Skip to main content
NCBI home page
Molecular Plant Pathology logoLink to Molecular Plant Pathology
. 2019 Jul 30;20(11):1491–1505. doi: 10.1111/mpp.12856

A MYST family histone acetyltransferase, MoSAS3, is required for development and pathogenicity in the rice blast fungus

Akanksha Dubey 1,†, Jongjune Lee 1,†, Seomun Kwon 2,5,†, Yong‐Hwan Lee 2,3,4,✉, Junhyun Jeon 1,4,✉
PMCID: PMC6804344  PMID: 31364260

Summary

Histone acetylation has been established as a principal epigenetic regulatory mechanism in eukaryotes. Sas3, a histone acetyltransferase belonging to the largest family of acetyltransferase, MYST, is the catalytic subunit of a conserved histone acetyltransferase complex. To date, the functions of Sas3 and its orthologues have been extensively studied in yeast, humans and flies in relation to global acetylation and transcriptional regulation. However, its precise impact on development and pathogenicity in fungal plant pathogens has yet to be elucidated. Considering the importance of Sas3 in H3K14 acetylation, here we investigate the roles of its orthologue in the rice blast fungus, Magnaporthe oryzae (Pyricularia oryzae). Unlike a previously reported Sas3 deletion in yeast, which led to no remarkable phenotypic changes, we found that MoSAS3 deletion alone had a profound effect on fungal growth and development, including asexual reproduction, germination and appressorium formation in M. oryzae. Such defects in pre‐penetration development resulted in complete loss of pathogenicity in the deletion mutant. Furthermore, genetic analysis of MoSAS3 and MoGCN5 encoding a Gcn5‐related N‐acetyltransferase family histone acetyltransferase suggested that two conserved components of histone acetylation are integrated differently into epigenetic regulatory mechanisms in the yeast and a filamentous fungus. RNA‐seq analysis of ΔMosas3 showed two general trends: many DNA repair and DNA damage response genes are up‐regulated, while carbon and nitrogen metabolism genes are down‐regulated in ΔMosas3. Our work demonstrates the importance of MYST family histone acetyltransferase as a developmental regulator and illuminates a degree of functional variation in conserved catalytic subunits among different fungal species.

Keywords: development, histone acetyltransferase, MoSAS3, pathogenicity, rice blast

Introduction

Histone modification is a conserved post‐translational modification (PTM) that plays a pivotal role in the epigenetic regulation of gene expression and chromatin in eukaryotes (Kouzarides, 2007). Modification of histones is catalysed by specialized histone‐modifying enzymes (HMEs) such as histone acetyltransferases/deacetylase (HATs/HDACs) and histone methyltransferases/demethylases (HMTs/HDMs) that function via addition or removal of acetyl and methyl residues from the histone proteins, respectively. Since the discovery of histone acetylation in 1964 (Allfrey and Mirsky, 1964; Allfrey et al., 1964), the antagonistic interplay of HATs and HDACs has been established as one of the principal epigenetic regulatory mechanisms.

HATs are one of the best‐characterized groups of HMEs, which use acetyl‐CoA as a substrate for the acetylation of lysine (K) residues within both the tail and globular domains of histones to poise the gene for active transcription (Dubey and Jeon 2017; Jeon et al., 2014; Sterner and Berger, 2000). However, as an increasing number of non‐histone targets of previously known HATs and HDACs are being discovered, a new classification placed these enzymes under the umbrella of lysine acetyltransferase (KATs) and deacetylases (KDACs) (Allis et al., 2007). Ranging from yeast to humans, several HATs have been identified that can be classified, based on the presence of conserved structural motifs, into five families including Gcn5‐related N‐acetyltransferase (GNAT) and MYST (members MOZ, MOF, MORF, Ybf2/Sas3, Sas2, Tip60 and HBO1) families (Jeon et al., 2014; Sterner and Berger, 2000).

