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Accepted Manuscript
The epigenetic reader SntB regulates secondary metabolism, development and
global histone modifications in Aspergillus flavus
Brandon T. Pfannenstiel, Claudio Greco, Andrew T. Sukowaty, Nancy P. Keller
YFGBI 3149
To appear in:
Fungal Genetics and Biology
Received Date:
Revised Date:
Accepted Date:
4 June 2018
13 August 2018
17 August 2018
Please cite this article as: Pfannenstiel, B.T., Greco, C., Sukowaty, A.T., Keller, N.P., The epigenetic reader SntB
regulates secondary metabolism, development and global histone modifications in Aspergillus flavus, Fungal
Genetics and Biology (2018), doi:
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The epigenetic reader SntB regulates secondary metabolism, development and
global histone modifications in Aspergillus flavus
Brandon T. Pfannenstiel 1, Claudio Greco2, Andrew T. Sukowaty2, Nancy P. Keller2, 3#
Department of Genetics, University of Wisconsin-Madison, Madison, WI, USA
Department of Medical Microbiology and Immunology, University of Wisconsin-
Madison, Madison, WI, USA
Department of Bacteriology, University of Wisconsin-Madison, Madison, WI, USA
Corresponding author
Key words:
Hyphal fusion
Kojic Acid
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 sntB 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. sntB developmental
phenotypes were linked with SntB regulation of NosA, a transcription factor regulating
the A. flavus cell fusion cascade.
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
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,
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
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 -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 laeA TXZ15.1, and pyrG auxotroph sntB 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 10 6 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 5days 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 UHPLCMS 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 10 6 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 antihistone 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-acetylhistone 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 sntB 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).
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
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 sntB strain (Fig. 2A).
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 sntB strain was likely
due to inhibition of NosA function, the sntB 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, laeA
(previously shown to be required for heterokaryon formation (Zhao et al., 2017), and
sntB auxotrophs. The deletion of sntB showed the same phenotype as the laeA
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).
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 (UHPLCHRMS) 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.
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 sntB 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 sntB 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.
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 sntB suggests that SntB might interact with the HDAC RpdA and
the demethylase KdmB.
3.6. Aflatoxin promoters show wild-type histone acetylation:
The A. nidulans sntB 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 sntB 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 sntB. 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).
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 sntB 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 sntB 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 sntB 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 sntB 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 sntB 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 sntB 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 sntB 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.
The authors would like to thank Dr. John Doebley for corn used in the colonization
assay, Kunlong Yang for infection guidance, and Jacob Hagen for technical assistance.
Funding was provided in part by NIH R01GM112739-01 to NPK. BTP was supported by
the Predoctoral Training Program in Genetics, funded by the National Institutes of
Health (5 T32 GM007133-40).
Albright, J.C., Henke, M.T., Soukup, A.A., McClure, R.A., Thomson, R.J., Keller, N.P.,
Kelleher, N.L., 2015. Large-scale metabolomics reveals a complex response of
Aspergillus nidulans to epigenetic perturbation. ACS Chem. Biol. 10, 1534–1541.
Amare, M.G., Keller, N.P., 2014. Molecular mechanisms of Aspergillus flavus secondary
metabolism and development. Fungal Genet. Biol. 66, 11–18.
Baker, L.A., Ueberheide, B.M., Dewell, S., Chait, B.T., Zheng, D., Allis, C.D., 2013. The
yeast Snt2 protein coordinates the transcriptional response to hydrogen peroxidemediated oxidative stress. Mol. Cell. Biol. 33, 3735–3748.
Bannister, A.J., Kouzarides, T., 2011. Regulation of chromatin by histone modifications.
Cell Res. 21, 381–95.
Bernreiter, A., Ramon, A., Fernández-Martínez, J., Berger, H., Araújo-Bazan, L.,
Espeso, E.A., Pachlinger, R., Gallmetzer, A., Anderl, I., Scazzocchio, C., Strauss,
J., 2007. Nuclear export of the transcription factor NirA is a regulatory checkpoint
for nitrate induction in Aspergillus nidulans. Mol. Cell. Biol. 27, 791–802.
