The hyphae-specific C₂H₂ transcription factor HscA regulates development, stress response, and mycotoxin production in Aspergillus species

ABSTRACT

The zinc cluster family is the largest group of transcription factors involved in regulating fungal growth, morphology, and differentiation in Aspergillus species. In this study, we investigated hyphae-specific zinc cluster transcription factors and characterized a novel hyphae-specific Cys₂His₂ zinc finger transcription factor, designated HscA, in the model fungus Aspergillus nidulans and the toxigenic fungus Aspergillus flavus. Phenotypic analyses demonstrated that HscA is essential for normal asexual and sexual development in A. nidulans and A. flavus. Deletion of hscA resulted in elevated sensitivity to cell wall stress agents and an ion depletion stressor. Moreover, the hscA null mutant exhibited decreased production of sterigmatocystin in A. nidulans and aflatoxin B1 in A. flavus. Conidial production in the kernel was decreased in the ΔhscA strain compared to the control in A. flavus. Overall, these results suggest that HscA plays a pivotal role in fungal development, stress tolerance, and mycotoxin production in Aspergillus species.

IMPORTANCE

Fungal growth and development are closely regulated by a variety of transcription factors. This study identified and characterized a hyphae-specific Cys₂His₂ zinc finger transcription factor in two Aspergillus species. HscA contains a Cys₂His₂ zinc finger domain and plays a crucial role in appropriate fungal development in A. nidulans and A. flavus. Particularly, HscA is involved in stress tolerance in both hyphal and conidial stages. We further demonstrated that HscA acts as a positive regulator of sterigmatocystin production in A. nidulans and is essential for proper aflatoxin B1 production in A. flavus. Additionally, our findings indicate that HscA is crucial for conidial formation in kernel assays, implying that HscA may function as a virulence factor. Overall, these findings enhance our understanding of mycotoxin production and fungal pathogenicity in Aspergillus species.

INTRODUCTION

Aspergillus spp. are ubiquitous filamentous fungi found in various climates worldwide (1). Some Aspergillus species can cause allergies, asthma, bronchiectasis, tuberculosis, and can even spread to the brain, kidneys, liver, or other organs (2, 3). Aspergillus-derived diseases particularly affect immunodeficient individuals and exhibit a high mortality rate, ranging from 40% to 90% (4). Several Aspergillus species produce harmful mycotoxins, including aflatoxins, sterigmatocystin, and ochratoxins, which, either directly or through contaminated food, can cause various diseases in humans and animals (5, 6). Therefore, understanding the life cycle of Aspergillus and the mechanisms of mycotoxin production is essential to reduce their detrimental effects on human health.

Aspergillus nidulans is a representative model organism for studies in genetics, fungal developmental biology, and secondary metabolism (7). The spores of A. nidulans typically exist in a dormant state in the air and begin to grow isotropically under favorable conditions. Through a series of cell cycles, polarized spores form germ tubes, elongate, and grow vegetatively (8). The hyphal cells remain interconnected in an undifferentiated state until they encounter environmental signals such as light, temperature, carbon, and nitrogen sources, as well as various genes and proteins (9, 10). When the fungal hyphae become capable of responding to these inducing signals, they undergo asexual or sexual development and form asexual (conidiophores) or sexual structures (cleistothecia) (11, 12).

Asexual development (conidiation) is the primary reproductive pathway in A. nidulans (13). The mechanisms of conidiation are initiated by various genetic regulators, including the Flb proteins, Nsd proteins, and Velvet proteins (13). Proper expression of brlA mediates the asexual developmental switch from vegetative growth and facilitates the formation of stalks and vesicles by regulating early developmental genes (9). Influenced by BrlA, AbaA facilitates the formation of metulae and phialides during the middle stage of development and coordinates the expression levels of genes such as wetA and vosA (14, 15). In the late stage of conidiation, WetA controls the proper formation of conidiophores and the maturation of conidia (16). These regulators of asexual reproduction also perform similar roles in the toxin-producing fungus Aspergillus flavus (17).

Transcription factors regulate chromatin structure and gene expression by recognizing specific DNA sequences (18). The zinc cluster family is the largest group of transcription factors that regulate fungal growth, morphology, and differentiation in Aspergillus (19). This family includes classical Gal4-like Zn(II)₂Cys₆ and Cys₂His₂ zinc finger types, as well as non-classical types such as RING-type, PHD-type, and LIM-type (20, 21). Among these, C₂H₂-type transcription factors are the second-largest group of zinc cluster proteins and exhibit pleiotropic functions in A. nidulans biology (22). For example, RocA is a C₂H₂ zinc finger transcription factor that negatively regulates asexual development by repressing the transcription of brlA and positively regulates sexual development by activating the expression of nsdC in A. nidulans (23). VadH, which contains four C₂H₂ zinc finger domains, is involved in the balance between asexual and sexual development and also contributes to osmotic stress tolerance in A. nidulans (24). Another C₂H₂ zinc finger protein, CrzA, plays essential roles in fungal growth, conidiation, and resistance to ion and osmotic stress in A. nidulans (25).

In a previous study, we analyzed conidia-specific and hyphae-specific genes of conidia-specific transcription factors in three Aspergillus species (22). In this study, we identified several zinc finger transcription factors among the hyphae-specific transcription factors and generated four deletion mutants that had not been previously studied in A. nidulans. Among these, we further investigated the roles of a C₂H₂-containing transcription factor, designated HscA (hyphae-specific Cys₂His₂ zinc finger A).