In fungi, histone acetylation/deacetylation was shown to be important for regulation of growth, asexual development, secondary metabolite production and pathogenicity (Ding et al., 2010; Jeon et al., 2014). Among the five families of HATs, the GNAT superfamily has been extensively studied. In particular, Gcn5 orthologues have been demonstrated to be required for asexual development in Aspergillus nidulans, dimorphism and virulence in Ustilago maydis, and regulation of autophagy in Magnaporthe oryzae (anamorph Pyricularia oryzae), respectively (Canovas et al., 2014; Gonzalez‐Prieto et al., 2014; Li et al., 2017). In addition, several studies investigated roles of Gcn5 and Sas2 orthologues in regulation of fungal secondary metabolism (Gomez‐Rodriguez et al., 2018; Zhang et al., 2018). Comparatively, however, the MYST family of HATs has not gained much attention. Lysine acetyltransferase 6 (KAT6) is an important member of the MYST family of HATs that catalyses acetylation of histone substrates H3K9, K14 and K23 (Huang et al., 2016). Several KAT6 homologues have been identified, including MOZ (monocytic leukemia zinc finger)/MORF (MOZ‐related factor) in humans (Huang et al., 2016), Sas3 (something about silencing) in yeast (Reifsnyder et al., 1996) and ENOK (enoki mushroom) in Drosophila (Scott et al., 2001). Phylogenetic analysis showed that the homologues Sas2 and Sas3 from yeast share homology with the human MOZ oncogene (Lafon et al., 2007) and other MYST members owing to the presence of a conserved domain containing an atypical zinc finger and a putative acetyl‐CoA binding motif (Takechi and Nakayama, 1999). These domains are important for the functioning of the MYST proteins. Mutant analysis of Sas3 through amino acid substitution in Saccharomyces cerevisiae revealed that both the acetyl‐CoA binding motif and the zinc finger motif are essential for HAT activity (Takechi and Nakayama, 1999). To date, functionally, MOZ/MORF have been implicated in leukaemogenesis (Rokudai et al., 2009, 2013), Sas3 in controlling global acetylation level in yeast in association with Gcn5 (Rosaleny et al., 2007), and ENOK in total global acetylation level in Drosophila (Feller et al., 2015; Huang et al., 2014). However, there is an evident lack of reports on the role of Sas3 in filamentous fungi and its impact on fungal development and pathogenicity.

In S. cerevisiae, Sas3, a member of the MYST family of HATs and the catalytic subunit of the conserved 400‐kDa NuA3 (nucleosomal acetyltransferase of histone H3) HAT complex, was originally discovered in a screen for mutations enhancing loss of silencing in Sir1 (HDAC) deletion mutant background (Reifsnyder et al., 1996). Sas3 of S. cerevisiae was shown to have in vitro HAT activity (Takechi and Nakayama, 1999) and specifically acetylate H3K9 and H3K14 in vivo (Durant and Pugh, 2006; Vicente‐Munoz et al., 2014). Being the catalytic subunit of the NuA3 complex, Sas3 is necessary for HAT activity as well as the integrity of the NuA3 complex (John et al., 2000). In S. cerevisiae, while the presence of a single H4‐specific HAT [Esa1 (essential SAS2‐related acetyltransferase)] is essential for cell viability (Clarke et al., 1999; Smith et al., 1998), catalytic subunits of HATs related to H3 acetylation appear to be dispensable (Georgakopoulos and Thireos, 1992) unless deleted in double mutant combinations (Howe et al., 2001). With an overlapping pattern of acetylation, NuA3 has been shown to function in close relation to Gcn5 in S. cerevisiae (Rosaleny et al., 2007). As a result, although the deletion of Sas3 alone does not elicit any phenotype, its combined deletion with Gcn5 causes complete loss of acetyltransferase activity, resulting in reduced global H3 acetylation and cell cycle arrest that is synthetically lethal (Howe et al., 2001).

In this study, we investigated the role of the Sas3 orthologue and its histone acetylation activity in the regulation of the growth, development and pathogenicity of M. oryzae (Couch and Kohn, 2002; Couch et al., 2005; Gomez‐Rodriguez et al., 2018; Grimaldi et al., 2006; Zhang et al., 2016), combining genetic and transcriptomic approaches. Magnaporthe oryzae, an ascomycete fungus, is a devastating plant pathogen known to cause rice blast disease (Dean et al., 2012; Talbot, 2003). It is a notorious fungal plant pathogen that infects a wide range of hosts and poses a threat to food security and rice production worldwide (Dean et al., 2012; Maciel et al., 2014; Martin‐Urdiroz et al., 2016; Wilson and Talbot, 2009). The fungus produces aerial conidiophores bearing conidia, which attach and germinate on the host surface to initiate fungal infection. Upon germination, the conidium produces a specialized single‐celled infection structure called an appressorium that breaches the plant surface usually protected by a cuticle layer and the cell wall (Dean, 1997; Talbot, 2003; Wilson and Talbot, 2009). Enormous turgor pressure generated inside the melanized appressorium drives the penetration hyphae into host cells (Wilson and Talbot, 2009). Such penetration is followed by transition into invasive hyphae, which requires suppression of plant immunity primarily through secretion of effector proteins (Molloy, 2010; Yan and Talbot, 2016).

