The epigenetic reader SntB regulates secondary metabolism, development and global histone modifications in Aspergillus flavus

Abstract

Due to the role, both beneficial and harmful, that fungal secondary metabolites play in society, the study of their regulation is of great importance. Genes for any one secondary metabolite are contiguously arranged in a biosynthetic gene cluster (BGC) and subject to regulation through the remodeling of chromatin. Histone modifying enzymes can place or remove post translational modifications (PTM) on histone tails which influences how tight or relaxed the chromatin is, impacting transcription of BGCs. In a recent forward genetic screen, the epigenetic reader SntB was identified as a transcriptional regulator of the sterigmatocystin BGC in A. nidulans, and regulated the related metabolite aflatoxin in A. flavus. In this study we investigate the role of SntB in the plant pathogen A. flavus by analyzing both ΔsntB and overexpression sntB genetic mutants. Deletion of sntB increased global levels of H3K9K14 acetylation and impaired several developmental processes including sclerotia formation, heterokaryon compatibility, secondary metabolite synthesis, and ability to colonize host seeds; in contrast the overexpression strain displayed fewer phenotypes. ΔsntB developmental phenotypes were linked with SntB regulation of NosA, a transcription factor regulating the A. flavus cell fusion cascade.

1. Introduction:

Aspergillus flavus has garnered the attention of many researchers due to its prevalence as a plant pathogen and potent producer of the highly carcinogenic mycotoxins, aflatoxins (Amare and Keller, 2014; Georgianna and Payne, 2009; Yu, 2012). Aflatoxin was first discovered as the causative agent of the Turkey X disease, where a shipment of contaminated peanuts decimated a large part of the UK poultry industry in 1960 (Nesbitt et al., 1962). Since its discovery, aflatoxin research has shown that it can cause chronic and acute disease in both domestic animals and humans, leading to a range of pathologies from liver disease and cancer, to death in extreme cases due to both toxicity and carcinogenicity issues (Williams et al., 2004; Wogan, 1992).

Aflatoxin is an example of a secondary metabolite, or natural product, which differ from primary metabolites in the regard that they are not required for growth in the lab and are often restricted to synthesis in specific tissues or certain life cycle stages (Lim and Keller, 2014). Despite their dispensable nature in the lab, they are thought to provide organisms an advantage in their ecological niche by helping to protect from biotic or abiotic stresses. For example, Aspergillus strains reduced in secondary metabolite production are, in general, more attractive to insect predation (Rohlfs, 2015) and aflatoxin provides a fitness advantage to A. flavus in confrontations with insects (Drott et al., 2017). Additionally, there are several examples of secondary metabolites as being involved in pathogenicity of both plants and animals (Scharf et al., 2014). Secondary metabolites are produced by biosynthetic machineries that are encoded by genes physically linked in the genome. The physical linking of these secondary metabolite biosynthetic gene clusters (BGCs) aids in the timely coordinated expression and repression of these genes to precisely control the production of these potent bioactive molecules. Regulation varies for each BGC, and they can be influenced by a range of regulators; from specific and globally acting DNA binding transcription factors, to cryptic regulation via epigenetic processes (Gacek and Strauss, 2012; Palmer and Keller, 2010).

The “epi” in epigenetics refers to features that are “on top of” the traditional genetic basis of inheritance and do not include changes to DNA sequences. The most studied form of epigenetic regulation that influences secondary metabolism is regulation through modifications of histone tails (Gacek and Strauss, 2012). Histone tails extend outward from the histone core, and are heavily modified post translationally (Bannister and Kouzarides, 2011). Modifications to these residues influence how tight the association is between DNA and histones, with the more tightly wound DNA being the most difficult for DNA binding transcription factor to access and activate. In general there are three types of proteins which influence the accessibility of chromatin to transcriptionally regulators through histone PTM; writers, erasers, and readers. These terms refer to the function of the protein, whether it places modifications on histone tails (writer), if it removes modifications on histone tails (eraser) or if it assists in the recognition of histone tails (reader).