RESULTS

Analysis of hyphae-specific zinc finger transcription factors in A. nidulans

Previous transcriptomic studies have identified conidia- or hyphae-specific genes in A. nidulans, A. fumigatus, and A. flavus (22). In this study, we analyzed hyphae-specific transcription factors across the three Aspergillus species and identified a total of 13 hyphae-specific zinc finger transcription factors: six GAL4-like Zn(II)₂Cys₆ transcription factors and seven Cys₂His₂ transcription factors (Table 1). Among them, five hyphae-specific zinc finger transcription factors had not been previously characterized. To investigate the roles of these five unstudied hyphae-specific zinc finger proteins, we constructed deletion mutants and examined their colony phenotypes in A. nidulans. As shown in Fig. 1A, the fungal colony of the ΔAN12029 mutant exhibited abnormal pigmentation compared to the wild-type strain. Based on these observations, we further analyzed the functions of AN12029 in A. nidulans.

TABLE 1.

List of zinc finger transcription factors significantly upregulated in the hyphae (H) compared to conidia (C) across three Aspergillus species (22)

TF family (n)	A. nidulans			A. fumigatus		A. flavus
	Gene	Name	log2FC(C/H)	Gene	log2FC(C/H)	Gene	log2FC(C/H)
Zn2Cys6 (6)	AN0902		−4.05	Afu1g15680	−3.61	AFLA_083820	−1.37
	AN1736		−3.87	Afu2g00880	−4.3	AFLA_104780	−2.81
	AN1848	nosA	−2.92	Afu6g07010	−2.93	AFLA_025720	−6.76
	AN3075	oefC	−4.35	Afu3g09670	−3.51	AFLA_085170	−1.18
	AN5170	rosA	−1.08	Afu6g07010	−2.93	AFLA_025720	−6.76
	AN5775		−3.85	Afu6g06535	−2.78	AFLA_037760	−8.00
Cys2His2 (7)	AN1652	msnA	−3.18	Afu4g09080	−4.27	AFLA_110650	−5.67
	AN2421	flbC	−2.63	Afu2g13770	−3.81	AFLA_137320	−1.63
	AN4263	nsdC	−2.31	Afu7g03910	−1.35	AFLA_131330	−1.34
	AN5583	aslA	−3.69	Afu4g11480	−6.76	AFLA_027460	−1.91
	AN5659		−8.65	Afu4g13600	−2.43	AFLA_052490	−6.93
	AN9492	amdХ	−1.22	Afu2g17220	−2.44	AFLA_002290	−1.87
	AN12029		−1.86	Afu6g01910	−2.96	AFLA_049760	−1.02

Fig 1.

Identification of HscA in A. nidulans. (A) Colony morphology of wild-type (WT) and deletion mutant strains point-inoculated on solid MM and incubated at 37°C for 5 days. (B) mRNA expression levels of hscA during the life cycle of A. nidulans using qRT-PCR. Time (h) indicates the duration of incubation in liquid culture or post-asexual developmental induction. The letter “C” indicates conidia. (C) Domain analysis of HscA. The lower panel shows the alignment of the C₂H₂ zinc-finger domain from A. nidulans HscA and its homologs from other Aspergillus species (Ani: Aspergillus nidulans, Afu: Aspergillus fumigatus, Afl: Aspergillus flavus, Aor: Aspergillus oryzae, Anig: Aspergillus niger). (D) Phylogenetic tree of HscA homologs identified in fungal species including A. niger CBS 513.88 (XP_001395971.2), A. flavus NRRL 3357 (XP_041140193.1), A. oryzae RIB40 (XP_001816961.3), A. fumigatus Af293 (XP_747886.1), Histoplasma capsulatum (QSS50878.1), Monascus purpureus (TQB77170.1), Penicillium expansum (XP_016593862.1), Saccharomyces cerevisiae s288c (NP_010539.1), Neurospora crassa OR74A (XP_959452.3), Trichoderma reesei QM6a (XP_006966625.1), and Fusarium graminearum PH-1 (XP_011320037.1). The phylogenetic tree was generated using MEGA X software based on the alignment data from ClustalW algorism and the maximum likelihood method with the Poisson correction model. The bootstrap consensus tree inferred from 1,000 replicates represents the evolutionary history of the taxa analyzed. Right panels present domains of HscA homologs in various fungal species.

Identification of HscA

The AN12029 gene encodes a protein that contains a hyphae-specific Cys₂His₂ zinc finger domain; therefore, we referred to it as HscA. To evaluate the specific expression of hscA in hyphae, we investigated its expression levels of hscA in the life cycle of A. nidulans using a quantitative reverse-transcription PCR (qRT-PCR) analysis. The hscA mRNA levels were high during vegetative growth and decreased after the onset of asexual development and in conidia (Fig. 1B). HscA consists of a single C₂H₂ zinc finger domain located at the C-terminus, similar to its homologs in other Aspergillus species (Fig. 1C). HscA is conserved in fungal species and contains a C₂H₂ zinc finger domain in the N-terminal region (Fig. 1D).

The role of hscA in fungal development in A. nidulans

To further investigate the roles of hscA in A. nidulans, we generated an hscA-complemented strain (C' hscA). Wild-type, ΔhscA, and C' hscA strains were point-inoculated onto solid minimal media containing 1% glucose (MMG) and incubated under dark or light conditions for 5 days. As shown in Fig. 2A, deletion of hscA did not affect fungal growth under either dark or light conditions. However, asexual spore production was significantly reduced in the ΔhscA mutant strain compared to the wild-type and C' hscA strains under both dark and light conditions (Fig. 2B). Additionally, the number of cleistothecia increased under both conditions (Fig. 2C). To further assess the role of HscA in sexual development, each strain was point-inoculated onto sexual media and incubated under dark conditions for 7 days (Fig. 2D). The cleistothecia produced by the ΔhscA strain were smaller in size compared to those formed by the wild-type and C' hscA strains (Fig. 2E). Overall, these results indicate that HscA is essential for appropriate fungal development in A. nidulans.