Bok, J.W., Chiang, Y.-M., Szewczyk, E., Reyes-Domingez, Y., Davidson, A.D.,
Sanchez, J.F., Lo, H.-C., Watanabe, K., Strauss, J., Oakley, B.R., Wang, C.C.C.,
Keller, N.P., 2009. Chromatin-level regulation of biosynthetic gene clusters. Nat.
Chem. Biol. 5, 462–464.
Cary, J.W., Bhatnagar, D., Linz, J.E., 2000. Aflatoxins: biological significance and
regulation of biosynthesis, in: Microbial Foodborne Diseases: Mechanisms of
Pathogenesis and Toxin Synthesis. Technomic Publishing Company, Inc., pp. 317–
Cary, J.W., Han, Z., Yin, Y., Lohmar, J.M., Shantappa, S., Harris-Coward, P.Y., Mack,
B., Ehrlich, K.C., Wei, Q., Arroyo-Manzanares, N., Uka, V., Vanhaecke, L.,
Bhatnagar, D., Yu, J., Nierman, W.C., Johns, M.A., Sorensen, D., Shen, H., De
Saeger, S., Diana Di Mavungu, J., Calvo, A.M., 2015a. Transcriptome Analysis of
Aspergillus flavus Reveals veA-Dependent Regulation of Secondary Metabolite
Gene Clusters, Including the Novel Aflavarin Cluster. Eukaryot. Cell 14, 983–97.
Cary, J.W., Harris-Coward, P.Y., Ehrlich, K.C., Di Mavungu, J.D., Malysheva, S. V., De
Saeger, S., Dowd, P.F., Shantappa, S., Martens, S.L., Calvo, A.M., 2014.
Functional characterization of a veA-dependent polyketide synthase gene in
Aspergillus flavus necessary for the synthesis of asparasone, a sclerotium-specific
pigment. Fungal Genet. Biol. 64, 25–35.
Cary, J.W., Harris-Coward, P.Y., Ehrlich, K.C., Mack, B.M., Kale, S.P., Larey, C., Calvo,
A.M., 2012. NsdC and NsdD affect Aspergillus flavus morphogenesis and aflatoxin
production. Eukaryot. Cell 11, 1104–11.
Cary, J.W., Uka, V., Han, Z., Buyst, D., Harris-Coward, P.Y., Ehrlich, K.C., Wei, Q.,
Bhatnagar, D., Dowd, P.F., Martens, S.L., Calvo, A.M., Martins, J.C., Vanhaecke,
L., Coenye, T., De Saeger, S., Di Mavungu, J.D., 2015b. An Aspergillus flavus
secondary metabolic gene cluster containing a hybrid PKS–NRPS is necessary for
synthesis of the 2-pyridones, leporins. Fungal Genet. Biol. 81, 88–97.
Chang, P.-K., Horn, B.W., Dorner, J.W., 2009. Clustered genes involved in
cyclopiazonic acid production are next to the aflatoxin biosynthesis gene cluster in
Aspergillus flavus. Fungal Genet. Biol. 46, 176–182.
Chang, P.-K., Scharfenstein, L.L., Li, R.W., Arroyo-Manzanares, N., De Saeger, S.,
Diana Di Mavungu, J., 2017. Aspergillus flavus aswA, a gene homolog of
Aspergillus nidulans oefC, regulates sclerotial development and biosynthesis of
sclerotium-associated secondary metabolites. Fungal Genet. Biol. 104, 29–37.
Christensen, S., Borrego, E., Shim, W.-B., Isakeit, T., Kolomiets, M., 2012.
Quantification of fungal colonization, sporogenesis, and production of mycotoxins
using kernel bioassays. J. Vis. Exp.
Clasquin, M.F., Melamud, E., Rabinowitz, J.D., 2012. LC-MS data processing with
MAVEN: a metabolomic analysis and visualization engine. Curr. Protoc.
Bioinforma. Chapter 14, Unit14.11.
Denisov, Y., Freeman, S., Yarden, O., 2011a. Inactivation of Snt2, a BAH/PHDcontaining transcription factor, impairs pathogenicity and increases autophagosome
abundance in Fusarium oxysporum. Mol. Plant Pathol. 12, 449–461.
