Role of the osaA Transcription Factor Gene in Development, Secondary Metabolism and Virulence in the Mycotoxigenic Fungus Aspergillus flavus

TL;DR

This study explores the role of the osaA gene in Aspergillus flavus, revealing its impact on fungal development, toxin production, and pathogenicity.

Contribution

The study identifies osaA as a key regulator of aflatoxin production, development, and virulence in Aspergillus flavus.

Findings

Deletion of osaA reduces aflatoxin B1 and cyclopiazonic acid production.

osaA is essential for normal seed colonization and fungal burden in corn kernels.

osaA influences cell-wall chitin content and oxidative stress sensitivity.

Abstract

Aspergillus flavus colonizes oil-seed crops, contaminating them with aflatoxins; highly carcinogenic mycotoxins that cause severe health and economic losses. Genetic studies may reveal new targets for effective control strategies. Here, we characterized a putative WOPR transcription factor gene, osaA, in A. flavus. Our results revealed that osaA regulates conidiation and sclerotial formation. Importantly, deletion of osaA reduces aflatoxin B1 production, while, unexpectedly, transcriptome analysis indicated upregulation of aflatoxin biosynthetic genes, suggesting post-transcriptional or cofactor-mediated regulation. Cyclopiazonic acid production also decreased in the absence of osaA. In addition, the osaA mutant exhibited upregulation of genes in the imizoquin and aspirochlorine clusters. Moreover, osaA is indispensable for normal seed colonization; deletion of osaA significantly reduced fungal burden in corn kernels. Aflatoxin content in seeds also decreased in the absence of osaA. Furthermore, deletion of osaA caused a reduction in cell-wall chitin content, as well as alterations in oxidative stress sensitivity, which could in part contribute to the observed reduction in pathogenicity. Additionally, promoter analysis of osaA-dependent genes indicated potential interactions with stress-responsive regulators, indicated by an enrichment in Sko1 and Cst6 binding motifs. Understanding the osaA regulatory scope provides insight into fungal biology and identifies potential targets for controlling aflatoxin contamination and pathogenicity.

Figures

Full text

1. Introduction

Aspergillus flavus is a ubiquitous filamentous fungus present in soil and decaying organic matter. It disseminates primarily through the formation of asexual spores called conidia, which are carried by wind, insects, and rain [1,2]. These conidia are highly adaptable and capable of germinating under a wide range of environmental conditions, and the resulting mycelial growth has been reported across temperatures spanning from 12 °C to 50 °C [3]. In addition to conidia, A. flavus produces sclerotia to persist in adverse environmental conditions [1,4]. Both conidia and sclerotia are important structures contributing to fungal dissemination and resilience [5].

From an ecological and agricultural perspective, A. flavus is particularly notorious as an opportunistic plant pathogen that colonizes oilseed crops such as corn, peanuts, cotton, sorghum, and tree nuts during both pre- and post-harvest [6,7]. In peanuts and cotton, dispersal occurs mainly through rain and soil, while in corn, infection typically takes place via silk colonization and kernel invasion by fungal mycelia [1]. The pathogen’s broad host range, environmental adaptability, and persistence mechanisms make it a formidable challenge for agricultural systems worldwide.

In addition to its agricultural impact, A. flavus poses a serious clinical threat, especially in immunocompromised individuals. It is the second leading cause of invasive aspergillosis (IA) after A. fumigatus. A. fumigatus is classified as a Risk Group 2 (RG-2) organism because it can cause opportunistic infections in humans, especially in immuno-compromised individuals [8]. A. flavus also contributes to pulmonary infections and systemic mycoses in patients undergoing chemotherapy, organ transplantation, or corticosteroid therapy [2]. This dual threat to plants and human health underscores the urgency of developing effective control strategies.

A major concern associated with A. flavus plant colonization is its ability to synthesize aflatoxins. Of these, aflatoxin B_1_ (AFB_1_) is the most toxic and has been classified by the World Health Organization as a Group 1A human carcinogen [9]. In addition to aflatoxins, A. flavus produces other secondary metabolites such as cyclopiazonic acid (CPA) and aflatrem [10,11]. Aflatoxin B_1_ (AFB_1_) is found contaminating crops for human consumption as well as in animal feed, and consequently in animal-derived products, posing a serious threat to both animal and human health [9]. In humans, aflatoxin-contaminated food can cause acute symptoms such as jaundice, abdominal pain, and liver failure, with outbreaks reaching fatality rates up to 40% [12,13]. Chronic exposure leads to immune suppression, growth stunting, and higher risks of cancers, particularly hepatocellular carcinoma, and increases susceptibility to infectious diseases like HIV/AIDS [12,14,15,16].

Beyond its severe health consequences, aflatoxin contamination also leads to significant economic losses, particularly in the United States and other developed countries. In the United States alone, aflatoxin contamination of corn results in annual losses ranging from USD $52 million to$1.68 billion, especially during years of drought and high heat [17]. The U.S. peanut industry lost 52.28 per metric ton in 2019, the worst aflatoxin year in recent history [[18](#B18-toxins-18-00023)]. According to the Farm Progress report, aflatoxin costs the U.S. peanut industry as much as 126 million each year [19]. Developed countries impose stringent regulatory limits on aflatoxin levels in food and feed products. While essential for safeguarding public health, these regulations can result in substantial economic losses when contamination exceeds the allowable thresholds [4]. Globally, it is estimated that approximately 25% of the world’s food crops are lost each year due to aflatoxin contamination [20]. Additional economic burdens arise from rejected agricultural exports, livestock losses, and the costs of testing and monitoring for aflatoxins [6]. Aflatoxin contamination costs Africa over USD 750 million annually, while compliance with EU aflatoxin regulations is estimated to cost food exporters USD 670 million each year, highlighting the substantial economic burden [21]. In contrast, many developing nations lack regulatory frameworks or effective enforcement mechanisms, resulting in higher rates of aflatoxin exposure. As a consequence, an estimated 4.5 billion people worldwide are at risk of chronic aflatoxin ingestion [22].

Efforts to manage A. flavus colonization and toxin contamination have met limited success. Current strategies, including chemical fungicides (e.g., azoles), biological control, and breeding for host resistance, often fall short due to variability in efficacy, environmental resistance, and evolving fungal populations [23]. Notably, the widespread use of azole fungicides in agriculture is believed to contribute to the emergence of azole-resistant fungal strains, thereby compromising both crop protection and clinical antifungal therapy [24]. These limitations necessitate the discovery of novel genetic targets and regulatory mechanisms to control the growth, development, mycotoxin production, and pathogenicity of A. flavus. A promising approach is the exploration of transcriptional regulators that could control developmental and metabolic pathways and possibly virulence. Among these, Gti1/Pac2 domain (also known as the WOPR domain) containing proteins are highly conserved fungal-specific transcriptional regulators that play a central role in controlling cell morphology, development, secondary metabolite production, and pathogenesis across diverse fungal species [25,26,27]. The WOPR domain has been shown to be important for DNA–protein interactions and contains two globular domains that are referred to as WOPRa and WOPRb [27,28]. This class of transcriptional regulators is typically separated into two main groups in fungi, which are named Gti1/Wor1/Ryp1/Mit1/Rge1/Ros1/Sge1 or Pac2, with several members of these groups having been functionally characterized in a variety of fungal species [27]. The name “WOPR” is derived from a group of functionally related transcription factors: Wor1 in Candida albicans, Pac2 in Schizosaccharomyces pombe, and Ryp1 in Histoplasma capsulatum, each of which is critical for development and virulence [26]. Candida albicans, Wor1 orchestrates the white-to-opaque phenotypic switch, a key process that regulates mating competence and influences tissue specificity during infection [29]. In S. pombe, Pac2 is thought to mediate nutritional signaling that activates ste11, a transcription factor essential for sexual development, whereas Gti1 was shown to function downstream of the Wis1-Sty1 MAPK pathway and to be essential for gluconate absorption [30,31]. In H. capsulatum, Ryp1 governs the dimorphic switch between yeast and mycelial forms and regulates the expression of hundreds of genes required for pathogenicity [32]. Notably, the best-characterized WOPR protein, Wor1 from C. albicans, binds to a conserved 9-nucleotide DNA motif (TTAAAGTTT). This motif is also recognized by its orthologs Ryp1 in H. capsulatum and Mit1 in Saccharomyces cerevisiae, reflecting the remarkable evolutionary conservation of WOPR domain–DNA interactions over an estimated 600 million to 1.2 billion years of fungal divergence [26,33].

