a pyrG auxotroph of the OE::zfpA strain, the pyrG marker was removed from the OE::zfpA strain by transforming TJW321.4 with a pyrG recycle construct. This construct consisted of two 1 kb DNA fragments from the upstream and downstream regions of the pyrG marker inserted in OE::zfpA. Transformants were grown on GMM containing 5‐Fluoroorotic Acid (1 mg/mL) and uracil/uridine. pyrG deleted OE::zfpA strains were confirmed by Southern blotting with both the P‐32 labelled 5' and 3' flanks of the pyrG recycle construct after digest with NcoI (TJW361.3, Figure S19).

For asaC deletion strains, two 1 kb DNA fragments immediately upstream and downstream of the asaC open reading frame (ORF) were amplified by PCR from A. flavus NRRL3357 genomic DNA and were fused to a 2 kb A. parasiticus pyrG fragment from pJW24 (Calvo et al. 2004) using double joint PCR (Bok et al. 2013). Fungal transformation of A. flavus TJES19.1 was performed following previously described approaches (Bok and Keller 2004). Transformants were confirmed for targeted replacement of the native locus by PCR (data not shown) and Southern blotting using PvuI digest for asaC with P‐32 labelled 5' and 3' flanks of each knock out construct to obtain TJW368.1 (Figure S20, left panel).

To create the double mutant OE::zfpA ΔasaC, TJW361.3 was transformed with an asaC deletion construct, and the double mutant TJW369.3 was confirmed by southern blotting as shown by asaC deletion confirmation (Figure S20, right panel). All of the primers for this study are listed in Table S3. DNA extraction, restriction enzyme digestion, gel electrophoresis, blotting, hybridization, and probe preparation were performed by standard methods (Sambrook 1989).

4.3. Assay for Conidia and Sclerotia Numeration

For density‐dependent conidia and sclerotia numeration, spores from the WT, OE::zfpA, and ΔzfpA strains were grown at three initial spore numbers per each plate: 102 spores, 104 spores, and 106 spores. Four replicate plates for each mutant density were prepared, totaling 36 plates for all three mutants. Each plate contained 8 mL of 1.6% agar GMM plus 2% sorbitol overlaid with 3 mL of 0.7% agar GMM plus 2% sorbitol containing the appropriate spore density. The plates were incubated at 30°C under continuous dark conditions for 7 days. Plugs of 1.5 cm in diameter were taken daily from both sets of plates and were homogenised in 5 mL of water containing 0.01% Tween 80. The suspensions were diluted in a 1:10 ratio in microcentrifuge tubes and counted with a spore counter (Cellometer X2, Nexcelom Bioscience LLC) and analysed using the software, Cellometer image cytometer version 3.2.1 to determine sporulation. Separate sets of culture plates were prepared for accurate sclerotia counts. To visualise the sclerotia clearly, the plates were sprayed thoroughly with 70% ethanol to remove conidia and aerial mycelia. The exposed sclerotia were collected, lyophilised, and weighed to determine sclerotia production (mg [dry weight] per plate). The same procedure was used for conidia count to examine the correlation between density‐dependent growth and 5,8‐diHODE production in the mutants.

4.4. Hyphal‐Branching Assessment

To evaluate hyphal branching using a 96‐well setup, approximately 500 conidia in 100 μL GMM with the designated treatment were inoculated into triplicate wells of the plate. When oxylipin treatment was involved, spores were treated with the desired concentration of 5,8‐diHODE with EtOH serving as the vehicle control before inoculation. The plate was incubated on a Nikon Eclipse Ti Inverted Microscope in a heated microscope enclosure (OKO Labs, Burlingame, CA) at 30°C for 15 h before imaging. Microscope frames were set on four germlings per well, and images were acquired every 15 min for 6 h using a Nikon Plan Fluor 20X Ph1 DLL objective out to 22 h post inoculation. Lateral branching of eight hyphae per condition at 20 h post inoculation was quantified manually using the NIS‐Elements AR Software package (Version 5.30) to measure the distance between each lateral branch and dividing the total distance from the first to last branch by the number of branches. The length of the leading hypha and the number of primary lateral branches per leading hypha were quantified and normalised to 100 μm per hypha.