The first example of epigenetic regulation of secondary metabolism came with the discovery that deletion of the histone deacetylase (HDAC) hdaA increased expression of the sterigmatocystin and penicillin BGCs and their respective products in Aspergillus nidulans (Shwab et al., 2007). Histone acetylation is common in areas of highly transcribed genes (Eberharter and Becker, 2002), so loss of a HDAC should equate to chromatin more accessible to transcriptional machinery and logically fit in with the noted higher transcription of the sterigmatocystin and penicillin BGCs (Shwab et al., 2007). Supporting an euchromatin role of acetylation, histone acetylation of the aflatoxin BGC (similar in gene content and regulation as the sterigmatocystin BGC) correlated with the activation of genes transcribed early, middle, and late in the biosynthesis of aflatoxin (Roze et al., 2007). In A. nidulans, two histone acetyltransferases, GcnE, part of the SAGA/Ada complex (H3K9Ac and H3K14Ac) (Nützmann et al., 2011) and EsaA (H4K12Ac) (Soukup et al., 2012), are associated with acetylation of sterigmatocystin BGC chromatin and overexpression of EsaA was able to partially restore a euchromatin state to the sterigmatocystin BGC in a heterochromatin repressive background (Soukup et al., 2012). Nevertheless, acetylation is not always a predictor of euchromatin as H3K4 acetylation is associated with heterochromatin formation in Schizosaccharomyces pombe (Xhemalce and Kouzarides, 2010) and H4K20 acetylation with heterochromatin formation in human cells (Kaimori et al., 2016).

In opposition to acetylation, heterochromatin formation and methylation of H3K9 were shown to decrease the expression of the sterigmatocystin BGC (Reyes-Dominguez et al., 2010). Loss of CclA, a member of the COMPASS complex required for H3K4 methylation, also decreased synthesis of a sterigmatocystin precursor but resulted in the up-regulation of gene expression and subsequent products of several otherwise cryptic BGCs in A. nidulans (Bok et al., 2009). Down regulation of the histone deacetylase RpdA in A. nidulans also resulted in expression of cryptic BGCs while also repressing sterigmatocystin and other commonly expressed BGCs (Albright et al., 2015). Together, these studies clearly show that the epigenetic regulation of secondary metabolism is multifaceted and complex.

A recent revitalization of a forward genetic screen identified an uncharacterized A. nidulans gene, sntB, encoding an epigenetic reader, as a transcriptional regulator of the sterigmatocystin BGC (Pfannenstiel et al., 2017). Deletion of this gene in A. flavus led to the loss of aflatoxin production, indicating a conserved role of SntB in regulation of these highly similar BGCs where sterigmatocystin is the penultimate precursor to aflatoxin (Keller and Adams, 1995; Pfannenstiel et al., 2017). The first sntB homolog was characterized in Saccharomyces cerevisiae (called Snt2 for its conserved SANT protein domain), and was found to interact with an eraser enzyme, the HDAC Rpd3 (RpdA homolog) (Baker et al., 2013). Snt2 protein interactions have been assessed in Schizzosaccharomyces pombe as well, however this study showed no overlap with those proteins identified in S. cerevisiae, indicating its specific role for Snt2 in these two yeasts may have diverged (Roguev et al., 2004). Lastly, a sntB homolog was identified as a regulator of virulence in Fusarium oxysporum, and respiration in F. oxysporum and Neurospora crassa (Denisov et al., 2011a, 2011b). Considering the wide array of phenotypes and functions that sntB homologs have in other species, we set forth to characterize this protein in A. flavus, initially following a hypothesis that SntB would be a global regulator of secondary metabolism in this species. We found this hypothesis to be valid with SntB regulating at minimum seven characterized A. flavus secondary metabolites. SntB also regulates global histone H3 acetylation and that its loss significantly modifies development and decreases pathogenicity on host seed.

2. Materials and Methods:

2.1. Strains and Culture Conditions:

Strains used in this study are listed in Table S1, and were grown on glucose minimal media (GMM) with additional supplements for auxotrophic strains unless otherwise noted. Solid and liquid cultures were grown in a light incubator at 30 °C. All strains were maintained as glycerol stocks at −80 °C. Strain TJW149 is used as our wild-type control unless otherwise noted.

2.2. Strain Construction:

sntB was overexpressed via transformation with PCR products generated from double joint PCR and the primers listed in Table S2 (van Leeuwen et al., 2015; Yu et al., 2004). The overexpression sntB strain, BTP107, was generated by amplifying a 1.0 kb fragment of the native promoter (OE:sntB 5’F and OE:sntB 5’R) and the first 1.0 kb of the open reading frame (OE:sntB 3’ F and OE:sntB 3’ R). These two fragments were fused with A. fumigatus pyrG::gpdA(p) in a double joint PCR reaction (Yu et al., 2004). TJES19.1 was transformed with this construct and the resulting transformant BTP107 was confirmed by Southern blot analysis, using the 5’ flanking region as a probe labeled with dCTP α-P32.

2.3. Radial Growth, Spore production, and Sclerotia Formation:

Radial growth was assessed by measuring the diameter from leading hyphae on either side of a 1,000 spore point inoculated plate of GMM every day for 7 days and the experiment triplicated. Spore production was assessed by creating a spore overlay. 5 mL of molten agar cooled to 56 °C containing 10⁶ spores/mL were inoculated and grown at 30 °C for 2 and 3 days. At the respective time point, a 15 mm core was taken and homogenized in 3 mL of 0.01% Tween 20. Spore solutions were enumerated via hemocytometer, and performed in triplicate.