In the model fungus Aspergillus nidulans, OsaA, a predicted WOPR domain-containing protein, was identified as a downstream regulator of the global regulator VeA, a member of the velvet complex that governs fungal development and secondary metabolism. OsaA contributes to the balance between asexual and sexual morphogenesis [34]. A more recent study in A. fumigatus demonstrated that the OsaA homolog plays a crucial role in regulating colony growth, conidiation, secondary metabolite production, and virulence in mouse models [35]. OsaA-deficient strains in A. fumigatus also showed impaired thermotolerance, reduced cell wall stability, and hypersensitivity to oxidative stress, traits essential for survival and infection [35].

Interestingly, A. flavus possesses an OsaA homolog that shares 68% sequence identity with its A. nidulans counterpart [34], but its role in this agriculturally relevant species remains uncharacterized. Given the established role of Gti1/Pac2 (WOPR) proteins in other fungi, we hypothesized that osaA functions as a major regulator in A. flavus, integrating development, mycotoxin biosynthesis, and virulence.

In the present study, we used a combination of genetic and transcriptomic analyses, as well as biochemical approaches and seed colonization bioassays to characterize the function of osaA in A. flavus. Our findings establish osaA as an important regulatory node in this fungus and highlight its potential as a molecular target for controlling fungal growth, aflatoxin production, and virulence, offering new avenues for integrated disease and toxin management in agriculture and health.

2. Results

2.1. OsaA in A. flavus Contains a Gti/Pac2 (WOPR) Domain Which Is Conserved with Orthologs in Other Aspergillus Species and Other Species in the Ascomycota phylum

The amino acid sequence of the A. flavus OsaA was obtained from FungiDB (accession number: F9C07_2071416). OsaA orthologous proteins were found by using NCBI BLASTP (version 2.17.0). Our phylogenetic analysis revealed that OsaA in Aspergillus spp. is conserved with other Gti1/Pac2 (WOPR) proteins in fungi but is most closely related to Gti1 proteins (Figure 1). Additionally, A. flavus OsaA and orthologous proteins contain a conserved WOPR domain in their N-terminal region. WOPR-domain proteins are a fungal-specific family of transcriptional factors that are involved in multiple biological processes [25]. Specifically, the predicted OsaA amino acid sequence contains the two conserved WOPR subdomains (12–87 residues and 166–187 residues), separated by a characteristic less conserved linker (Figure S1).

2.2. osaA Regulates Development in A. flavus

Recently, it was shown that osaA plays a significant role in the morphogenesis of the opportunistic human pathogen A. fumigatus [35]. Moreover, in the model fungus A. nidulans, osaA regulates development primarily by suppressing the formation of sexual structures [34]. To understand the role of osaA in the agriculturally relevant fungus A. flavus, osaA deletion strain (TMR1.1) and osaA complementation strain (TFEH 8.1) were constructed by previously described methods [36,37]. The osaA deletion strain (ΔosaA), where the osaA gene was replaced with the pyrG marker, was confirmed by PCR, yielding the expected 3.37 kb PCR product (Figure 2A,C). The complementation strain (Com), where the osaA allele was reintroduced into the osaA mutant, was also confirmed by PCR, obtaining a 3.304 kb target size DNA band (Figure 2B,D). Expression analysis of osaA in the wild type, deletion, and complementation strains was carried out by qRT-PCR. As expected, the ΔosaA lost osaA expression, while the complementation strain presented osaA expression levels similar to those of the wild type (Figure 2E).

In the present study, we observed that colony diameter was reduced (14.11%) in ΔosaA compared to the wild-type control on day 7 (Figure 3A,B), indicating that osaA is required for normal growth in A. flavus. Our study also showed that conidial production was significantly increased in the osaA deletion strain compared to the control (Figure 3C), revealing the role of osaA as a repressor of conidiation in this fungus.

In addition, osaA also influences sclerotial formation in A. flavus. Specifically, we found that osaA is essential for normal sclerotial development. In point-inoculated cultures, osaA deletion strains produced significantly less sclerotia than the controls on day 14, while in top-agar cultures, where crowdedness is a factor, sclerotial formation was completely abolished in the mutant (Figure 4).

2.3. Secondary Metabolism Is Influenced by osaA

Development and secondary metabolite production are genetically linked processes in Aspergillus species [5]. In A. fumigatus, secondary metabolite production was influenced by osaA [35]. Therefore, it is possible that osaA also regulates the production of secondary metabolites in A. flavus. In this study, we evaluated the role of osaA in regulating biosynthesis of aflatoxin B_1_ and CPA in A. flavus. Our TLC analysis showed that AFB_1_ production dramatically decreased in the osaA deletion strain compared to the wild type (Figure 5A). Densitometry of TLC band intensity indicated that such reduction was approximately 95.1% with respect to the wild-type control (Figure 5B). Additional LC–MS analysis also confirmed that AFB_1_ levels were significantly reduced, approximately 84.62% in the absence of osaA with respect to the wild type (Figure 5C), supporting that osaA positively regulates the biosynthesis of AFB_1_. Moreover, the LC–MS analysis also showed that production of CPA significantly decreased approximately 54.65% in the osaA deletion mutant compared to the control (Figure 5D), suggesting that osaA positively regulates biosynthesis of CPA. Complementation of the deletion mutant with the osaA wild-type allele rescued AFB_1_ and CPA production to levels similar to those in the wild type.

2.4. osaA Affects Sensitivity to Environmental Stresses in A. flavus

Fungi need to adapt to survive environmental stresses. It has been shown that osaA influences A. fumigatus to resist high temperature and oxidative stress [35]. Our results show that A. flavus osaA also plays a role in temperature and oxidative stress sensitivity. Fungal strains were grown at a range of temperatures, 25 °C, 30 °C, 37 °C, and 42 °C (Figure S2). Percentages of growth rate change were calculated with respect to the colony diameter observed at 30 °C. In the absence of osaA, colony growth reduction was significantly higher than in the controls at 25 °C, indicating that osaA is relevant for cold resistance. In addition, all the strains tested showed the highest growth reduction at 42 °C. Interestingly, ΔosaA slightly recovers colony growth at 37 °C.