Fig 2.

Phenotypic analysis of the ΔhscA mutant strains in A. nidulans. (A) Colony morphology of wild-type (TNJ36), ΔhscA (TYE49.1), and C’ hscA (TYE56.1) strains point-inoculated on solid MM and incubated for 5 days under dark or light conditions. Bottom panels show close-up views of the colony center (bar = 0.25 µm). (B) Quantitative analysis of the number of asexual spores shown in panel A (**P ≤ 0.01). (C) Quantitative analysis of the number of cleistothecia shown in panel A (***P ≤ 0.001). (D) Colony morphology of wild-type (TNJ36), ΔhscA (TYE49.1), and C’ hscA (TYE56.1) strains point-inoculated on solid sexual media and incubated for 7 days under dark conditions. (E) Measurement of cleistothecia size in wild-type, ΔhscA, and C’ hscA strains shown in panel D (***P ≤ 0.001).

The role of HscA in cell wall integrity and ion depletion stresses in A. nidulans

To investigate the roles of HscA in hyphal stress responses, we examined the tolerance of wild-type, ΔhscA, and C' hscA strains to various stress conditions. Approximately 10⁵ conidia of each strain were point-inoculated on minimal media supplemented with agents causing cell wall integrity stress (Congo Red and Calcofluor White), ion depletion stress (EDTA), osmotic stress (NaCl and KCl), oxidative stress (H₂O₂), and cell membrane integrity stress (SDS). The plates were incubated at 37°C for 5 days. The hscA deletion strains exhibited increased sensitivity against Congo Red, Calcofluor White, and EDTA (Fig. 3A). Deletion of hscA resulted in decreased colony diameters under these conditions (Fig. 3B). In contrast, no significant differences were observed between wild-type, C' hscA, and ΔhscA strains in response to NaCl, KCl, H₂O₂, and SDS (Fig. S1). These results suggest that HscA plays a crucial role in stress responses related to cell wall integrity and ion depletion in hyphae.

Fig 3.

Function of HscA in stress responses in A. nidulans. (A) Colonies of wild-type (WT; TNJ36), ΔhscA (TYE49.1), and C’ hscA (TYE56.1) strains exposed to cell wall integrity and ion depletion stresses. Each strain was point-inoculated on solid MM containing various stress reagents, including Congo Red, Calcofluor White, and EDTA, and incubated at 37°C for 5 days. (B) Relative growth rates of the indicated strains under stress conditions. The relative growth rate was calculated based on colony diameter and normalized to that of colonies grown without stress agents (**P ≤ 0.01 and ***P ≤ 0.001). (C) Trehalose content in conidia of WT, ΔhscA, and C’ hscA strains (**P ≤ 0.01). (D) Conidial viability of WT, ΔhscA, and C’ hscA strains (***P ≤ 0.001). (E) Oxidative, thermal, and UV stress tolerance of wild-type, ΔhscA, and C’ hscA mutant conidia (**P ≤ 0.01 and ***P ≤ 0.001).

To further investigate the roles of HscA in conidial stress tolerance, we measured the amount of conidial trehalose, which plays a key role in stress tolerance in fungi (26). The ΔhscA conidia contained a lower amount of trehalose compared to wild-type and C' hscA conidia (Fig. 3C). Conidial viability of the ΔhscA conidia was also decreased compared with that of the wild-type and C' hscA conidia (Fig. 3D). Furthermore, the ΔhscA conidia were more sensitive to oxidative, thermal, and UV stresses compared to wild-type and C' hscA conidia (Fig. 3E). These results demonstrate that HscA plays a crucial role in stress responses in both hyphae and conidia in A. nidulans.

The function of hscA in sterigmatocystin production in A. nidulans

To determine whether hscA influences sterigmatocystin production in A. nidulans, we extracted secondary metabolites from wild-type, ΔhscA, and C' hscA strains. Each sample was spotted onto TLC plates with sterigmatocystin as a standard. Deletion of the hscA strain produced less sterigmatocystin compared to the wild-type and C' hscA strains (Fig. 4A and B). Additionally, the mRNA level of aflR, the primary regulator of sterigmatocystin biosynthesis, was downregulated in the hscA null mutant (Fig. 4C). Furthermore, in the hscA-overexpressing strain (OEhscA), sterigmatocystin production was higher than in the control under inducing conditions (Fig. S2). These results indicate that HscA functions as an activator of sterigmatocystin production in A. nidulans.

Fig 4.

Effect of HscA on sterigmatocystin production in A. nidulans. (A) Thin-layer chromatography (TLC) analysis of sterigmatocystin in wild-type (TNJ36), ΔhscA (TYE49.1), and C’ hscA (TYE56.1) strains. The black arrow indicates the sterigmatocystin (ST) band. (B) Relative intensity of sterigmatocystin bands shown in panel A (**P ≤ 0.01). The relative band intensity of sterigmatocystin was quantified using ImageJ software. (C) mRNA expression levels of aflR in the indicated strains (*P ≤ 0.05).