Here we examined the functions of MoSAS3 as histone acetyltransferase and its impact on fungal development and pathogenesis in the rice blast fungus, compared to the yeast counterpart. Furthermore, we analysed the transcriptome of the wild‐type and the deletion mutant in order to gain insight into the role of MoSAS3.

Results

Identification and domain analysis of MoSAS3

To identify orthologues of S. cerevisiae Sas3, we ran a BLAST search using Sas3 sequences as a query against the NCBI nr database (Altschul et al., 1990). Through the database search, we retrieved amino acid sequences of its orthologues from diverse organisms, including M. oryzae. A gene encoding the Sas3 orthologue in M. oryzae was designated as MoSAS3 (MGG_04615). Comparison of the domain architecture using the InterPro database (Finn et al., 2017) revealed that the Sas3 orthologues possess the signature MYST‐type HAT domain in common (Fig. 1A). Close examination of MYST domains showed that the central core‐binding region to acetyl‐CoA with flanking N‐ and C‐terminal regions harbouring a C2H2 zinc‐binding domain and helix‐turn‐helix DNA‐binding motif are well conserved in all the Sas3 orthologues we examined (Lafon et al., 2007; McCullough and Marmorstein, 2016; Yan et al., 2000). Compared to the Sas3 of S. cerevisiae and Candida albicans, the Sas3 orthologues of filamentous ascomycetes, Aspergillus nidulans, Neurospora crassa, M. oryzae, Gibberella zeae and Fusarium oxysporium contain an additional PHD finger domain, which is known to be involved in recognition of H3K4me3 (Musselman and Kutateladze, 2011). Interestingly, linker histone (H1/H5) domain, which is essential for the binding of linker histone H1 or H5 to the nucleosome, was found only in human and fly.

Figure 1.

Figure 1

Open in a new tab

MoSAS3 as a histone acetyltransferase. (A) Domain architecture of MoSAS3 and its orthologues. (B) Localization of MoSAS3 in Magnaporthe oryzae spores. Green fluorescence in conidia expressing MoSAS3‐GFP (middle panel) and DAPI staining pattern (bottom panel). (C) Quantification of western blot signal for H3K14 acetylation, relative to H3. The western blot image was quantified using ImageJ software (https://imagej.nih.gov/ij/index.html).

Since Sas3 of S. cerevisiae is a component of NuA3 HAT complex, it was expected that MoSAS3 should be localized to the nucleus of M. oryzae. PSORT analysis predicted nucleus localization of MoSAS3 (95.7%) (Horton et al., 2007). To test this, we made a translational fusion construct between MoSAS3 and green fluorescence protein (eGFP) under the control of the native MoSAS3 promoter, and introduced the construct into the wild‐type strain KJ201. Indeed, we were able to detect green fluorescence signal from the fungal nucleus under the epifluorescence microscope, although a faint/weak signal appears to be present also in the cytoplasm (Fig. 1B). Conservation of protein domains and localization strongly suggests that MoSAS3 is an evolutionarily conserved histone acetyltransferase.

MoSAS3 deletion impairs vegetative growth and asexual spore production

To investigate the functions of MoSAS3 in the development and pathogenicity of M. oryzae, we generated a gene deletion mutant using double‐joint PCR and protoplast transformation (Materials and Methods, and Fig. S1, see Supporting Information) and compared it to the wild‐type strain KJ201 for changes in histone acetyltransferase activity and phenotypes. Our western blot analysis using antibody raised against acetylated H3K14 showed that deletion of MoSAS3 gene results in about 48% reduction in H3K14ac level (Figs 1C and S2A, see Supporting Information). Quantification of H3K14ac through an independent method consistently showed over 40% decreases upon deletion of MoSAS3 (Fig. S2B, see Supporting Information), lending support to the role of MoSAS3 as a histone acetyltransferase. When the mutant was grown on oatmeal agar plates, it exhibited highly diminished growth (c.60% of the wild‐type) (Fig. 2A,B). The mutant colonies showed not only reduced radial growth but also a decrease in aerial hyphae masses compared to the wild‐type strain. Additionally, the mutant was more pigmented than the wild‐type (Fig. 2A).