With respect to sensitivity to oxidative stress, ΔosaA showed an increase in tolerance to menadione with respect to the control strains at 0.6 mM, a menadione concentration where the control strains were unable to grow (Figure S3A). On the other hand, the ΔosaA strain presented increased sensitivity to H_2_O_2_ with respect to the controls (Figure S3B); ΔosaA colonies showed a further colony growth reduction at 0.3% with respect to the wild type and complementation strains, and growth was completely abolished at 0.4% in the absence of osaA.

In cell wall stress tests, ΔosaA showed drastically higher sensitivity to Calcofluor white (29.48%) compared to the wild type (5.57%) (Figure S4). Absence of osaA only caused a slight effect in the growth reduction resulting from Congo red supplementation (33.58%% growth reduction in the wild type and 30.76% in the mutant). These findings suggest a role of osaA in cell wall integrity and possibly composition. To further evaluate this possibility, in this study, cell wall components were also analyzed in each strain.

2.5. osaA Is Necessary for Normal Cell-Wall Composition in A. flavus

Based on the observed osaA-dependent change to environmental stress sensitivity, we hypothesized that they could be, at least in part, due to alterations in cell wall composition in the ΔosaA strain. Fungal cell walls are a crucial structure providing support, protection, and regulating interactions with the environment. In A. fumigatus, cell wall stability was also decreased in the absence of the osaA homolog [35]. Our cell wall composition analysis revealed that chitin content in A. flavus osaA deletion strain was significantly decreased, 38.39%, compared to the control strains. However, mannoprotein and glucan content in the osaA deletion strain were not statistically significantly different with respect to those in the wild type (Figure S5).

2.6. osaA Is Indispensable for A. flavus Seed Infection

Previously, it was shown that A. fumigatus osaA was important in the infection of immunocompromised animals [35]. Based on this and the fact that osaA affects multiple aspects of A. flavus biology, it is likely that osaA could also have a role in plant seed infection and colonization. To test this hypothesis, viable B73 corn seeds were infected with the wild type, ΔosaA, and osaA complementation strain. Cultures were photographed after 7 days of incubation (Figure 6A). In this experiment, levels of ergosterol were used as an indicator of fungal burden present in the infected plant tissue (Figure 6B) (expression of putative ergosterol biosynthetic genes was unaffected by the osaA deletion, Table S2). Seeds infected with the ΔosaA mutant strain contained significantly less ergosterol than seeds infected with the control strains. Importantly, the absence of osaA resulted in a statistically significant decrease in AFB_1_ and AFB_2_ production in viable seeds infected with the ΔosaA strain compared with the controls (Figure 6C,D).

2.7. Global Transcriptional Changes Induced by osaA Deletion

RNA-Seq analysis was performed to investigate the osaA-dependent transcriptome in A. flavus. Our analysis revealed substantial transcriptome alterations in the ΔosaA mutant, with 1488 differentially expressed genes (DEGs) compared to the wild type; 546 upregulated and 942 downregulated genes (Figure 7). Volcano plots indicated that most transcriptional changes were directly attributable to the absence of osaA in the mutant.

2.8. Gene Ontology and Heatmap Analyses Results

2.8.1. Downregulated Genes

Gene Ontology (GO) and functional enrichment analyses of downregulated genes in the ΔosaA mutant revealed significant enrichment of several biological categories (Table S3). Most notably, genes associated with the apoplast (96 out of 470 genes, adjusted p-value < 4 × 10^−14^, odds ratio = 3.0) and apoplastic effectors (38 out of 146 genes, adjusted p-value < 8 × 10^−8^, odds ratio = 3.9) were significantly downregulated. Additionally, genes related to membrane components (GO:0016020; 224 out of 1953 genes, adjusted p-value = 2.8 × 10^−5^, odds ratio = 1.5) and extracellular regions (GO:0005576; 27 out of 127 genes, adjusted p-value = 9.5 × 10^−4^, odds ratio = 3.0) showed reduced expression. Cytoplasmic effectors were also downregulated (19 out of 92 genes, adjusted p-value = 0.021, odds ratio = 2.9). Other significantly downregulated functional categories included cutinase activity (GO:0050525; 4 out of 5 genes, adjusted p-value = 0.025, odds ratio = 44.2) and the pentose phosphate pathway (9 out of 28 genes, adjusted p-value = 0.029, odds ratio = 5.2) (Table S3).

2.8.2. Upregulated Genes

Strikingly, our analysis revealed a strong enrichment of secondary metabolism genes among upregulated DEGs in the ΔosaA (Table S3), except for two of the clustered aflatrem genes, which were downregulated (Table S2). Most prominently, genes belonging to the aflatoxin biosynthetic cluster (SMURF cluster 54; 26 out of 30 genes, adjusted p-value < 1.9 × 10^−7^, odds ratio = 134.2) (Table S3) are highly upregulated (Figure 8, Table S4). Intriguingly, this transcriptional upregulation of aflatoxin biosynthetic genes stands in contrast to the reduced aflatoxin production observed in the ΔosaA (Figure 5). Moreover, genes of the cyclopiazonic acid gene clusters were significantly upregulated in ΔosaA (Table S3 and Figure S6). The upregulation of CPA genes in ΔosaA with respect to the control also contrasts with LC–MS results showing a reduction in CPA in the mutant (Figure 5). Other secondary metabolite gene clusters with significant upregulation included aspirochlorine (8 out of 15 genes, adjusted p-value = 4.8 × 10^−5^, odds ratio = 22.8) and imizoquins (8 out of 9 genes, adjusted p-value < 1.2 × 10^−7^ odds ratio = 159.7) (Table S3 and Figure S6).

Additional upregulated categories includes monooxygenase activity (GO:0004497; 23 out of 152 genes, adjusted p-value = 3.1 × 10^−4^, odds ratio = 3.62), phosphopantetheine binding (GO:0031177; 11 out of 41 genes, adjusted p-value = 7.1 × 10^−4^, odds ratio = 7.3), O-methyltransferase activity (GO:0008171; 9 out of 28 genes, adjusted p-value = 9.3 × 10^−4^, odds ratio = 9.47), cytochrome P450s (6 out of 20 genes, adjusted p-value = 0.027, odds ratio = 8.5), iron ion binding (GO:0005506; 20 out of 151 genes, adjusted p-value = 0.0059, odds ratio = 3.0), and oxidoreductase activity (GO:0016705; 17 out of 116 genes, adjusted p-value = 0.0059, odds ratio = 3.4), all of which are commonly associated with secondary metabolism (Table S3).

2.9. Other osaA-Dependent DEGs

2.9.1. Genes Involved in Transmembrane Transporter Activity

Transcriptome analysis revealed that osaA deletion significantly altered the expression of genes associated with transmembrane transporter activity (Figure S7). In the osaA mutant, several transmembrane transporter genes exhibited marked downregulation while others were upregulated concurrently. These transporter sets encompass diverse families, such as the major facilitator superfamily (MFS), amino acid transporters, and multidrug resistance proteins. Among the downregulated transporters are F9C07_2149740, which encodes an inorganic phosphate transporter; F9C07_2280468, encoding a sugar transporter; F9C07_2282850, predicted to function as a siderochrome–iron transporter; and F9C07_2236497, encoding an MFS multidrug transporter, among others. Those that were upregulated included the MFS transporter (F9C07_2285417), sugar transporter (F9C07_7897), amino acid transporter (F9C07_2232111), and others. These unbalanced transport systems likely have detrimental effects that could affect growth, development, secondary metabolism, and virulence.