4.5. RNA Extraction and Northern Blotting

Conidia were inoculated at 106 spores/mL into 50 mL GMM and incubated at 37°C and 250 rpm for 18 h. Tissue was collected into sterile Miracloth, flash frozen, and lyophilized. Total RNA was extracted using QIAzol lysis reagent (Qiagen) per manufacturer's protocol with the addition of a phenol:chloroform:isoamyl alcohol (25:24:1) extraction step before RNA precipitation. RNA purity and concentration were assessed by nanodrop. Approximately 12 μg of each sample was run in a 1.3% agarose 1.5% formaldehyde gel and transferred to an Amersham Hybond N+ Membrane. Membranes were hybridised with dCTP‐αP32 labelled probes complementary to a 1 kb region of the gene of interest lacking any predicted introns.

4.6. Microscopic Examination of Sclerotia

To visualise sclerotia growth, fungal strains were cultured on GMM plates with a spore density of 102 and incubated in the dark at 30°C. Sorbitol was added to the media to induce sclerotia production. After 7 days, the culture plates were examined under an Olympus dissecting microscope (Leeds Precision Instruments Inc., MN) and images of the plates were captured.

4.7. Oxylipin and Secondary Metabolite Extractions

For the extraction of 5,8‐diHODE from the NRRL3357 strain, a volume of 50 mL of GMM was inoculated with spores at a concentration of 106 spores/mL. The inoculated medium underwent shaking at 150 rpm for 3 days in darkness at a temperature of 30°C. Following the incubation period, the culture was filtered through a Mira cloth to separate fungal tissue from supernatant. The tissue was then transferred into a 50 mL tube, to which approximately 15 mL of distilled water was added for homogenisation. The homogenised mixture was transferred into a glass bottle and 50 mL of ethyl acetate was added. Similarly, the supernatant of the culture was transferred into a glass bottle, and 100 mL of ethyl acetate was added and left to stand overnight. The ethyl acetate layers were separated using a separation funnel and evaporated using a Buchi Rotovap R‐210. The crude extracts obtained were then resuspended using 2–3 mL of 100% methanol (Sigma‐Aldrich, catalogue number: 34860–4L‐R) and transferred into pre‐measured glass vials. Methanol was evaporated and the dried weight of the crude extracts for all replicates was measured, and each sample was dissolved to 1 mg/mL concentration with HPLC‐grade methanol prior to UHPLC–HRMS analysis.

For density‐dependent 5,8‐diHODE and secondary metabolite extractions, different initial spore concentrations of mutants were grown using 60 × 15 mm petri dishes under the same conditions as described earlier for density‐dependent conidial/sclerotial counts. The cultures were incubated at 30°C. After 7 days of incubation, the whole plate culture was homogenised in 15 mL of distilled water, and the homogenate mixture was transferred into glass bottles. Crude extracts were obtained using the same procedure outlined above. The extracts from 7 days growth cultures were subsequently utilised for UHPLC–HRMS analysis of secondary metabolites.

4.8. UHPLC–HRMS Analysis

UHPLC–HRMS was performed on a Thermo Fisher Scientific‐Vanquish UHPLC system coupled with a Thermo Q‐Exactive HF hybrid quadrupole‐orbitrap high‐resolution mass spectrometer equipped with a HESI ion source (Thermo Fisher Scientific, Waltham, MA, USA). Each spectrum was obtained in both negative and positive ionisation modes using an m/z range of 100 to 1500. A Waters XBridge BEH‐C18 column (2.1 × 100 mm, 1.7 μm) (Waters, Milford, MA, USA) was used with 0.05% formic acid in acetonitrile (organic phase) and 0.05% formic acid in water (aqueous phase) as solvents at a flow rate of 0.2 mL/min. Ten μL of samples was injected. A 20‐min reverse‐phase gradient condition was used, starting at 5% organic with a linear increase to 98% for 10 min, holding at 98% organic for 5 min, decreasing back, and holding at 5% organic for 5 min for a total of 20 min. For feature detection and characterisation, UHPLC–MS/MS RAW files were converted to mzXML format (centroid mode) using RawConverter (v.1.2.0.1) (The Scripps Research Institute, San Diego, CA, USA) followed by analysis using the MAVEN2 software (Calico Life Sciences, South San Francisco, CA, USA) (Seitzer et al. 2022).