Sclerotia formation was assessed on GMM+1.2 M sorbitol. Five milliliters of a 1,000 spores/mL of molten GMM+1.2 M sorbitol top agar was poured on 10 cm plates containing GMM+1.2 M sorbitol. These were incubated in the dark for 5 days, washed with 70% ethanol, sclerotia scrapped off and weighed. Experiment was performed with four replicates.

2.4. RNA Extraction and semi quantitative PCR:

RNA was extracted from lyophilized tissue, and total RNA was then isolated using Trizol (Invitrogen). To extract RNA from mycelia, strains were grown in 50 mL liquid cultures, shaking at 250 RPM and 30 °C, at a concentration of 10⁶ spores/mL. Cultures were filtered through Miracloth (CalBioChem), and lyophilized. When examining gene expression during sclerotia development, 500 mL liquid cultures of GMM was grown at 30 °C at 250 RPM for 24 hours, then equal amounts of mycelia was transferred to GMM+2% sorbitol plates. These were stored at 30 °C in a dark environment for the indicated amount of days, flash frozen in liquid nitrogen and lyophilized.

For semi quantitative PCR, RNA was quantified and 10 μg was DNaseI treated before cDNA synthesis using iScript (Bio-Rad). 50 ng was used in each PCR reaction, with the ubiD gene used as a loading control. Primers used in semi-qPCR are listed in Table S2.

2.5. Heterokaryon Formation:

Hyphal fusion and heterokaryon formation was assessed as previously reported (Zhao et al., 2017). Briefly, the pyrG auxotroph TJES19.1, argB auxotroph TJES20.1, pyrG auxotroph ΔlaeA TXZ15.1, and pyrG auxotroph ΔsntB TXZ23.1 were grown on GMM with supplements. Conidia were collected and the pyrG auxotrophs were mixed pairwise in equal numbers to TJES20.1. 10⁷ conidia were spotted on GMM with arginine (1 g/L), uracil (5 mM), and uridine (5 mM) for five days at 29 °C. Conidia were collected, and 10⁵ conidia were spread on GMM with 0.25% Triton X-100 with no added supplement. Colonies were counted after 3 days growth at 29 °C. This experiment was performed in triplicate and repeated twice.

2.6. Pathogenicity Assays:

Pathogenicity assays were conducted as previously described (Christensen et al., 2012), with some modifications. Corn kernels (Blue River organic hybrid) were washed with ethanol for 5 min, rinsed with sterile water, and shaken in bleach for 10 min. After rinsing three times in sterile water, kernels were dried on sterile paper towels. Kernels were wounded with a sterile needle in their embryo. Four kernels were placed in sterile scintillation vials, weighed, and inoculated with 200 μL of a 10⁶ spores/mL solution. Vials were briefly vortexed to coat kernels in spore mixture. Kernels wounded and inoculated with 0.01% Tween 20 were used as a negative, mock control. Five replicates were used for each treatment, and the entire experiment was repeated twice. After caps of scintillation vials were loosened, the vials were placed in a chamber with wet paper towels, and covered with plastic wrap. This humidity chamber was kept at 29 °C for 5-days in a 12-hour light and dark cycle.

Following the 5-day incubation, 2.5 mL of methanol was added to the kernels and they were vortexed to remove spores. 100 μL was removed, and the conidia were enumerated via a hemocytometer. 5 mL of chloroform was then added to the kernels, and vials were kept in the dark overnight for extraction of aflatoxin. 3 mL of the methanol:chloroform mixture from the kernels was removed and dried in vacuum. Samples were resuspended in 1 mL of a 50:40:10 mixture of water, methanol, and acetonitrile. Aflatoxin B₁ was quantified using a PerkinElmer Flexar instrument equipped with a Zorbax Eclipse XDB-C18 column (Agilent) (150 mm × 4.6mm; 5 μm pore size). Aflatoxin B1 was detected with a Flexar fluorescent light (FL) detector (PerkinElmer) with the excitation wavelength set to 365 nm and the emission wavelength set to 455 nm. The column was run isocratically, with a 50:40:10 mixture of water: methanol: and acetonitrile run at 1.5 mL/min.