2.9.2. Genes Involved in Oxidoreductase Activity

RNA-seq analysis also revealed broad transcriptional changes in oxidoreductase-related genes in the absence of osaA. Among these, F9C07_2282817, encoding a NADP-dependent oxidoreductase domain-containing protein, was significantly upregulated (Table S2). This enzyme converts NADPH to NADP^+^, which limits the availability of NADPH.

2.9.3. Developmental Genes

Transcriptome analysis of the osaA mutant revealed differential expressions of key developmental regulators (Table S2). brlA (F9C07_2279377) and flbD (F9C07_2279158), essential for conidiophore formation, were upregulated in the osaA mutant, consistent with the observed increase in conidiation (Figure 3). In contrast, atfA (F9C07_2277799), atfB (F9C07_9086), sfgA (F9C07_10617), hogA (F9C07_2156698), flbC (F9C07_2282647), and fluG (F9C07_5280) were downregulated in the osaA mutant. AtfA and AtfB are bZIP transcription factors involved in development and stress response [39], while SfgA is a developmental repressor [40]. HogA also modulates conidiation [41], and FlbC and FluG function upstream as developmental activators [42].

2.9.4. Superoxide Dismutase and Catalase Genes

Transcriptome analysis revealed that the deletion of osaA in A. flavus altered the expression of oxidative stress-related genes. Superoxide dismutase was upregulated, whereas catalase was downregulated in the osaA mutant. F9C07_2286715, encoding a superoxide dismutase, was upregulated by log_2_fold-changes of + 0.58. On the other hand, F9C07_2285585, F9C07_2284865, F9C07_2237005, and F9C07_2277540, annotated as putative catalase, catalase-like domain-containing protein, heme peroxidase, and catalase, respectively, were downregulated by log_2_ fold-changes of −1.1, −1.4, −1.5, and −1.6, respectively (Table S2).

2.9.5. Chitin Synthase Gene

Transcriptome analysis also revealed that deletion of osaA in A. flavus decreased the expression of putative chitin synthase genes. For example, F9C07_366 and F9C07_2286842, annotated as genes encoding chitin synthase and chitin synthesis regulation, respectively, were downregulated by log_2_ fold-changes of −1.2 and −2.8, respectively, in the osaA mutant (Table S2).

2.10. Motif Enrichment Analysis in Promoters of Differentially Expressed Genes

To identify potential transcription factor binding sites involved in osaA-mediated regulation, we performed motif enrichment analysis on the 1000 bp upstream regions of all DEGs in the ΔosaA vs. WT comparison. Using the MEME suite’s Simple Enrichment Analysis (SEA) with the JASPAR 2022 fungi non-redundant database and a p-value threshold of 1 × 10^−10^, we identified significant enrichment of two similar motifs: MA0382.2 (SKO1 binding motif; consensus sequence DNHDATGACGTAATWDN; p-value = 1.5 × 10^−16^; enrichment ratio = 1.5) and MA0286.1 (CST6 binding motif; consensus sequence RTGACGTMA; p-value = 6.6 × 10^−14^; enrichment ratio = 2.3) (Figure 9). Among the DEGs, 347 contained the SKO1 motif, and 110 contained the CST6 motif, with 112 genes containing both motifs.

Additionally, de novo motif discovery using STREME identified three significant motifs in the promoters of DEGs. The first motif closely resembled the SKO1 binding site (MA0382.2) identified with SEA, while the second showed similarity to the MA0403.2 motif in the JASPAR database (Figure 9). The third motif did not match any known fungal transcription factor binding sites, potentially representing a novel regulatory element associated with osaA-mediated gene expression.

3. Discussion

Novel regulatory genes and their products that govern fungal development, secondary metabolism, and pathogenesis could serve as targets for control strategies to effectively decrease A. flavus colonization and aflatoxin contamination. Gti1/Pac2 (WOPR) domain proteins represent a class of fungal-specific transcription factors involved in regulating those cellular processes [25]. In our study, we identified that in the agriculturally important fungus A. flavus, OsaA contains a typical WOPR domain in its N-terminal region, composed of two conserved subdomains separated by a short linker. Our results also support that the Aspergillus OsaA protein is phylogenetically related to the Gti1/Wor1/Ryp1/Mit1/Rge1/Ros1/Sge1 proteins described in other fungal species [26]. Structural analysis of another study shows WOPR subdomains are tightly connected by a β-sheet, with the linker extending outward from the DNA, and the β-strands of each subdomain interdigitating to form a stable structure [25].

In studies using the model filamentous fungus A. nidulans, the osaA homolog has been shown to regulate development by repressing sexual reproduction downstream of the global regulator VeA [34]. Similarly, osaA plays a key role in A. fumigatus morphology. Absence of osaA significantly reduces colony growth, germination rate, and conidial production in this opportunistic human pathogen [35]. In contrast, our study revealed that loss of osaA in A. flavus leads to increased conidial production. Consistent with these phenotypes, transcriptome data revealed upregulation of key conidiophore developmental genes in the osaA mutant. Notably, brlA (F9C07_2279377) and flbD (F9C07_ 2279158), which are central developmental regulators, were upregulated. BrlA, is a C_2_H_2_ zinc finger transcription factor, essential for initiating conidiophore formation, while FlbD acts upstream to activate brlA expression [39,43]. These gene expression patterns support the observed increase in conidiation in the osaA mutant.

In addition to conidiation, osaA influences sclerotial formation in A. flavus. osaA is indispensable for normal sclerotial production rates. The colony of the osaA deletion strain produced significantly fewer sclerotia. This effect was more dramatic on top-agar inoculated cultures, where sclerotial formation is completely abolished in the absence of osaA. This dramatic reduction may be due to high spore density and, therefore, a crowding effect in the latter cultures, leading to an accentuated competition. In addition, our transcriptome data showed that transcription factor genes atfA (F9C07_2277799) and atfB (F9C07_9086) were downregulated in the osaA mutant. These genes positively affect sclerotial formation [44]; therefore, a reduction in atfA and atfB expression in the osaA deletion mutant further contributes to the observed decrease in sclerotial production.