Figure 2.

Figure 2

Open in a new tab

Growth and asexual reproduction of wild‐type Magnaporthe oryzae (KJ201, WT), ΔMosas3 (KO) and complementation strain (Comp, MoSAS3c). (A) Colony morphology and pigmentation on oatmeal agar plates. (B) Growth rate measured in colony diameter at 9 days post‐inoculation. (C) Asexual sporulation measured in the number of spores produced from individual strains. (D) Conidiophore development. Mycelial mat was scraped off and then a small agar block (1 × 1 cm) was excised and incubated for 8 h under constant fluorescence light to induce sporulation before being observed under a microscope. Asterisks indicate statistically significant differences (P < 0.001, Tukey HSD test).

Counting the number of asexual spores from agar plates showed that the mutant is not capable of producing as many spores as the wild‐type (Fig. 2C). Examination of conidiophore development in agar plugs revealed that such dramatic reduction in asexual spore production is attributed largely to a decrease in the number of conidiophores in ΔMosas3, which is consistent with a decrease in aerial hyphae masses (Fig. 2D). Introduction of the MoSAS3 gene back into the deletion mutant (MoSAS3c) was able to restore the mutant phenotypes to the wild‐type level (Fig. 2), indicating that targeted deletion of MoSAS3 is responsible for the defects in growth and asexual reproduction.

Germination and appressorium formation of ΔMosas3

Next, we monitored pre‐penetration development involving germination and appressorium formation of the mutant strain compared to wild‐type. While in the wild‐type and the complementation strain, more than 95% of conidia germinated and c.96% of germinated conidia were able to form an appressorium, only 34% of the total conidia germinated, of which only 20% formed an appressorium in ΔMosas3 (Fig. 3A,B). Furthermore, deformed conidia were found more frequently in the deletion mutant and such deformed ones, if they germinated, tended to form a longer germ tube and an irregular appressorium (Fig. 3B).

Figure 3.

Figure 3

Open in a new tab

Defect in conidial germination and appressorium formation of Magnaporthe oryzae ΔMosas3. (A) Quantitative analysis of spore germination and appressorium formation in wild‐type (KJ201), ΔMosas3 and complementation strain (MoSAS3c) after 8 h of incubation on hydrophobic surface (glass coverslip) at 25 °C. Percentages of ungerminated, germinated and appressorium‐forming conidia were counted under a light microscope. (B) Representative image for germination and appressorium formation of different strains on hydrophobic surface (plastic coverslip). (C) Spore viability test using FUN‐1 staining. Dead spores were prepared by subjecting them to microwave treatment for 1 min. Live cells localize dye to vacuoles, while dead cells display diffuse signals throughout the cytoplasm.

To understand why the mutant conidia are defective in germination, we checked the viability of the mutant conidia using the FUN‐1 staining method (Essary and Marshall, 2009; Millard et al., 1997). In metabolically active cells, FUN‐1 dye, which is visible as red fluorescence, concentrates at intravacuolar structures. In contrast, red fluorescence signals are diffuse in the whole cell in metabolically inactive cells. Our preliminary experiments showed that treatment of wild‐type conidia with non‐permissible temperature (65 °C) for 15 min could render the conidia inviable and completely block germination (Fig. 3C, KJ201 panels). Unlike conidia from wild‐type and complementation strains displaying punctuated fluorescence signals, the FUN‐1 staining pattern of the mutant conidia resembled that of the negative control (heat‐killed spores), suggesting that MoSAS3 is essential for production of viable conidia.

MoSAS3 is required for fungal pathogenicity

In order to assess the degree to which MoSAS3 is involved in pathogenicity, conidial suspensions prepared from plate cultures of wild‐type, ΔMosas3 and the complemented strain MoSAS3c were sprayed onto a susceptible rice cultivar, Nakdongbyeo. Disease symptom development was evaluated for inoculated plants 7 days post‐inoculation (dpi). While the wild‐type strain KJ201 and MoSAS3c produced typical lesions, including dark brown spots and yellowing of leaves, almost no lesions were observed on the leaves spray‐inoculated with a spore suspension of ΔMosas3 (Fig. 4A). It stands to reason that such loss of pathogenicity in the mutant is accountable for the observed defects in pre‐penetration development.