Effect of deletion of AflhscA on fungal development in A. flavus

In A. nidulans, HscA is required for proper asexual and sexual development. To examine the role of A. flavus hscA (AflhscA) in growth and development, the control, AflhscA deletion mutant (ΔAflhscA), and AflhscA complemented strains (C' AflhscA) were point-inoculated onto solid MMYE medium. As shown in Fig. 5A, the ΔAflhscA strains exhibited larger colonies than the control and C' AflhscA strains, and the conidiophores of the ΔAflhscA strains were significantly smaller than those of the control and C' AflhscA strains. Moreover, the deletion of AflhscA led to a reduction in the number of conidiospores (Fig. 5B), implying that AflHscA is required for appropriate colony growth and asexual development. We then examined the function of AflhscA in sexual development. After incubating control, ΔAflhscA, and C' AflhscA strains onto sexual media, the number of sclerotia which act as sexual structures in A. flavus was counted. The ΔAflhscA strains produced significantly fewer sclerotia compared to the control and C' AflhscA strains (Fig. 5C and D). Collectively, these results indicate that AflHscA is essential for normal sexual development in A. flavus.

Fig 5.

Phenotypic analysis of the ΔAflhscA mutant strains. (A) Colony morphology of control (TTJ6.1), ΔAflhscA (TKH1.1), and C′ AflhscA (TKH2.1) strains point-inoculated on solid MMYE and incubated at 37°C for 5 days under light conditions. Middle panels show close-up views of the central area of each colony (scale bars = 250 µm). Bottom panels display conidiophore images of control, ΔAflhscA, and C′ AflhscA strains. (B) Under light conditions, colony diameter was measured to assess fungal growth, and the total number of conidia per plate was quantified to evaluate asexual development. Error bars represent standard deviation. Statistical differences between control and mutant strains were estimated using Student’s t-test (*P < 0.05; **P < 0.01). (C) Control, ΔAflhscA, and C′ AflhscA strains were point-inoculated on solid MMYE media and incubated at 37°C for 7 days under dark conditions. After inoculation, plates were washed with 100% ethanol, and sclerotia were counted. (D) Quantitative analysis of sclerotia as an indicator of sexual development. All experiments were conducted in at least triplicate. Error bars indicate standard deviation. Statistical differences between control and deletion mutant strains were estimated using Student’s t-test (*P < 0.05).

The role of AflHscA in stress tolerance

Given that deletion of hscA affects stress tolerance in A. nidulans, we examined whether AflHscA functions in a similar manner. Control, ΔAflhscA, and C' AflhscA strains were point-inoculated on solid MMYE media containing various stressors including Congo Red, EDTA, SDS, and KCl. The plates were incubated at 37°C for 5 days (Fig. 6A). The ΔAflhscA strains showed increased sensitivity to Congo Red, EDTA, and SDS (Fig. 6B), but not to osmotic stressors (Fig. S2). We further examined the amount of trehalose in conidia and stress resistance. We found that deletion of AflhscA resulted in reduced trehalose content in conidia (Fig. 6C) and increased sensitivity to oxidative stress (Fig. 6D). Overall, these findings suggest that AflHscA plays an important role in stress tolerance in both hyphae and conidia of A. flavus.

Fig 6.

Phenotypes of the ΔAflhscA mutant under various stress conditions in A. flavus. (A) Colonies of control, ΔAflhscA, and C′ AflhscA strains subjected to cell wall integrity and ion depletion stresses. Each strain was point-inoculated on solid MM containing various stress reagents, including Congo Red, EDTA, and SDS, and incubated at 37°C for 5 days. (B) Relative growth rates of the designated strains under stress conditions. The relative growth rate was calculated by measuring the diameter of the fungal colonies and normalized to the diameter of fungal colonies grown without stress agents (***P ≤ 0.001). (C) Trehalose content in the conidia of control, ΔAflhscA, and C′ AflhscA strains (**P ≤ 0.01). (D) Oxidative stress tolerance of control, ΔAflhscA, and C′ AflhscA mutant conidia (***P ≤ 0.001).

The function of AflHscA in aflatoxin B1 production

To investigate whether AflHscA influences aflatoxin B1 production, control, ΔAflhscA, and C' AflhscA strains were inoculated into liquid CM medium and incubated at 30°C for 7 days under dark conditions. Following this, aflatoxin B1 was extracted from each sample, and the amount of aflatoxin B1 was analyzed using thin-layer chromatography. We found that the ΔAflhscA mutant strain produced a lesser amount of aflatoxin B1 compared to the control or complemented strains (Fig. 7). These results suggest that AflHscA is crucial for proper aflatoxin B1 production.

Fig 7.

Effect of AflHscA on aflatoxin B1 production in A. flavus. (A) Thin-layer chromatography (TLC) analysis of aflatoxin B1 in control, ΔAflhscA, and C′ AflhscA strains. The black arrow indicates aflatoxin B1 (AFB1). (B) Relative intensity of the aflatoxin B1 bands shown in panel A (*P ≤ 0.05). The relative band intensity of aflatoxin B1 was quantified using ImageJ software.

The role of AflHscA in pathogenicity

To investigate the role of AflHscA in fungal pathogenicity, a kernel infection assay was performed. Conidia of the control, ΔAflhscA, and C' AflhscA strains were infected onto maize kernels and cultured at 30°C for 7 days under dark conditions (Fig. 8A). After incubation, the ΔAflhscA strain produced fewer conidia compared to the control and C’ AflhscA strains (Fig. 8B). However, in the ΔAflhscA mutant, aflatoxin B1 production was lower than in the control strain, but the difference was not statistically significant (data not shown). Overall, these results suggest that AflHscA is required for proper fungal development and contributes to plant pathogenesis.

Fig 8.