Denisov, Y., Yarden, O., Freeman, S., 2011b. The transcription factor SNT2 is involved
in fungal respiration and reactive oxidative stress in Fusarium oxysporum and
Neurospora crassa. Physiol. Mol. Plant Pathol. 76, 137–143.
Drott, M.T., Lazzaro, B.P., Brown, D.L., Carbone, I., Milgroom, M.G., 2017. Balancing
selection for aflatoxin in Aspergillus flavus is maintained through interference
competition with, and fungivory by insects. Proc. R. Soc. B Biol. Sci. 284,
Eberharter, A., Becker, P.B., 2002. Histone acetylation: a switch between repressive
and permissive chromatin. EMBO Rep. 3, 224–9.
Forseth, R.R., Amaike, S., Schwenk, D., Affeldt, K.J., Hoffmeister, D., Schroeder, F.C.,
Keller, N.P., 2013. Homologous NRPS-like Gene Clusters Mediate Redundant
Small-Molecule Biosynthesis in Aspergillus flavus. Angew. Chemie Int. Ed. 52,
Gacek, A., Strauss, J., 2012. The chromatin code of fungal secondary metabolite gene
clusters. Appl. Microbiol. Biotechnol. 95, 1389–1404.
Georgianna, D.R., Payne, G.A., 2009. Genetic regulation of aflatoxin biosynthesis: from
gene to genome. Fungal Genet. Biol. 46, 113–125.
He, M., Xu, Y., Chen, J., Luo, Y., Lv, Y., Su, J., Kershaw, M.J., Li, W., Wang, J., Yin, J.,
Zhu, X., Liu, X., Chern, M., Ma, B., Wang, J., Qin, P., Chen, W., Wang, Y., Wang,
W., Ren, Z., Wu, X., Li, P., Li, S., Peng, Y., Lin, F., Talbot, N.J., Chen, X., 2018.
MoSnt2-dependent deacetylation of histone H3 mediates MoTor-dependent
autophagy and plant infection by the rice blast fungus Magnaporthe oryzae.
Autophagy 15548627.2018.1458171.
Horn, B.W., Moore, G.G., Carbone, I., 2009. Sexual reproduction in Aspergillus flavus.
Mycologia 101, 423–9.
Kaimori, J.-Y., Maehara, K., Hayashi-Takanaka, Y., Harada, A., Fukuda, M., Yamamoto,
S., Ichimaru, N., Umehara, T., Yokoyama, S., Matsuda, R., Ikura, T., Nagao, K.,
Obuse, C., Nozaki, N., Takahara, S., Takao, T., Ohkawa, Y., Kimura, H., Isaka, Y.,
2016. Histone H4 lysine 20 acetylation is associated with gene repression in human
cells. Sci. Rep. 6, 24318.
Keller, N.P., Adams, T.H., 1995. Analysis of a mycotoxin gene cluster in Aspergillus
nidulans. SAAS Bull. Biochem. Biotechnol. 8, 14–21.
Khalid, S., Baccile, J.A., Spraker, J.E., Tannous, J., Imran, M., Schroeder, F.C., Keller,
N.P., 2018. NRPS-Derived Isoquinolines and Lipopetides Mediate Antagonism
between Plant Pathogenic Fungi and Bacteria. ACS Chem. Biol. 13, 171–179.
Lan, H., Sun, R., Fan, K., Yang, K., Zhang, F., Nie, X.Y., Wang, X., Zhuang, Z., Wang,
S., 2016. The Aspergillus flavus histone acetyltransferase AflGcnE regulates
morphogenesis, aflatoxin biosynthesis, and pathogenicity. Front. Microbiol. 7, 1324.
Lebar, M.D., Cary, J.W., Majumdar, R., Carter-Wientjes, C.H., Mack, B.M., Wei, Q.,
Uka, V., De Saeger, S., Diana Di Mavungu, J., 2018. Identification and functional
analysis of the aspergillic acid gene cluster in Aspergillus flavus. Fungal Genet.
Biol. 116, 14–23.
Lim, F.Y., Keller, N.P., 2014. Spatial and temporal control of fungal natural product
synthesis. Nat. Prod. Rep. 31, 1277–86.