4.9. Statistical Analysis

All statistical analyses were performed using GraphPad Prism version 10.2.0 software. Statistical analysis was performed using one‐way or two‐way ANOVA with the Tukey's multiple comparisons test when comparing mean values of quantities to each other and when comparing quantities at different spore densities. A p‐value of 0.05 or lower was considered statistically significant.

4.10. PCA and Volcano Plots

Interactive PCA was performed by XCMS v. 3.7.1 software (Gowda et al. 2014) based on the triplicate analyses in positive ion mode on the Thermo Q‐Exactive system. The loadings threshold was set to 1000 and the Log scaling option was chosen to plot the PCA. Volcano plots were constructed based on the comparison of chemical features between mutant and WT strains by XCMS v. 3.7.1 software in triplicates (Gowda et al. 2014). The intensity threshold was set to 5 × 105, removing 20,479 and 16,382 features from OE::zfpA and ΔzfpA strains, respectively. From 15,203 significant features in the ΔzfpA strain, 1201 of them were upregulated and 1702 were downregulated. In the OE::zfpA strain, 2132 features were upregulated and 3633 features were downregulated from the total of 14,116 significant features.

4.11. CAS Assay

CAS assay plates were prepared as described previously (Raffa et al. 2021; Yoon et al. 2010). A sterile razor was used to remove half of the CAS media, and 10 mL of warm molten GMM was then inoculated with 1 × 106 spores/plate and poured into the vacant space. The plates were then allowed to solidify and incubated for 5 days at 30°C in the dark. The zone of activity was quantified in triplicates by measuring the distance of clearance between the growing mycelia and the edge of the Fe–CAS or Cu–CAS complex with a ruler. The distance of clearance is defined as the distance in colour change due to Fe/Cu scavenging resulting in the release of the CAS dye from blue to orange.

Author Contributions

Benjamin Otoo: data curation, formal analysis, investigation, writing – original draft, writing – review and editing, validation, visualization. Dante G. Calise: investigation, validation, writing – review and editing, data curation. Sung Chul Park: investigation, validation, writing – review and editing, data curation. Jin Woo Bok: investigation, validation, writing – review and editing. Nancy P. Keller: conceptualization, funding acquisition, methodology, supervision, writing – review and editing. Mira Syahfriena Amir Rawa: conceptualization, data curation, investigation, formal analysis, validation, writing – review and editing, writing – original draft, visualization.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Table S1. List of A. flavus known secondary metabolites linked to known BGCs assessed in this study.

Table S2. Strains of A. flavus used in this study.

Table S3. Primers used to construct strains in this study.

Figure S1. ZfpA mutant germination.

Figure S2. MS/MS spectra of aflavinine, hydroxyaflatrem, oxyasparasone A, and kotanin from this study.

Figure S3. MS/MS spectra of aflatoxin B1, ditryptophenaline, α‐cyclopiazonic acid, and leporin B from this study.

Figure S4. Graph bars of production of ditryptophenaline, α‐cyclopiazonic acid, and leporin B at three different population densities in WT, ΔzfpA, and OE::zfpA strains in Log10 height intensity.

Figure S5. Tentative molecular formula of precursor ion peak 1 and its MS/MS spectrum.

Figure S6. Tentative molecular formula of precursor ion peak 2 and its MS/MS spectrum.

Figure S7. Tentative molecular formula of precursor ion peak 3 and its MS/MS spectrum.

Figure S8. Tentative molecular formula of precursor ion peak 4 and its MS/MS spectrum.

Figure S9. Tentative molecular formula of precursor ion peak 5 and its MS/MS spectrum.

Figure S10. Tentative molecular formula of precursor ion peak 6 and its MS/MS spectrum.

Figure S11. Tentative molecular formula of precursor ion peak 7 and its MS/MS spectrum.

Figure S12. Tentative molecular formula of precursor ion peak 8 and its MS/MS spectrum.

Figure S13. Tentative molecular formula of precursor ion peak 9 and its MS/MS spectrum.

Figure S14. Tentative molecular formula of precursor ion peak 10 and its MS/MS spectrum.

Figure S15. Tentative molecular formula of precursor ion peak 11 and its MS/MS spectrum.

Figure S16. MS/MS spectrum of ferriaspergillin.