Maize kernel infection assay. (A) Phenotypes of uninfected corn seeds (Mock) and corn seeds infected by control, ΔAflhscA, and C′ AflhscA strains after incubation at 30°C for 7 days under dark conditions. The bottom row shows enlarged views of each maize seed. (B) Quantitative analysis of conidia production from each infected maize kernel. This experiment was performed in triplicate for each strain. Statistical differences between control and deletion mutant strains were assessed using Student’s t-test (*P < 0.05).

DISCUSSION

Transcription factors have been reported to play various roles in fungal growth, development, secondary metabolite production, and pathogenicity (19, 21). In a previous study, we analyzed mRNA expression in hyphae and conidia and performed RNA-seq analysis to identify hyphae- or conidia-specific genes (22). We also previously investigated the function of the SscA transcription factor in three Aspergillus species (22, 27, 28). In this study, we focused on genes with higher mRNA expression in hyphae compared to conidia and investigated genes encoding proteins belonging to the zinc finger family (Table 1). Among the 13 transcription factors identified, the functions of seven (NosA, OefC, RosA, MsnA, FlbC, NsdC, AmdX, and AslA) have been previously studied (29, 30). Interestingly, these transcription factors which exhibit higher mRNA expression in hyphae than in conidia are involved in various aspects of fungal biology. For instance, MsnA is known to participate in stress responses and developmental regulation in hyphae (31). FlbC is a key transcription factor that regulates the expression of brlA, a transcription factor that initiates asexual development (32). Additionally, NosA, RosA, and NsdC have been implicated in sexual reproduction (33–35). AslA promotes normal asexual development and represses sterigmatocystin biosynthesis in A. nidulans (36). Furthermore, AslA is also associated with osmotic stress resistance and vacuolar transporters (37). In this study, we further examined five transcription factors whose functions remain unknown (Fig. 1A). Among them, the deletion mutant of AN12029 showed significant phenotypic differences, leading us to designate this gene as hscA. We subsequently conducted a detailed investigation of hscA in A. nidulans and A. flavus.

In this study, we investigated the function of hscA in the growth and development of A. nidulans and A. flavus. In A. nidulans, the hscA deletion mutant did not affect growth but resulted in a reduction in conidia production, an increase in the number of sexual reproductive structures, and a decrease in cleistothecia size (Fig. 2). In A. flavus, the hscA deletion mutant exhibited increased colony growth but showed abnormal conidiophore structures, along with a decrease in the production of both conidia and sclerotia (Fig. 5). These findings suggest that HscA plays a crucial role in the production of both asexual and sexual reproductive structures, although its specific role may differ between Aspergillus species.

HscA was also found to play a similar role in stress responses in both A. flavus and A. nidulans (Fig. 3 and 6). The hscA deletion mutants of both species exhibited reduced growth on media containing cell wall inhibitors or EDTA, indicating their involvement in stress sensitivity. Furthermore, the hscA deletion mutants exhibited decreased trehalose contents and increased sensitivity to oxidative stress in the conidia of both Aspergillus species. These findings suggest that HscA plays a crucial role in stress responses in both hyphae and asexual spores and that this function appears to be well conserved across Aspergillus species.

Aspergillus species produce various mycotoxins, including sterigmatocystin, aflatoxins, and ochratoxins (5). Notably, sterigmatocystin produced by A. nidulans and aflatoxin B1 produced by A. flavus share similar chemical structures and biosynthetic pathways (38). In this study, we found that hscA deletion reduced sterigmatocystin production in A. nidulans (Fig. 4). Similarly, the deletion of hscA in A. flavus led to decreased aflatoxin B1 production (Fig. 7), suggesting that HscA may play a similar role in the biosynthesis of both sterigmatocystin and aflatoxin B1. However, results from the kernel assay indicated that aflatoxin B1 production in the hscA deletion mutant of A. flavus was comparable to that of the control strain. This suggests that under certain environmental conditions, the reduction in aflatoxin B1 production caused by hscA deletion may be compensated. Therefore, while HscA plays an important role in aflatoxin B1 biosynthesis, it is not essential for its production.

In summary, we investigated hyphae-specific zinc finger transcription factors and characterized a hyphae-specific C₂H₂-type zinc finger transcription factor, HscA, in the model organism A. nidulans and the toxigenic fungus A. flavus. Our phenotypic analyses demonstrate that HscA is essential for regulating fungal differentiation, tolerating environmental stresses, and producing mycotoxins. Further studies will provide deeper insights into the genetic regulatory mechanisms of HscA in A. nidulans, and additional research will help clarify its conserved roles in other Aspergillus species.

MATERIALS AND METHODS

Strains, media, and culture conditions

Fungal strains used in this study are listed in Table 2. For general purposes, A. nidulans strains were cultured on liquid or solid minimal media with 1% glucose (MMG) at 37°C. To induce sexual development, A. nidulans strains were grown on solid sexual media at 37°C (39). Each A. flavus strain was cultured on MMG supplemented with 0.1% yeast extract (MMYE) at 37°C. For auxotrophic strains, uridine/uracil or pyridoxine was added to the medium as required. Escherichia coli DH5α cells were grown in Luria-Bertani (LB) medium supplemented with ampicillin (100 µg/mL) for plasmid amplification.

TABLE 2.