Since development and secondary metabolism are found to be genetically linked in Aspergillus species [5], we also investigated the possible role of osaA in regulating the production of natural compounds in A. flavus, particularly mycotoxins. Our findings demonstrate that osaA positively regulates the biosynthesis of both AFB_1_ and CPA in A. flavus. Unexpectedly, most aflatoxin biosynthetic genes and genes of CPA gene clusters were upregulated in the osaA mutant compared to the control. This contrasts with chemical analysis results, which showed a clear reduction in AF, as well as CPA production, in the mutant. This discrepancy suggests the presence of a feedback mechanism, where the fungal system attempts to compensate for reduced mycotoxin levels by upregulating gene expression. The deletion of osaA may impair post-transcriptional regulation, preventing proper translation, processing, or localization of AF biosynthetic enzymes. Aflatoxin biosynthesis and transport are known to involve specialized vesicles called aflatoxisomes [45]. In Aspergillus parasiticus, the vesicle-vacuole system is essential for converting sterigmatocystin (ST) to aflatoxin B_1_ and compartmentalizing the toxin [46]. Key enzymes such as Nor-1, Ver-1, and OmtA are synthesized in the cytoplasm and transported to vacuoles via the cytoplasm-to-vacuole targeting (Cvt) pathway [46]. More recently, lipid droplets have also been implicated in aflatoxin biosynthesis and export [47]. Given the connection between aflatoxisome function and membrane transport, defects in transporter gene expression in the osaA mutant may further impair AF production. Expression of several transmembrane transporter genes was altered in the osaA mutant. These widespread changes in transporters could restrict the availability of key nutrients and cofactors necessary for aflatoxin biosynthesis, including phosphate, iron ions, and sugar. Furthermore, aflatoxin biosynthesis requires NADPH as a cofactor, especially for the cytochrome P450 monooxygenase OrdA, which catalyzes the conversion of O-methylsterigmatocystin (OMST) and dihydro-OMST into aflatoxins B_1_, B_2_, G_1_, and G_2_ [48]. In the osaA mutant, we observed upregulation of a gene (F9C07_2282817) encoding NADP-dependent oxidoreductase domain-containing protein (Table S2), which likely catalyzes the conversion of NADPH to NADP^+^. This thereby decreases the availability of NADPH required for aflatoxin biosynthetic enzymes, which would further contribute to the observed reduction in AF production. These plausible causes will be the subject of future studies.

Additionally, downregulation of atfA (F9C07_2277799) and atfB (F9C07_9086) may partially explain the outcome of reduction in aflatoxin production despite the upregulation of aflatoxin biosynthetic genes. The bZIP transcription factors were demonstrated as co-regulators of aflatoxin biosynthesis and oxidative stress in A. parasiticus [49,50]. The reduced expression of bZIP transcription factors may compromise the fungus’s ability to manage oxidative stress, which in turn can negatively influence aflatoxin biosynthesis.

Expression of other secondary metabolite genes, including genes in the imizoquin and aspirochlorine biosynthetic clusters, was upregulated by osaA. Imizoquin possesses reactive oxygen species (ROS)-quenching properties, which help to maintain ROS homeostasis and promote spore germination in the A. flavus [51]. In A. flavus, the imizoquin biosynthetic pathway is enhanced by specific oxylipins, contributing to the inhibition of other metabolic pathways such as aflatoxin biosynthesis [52]. Additionally, genes within the aspirochlorine biosynthetic cluster were also upregulated in the osaA mutant. Aspirochlorine is a halogenated epidithiodiketopiperazine mycotoxin with antibiotic activity, also known as antibiotic A30641 [53]. The increased expression of this cluster in the osaA mutant raises the possibility that osaA could contribute to antifungal defense and may represent a promising target for antifungal drug development.

Production of secondary metabolites, including mycotoxins, is also influenced by environmental stressors. A. flavus withstands various biotic and abiotic stresses, including fluctuations in temperature and oxidative stress. This fungus can grow and produce aflatoxins over a wide temperature range, with thermal tolerance being an important factor influencing its pathogenicity and toxin biosynthesis [1,54]. As mentioned above, oxidative stress also plays a critical role in regulating aflatoxin production, as A. flavus responds to reactive oxygen species through complex regulatory pathways that modulate secondary metabolite biosynthesis [1,55]. The possible role of osaA in A. flavus in oxidative stress response was evaluated in our study. The osaA mutant showed increased sensitivity to hydrogen peroxide. However, this strain presented greater resistance to menadione compared to control strains. In Saccharomyces cerevisiae, rad9 mutants showed normal resistance to menadione but were up to 100-fold more sensitive to H_2_O_2_ [56], indicating distinct features in the cellular responses [57]. Menadione generates superoxide radicals, which can cause oxidative damage. If the mutation enhances superoxide dismutase activity, the mutant could exhibit increased resistance to menadione. However, in the case of hydrogen peroxide, its detoxification relies on catalases [58,59,60]. Our transcriptome indicated that the catalase gene (F9C07_2237005) was significantly downregulated in the osaA mutant, which could contribute to the increased sensitivity to hydrogen peroxide in this strain. Conversely, superoxide dismutase gene (F9C07_2286715) expression was slightly upregulated, and could contribute to the increased resistance to menadione observed in the osaA mutant. Future enzymatic assays will be carried out to further assess these effects. Several studies have demonstrated that certain aflatoxin inhibitors reduce AFB_1_ production by modulating antioxidant activity. For example, ascorbic acid and cinnamaldehyde significantly decreased AFB_1_ levels while increasing superoxide dismutase activity [49,55,61]. Enhanced superoxide dismutase activity in ΔosaA could contribute to a reduction in aflatoxin biosynthesis. In addition, downregulation of Pentose Phosphate Pathway (PPP) genes in osaA mutant can also weaken both oxidative stress tolerance. The PPP is crucial for producing NADPH, which acts as a reducing equivalent to maintain the redox balance inside fungal cells. NADPH is required for the function of antioxidant enzymes such as catalase. When PPP genes are downregulated, the NADPH pool decreases, leading to increased sensitivity to reactive oxygen species (ROS) [62]. Since osaA mutant presents downregulation of PPP genes, it is expected that this strain is unable to efficiently neutralize ROS.

In addition to oxidative stress, we also assessed osaA-dependent temperature sensitivity. Both low and high temperatures can affect A. flavus growth [54,63]. We found that the decrease in growth rate in the A. flavus osaA mutant compared to the wild type was more pronounced at a lower temperature (25 °C) than that observed at 30 °C. All strains exhibited growth at 42 °C. This contrasts with findings in A. fumigatus, where osaA is specifically required for growth at elevated temperatures (42 °C) [35].

The changes in sensitivity to environmental stresses could also be influenced by alterations in the fungal cell wall [64]. The fungal cell wall is a rigid, dynamic structure essential for fungal viability [65]. In A. fumigatus, deletion of osaA compromises cell wall stability, further implicating its role in stress adaptation [35]. Thus, the increase in cold sensitivity observed in the A. flavus ΔosaA strain could be the result of possible changes in its cell wall composition and integrity. We found A. flavus osaA influenced cell wall function, further reducing colony growth in the osaA mutant when exposed to the perturbing cell wall agents, particularly Calcofluor, a chitin-binding agent. Moreover, our study revealed a significant reduction in chitin content in the osaA mutant with respect to the wild type, which suggests that osaA positively regulates chitin synthesis in A. flavus. Furthermore, the chitin synthase gene (F9C07_366) is downregulated in the osaA mutant, which could contribute to the decreased chitin level in this strain’s cell wall. However, mannoprotein and glucan levels in the cell wall were unaffected by osaA.

Reduced chitin and weakened cell wall integrity in the osaA mutant, along with other affected processes, likely decrease its virulence. We investigated the role of osaA in seed infection. Our results show that osaA is required for wild-type levels of seed colonization, as its deletion reduces fungal burden in corn kernels. Our transcriptome analysis indicated downregulation of apoplast, apoplastic effectors, and cytoplasmic effector genes in the ΔosaA, which could lead to reduced pathogenicity in the seed [66,67]. Additionally, the downregulation of pentose phosphate pathway genes in the osaA mutant and its effect on oxidative stress could lead to impaired infection ability and attenuated virulence. Importantly, the absence of osaA led to a marked decrease in AFB_1_ and AFB_2_ production also during infected seeds.