Melamud, E., Vastag, L., Rabinowitz, J.D., 2010. Metabolomic Analysis and
Visualization Engine for LC−MS Data. Anal. Chem. 82, 9818–9826.
Nesbitt, B.F., O’Kelly, J., Sargeant, K., Sheridan, A.N.N., 1962. Aspergillus flavus and
Turkey X Disease: Toxic metabolites of Aspergillus flavus. Nature 195, 1062–1063.
Nicholson, M.J., Koulman, A., Monahan, B.J., Pritchard, B.L., Payne, G.A., Scott, B.,
2009. Identification of two aflatrem biosynthesis gene loci in Aspergillus flavus and
metabolic engineering of Penicillium paxilli to elucidate their function. Appl. Environ.
Microbiol. 75, 7469–81.
Nützmann, H.-W., Reyes-Dominguez, Y., Scherlach, K., Schroeckh, V., Horn, F.,
Gacek, A., Schümann, J., Hertweck, C., Strauss, J., Brakhage, A.A., 2011.
Bacteria-induced natural product formation in the fungus Aspergillus nidulans
requires Saga/Ada-mediated histone acetylation. Proc. Natl. Acad. Sci. U. S. A.
108, 14282–7.
Palmer, J.M., Keller, N.P., 2010. Secondary metabolism in fungi: does chromosomal
location matter? Curr. Opin. Microbiol. 13, 431–436.
Palmer, J.M., Perrin, R.M., Dagenais, T.R.T., Keller, N.P., 2008. H3K9 methylation
regulates growth and development in Aspergillus fumigatus. Eukaryot. Cell 7,
Pfannenstiel, B.T., Zhao, X., Wortman, J., Wiemann, P., Throckmorton, K., Spraker,
J.E., Soukup, A.A., Luo, X., Lindner, D.L., Lim, F.Y., Knox, B.P., Haas, B., Fischer,
G.J., Choera, T., Butchko, R.A.E., Bok, J.-W., Affeldt, K.J., Keller, N.P., Palmer,
J.M., 2017. Revitalization of a forward genetic screen identifies three new
regulators of fungal secondary metabolism in the genus Aspergillus. MBio 8,
Reyes-Dominguez, Y., Bok, J.W., Berger, H., Shwab, E.K., Basheer, A., Gallmetzer, A.,
Scazzocchio, C., Keller, N., Strauss, J., 2010. Heterochromatic marks are
associated with the repression of secondary metabolism clusters in Aspergillus
nidulans. Mol. Microbiol. 76, 1376–1386.
Roguev, A., Shevchenko, A., Schaft, D., Thomas, H., Stewart, A.F., Shevchenko, A.,
2004. A comparative analysis of an orthologous proteomic environment in the
yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe. Mol. Cell.
Proteomics 3, 125–132.
Rohlfs, M., 2015. Fungal secondary metabolite dynamics in fungus-grazer interactions:
novel insights and unanswered questions. Front. Microbiol. 5, 788.
Roze, L. V., Arthur, A.E., Hong, S.-Y., Chanda, A., Linz, J.E., 2007. The initiation and
pattern of spread of histone H4 acetylation parallel the order of transcriptional
activation of genes in the aflatoxin cluster. Mol. Microbiol. 66, 713–726.
Saruwatari, T., Yagishita, F., Mino, T., Noguchi, H., Hotta, K., Watanabe, K., 2014.
Cytochrome P450 as dimerization catalyst in diketopiperazine alkaloid
biosynthesis. ChemBioChem 15, 656–659.
Scharf, D.H., Heinekamp, T., Brakhage, A.A., 2014. Human and plant fungal pathogens:
the role of secondary metabolites. PLoS Pathog. 10, e1003859.
Shwab, E.K., Bok, J.W., Tribus, M., Galehr, J., Graessle, S., Keller, N.P., 2007. Histone
deacetylase activity regulates chemical diversity in Aspergillus. Eukaryot. Cell 6,
Smith, C.A., Want, E.J., O’Maille, G., Abagyan, R., Siuzdak*, G., 2006. XCMS:
processing mass spectrometry data for metabolite profiling using nonlinear peak
alignment, matching, and identification. Anal. Chem. 78, 779–787.