Aspergillus strains used in this study

Strain name	Relevant genotype	Reference
A. nidulans
FGSC4	Wild type, veA+	FGSCa
RJMP1.59	pyrG89; pyroA4, veA+	(40)
TNJ36	pyrG89; AfupyrG +; pyroA4, veA+	(32)
THS30	pyrG89; AfupyrG +	(41)
TYE1.1~3	pyrG89; pyroA4; ∆AN0902::AfupyrG+, veA+	This study
TYE46.1~3	pyrG89; pyroA4; ∆AN5775::AfupyrG+, veA+	This study
TYE47.1~3	pyrG89; pyroA4; ∆AN5659::AfupyrG+, veA+	This study
TYE48.1~3	pyrG89; pyroA4; ∆AN1736::AfupyrG+, veA+	This study
TYE49.1~3	pyrG89; pyroA4; ∆AN12029::AfupyrG+, veA+	This study
TYE56.1~2	pyrG89; pyroA::hscA(p):: hscA::FLAG3x::pyroAb; ∆ hscA::AfupyrG+, veA+	This study
TYE123.1	pyrG89; AfupyrG+; pyroA::alcA(p):: hscA::FLAG3x::pyroAb, veA+	This study
A. flavus
NRRL 3357	Wild type (WT)	FGSCa
NRRL 3357.5	pyrG−	(42)
TTJ6.1	pyrG−; ΔAflpyrG::AfupyrG+	(43)
TKH1.1-2	pyrG−; ΔAflhscA::AfupyrG+	This study
TKH2.1-2	pyrG−; AflhscA(p)::AflhscA::FLAG4x::ptrA; ΔAflhscA::AfupyrG+	This study

^a Fungal Genetic Stock Center.

^b The 3/4 pyroA marker allows targeted integration at the pyroA locus.

Generation of gene deletion mutants

The oligonucleotide primers used in this study are detailed in Table 3. The double-joint PCR (DJ-PCR) method was used to generate gene deletion mutant strains (44). The 5′ and 3′ flanking regions of each gene were amplified using the primer pairs 5′DF/3′ tail or 5′tail/3′ DR, respectively, with A. nidulans FGSC4 genomic DNA (gDNA) or A. flavus NRRL 3357 gDNA as templates. The A. fumigatus pyrG (AfupyrG) marker was amplified with the primer pair OHS1542/OHS1543 using A. fumigatus AF293 gDNA as a template. The final PCR constructs for each gene deletion were amplified with the primer pair 5′ NF/3′ NR of the corresponding gene. The deletion cassettes were introduced into A. nidulans RJMP1.59 or A. flavus NRRL 3357.5 protoplasts, which were generated using Vinoflow FCE lysing enzyme (Novozymes, Bagsvaerd, Denmark) (45, 46). Each resulting strain was independently isolated and finally verified through PCR, restriction enzyme treatment, and qRT-PCR analyses.

TABLE 3.