To further gain insight into the osaA’s regulatory role, a promoter analysis was conducted using known fungal transcription factors' binding motifs from the JASPAR database. Two motifs were MA0382.2 (SKO1 binding motif) and MA0286.1 (CST6 binding motif), which were identified by motif enrichment analysis on the 1000 bp upstream regions of all DEGs in the ΔosaA vs. wild type comparison. The SKO1 protein that is associated with the MA0382.2 motif seems to operate in response to osmotic stress and cell wall damage. In Aspergillus cristatus, the SKO1 protein, a homolog of the yeast SKO1p, functions as a transcription factor involved in regulating osmotic stress response, similar to its role in S. cerevisiae [68]. This further suggests that OsaA, perhaps in cooperation with a SKO1 homolog in A. flavus, may regulate stress-responsive pathways via transcriptional control. With respect to CST6, a recent study has demonstrated that this transcription factor is involved in azole susceptibility in Candida glabrata, showing that CST6 regulates a broad gene network, including genes involved in respiration, cell-wall, and adhesins in C. glabrata. It is possible that OsaA could also be associated with antifungal drug susceptibility in A. flavus [69]. Future studies will focus on possible interactions between these relevant regulatory factors.

In summary, osaA functions as a central regulator integrating fungal development, environmental stress management, secondary metabolite production, and virulence in A. flavus. These findings provide valuable insights into the regulatory networks underlying fungal pathogenicity and identify osaA as a promising target for future antifungal strategies.

4. Materials and Methods

4.1. Sequence Analysis

The deduced amino acid and nucleotide sequences used in the phylogenetic analysis were obtained from FungiDB (https://fungidb.org/fungidb/app/; version: Release 68, 7 May 2024). All gene and protein accessions are listed in Table 1.

Corresponding protein accession numbers from NCBI were found by conducting a blastP analysis using sequences obtained from FungiDB. Using Geneious software (version 2023.0.1), a MAFFT multisequence alignment (Version 1.2.2) was conducted using the default parameters to align the protein sequences [70]. Subsequently, a maximum-likelihood tree was constructed using PhyML (version 3.3.20180621) with a LG substitution model and 1000 bootstrap replicates [71].

4.2. Strains and Culture Conditions

The A. flavus strains utilized in this study were the CA14 pyrG-1control strain, osaA deletion strain (TMR1.1), and osaA complementation strain (TFEH 8.1) (Table 2). All strains were grown on commercial PDA (Potato Dextrose Agar) medium (BD Difco™, Franklin, NJ, USA) unless otherwise indicated. Fungal strains were maintained in 30% glycerol stocks at −80 °C in a freezer (Forma Scientific Bio-Freezer, Model 8358, Marietta, OH, USA).

4.3. Generation of the osaA Deletion Strain

The ΔosaA strain was generated by a gene replacement strategy described by [35]. The deletion DNA cassette was generated by fusion PCR. The 5′ untranslated region (1.854 kb) and 3′ untranslated region (1.778 kb) flanking the osaA coding sequence were amplified from A. flavus genomic DNA using primer pairs 3011/3012 and 3013/3014, respectively (Table S1). The A. fumigatus pyrG selection marker was amplified from plasmid p1439 [72] with primers 3015 and 3016. These three fragments were fused via PCR using primers 3017 and 3018 to generate the knockout cassette. The construct was introduced into A. flavus CA14_pSL82 (pyrG−, niaD+, ptrA^S^, Δku70) the protoplasts via polyethylene glycol-mediated transformation. Successful deletion of the osaA coding region and replacement with pyrG was confirmed by diagnostic PCR using primers 3011 and 2846, resulting in the ΔosaA strain (TMR1.1).

4.4. Generation of the osaA Complementation Strain

The complementation strain (TFEH8.1) was constructed by reintroducing the wild-type osaA allele into the ΔosaA strain at the same locus. To generate the complementation cassette, four DNA fragments were PCR-amplified: the osaA coding sequence along with a 2 kb upstream region (5′UTR) using primers 3065 and 3066; the trpC terminator using primers 2869 and 2870; the ptrA selection marker using primers 3067 and 3068; and the A. fumigatus pyrG gene from plasmid p1439 using primers 2871 and 2872. These four fragments were fused in a single PCR reaction using primers 3063 and 3064. The resulting fusion cassette was introduced into the ΔosaA strain TMR1.1 via polyethylene glycol-mediated transformation [36]. Transformants were screened and verified by diagnostic PCR using primers 3063 and 3047. The selected transformant was designated as TFEH8.1.

4.5. Morphological Analysis

4.5.1. Colony Growth

To assess the role of osaA in vegetative colony growth, A. flavus CA14 wild type, ΔosaA, and osaA complementation strain were point-inoculated onto commercial PDA medium (BD Difco™, Franklin, NJ, USA) and incubated at 30 °C in a dark incubator (Thermo Scientific Heraeus^®^ BBD6220, Waltham, MA, USA). Colony growth was measured as colony diameter at 7 days post-inoculation. All experiments were performed in triplicate.

4.5.2. Conidial Production

To investigate the role of osaA in conidiation, A. flavus CA14 wild type, ΔosaA, and osaA complementation strain were point-inoculated onto PDA medium and incubated at 30 °C in the dark. Conidiation was evaluated at 7 days post-inoculation. Agar cores (10 mm in diameter) were collected from regions 5 mm and 20 mm away from the colony center. The samples were homogenized in sterile water, and conidia were quantified using a haemocytometer (Hausser Scientific, Horsham, PA, USA) under Nikon Eclipse E-400 bright-field microscopy(Nikon, Inc., Melville, NY, USA). Additionally, close-up images of conidiophores from both regions were captured using a Leica dissecting microscope (Leica Microsystems, Inc., Buffalo Grove, IL, USA) at 32× magnification.

4.5.3. Sclerotial Development

To determine the role of osaA in sclerotial production, A. flavus CA14 wild type, ΔosaA and osaA complementation strain were both point-inoculated and top-agar inoculated (10^6^ spores/mL) onto Wickerham agar medium (Per liter: 2.0 g yeast extract, 3.0 g peptone, 5.0 g corn steep solids, 2.0 g dextrose, 30.0 g sucrose, 2.0 g NaNO_3_, 1.0 g K_2_HPO_4_·3H_2_O, 0.5 g MgSO_4_·7H2O, 0.2 g KCl, 0.1 g FeSO_4_·7H_2_O (10-fold the original recipe), and 15.0 g agar per liter [pH 5.5]). Cultures were incubated at 30 °C in the dark for 14 days. Following incubation, cultures were washed with 70% ethanol to remove conidia and enhance sclerotial visibility. Micrographs were captured using a Leica MZ75 dissecting microscope equipped with a DC50LP camera (Leica Microsystems, Inc., Buffalo Grove, IL, USA) at 12.5× and 32× magnifications.