Figure S17. Chrome azurol S (CAS) assay assessing the metal chelation (distance of clearance) in WT, ΔzfpA, OE::zfpA, and OE::zfpA ΔasaC, and ΔasaC strains in triplicates.

Figure S18. Bar chart of Leporin A production in the WT, ΔzfpA, OE::zfpA, OE::zfpA ΔasaC, and ΔasaC strains.

Figure S19. Southern confirmation of pyrG recycled OE::zfpA mutants in A. flavus .

Figure S20. Southern confirmation of ΔasaC (TJW368) and OE::zfpA ΔasaC mutants (TJW369) in A. flavus.

EMI-27-e70100-s001.docx (74MB, docx)

Acknowledgements

We thank all members of the Keller lab for their support and their helpful discussions of the research results and manuscript. This study was supported by the National Institutes of Health R01 1R01AI150669‐01A1 and Hatch act fund WIS03041 awarded to Nancy P. Keller for salary support for Benjamin Otoo, Jin Woo Bok, Dante Calise, and Sung Chul Park, and NIH 5R01AT009143‐20 awarded to Nancy P. Keller for salary support for Mira Syahfriena Amir Rawa.

Funding: This work was supported by National Institutes of Health 1R01AI150669‐01A1 and 5R01AT009143‐20; Hatch Act Fund WIS03041.

Contributor Information

Nancy P. Keller, Email: npkeller@wisc.edu.

Mira Syahfriena Amir Rawa, Email: bintiamirraw@wisc.edu.

Data Availability Statement

The data that supports the findings of this study are available in the Supporting Information of this article.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Table S1. List of A. flavus known secondary metabolites linked to known BGCs assessed in this study.

Table S2. Strains of A. flavus used in this study.

Table S3. Primers used to construct strains in this study.

Figure S1. ZfpA mutant germination.

Figure S2. MS/MS spectra of aflavinine, hydroxyaflatrem, oxyasparasone A, and kotanin from this study.

Figure S3. MS/MS spectra of aflatoxin B1, ditryptophenaline, α‐cyclopiazonic acid, and leporin B from this study.

Figure S4. Graph bars of production of ditryptophenaline, α‐cyclopiazonic acid, and leporin B at three different population densities in WT, ΔzfpA, and OE::zfpA strains in Log10 height intensity.

Figure S5. Tentative molecular formula of precursor ion peak 1 and its MS/MS spectrum.

Figure S6. Tentative molecular formula of precursor ion peak 2 and its MS/MS spectrum.

Figure S7. Tentative molecular formula of precursor ion peak 3 and its MS/MS spectrum.

Figure S8. Tentative molecular formula of precursor ion peak 4 and its MS/MS spectrum.

Figure S9. Tentative molecular formula of precursor ion peak 5 and its MS/MS spectrum.

Figure S10. Tentative molecular formula of precursor ion peak 6 and its MS/MS spectrum.

Figure S11. Tentative molecular formula of precursor ion peak 7 and its MS/MS spectrum.

Figure S12. Tentative molecular formula of precursor ion peak 8 and its MS/MS spectrum.

Figure S13. Tentative molecular formula of precursor ion peak 9 and its MS/MS spectrum.

Figure S14. Tentative molecular formula of precursor ion peak 10 and its MS/MS spectrum.

Figure S15. Tentative molecular formula of precursor ion peak 11 and its MS/MS spectrum.

Figure S16. MS/MS spectrum of ferriaspergillin.

Figure S17. Chrome azurol S (CAS) assay assessing the metal chelation (distance of clearance) in WT, ΔzfpA, OE::zfpA, and OE::zfpA ΔasaC, and ΔasaC strains in triplicates.

Figure S18. Bar chart of Leporin A production in the WT, ΔzfpA, OE::zfpA, OE::zfpA ΔasaC, and ΔasaC strains.

Figure S19. Southern confirmation of pyrG recycled OE::zfpA mutants in A. flavus .

Figure S20. Southern confirmation of ΔasaC (TJW368) and OE::zfpA ΔasaC mutants (TJW369) in A. flavus.

EMI-27-e70100-s001.docx (74MB, docx)

Data Availability Statement

The data that supports the findings of this study are available in the Supporting Information of this article.

Articles from Environmental Microbiology are provided here courtesy of Wiley

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