Oligonucleotides used in this study

Name	Sequence (5′ → 3′)a	Purpose
OHS1542	CCTGGTCTTTGGTTTGGTACACC	5′ AfupyrG marker_F
OHS1543	CGACTGGCAGGAGATGATCC	3′ AfupyrG marker_R
OHS0214	GAGGCATGGCATTGGCTTTG	5′ AN0902 DF
OHS0216	GGCTTTGGCCTGTATCATGACTTCA CGAGGTTGAGAACTCGAGCCTTC	3′ AN0902 with AfupyrG tail
OHS0217	TTTGGTGACGACAATACCTCCCGAC CGACGTGGAACGTTTAATTGGC	5′ AN0902 with AfupyrG tail
OHS0215	TGGTTGCGGTGGTTGAGGAAG	3′ AN0902 DR
OHS0218	GAGCTATAACCCTTGTCGATGGC	5′ AN0902 NF
OHS0219	GAGGAAGGAGTGTGCGGTGTC	3′ AN0902 NR
OHS0441	GGGCATCCCAGTGCTACTTT	5′ AN0902 RT_F
OHS0442	TTCTCCGCAGCAACCCTATC	3′ AN0902 RT_R
OHS1614	GGC CTA TGT GGT CCT CGA	5′ AN1736 DF
OHS1574	GCTAACCCTCTTACCGCAGT	3′ AN1736 with AfupyrG tail
OHS1575	GGCTTTGGCCTGTATCATGACTTCA GGTTCAAGCGTGTCGAACTAG	5′ AN1736 with AfupyrG tail
OHS1576	TTTGGTGACGACAATACCTCCCGAC GTGCTGTTTACCAGGCGAG	3′ AN1736 DR
OHS1577	GATGATGCGGTTTCGACCG	5′ AN1736 NF
OHS1578	CACTATCTCGCAACGAACGG	3′ AN1736 NR
OHS1579	GCGAACCCTAATGCCAGG	5′ AN1736 RT_F
OHS1580	CTGGTGGAGAACCAGCCATA	3′ AN1736 RT_R
OHS1581	GTTGGTGAGCTCTGGTGAAC	5′ AN5775 DF
OHS1582	GAGAGACCAGGCCTCGAG	3′ AN5775 with AfupyrG tail
OHS1583	GGCTTTGGCCTGTATCATGACTTCA GAGTCGCGAGCTGGTCT	5′ AN5775 with AfupyrG tail
OHS1584	TTTGGTGACGACAATACCTCCCGAC GAGTCGCGAGCTGGTCT	3′ AN5775 DR
OHS1585	GTATTCTGCGACGCCGTG	5′ AN5775 NF
OHS1586	ATCTCTGAAGCGAGCTCCT	3′ AN5775 NR
OHS1587	GGACTATCGACTGGAGGTCC	5′ AN5775 RT_F
OHS1588	CGAGCCCACTATCCGAGTAT	3′ AN5775 RT_R
OHS1589	GGCTGGGATAGACCTGCTTT	5′ AN5659 DF
OHS1590	GGATCAGGCTCGTGGTCT	3′ AN5659 with AfupyrG tail
OHS1591	GGCTTTGGCCTGTATCATGACTTCA GTGCACAGCTGTACCAACC	5′ AN5659 with AfupyrG tail
OHS1592	TTTGGTGACGACAATACCTCCCGAC GTGCACAGCTGTACCAACC	3′ AN5659 DR
OHS1593	GCGAAGTAGTAGAGAGCTGCG	5′ AN5659 NF
OHS1594	GACCTTCCAGGTAGGTCCAG	3′ AN5659 NR
OHS1595	CAGTGCAGGAACCAATACAAGC	5′ AN5659 RT_F
OHS1596	CGTCGAAGATACTGCCAGGT	3′ AN5659 RT_R
OHS1597	TGACGAGATGGAACCGGAAT	5′ AN12029 DF
OHS1598	CATGGACAGTCCGCTGTC	3′ AN12029 with AfupyrG tail
OHS1599	GGCTTTGGCCTGTATCATGACTTCA CAAGGGTCGCCAGGTTTG	5′ AN12029 with AfupyrG tail
OHS1600	TTTGGTGACGACAATACCTCCCGAC CAAGGGTCGCCAGGTTTG	3′ AN12029 DR
OHS1601	CCTGCTCTTCCTGCAAGTG	5′ AN12029 NF
OHS1602	GTATCTGACCGCTCGCTG	3′ AN12029 NR
OHS1603	CATCGAAGTGCCACCAGTC	5′ AN12029 RT_F
OHS1604	CACCGGCTCATATAGTGGGA	3′ AN12029 RT_R
OHS1812	AATT GAATTC CAAGATCATGGACAGTCCGC	5′ hscA with promoter and EcoR1
OHS1813	AATT GAATTC CTCGCCGCTCTCAACCTT	3′ hscA with EcoR1
OHS2748	AATT GCGGCCGC ATGGGGACGGGCCTCG	5′ hscA with Not1
OHS1805	AATT GCGGCCGC CTCGCCGCTCTCAACCTT	5′ hscA with Not1
OHS0044	GTAAGGATCTGTACGGCAAC	5′ actin RT_F
OHS0045	AGATCCACATCTGTTGGAAG	3′ actin RT_R
OHS0580	CAAGGCATGCATCAGTACCC	5′ brlA RT_F
OHS0581	AGACATCGAACTCGGGACTC	3′ brlA RT_R
OHS0779	ATTGACTGGGAAGCGAAGGA	5′ abaA RT_F
OHS0780	CTGGGCAGTTGAACGATCTG	3′ abaA RT_R
OHS0599	GCGCGAAGAAGACTTCAAC	5′ aflR RT_F
OHS0600	TGCAATAACTGCCGACGAC	3′ aflR RT_R
OHS3274	CCTGTCAACCACGTTGTCGG	5′ AflhscA DF
OHS3275	GGCTTTGGCCTGTATCATGACTTCA GAACCGCAAGAGCAGCCA	3′ AflhscA with AfupyrG tail_R
OHS3276	TTTGGTGACGACAATACCTCCCGAC CCCTTGATCTCGAGCGAC	5′ AflhscA with AfupyrG tail_F
OHS3277	CTTCTGGTCCATCCTGGC	3′ AflhscA DR
OHS3278	TCCCGTTCCATTTGCTCG	5′ AflhscA nested NF
OHS3279	TCCAACAAGCCGACGCAG	3′ AflhscA nested NR
OHS3280	CGATCCGACCTCGTCCTAC	5′ AflhscA RT_F
OHS3281	GTCGCCGAATTTCGATGGAG	3′ AflhscA RT_R
OHS3318	AATT GCGGCCGC CTCCGTACTTACTCTCGC	5′ AflhscA with promoter and NotI
OHS3319	AATT GCGGCCGC GCCATGGTCCAGGTCTTT	3′ AflhscA NotI

^a Tail sequences are shown in italics. Restriction enzyme sites are in bold.

Construction of hscA-complemented strains

To generate the ΔhscA complemented strain in A. nidulans, the wild-type hscA gene region, including its predicted promoter, was amplified using the primer pair OHS1812/OHS1813, digested with EcoRI, and cloned into pHS13 (47). The resulting plasmid, pYE13.1 was then introduced into the recipient ΔhscA strain TYE49.1, yielding TYE56.1~2. The complemented strains were screened by PCR and confirmed by quantitative reverse transcription- PCR (qRT-PCR).

To generate AflhscA-complemented strains in A. flavus, the promoter and open reading frame (ORF) region of AflhscA was amplified using primers OHS3318 and OHS3319. The PCR product and pYES1 (27) were digested with NotI and cloned. The resulting plasmid, pKH1.1, was transformed into the ΔAflhscA mutant, yielding strains TYE56.1~2. Ultimately, strain TKH2.1-2 was obtained as a complemented strain and verified through PCR, qRT-PCR, and phenotypic analyses.

Nucleic acid isolation and qRT-PCR analysis

To isolate gDNA, approximately 10⁶ conidia of each strain were inoculated into 2 mL of liquid MMG supplemented with 0.5% yeast extract and incubated at 37°C for 24 h. The mycelial mat was harvested, and squeeze-dried, and genomic DNA was extracted as previously described (46).

For total RNA preparation, fresh conidia were inoculated into liquid MM or MMYE and incubated at 37°C for 12, 18, and 24 h. To induce developmental stages, mycelia grown for 18 h were harvested, washed, and transferred to solid MM or MMYE, and incubated at 37°C under either light or dark conditions. Samples were collected at designated time points and stored at −80°C until RNA extraction (48).