4.6. Aflatoxin B1 Production Analysis

To evaluate whether osaA regulates AFB_1_ biosynthesis, A. flavus CA14 wild type, ΔosaA, and osaA complementation strain were point-inoculated on PDA medium and incubated in the dark at 30 °C for 7 days. Three 16 mm-diameter cores were collected approximately 5 mm from the colony center. AFB_1_ was extracted using 5 mL of chloroform per sample. The extracts were evaporated to dryness and subsequently resuspended in 200 µL of chloroform. AFB_1_ detection was performed via thin-layer chromatography (TLC) using silica-precoated Polygram Sil G/UV254 TLC plates (Macherey-Nagel, Bethlehem, PA, USA) and a solvent system composed of toluene:ethyl acetate:formic acid (50:40:10). After air-drying, the TLC plates were sprayed with a 12.5% aluminum chloride (AlCl_3_) solution in ethanol, baked at 80 °C for 10 min, and visualized under UV light at 375 nm. Aflatoxin B_1_ standard was obtained from Sigma-Aldrich (St. Louis, MO, USA). Samples were also analyzed to identify other secondary metabolites by LC–MS (Section 4.7) and to further quantify Aflatoxin B_1_ by UPLC with fluorescence detection (Section 4.10.2).

4.7. Secondary Metabolite Analysis by LC–MS

A. flavus secondary metabolites in the extracts were analyzed on a Waters Acquity UPLC system (Waters Corporation, Milford, MA, USA) coupled to a Waters Xevo G2 XS QTOF mass spectrometer(Waters Corporation, Milford, MA, USA). Extract injections (1 µL) were separated on a Waters BEH C18 1.7 µm, 2.1 × 50 mm column with the following gradient solvent system: (0.5 mL/min, solvent A: 0.1% formic acid in water; solvent B: 0.1% formic acid in acetonitrile): 5% B (0–1.25 min), gradient to 25% B (1.25–1.5 min), gradient to 100% B (1.5–5.0 min), 100% B (5.0–7.5 min), then column equilibration to 5% B (7.6–10.1 min). The Z-spray ionization source was run in ESI+ mode using MassLynx 4.2 software with the following settings: source temperature: 100 °C, desolvation temperature: 250 °C, desolvation gas flow: 600 L/h, cone gas flow: 50 L/h, capillary voltage: 3.0 kV, sampling cone voltage: 40 V. Analyses were performed in sensitivity and continuum mode, with a mass range of m/z 50–1200 and a scan time of 0.1 s. A data-independent acquisition method with elevated collision energy (MS^E^) was used with 6 eV low energy and a high energy ramp from 15 to 45 eV. Mass data were collected from 2.0 to 6.0 min. then imported, analyzed, and quantified on Waters UNIFI 1.9.4 software using “Quantify Assay Tof 2D” analysis method with lock mass corrected by UNIFI. Aflatoxin B_1_ and CPA standards were purchased from Sigma-Aldrich (St. Louis, MO, USA). Metabolite content is expressed in ppb (ng/g samples).

4.8. Environmental Stress Tests

4.8.1. Temperature Sensitivity

To assess the role of osaA in temperature sensitivity, A. flavus CA14 wild type, ΔosaA, and osaA complementation strain were point-inoculated on PDA medium and incubated in the dark at 25 °C, 30 °C, 37 °C, and 42 °C for 5 days. Colony diameters were measured, and growth was assessed across three biological replicates. The percentage reduction in growth was calculated by comparing colony diameters at 30 °C to those at 25 °C, 37 °C, and 42 °C.

4.8.2. Oxidative Stress Sensitivity

To evaluate the potential role of osaA in oxidative stress sensitivity, A. flavus CA14 wild type, ΔosaA, and complementation strain were point-inoculated on PDA plates supplemented with 0.4, 0.5, and 0.6 mM menadione and incubated at 30 °C in the dark for 3 days. Colony growth was measured, and the percentage of growth reduction was calculated relative to control plates without menadione. Experiments were performed in triplicate. Additionally, oxidative stress sensitivity was tested using hydrogen peroxide. The same strains were point-inoculated on PDA medium supplemented with 0.1%, 0.2%, 0.3%, and 0.4% hydrogen peroxide and incubated at 30 °C for 3 days. Four biological replicates were included. Growth inhibition was calculated by comparing colony diameters from treated versus untreated conditions.

4.8.3. Cell Wall Stress Test

To elucidate possible alterations of cell wall integrity due to the absence of osaA, wild type (WT), deletion osaA (ΔosaA), and complementation osaA (Com) were point-inoculated on PDA medium supplemented with Calcofluor white (0.1 mg/mL) or Congo Red (0.3 mg/mL) [73] and incubated at 30 °C for 3 days in the dark. The experiments were carried out with three replicates. Growth inhibition was calculated by comparing colony diameters from treated versus untreated conditions.

4.9. Cell Wall Chemical Analysis

To measure the levels of cell wall components chitin, mannoprotein, and glucan in the A. flavus CA14 wild type, ΔosaA, and osaA complementation strain cell walls, a previously described protocol [74] was followed with minor modifications. Briefly, A. flavus strains were inoculated into 50 mL of liquid commercial PDB (BD Difco™, Franklin, NJ, USA) medium (10^6^ spores/mL) and incubated at 30 °C for 48 h at 250 rpm. Mycelia were harvested using Miracloth (Calbiochem, San Diego, CA, USA), washed three times with sterile distilled water, and stored at −20 °C.

For cell wall analysis, frozen mycelia were resuspended in 1 mL of cell wall buffer (2% SDS in 50 mM Tris-HCl, pH 7.5, supplemented with 100 mM Na-EDTA, 40 mM β-mercaptoethanol, and 1 mM PMSF) and boiled for 15 min to remove unbound proteins and soluble sugars. After boiling, samples were washed three times with sterile ddH_2_O and lyophilized overnight.

Approximately 40 mg of lyophilized mycelia per strain (three replicates each) were treated with 3% NaOH at 75 °C for 1 h. Samples were centrifuged at 15,000× g for 15 min to separate the soluble and insoluble fractions. The supernatant was collected and used for the analysis of mannoprotein and any soluble glucans that may be present.

The remaining pellet was digested with 96% formic acid at 100 °C for 4 h. After evaporation of formic acid, the residues were resuspended in 1 mL of sterile ddH_2_O for chitin and insoluble glucan present.

Quantification of chitin, mannoprotein, and glucan was performed as described in [75,76,77] and measured absorbance at 520 nm, 560 nm, and 490 nm, respectively, using an Epoch spectrophotometer (BioTek, Winooski, VT, USA).

4.10. Virulence Studies by Seed Infection Assay

4.10.1. Kernel Screening Assay

A Kernel Screening Assay (KSA) was performed on corn seeds from the aflatoxin-sensitive B73 corn line as a plant model, as previously described in [78]. Conidiospores from the CA14 wild-type control, ΔosaA, and osaA complementation strain were cultivated on 2 × V8 agar (100 mL V8 juice, 40 g agar per liter of medium, and pH 5.2) and incubated at 30 °C in the light for 7 days. Spores were collected in 25 mL of water in aseptic conditions and quantified using an Olympus Automated Cell Counter Model R1 (Olympus Corporation, Shinjuku, Tokyo, Japan). Undamaged B73 seeds of a relatively similar size were collected and surface sterilized using 10% bleach. The seeds were infected with spores of the A. flavus strains by being soaked in 10 mL of 1.0 × 10^4^ spores/mL spore suspension for each strain. Uninfected B73 seeds, serving as control (mock), were also treated with water only, in a manner similar to the seeds infected with A. flavus spores. The falcon tubes containing the seeds and spore suspensions were rocked for 3 min to ensure equal distribution of inoculum throughout the seeds. The inoculum was drained from the falcon tubes, and the seeds were transferred to sterile Petri plate lids (60 × 15 mm). These were placed onto larger trays containing 3 MM Whatman filter paper and a 50 mL sterile water reservoir to provide humidity. The experiment was performed in replicates of six for each strain. The cultures were then incubated at 30 °C under dark conditions for 7 days. The seeds were then photographed and harvested by flash freezing the seeds in liquid nitrogen. Prior to LC–MS analysis, the seeds were pulverized using a SPEX SamplePrep 2010 Geno/Grinder (SPEX SamplePrep, Metuchen, NJ, USA) prior to lyophilization and storage at −80 °C.