2.7. Secondary Metabolite Analysis:

For secondary metabolite analysis, strains were point inoculated and grown at 29 °C on PDA media grown on 25 mL of PDA in a 90 mm petri dish wrapped with Parafilm at 29 °C. After 12 days, whole agar plates were blended and soaked in ethyl acetate (100 mL). After 2 hours, the solid was removed using vacuum filtration and the organic layer was separated. The aqueous layer was extracted with ethyl acetate (2 × 25 mL). The combined organic phases were dried over anhydrous magnesium sulfate and concentrated under reduced pressure. The crude extracts were resuspended in acetonitrile (10 mg/mL) and filtered through an Acrodisc syringe filter with a nylon membrane (Pall Corporation) (0.45 μm pore size). Ultra-high-performance high resolution mass spectrometry (UHPLC-HRMS) was then performed on a Thermo Scientific-Vanquish UHPLC system connected to a Thermo Scientific Q Exactive Orbitrap mass spectrometer in ES⁺ mode between 200 m/z and 1000 m/z to identify metabolites. A Zorbax Eclipse XDB-C18 column (2.1 × 150 mm, 1.8 μm particle size) was used with a flow rate of 0.2 ml/min for all samples. LCMS grade water with 0.5% formic acid (solvent A) and LCMS grade acetonitrile with 0.5% formic acid (solvent B) were used with the following gradient 0 min, 20% Solvent B; 2 min, 20% Solvent B; 15 min, 95% Solvent B; 20 min, 95% Solvent B; 20 min, 20% Solvent B; 25 min, Solvent B. Nitrogen was used as the sheath gas. Data acquisition and procession for the UHPLC-MS were controlled by Thermo Scientific Xcalibur software. Files were converted to the .mzXML format using MassMatrix MS Data File Conversion, and analyzed in MAVEN and XCMS (Clasquin et al., 2012; Melamud et al., 2010; Smith et al., 2006).

For assessing production of kojic acid, strains were point inoculated on Kojic Acid Media (KAM) as previously published (Chang et al., 2017). Strains were incubated at 29 °C for 10 days and photographed.

2.8. Western analysis of histone modifications:

50 mL cultures of liquid YES media were inoculated with 10⁶ spores/mL, and incubated at 30 °C shaking at 200 RPM for 72 hours. Nuclear extracts were isolated as previously described, with the exception that the final extracts were resuspended in ChIP sonication buffer (Bernreiter et al., 2007; Palmer et al., 2008). Approximately 50 μg of protein were run on 12% Bis-tris gel and transferred to a PVDF membrane. Histones and their modifications were detected using the anti-acetyl-histone H4 (H4K5K8K12K16Ac) (Millipore Sigma, 06–866), anti-H3K4me3 (Active Motif, Cat#39916, 1:2000), anti-acetyl-histone H3 (Millipore Sigma, 06–599, 1:5000), and anti-histone H3 (abcam, ab1791, 1:5000). A secondary goat anti-rabbit alkaline phosphatase conjugated antibody (Pierce, #31342) was used for detection with PierceTM ECL Plus Western Blotting Substrate (Thermo Scientific, #32132) as substrate.

2.9. Chromatin Immunoprecipitation:

50 mL cultures of liquid YES media were inoculated with 10⁶ spores/mL, and incubated at 30 °C shaking at 200 RPM for 36 hours. Strains were grown in duplicate. Chromatin immunoprecipitation (ChIP) was carried out as previously described (Bernreiter et al., 2007). Immunoprecipitation was performed with anti-acetyl-histone H4, and anti-histone H4 (Abcam, ab10158), anti-acetyl-histone H3, and anti-histone H3. 5 μL of anti-acetyl-histone H4 and two μg of anti-histone H4, anti-acetyl-histone H3, and anti-histone H3 were used with 200 mg of total protein in each ChIP experiment. Quantification of precipitated DNA was measured by qPCR using iQ SYBR Green Supermix (Bio-Rad, Cat #170–8882) according to manufacturer’s instructions, each PCR was performed in triplicate. Relative amounts of DNA were calculated by normalizing immunoprecipitated DNA to input DNA. DNA precipitated with anti-acetyl-histone H4 was normalized to amount of DNA precipitated from the anti-histone H4, and same for anti-acetyl-histone H3 and anti-histone H3. qPCR primers are listed in Table S2.

2.10. Statistical Analysis:

GraphPad Prism software (La Jolla, CA, United States) was used for statistical analysis. Statistically significant differences were determined by ANOVA and P < 0.05. The error bars in all figures indicate the standard error of the mean.