Figure 4.

Figure 4

Open in a new tab

Dissection of pathogenicity defect in MoSAS3 deletion mutant. (A) Lesion development in rice leaves 7 days post‐spray inoculation. (B) Lesion development in wound‐inoculated rice leaves 5 days post‐inoculation. Distilled water was used as a mock experiment. (C) Observation of invasive growth using rice sheath assay. Invasive growth in rice sheath cells was monitored under microscope at 48 h post‐inoculation.

To determine if the mutant retains the ability to grow inside host plants, we performed wound‐inoculation by placing spore suspensions on wound sites made by pressing pipette tips against host leaves (Fig. 4B). This allows the fungus to gain direct access to the host cells without appressorium‐mediated penetration. Again, even in wound‐inoculation, the mutant was not able to cause disease symptoms, suggesting that MoSAS3 is required for invasive growth as well. In line with these results, rice sheath assay of the mutant showed that appressoria of ΔMosas3 were rarely able to penetrate plant cells with poorly growing invasive hyphae. Furthermore, in many cases there was either an abundant accumulation of dark granules or discolouration of plant cells, possibly suggesting inability of the mutant to suppress the initial defence responses of host plants.

MoSAS3 does not genetically interact with MoGCN5

In S. cerevisiae it was shown that acetylation of histone H3 is accomplished through the combined activity of two enzymes, Sas3 and Gcn5, the latter of which is a GNAT family histone acetyltransferase. Due to the overlapping pattern of acetylation via two enzymes, deletion of individual genes has a negligible effect, instead lack of both genes is known to cause synthetic lethality in yeast. Our data show that the single deletion of MoSAS3 alone resulted in significant growth and developmental defects. This observation prompted us to investigate if such interaction is also conserved in M. oryzae.

Using dbHiMo, a web‐based database for histone‐modifying enzymes (http://hme.riceblast.snu.ac.kr/), we identified two genes (MGG_03677 and MGG_11716) encoding the putative Gcn5 orthologue in the M. oryzae genome and showed that two genes are probably the result of a relatively recent duplication event (Choi et al., 2015). A previous work investigating the role of GCN5 in light‐induced autophagy designated MGG_03677 and MGG_11716 as MoGCN5 and MoGCN5b, respectively (Zhang et al., 2017). Pairwise sequence alignment of MoGCN5 or MoGCN5b with S. cerevisiae Gcn5 using Needle (https://www.ebi.ac.uk/Tools/psa/emboss_needle/) indicated that MoGCN5 is more similar (56.6% identity and 71.3% similarity) to yeast Gcn5 than MoGCN5b (50.5% identity and 66.2% similarity). Moreover, deletion of MoGCN5b had no detectable effect, in contrast to deletion of MoGCN5 rendering the fungus defective in growth, sporulation and pathogenicity (Figs 5 and S3, see Supporting Information). Sequence similarity, evolutionary history and phenotypes of deletion mutants suggested that MoGCN5 is a functional, bona fide histone acetyltransferase. Based on this, we tested if there is genetic interaction between MoSAS3 and MoGCN5 by generating a strain where both genes are deleted. Comparative analysis of different mutant strains showed that vegetative growth and asexual sporulation of the double deletion mutant (ΔMosas3/ΔMogcn5) are comparable to those of ΔMosas3, indicating no genetic interaction of the kind that is found between the two genes in yeast (Fig. 5).

Figure 5.

Figure 5

Open in a new tab

Test for genetic interaction between MoSAS3 and MoGCN5. (A) Comparison of growth on oatmeal agar plates for ΔMosas3, ΔMogcn5 and the double deletion mutant (ΔMosas3/ΔMogcn5). (B) Growth of individual strains measured as colony diameter. (C) Asexual sporulation measured as the number of spores produced from individual strains. Different letters in bar plots indicate significantly different mean values (Tukey HSD test, P < 0.001).