4.10.2. Extraction and Aflatoxin Analysis from Seeds

Lyophilized ground corn powder (100 mg) was transferred into a 2 mL Eppendorf tube with methanol (1 mL) and vortexed vigorously for 15 s. Tubes were secured onto an orbital platform shaker (Solaris 2000, Thermo Fisher Scientific) then rotated at 200 rpm, at RT and in darkness, approximately 22 h. Next, tubes were centrifuged for 5 min at 14,000 rpm to remove particulate, and a portion of the particulate-free extract (1 mL) was transferred to a new tube for storage and analysis. The aflatoxin-containing solution was analyzed (1 µL injections) using a Waters ACQUITY UPLC system (40% methanol in water, BEH C18 1.7 μm, 2.1 mm × 50 mm column) with fluorescence detection (Ex = 365 nm, Em = 440 nm). Samples were diluted if the aflatoxin signal saturated the detector. Analytical standards (Sigma-Aldrich, St. Louis, MO, USA) were used to identify and quantify aflatoxins AFB_1_ and AFB_2_. Aflatoxin content was expressed in ppb (ng/g corn dry weight).

4.10.3. Ergosterol Extraction and Analysis

Lyophilized ground corn (50 mg) was transferred to 15 mL Falcon tubes. Alcoholic potassium hydroxide (KOH) was prepared: 25 g KOH was dissolved in 35 mL of water, then 100% ethanol was added for a total volume of 100 mL. 3 mL of the alcoholic KOH solution was added to each sample tube, the mixture was vortexed for 1 min, and then incubated at 85 °C in a water bath for 1.5 h. The samples were then cooled to room temperature, and distilled water (1 mL) and hexane (3 mL) were added to the tubes and vortexed vigorously for 3 min to extract ergosterol. The hexane layers were carefully transferred to clean 4 mL glass vials and concentrated via SpeedVac (Savant, Thermo Scientific). The concentrated extracts were redissolved in methanol (500 µL) for analysis. The redissolved solution was analyzed (1 µL injections) using a Waters ACQUITY UPLC system (95% methanol in water, BEH C18 1.7 μm, 2.1 mm × 50 mm column) with UV detection (λ = 282 nm). An analytical standard of ergosterol (Sigma-Aldrich, St. Louis, MO, USA) was used for quantification. Ergosterol content is reported as µg/g corn dry weight.

4.11. osaA Expression Analysis

Plates containing 25 mL of PDB were inoculated with 10^6^ spores/mL of A. flavus CA14 wild type, ΔosaA, and osaA complementation strain, and incubated in the dark at 30 °C for 72 h. Total RNA was extracted from lyophilized mycelial samples using TRIsure™ (Meridian Bioscience, Bioline, Cincinnati, OH, USA) according to the manufacturer’s instructions. For gene expression analysis, 1 µg of total RNA was treated with the Ambion TURBO DNA-free™ Kit (Thermo Fisher Scientific, Waltham, MA, USA) to remove genomic DNA. First-strand cDNA was synthesized using Moloney murine leukemia virus (MMLV) reverse transcriptase (Promega, Madison, WI, USA).

Quantitative real-time PCR (qRT-PCR) was carried out using either the Bio-Rad CFX96 Real-Time PCR System or the Applied Biosystems 7000 system. Reactions were prepared using either iQ SYBR Green Supermix (Bio-Rad, Hercules, CA, USA) or SYBR Green JumpStart Taq ReadyMix (Sigma-Aldrich, St. Louis, MO, USA) for fluorescence detection. The primer pairs 3071 and 3072 were used for target gene amplification. Expression levels were normalized to 18S rRNA as an internal control. Relative expression was calculated using the 2^−ΔΔCT^ method [38].

4.12. Transcriptome Analysis

4.12.1. RNA Extraction

Plates containing 25 mL of liquid potato dextrose broth (PDB) were inoculated with 10^6^ spores/mL of A. flavus CA14 wild type, ΔosaA, and osaA complementation strain, incubated in the dark at 30 °C for 3 days. Following incubation, the mycelia were harvested, frozen in liquid nitrogen, and lyophilized. Total RNA was extracted from the lyophilized mycelia using the RNeasy Plant Mini Kit (Qiagen, Germantown, MD, USA) according to the manufacturer’s instructions.

4.12.2. RNA Sequencing

RNA sequencing generated an average of 56 million 150 bp paired-end reads per sample. Adapters and low-quality sequences were removed from the reads using fastp [79]. Trimmed reads were aligned to the A. flavus NRRL 3357 genome (GCA_009017415.1) using STAR (version 2.7.10b) with the following settings “--alignIntronMax 1000 --twopassMode Basic --quantMode GeneCounts”. The forward-stranded gene-level read counts output by STAR were used as input for differential expression analysis. Differentially expressed genes (DEGs) between the samples were identified using DESeq2 (version 1.36.0) [80]. Log2 fold changes and p-values were estimated using the lfcShrink function with type = “ashr” [81] and alpha = 0.05 Results were filtered to retain genes with adjusted p-value < 0.05 and absolute log2 fold change > 1. Functional enrichment analysis of DEGs was performed using the enrichment function in the BC3NET R package version 1.0.4 [82] which implements a one-sided Fisher’s exact test. False discovery rate was controlled using the p.adjust R function with method = “fdr” [83]. Heatmaps were created using the tidyHeatmap R package [84] with regularized log-transformed counts and row scaling enabled. All heatmaps display only genes differentially expressed in the deletion vs. wild-type comparison.

Transcription factor binding site enrichment in promoter sequences was analyzed using the MEME Suite (version 5.5.7) [85]. The sequence of 1000 base pairs upstream of the coding start site of each DEG was used as the input, and the corresponding upstream region from all other genes was used as the background. Known transcription factors from the JASPAR non-redundant fungi 2022 database were used with the Simple Enrichment Tool from MEME with an e-value threshold of 1 × 10^−10^. De Novo motif analysis was also performed using STREME from the MEME suite with default options, the same input and background sequences as above, and an e-value threshold of 1 × 10^−10^. Motifs discovered using STREME were compared to known motifs using the Tomtom tool in MEME.

4.13. Statistical Analysis

Statistical analysis was applied to analyze all quantitative data in this study, utilizing analysis of variance (ANOVA) in conjunction with a Tukey multiple-comparison test using a p-value of <0.05 for samples that are determined to be significantly different unless otherwise indicated.