For total RNA extraction, each sample was homogenized using a Mini-Bead Beater (BioSpec Products Inc., Bartlesville, OK, USA) with 1 mL of TRIzol reagent (Geneall, Seoul, South Korea) and 0.3 mL of glass beads (Daihan Scientific, Wonju, South Korea). After centrifugation, the supernatant was transferred to a new tube and mixed with an equal volume of cold isopropanol. Following a second centrifugation and removal of the supernatant, the RNA pellet was washed with 70% ethanol treated with diethyl pyrocarbonate (DEPC, Bioneer, Daejeon, South Korea). cDNA was synthesized from total RNA using reverse transcriptase (Promega, Madison, WI, USA). qRT-PCR assay was performed using the iTaq Universal SYBR Green Supermix (Bio-Rad, Hercules, CA, USA) on a CFX96 Touch Real-Time PCR system (Bio-Rad, USA). The β-actin gene was used as a control.

Stress sensitivity assay

For A. nidulans, approximately 10⁵ conidia of each strain were point-inoculated onto solid MM supplemented with various stress-inducing agents and incubated at 37°C for 5 days. The following stress agents were used: cell wall integrity stress: 15 and 30  µg/mL Congo Red (Sigma, St Louis, MO, USA), 2.5 and 5.0 µg/mL Calcofluor White (Thermo Fisher, Waltham, MA, USA); ion depletion stress: 1.5 and 3.0 mM ethylenediaminetetraacetic acid (EDTA; Bioneer, Daejeon, South Korea). Relative growth rates were calculated using the following formula: relative growth rate = fungal colony diameter with stress agent/fungal colony diameter without stress agent. Each experiment was performed in triplicate per strain.

For A. flavus, each strain was point-inoculated onto solid MMYE medium containing 75 µg/mL Congo Red (Sigma, USA), 2.5 mM EDTA (Bioneer, South Korea), and 0.012% sodium dodecyl sulfate (SDS), and incubated at 37°C under dark conditions for 5 days. After incubation, colony sizes (colony diameter, cm) were measured. All experiments were performed in triplicate.

Sterigmatocystin extraction and thin-layer chromatography (TLC) analysis

For sterigmatocystin extraction from A. nidulans, approximately 10⁵ conidia of each strain were inoculated into 5 mL of liquid complete medium (CM) and cultured at 30°C for 7 days. After incubation, 5 mL of CHCl₃ (Duksan, Seoul, Republic of Korea) was added to each culture, and the samples were centrifuged to separate the organic and aqueous phases. The organic phase was transferred to clean glass vials and evaporated in an oven for 24 h. The dried residue was resuspended in 50 µL of CHCl₃ and spotted onto a TLC silica gel plate with a fluorescence indicator (Kieselgel 60, 0.25 mm; Merck, Rahway, NJ, USA). The TLC plate was resolved in toluene:ethyl acetate:acetic acid (8:1:1, vol/vol) and treated with 1% aluminum hydroxide hydrate (Sigma, St Louis, MO, USA). The developed plates were exposed to ultraviolet illumination at 366 nm and photographed. Spot intensity of sterigmatocystin was quantified using ImageJ software. Experiments were performed in triplicate for each strain.

Aflatoxin B1 extraction and TLC analysis

For aflatoxins B1 extraction from A. flavus, approximately 2 × 10⁶ of each strain were inoculated into 5 mL of liquid CM and incubated at 30℃ under dark conditions for 7 days. After incubation, 5 mL of chloroform (Duksan, Ansan, South Korea) was added, and the samples were vigorously mixed using a Voltex mixer. The mixture was centrifuged to separate the aqueous phase and organic phase. The organic phase was collected, filtered through filter paper, and mixed with distilled water (ddH₂O). Following vortexing and centrifugation, the lower layer was collected and treated with Na₂SO₄, and 2 mL of chloroform was added, and the mixture was vortexed and centrifuged again. The final organic phase was evaporated overnight in an oven. Chloroform (100 µL) was added to dissolve the dried samples, and the prepared samples were loaded onto a TLC silica gel plate (Kieselgel 60, 0.25 mm; Merck KGaA, Darmstadt, Germany). The TLC plate was placed in a chamber containing a chloroform: acetone (9:1, vol/vol) solvent. Aflatoxin B1 bands were visualized under UV light at 366 nm.

Kernel assay

To assess the fungal pathogenicity of each strain, two-day-old conidia were harvested using ddH₂O containing 0.02% Triton X-100 (Sigma, USA), and diluted to a concentration of 2 × 10⁶ conidia/mL. Kernels were washed with 70% ethanol for 5 min using a RotoBot Mini Programmable Rotator (Benchmark Scientific, Sayreville, NJ, USA), followed by treatment with 6% sodium hypochlorite (Samchun, Pyeongtaek, Republic of Korea) for 10 min. The kernels were then rinsed with ddH₂O for 5 min; this process was repeated five times. After drying, each maize kernel was infected with conidia from each strain. Inoculated kernels were incubated at 30℃ incubator under light conditions for 7 days. To quantify conidia production from infected kernels, a hemocytometer was used.

Microscopy

Colony photographs were captured using a Pentax MX-1 digital camera. Photomicrographs were captured using a Leica DM500 microscope equipped with a Leica ICC50 E camera and Leica Application Suite X software.

Statistical analysis

Statistical differences between wild-type or control strains and mutant strains were evaluated using Student’s unpaired t-test. Data are presented as mean ± standard deviation (SD). P values < 0.05 were considered statistically significant (*P ≤ 0.05; **P ≤ 0.01; ***P ≤ 0.001).

SUPPLEMENTAL MATERIAL

The following material is available online at https://doi.org/10.1128/msphere.00254-25.

Supplemental figures. msphere.00254-25-s0001.pdf.

Fig. S1 and S2.

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