3. Results:

3.1. SntB is required for sexual stage development in Aspergillus flavus:

SntB has been previously characterized in the filamentous fungi N. crassa and F. oxysporum and found to regulate asexual and sexual spore formation in these species (Denisov et al., 2011a). To more thoroughly investigate any impact of this protein on spore development in A. flavus, an overexpression sntB strain was constructed (Fig. S1 and S2) to compare to wild type and the previously constructed ΔsntB strain (Pfannenstiel et al., 2017). Because A. flavus transformants often display a marker gene effect, we assessed the mutant strains for impaired growth on arginine, uracil, and uridine, but did not observe any defect (data not shown). Point inoculation of these strains showed a reduction in radial growth in the deletion strain (Fig. 1A&B). Asexual development was determined to not be impacted by sntB in A. flavus using an overlay culture after two and three days (Fig. 1C and data not shown).

Figure 1:

Growth phenotypes of sntB mutants. A) Stains were point inoculated on GMM and grown at 30 °C for 7 days under constant light. B) Radial growth of plates was measured, all plates were grown in triplicate. C) Conidia were enumerated from a core taken from two day old overlay cultures on GMM. D) Sclerotia dry weight measured from overlay cultures grown on GMM+2% sorbitol grown in the dark at 30 °C for six days. E) Images of plates analyzed in D. Plates were washed with 70% EtOH to remove conidia and allow for visualization of sclerotia. P-value **p < 0.01.

In N. crassa the deletion of the sntB homolog led to a block in perithecium formation, the sexual spore containing structure (Denisov et al., 2011a). Similar to N. crassa, A. flavus is heterothallic, requiring two mating types to undergo sexual development, however the production of sexual spores can take six to eleven months (Horn et al., 2009). These sexual spores are produced in sclerotia, which are black over-wintering bodies. Unlike the ascospores, sclerotia are produced within a week and easily counted. Both the deletion and overexpression strains were greatly decreased in sclerotia production as compared to wild type, with the overexpression able to produce a small amount while no dry weight was able to be measured for the deletion strain (Fig. 1D&E).

3.2. SntB regulates sclerotia development through nosA and heterokaryon formation:

We were interested to further explore the mechanisms of the severe reduction of sclerotia production in the sntB mutants. Sclerotia development is regulated by several transcription factors which orchestrate sexual development, including nsdD (never in sexual development), nsdC, sclR (sclerotium regulator) and nosA (number of sexual spores) (Cary et al., 2012; Chang et al., 2017; Zhao et al., 2017). An examination of transcript patterns showed that nosA was the most down regulated in both the OE::sntB strain and the ΔsntB strain (Fig. 2A).

Figure 2:

Loss of sclerotia in the deletion mutant is through transcriptional regulation of nosA. A) Semi-qPCR analysis of gene expression from three time points, vegetative growth at 24 hours, and then RNA from mycelia that was transferred to small GMM+2% sorbitol plates after one and three days. ubiD is used as a loading control. B) The number of heterokaryotic conidia was enumerated for three heterokaryotic crosses. Positive control was TJES19.1 (pyrG-) and TJES20.1, negative control ΔlaeA crossed with TJES20.1, and ΔsntB strain crossed with TJES20.1. P-value **p < 0.01.

Deletion of nosA yields a strain unable to produce sclerotia (Zhao et al., 2017). Sclerotial formation requires NosA positive regulation of a cell fusion cascade required for hyphal anastomosis (Zhao et al., 2017). Hyphal anastomosis, or hyphal fusion, is a requirement for several biological processes including sclerotial and heterokaryon formation. To further test the hypothesis that sclerotia loss in the ΔsntB strain was likely due to inhibition of NosA function, the ΔsntB mutant was assessed for its ability to undergo hyphal fusion through a heterokaryon formation assay. Heterokaryon formation is easily detected by mixing conidia of different auxotrophies (in this case pyrG and argB auxotrophs) and observing if colonies can grow on media containing no supplements. Heterokaryon formation was compared between wild type, ΔlaeA (previously shown to be required for heterokaryon formation (Zhao et al., 2017), and ΔsntB auxotrophs. The deletion of sntB showed the same phenotype as the ΔlaeA strain, an inability to form heterokaryons (Fig. 2B). Thus, it appeared that the loss of sclerotia in the sntB deletion strain is at least in part due to the loss of hyphal fusion through down regulation of nosA.

3.3. Seed colonization is impaired in sntB deletion strains:

sntB was identified in the plant pathogen F. oxysporum in a genetic screen for decreased virulence on muskmelon (Denisov et al., 2011a). Considering this finding as well as the previous study showing NosA to be an A. flavus virulence factor on corn seed (Zhao et al., 2017), we examined if loss or overexpression of sntB affected colonization of host seed. Corn kernels were wounded and inoculated with spores from wild type, deletion, and overexpression strains. The deletion of sntB led to a significant decrease in its ability to colonize seed as determined by near inability to produce spores on wounded seed (Fig. 3A-B). Aflatoxin production was similarly decreased in the deletion strain, matching the results seen from previous publications (Fig. 3C) (Pfannenstiel et al., 2017). The overexpression did not show any statistically significant deviation from the wild type in either assay (Fig. 3).