Soukup, A.A., Chiang, Y.-M., Bok, J.W., Reyes-Dominguez, Y., Oakley, B.R., Wang,
C.C.C., Strauss, J., Keller, N.P., 2012. Overexpression of the Aspergillus nidulans
histone 4 acetyltransferase EsaA increases activation of secondary metabolite
production. Mol. Microbiol. 86, 314–330.
Strauss, J., Reyes-Dominguez, Y., 2011. Regulation of secondary metabolism by
chromatin structure and epigenetic codes. Fungal Genet. Biol. 48, 62–69.
Terabayashi, Y., Sano, M., Yamane, N., Marui, J., Tamano, K., Sagara, J., Dohmoto,
M., Oda, K., Ohshima, E., Tachibana, K., Higa, Y., Ohashi, S., Koike, H., Machida,
M., 2010. Identification and characterization of genes responsible for biosynthesis
of kojic acid, an industrially important compound from Aspergillus oryzae. Fungal
Genet. Biol. 47, 953–961.
Umemura, M., Nagano, N., Koike, H., Kawano, J., Ishii, T., Miyamura, Y., Kikuchi, M.,
Tamano, K., Yu, J., Shin-ya, K., Machida, M., 2014. Characterization of the
biosynthetic gene cluster for the ribosomally synthesized cyclic peptide ustiloxin B
in Aspergillus flavus. Fungal Genet. Biol. 68, 23–30.
van Leeuwen, J., Andrews, B., Boone, C., Tan, G., 2015. Rapid and efficient plasmid
construction by homologous recombination in yeast. Cold Spring Harb. Protoc.
2015, pdb.prot085100.
Williams, J.H., Phillips, T.D., Jolly, P.E., Stiles, J.K., Jolly, C.M., Aggarwal, D., 2004.
Human aflatoxicosis in developing countries: a review of toxicology, exposure,
potential health consequences, and interventions. Am. J. Clin. Nutr. 80, 1106–
Wogan, G.N., 1992. Aflatoxins as risk factors for hepatocellular carcinoma in humans.
Cancer Res. 52, 2114s–2118s.
Xhemalce, B., Kouzarides, T., 2010. A chromodomain switch mediated by histone H3
Lys 4 acetylation regulates heterochromatin assembly. Genes Dev. 24, 647–52.
Yu, J.-H., Hamari, Z., Han, K.-H., Seo, J.-A., Reyes-Domínguez, Y., Scazzocchio, C.,
2004. Double-joint PCR: a PCR-based molecular tool for gene manipulations in
filamentous fungi. Fungal Genet. Biol. 41, 973–981.
Yu, J., 2012. Current understanding on aflatoxin biosynthesis and future perspective in
reducing aflatoxin contamination. Toxins. 4, 1024–1057.
Zhao, X., Spraker, J.E., Bok, J.W., Velk, T., He, Z.-M., Keller, N.P., 2017. A Cellular
Fusion Cascade Regulated by LaeA Is Required for Sclerotial Development in
Aspergillus flavus. Front. Microbiol. 8, 1925.
Figure Legends
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 <
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 laeA crossed with TJES20.1, and sntB strain
crossed with TJES20.1. P-value **p < 0.01.
Figure 3: SntB is required for pathogenicity of corn. A) corn kernels were inoculated
with 2x105 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.
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.
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.
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 Photoshop TM, 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.
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
Fig. S1: Southern blot analysis of sntB overexpression strain. The argB gene and A.
nidulans gpdA(p) was placed in front of the sntB open reading frame. WT,
parental wild-type control. Schematic of Southern design is below the blot. The 5’
flank of the overexpression construct was used as a probe. * indicates correct
transformants (1 and 5).
Fig. S2: SntB transcriptionally regulates the aflatoxin BGC. Semi-qPCR of wild type,
deletion, and overexpression sntB strains grown in YES media for 40 hours.
Primers used are in Table S2. ubiD is used as a loading control.
Fig. S3: MS spectra of the identified known secondary metabolites from A. flavus under
ESI positive mode.
Table 1: Protein Interactors of SntB Homologs
A. flavus Homolog
AFLA_092360 (RpdA)
AFLA_006240 (KdmB)
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
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