Figure 3:

SntB is required for pathogenicity of corn. A) corn kernels were inoculated with 2×10⁵ spores and kept under a 12 hour light and dark cycle for five days. Each strain was inoculated with 5 reps, including a mock control which was kernels inoculated with 0.01% Tween 20. B) Spores were removed from surface of kernels and enumerated using a hemocytometer. Spore counts were corrected by weight of corn. C) Relative aflatoxin production of mutants compared to wild type. P-value, **p < 0.01, ***p < 0.001.

3.4. SntB is a global regulator of secondary metabolism:

Considering the global impact deletion or down regulation of erasers and writers have on the secondary metabolome in fungi (Gacek and Strauss, 2012), we were interested if SntB regulation of secondary metabolism extended beyond aflatoxin in A. flavus. The abundance of known secondary metabolites in A. flavus was assessed using Ultra High Performance Liquid Chromatography paired with a High Resolution Mass Spectrometer (UHPLC-HRMS) from 12 day old cultures grown on PDA media. Both sntB mutants were compared to wild type using the program XCMS and visualized using volcano plots (Fig. 4) (Smith et al., 2006). The deletion strain shows a greater number of metabolites with changes in abundance than the overexpression strain when compared to wild type, thus presenting the most unique profile of the three strains (Fig. 4). Red dots indicate masses matching known secondary metabolites produced in A. flavus.

Figure 4:

Deletion of sntB leads to a greater change in secondary metabolite profile than overexpression. Metabolites were extracted from twelve-day-old cultures on PDA, run on a UHPLC-HRMS, and analyzed via XCMS. Experiment was completed in triplicate. A) Comparison of metabolite extracts from sntB deletion and wild type. Each dot represents a peak called by XCMS. The (–)log10 of the pvalue is plotted on the y-axis, with a gray dashed line indicating where the pvalue is equal to 0.05, values higher on the y-axis indicating higher statistical significance. Log2 of the fold change is on the x-axis, with values in the right half more abundant in the deletion strain, and values on the left half more abundant in the wild type. Red dots indicate known final products that were detected by the program including aflavarin, aflatoxin, asparasone A, ditryptophenaline, and leporin B. B) Same analysis comparing the overexpression of sntB to wild type.

The A. flavus genome has been predicted to encode for 56 putative BGCs, and to date fourteen BGCs have been linked to production of specific secondary metabolites; aflatrem (two clusters) (Nicholson et al., 2009), aflatoxins (Cary et al., 2000), aflavarin (Cary et al., 2015a), asparasone A (Cary et al., 2014), aspergillic acid (Lebar et al., 2018), cyclopiazonic acid (Chang et al., 2009), ditryptophenaline (Saruwatari et al., 2014), imizoquin (Khalid et al., 2018), kojic acid (Terabayashi et al., 2010), leporin B (Cary et al., 2015b), the piperazines (two clusters) (Forseth et al., 2013), and ustiloxin B (Umemura et al., 2014). Visualization using MAVEN identified seven of these metabolites, with six of them exhibiting differences in abundance (Fig. 5, Table S3, Fig. S3). Cyclopiazonic acid did not show any statistical difference between wild type and the sntB mutant strains (data not shown). The ΔsntB strain could not produce several secondary metabolites detected in the wild type strain grown on PDA medium, including aflavarin, aflatoxin B1, asparasone A, and aflatrem. Additionally, when grown on Kojic Acid Media (KAM), the deletion strain could not produce kojic acid, which is the orange pigment seen in the wild type and overexpression strains (Fig. 5B) (Terabayashi et al., 2010). On the other hand, the ΔsntB strain produced higher amounts of ditryptophenaline and leporin B (Fig. 5A–C). The overexpression strain showed a similar secondary metabolite profile as the wild type with the exception of a significant decrease in aflatrem (Fig. 5A). Taken together, SntB is a positive regulator of aflavarin, aflatoxin B₁, asparasone A, aflatrem, and kojic acid, and a negative regulator of ditryptophenaline and leporin B as grown under conditions described here.

Figure 5:

SntB is a global regulator of secondary metabolism in A. flavus. A) Individual graphs of known secondary metabolites produced by A. flavus detected via LCHRMS. Average peak area and standard error of mean was calculated from three biological repetitions. B) Wild type and sntB mutant strains grown on KAM, where kojic acid production is lost in the deletion strain. C) Structure of secondary metabolite analyzed. P-value **p < 0.01, n.d.-not detected.

3.5. SntB regulates global histone modifications:

Although epigenetic readers do not place or remove PTM on histone tails, they do control these modifications through interactions with the cognate eraser and writer enzymes. SntB homologs in S. cerevisiae and S. pombe have been shown to interact with enzymes such as the histone deacetylase Rpd3 in S. cerevisiae and the histone demethylases Lid2 and Jmj3 in S. pombe leading to changes in histone acetylation or methylation (Table 1) (Baker et al., 2013; Roguev et al., 2004). Considering the yeast studies, we hypothesized that loss of sntB in A. flavus would engender discernable global histone modifications. There is no measurable change in hyperacetylation of histone H4 (P=0.22) (Fig. 6A&B), while there is a moderate increase in histone H3K4me3 (P=0.09) (Fig. 6A&C), and a larger increase in histone H3 acetylation of lysines 9 and 14 (P=0.04) (Fig. 6A&D). The increase in H3K9K14 dual acetylation and H3K4me3 levels in ΔsntB suggests that SntB might interact with the HDAC RpdA and the demethylase KdmB.

Table 1:

Protein Interactors of SntB Homologs

Species	Interactor	A. flavus Homolog
Saccharomys cerevisiae	Rpd3	AFLA_092360 (RpdA)
	Ecm5	No
Schizosaccharomyes pombe	Lid2	AFLA_006240 (KdmB)
	Ash2	AFLA_089250
	Sdc1	AFLA_075050
	Jmj3	No

Figure 6:

SntB regulates global levels of histone modifications. A) Levels of histone modifications were assessed by western blot from cultures grown for 72 hours in YES media. B) The relative intensity of the bands corresponding to histone H4 hyper acetylation (H4Ac) were calculated in Adobe PhotoshopTM, and standardized to the loading control, histone H3. Values were normalized to wild type. The same analysis was done for C (H3K4me3) and D (H3K9K14Ac). The deletion of sntB showed an increase in histone H3 acetylation, and a minimal increase in H3K4me3. Histone H3 is used as a loading control.

3.6. Aflatoxin promoters show wild-type histone acetylation:

The A. nidulans ΔsntB strain was unable to produce sterigmatocystin in part through down regulation of the sterigmatocystin regulatory gene, aflR (Pfannenstiel et al., 2017). Semi-quantitative PCR indicated a similar regulation in A. flavus where the aflR gene is not expressed as highly in the deletion strain (Fig. S2). The global change in histone H3K9K14Ac levels in the ΔsntB strain (Fig. 6) coupled with two previous studies associating histone H4 hyperacetylation (Roze et al., 2007) and H3K9 acetylation (Lan et al., 2016) with aflatoxin production, suggested there may be acetylation modifications in aflatoxin gene promoters of ΔsntB. Two of the same promoters examined in the H4 acetylation study (Roze et al., 2007), aflR and aflM as well as the control ubiD promoter, were examined using chromatin immunoprecipitation (ChIP) followed by qPCR under aflatoxin inducing conditions. Levels of H4 and H3, H4 hyperacetylation (H4Ac) and H3 acetylation (H3K9K14Ac) were measured. Contrary to expectations, there is no significant change in histone acetylation levels at H3 or H4 in either mutant compared to wild type (Fig. 7).

Figure 7:

Histone acetylation levels are not changed in sntB mutants at aflatoxin gene cluster promoters. Histone H4, H4Ac, H3, and H3K9K14Ac occupancy levels in wild type, deletion, and overexpression sntB mutants at the aflatoxin BGC after 36 hours of growth in YES media. Promoter regions of aflM and aflR were tested to represent the aflatoxin BGC. ubiD was chosen as an out of cluster control. Error bars represent standard error of mean, which was calculated from biological duplicates.

4. Discussion:

Understanding how secondary metabolism is regulated by chromatin remodeling is both an area of pharmaceutical promise and, in the case of fungal pathogens such as A. flavus, a process to better understand pathways required for toxin synthesis and other virulence attributes. To date, research concerning epigenetic regulation of BGCs has focused on up or down regulating histone modifying enzymes through chemical or genetic means, specifically erasers and writers which actively place or remove PTM on histone tails.

SntB was recently identified in a genetic screen for novel regulators of sterigmatocystin synthesis in A. nidulans, all of which including SntB were found to regulate the similar metabolite, aflatoxin, in A. flavus (Pfannenstiel et al., 2017). We were interested in identifying the function of this particular protein and its role in A. flavus fungal biology, thinking it could uncover clues on how the mycotoxin aflatoxin is regulated as well as provide an in-depth view of the effects of a reader protein on global secondary metabolism synthesis and global histone modifications.

Not surprisingly, considering the effect of sntB loss in the filamentous fungi F. oxysporum and N. crassa, deletion of sntB lead to a pleiotropic response in A. flavus with multiple developmental aberrancies. The most striking result was the complete loss of sclerotia production in the ΔsntB strain (Fig. 1D&E). Also, although the OE::sntB presented nearly wild type responses in other physiological parameters examined in this study, this strain was also affected in sclerotia production. Examination of gene expression of several known transcription factors governing the regulation of sclerotia production revealed nosA transcript was greatly decreased in both mutants (Fig 2A). NosA is required for hyphal fusion, a prerequisite for the formation of sclerotia and heterokaryons, aflatoxin biosynthesis, and virulence on corn (Zhao et al., 2017). Taken together, the findings that the ΔsntB strain was unable to form heterokaryons, to produce aflatoxin in vitro, and was decreased in ability to colonize host seed supports a role for a NosA mediated cascade in directing the effects of SntB on A. flavus biology and pathogenicity.

Because deletion of histone writers and erasers have had global impacts on fungal secondary metabolism (Strauss and Reyes-Dominguez, 2011), we were interested to see if additional secondary metabolites which could be regulated by this reader. Of the approximately dozen characterized A. flavus natural products, UHPLC-HRMS revealed that the loss of SntB leads to a large difference in the secondary metabolome as measured from extracts of strains grown on PDA media (Fig. 4). Five metabolites were decreased in the ΔsntB strain and two, ditryptophenaline and leporin B, were greatly up-regulated (Fig. 5A). Unlike ditryptophenaline which A. flavus can produce under certain inducing conditions (Saruwatari et al., 2014), leporin B synthesis is cryptic and requires overexpression of the BGC specific transcription factor to obtain high levels of this iron coordinating secondary metabolite (Cary et al., 2015b). Interestingly, the overexpression of the leporin BGC led to a decrease in sclerotia production, similar to what was seen in the ΔsntB mutant (Cary et al., 2015b) (Fig. 1&2). Several of the other metabolites are positively correlated with sclerotia production (Chang et al., 2017), and the loss of sclerotia may explain why some of these compounds were not produced in the ΔsntB strain although does not explain the OE::sntB results.

SntB homologs in S. cerevisiae and S. pombe have interacting proteins identified previously, for which there are several homologs in A. flavus (Table 1). This includes a homolog for a HDAC (RpdA) and a H3K4me3 demethylase (KdmB). Our hypothesis was that loss of sntB could lead to global changes in histone PTM it is responsible for removing or placing. In fact, we see a moderate increase in H3K4me3 levels, and a larger increase in H3K9K14Ac levels in the deletion mutant (Fig. 6). This matches recent results, that loss of sntB in Magnaporthe orzyae led to an increase in H3 acetylation (He et al., 2018). These results support a model in which SntB may interact with RpdA or KdmB, however further studies must be conducted to confirm these interactions. Interestingly, if these interactions do occur it would seem that SntB is part of a complex which is a hybrid to what is seen in S. cerevisiae and S. pombe. The aflatoxin/sterigmatocystin BGCs are some of the best studied clusters, particularly in regards to epigenetic regulation. We thought it possible that the aflatoxin cluster would be silenced in the ΔsntB strain through chromatin regulation due to the changed global acetylation pattern (Fig. 6) and decreased aflR expression (Fig.S2). Also, two previous studies have linked histone 4 hyperacetylation (Roze et al., 2007) and H3K9 acetylation (Lan et al., 2016) to active aflatoxin gene expression and subsequent metabolite production. However, there was no difference in H4Ac or H3Ac at either aflR or aflM promoters in ΔsntB suggesting that the loss of aflatoxin is not due directly to chromatin remodeling of the aflatoxin BGC (Fig. 7). This supports the above discussed role of NosA and hyphal anastomosis in aflatoxin synthesis (Zhao et al., 2017) as possibly the dominant reason for diminished aflatoxin loss in the deletion mutant.

Understanding the mechanism of epigenetic regulation of secondary metabolite BGCs can be aided by investigating the functions of epigenetic reading proteins. These are the proteins which give specify and direction to the writing and erasing enzymes. Reading protein domains recognize specific modifications of histones. SntB contains four of these domains, each of which may recognize four distinct histone modifications, a topic of further exploration in our lab. By exploring the roles of reader proteins in fungal secondary metabolism, it is predictable that not only will the research community identify cryptic metabolites but also learn more about the chromatin context of BGCs.

Highlights:

• SntB has a conserved pleiotropic response to genetic manipulation in filamentous fungi

• SntB regulates sclerotia and heterokaryon formation, likely through transcriptional regulation of nosA

• Epigenetic reading proteins can be master regulators of secondary metabolism

• SntB regulates global levels of histone H3 acetylation