Identifying MoSAS3‐dependent changes in gene expression of M. oryzae

To delve into the roles of MoSAS3 in M. oryzae, we compared the transcriptome of ΔMosas3 with that of wild‐type strain by performing RNA‐seq on three biological replicates each for wild‐type and the mutant grown in complete media. Our RNA‐seq analysis showed that about 14.1% of genes (n = 1810) are differentially expressed in the mutant compared to the wild‐type, with statistically significant differences and a two‐fold expression change cut‐off. Among those differentially expressed genes (DEGs), about 44.9% (n = 813) and 54.1% (n = 997) were up‐regulated and down‐regulated in the mutant strain, respectively (Tables S1 and S2, see Supporting Information). Several genes that displayed transcriptional differences by RNA‐seq were validated through qRT‐PCR experiments (Fig. S4, see Supporting Information). Similar patterns were detected by RNA‐seq and qRT‐PCR (Spearman's rank‐order correlation = 0.87), indicating that we have accurately detected gene expression changes in our experiments.

Altered expression profile of ΔMosas3 resembles that of ΔMortt109

To gain insight into transcriptomic changes associated with absence of MoSAS3, we performed gene ontology (GO) term analysis to quantify the enrichment of genes associated with biological processes. Our GO term analysis showed two general trends among DEGs: (i) up‐regulated genes were enrichment with functions including cellular response to stress, DNA metabolic process and DNA repair (Figs 6A and S5, see Supporting Information), and (ii) unlike the up‐regulated genes, the down‐regulated genes displayed enrichment with genes implicated in nutrient transport and diverse metabolic processes including nitrogen utilization, organic acid metabolism and vitamin metabolism (Figs 6B and S6, see Supporting Information). Recently, we demonstrated that a fungal‐specific histone acetyltransferase, MoRtt109, is required for proper regulation of the cell cycle as well as DNA damage repair and responses (Kwon et al., 2018). We noticed that transcriptional output of the mutant bears a close resemblance to that of ΔMortt109. Comparison of DEGs showed that there is highly significant overlap (n = 1006) between DEGs of ΔMosas3 (n = 1810) and ΔMortt109 (n = 1641), confirming our observation (hypergeometric test, P < 0.001) (Fig. S7, see Supporting Information). However, expression of MoRtt109 itself was marginally up‐regulated (2.04‐fold change, P < 0.01) in ΔMosas3, suggesting that similarity in expression profiles is created by different mechanisms in the two mutants.

Figure 6.

Figure 6

Open in a new tab

Summary of GO enrichment analysis (biological process) for selected terms. The whole GO enrichment analysis is available in the Supporting Information. GO terms that are enriched with up‐regulated and down‐regulated genes are shown in (A) and (B), respectively (Fisher's exact test, P < 0.01). The area of the circle represents the number of genes assigned to the particular GO term. The colour of the circle indicates the proportion of genes assigned to the GO term in our dataset among the total number of genes having that GO term in the genome.

Deletion of MoSAS3 alters expression of genes involved in cellular metabolism

In addition to DNA repair and damage response genes, some of the metabolic genes were up‐regulated in the mutant as well. One such example is trihydroxynaphthalene reductase (3HNR, MGG_07216), which is one of the catalytic enzymes for melanin biosynthesis. In the mutant, expression of 3HNR increased by nearly 45‐fold, and this might underlie the highly melanized mutant colonies in plate cultures, although expression of other melanin biosynthesis genes was not differentially expressed in submerged cultures (Fig. 2A). However, metabolic gene expression was globally down‐regulated in the mutant relative to the wild‐type. Detailed analysis of those metabolic genes revealed that many of the carbon metabolism genes involved in glycolysis and the citrate cycle were down‐regulated in the mutant (Tables S3 and S4, see Supporting Information). Furthermore, nitrogen utilization genes such as NADP‐specific glutamate dehydrogenase (MGG_08074), nitrate reductase (MGG_06062), nitrite reductase (MGG_00634) and nitrilase (MGG_03280) were down‐regulated (Table S5, see Supporting Information). Together with down‐regulation of genes involved in nutrient transport, our RNA‐seq data indicate that global down‐regulation of primary metabolism accounts for most, if not all, of the growth defects observed in ΔMosas3.

Since transcription factors (TFs) are generally responsible for large‐scale changes in transcriptional output of the genome, we next investigated changes in the expression of genes encoding these regulatory proteins using the Fungal Transcription Factor Database (http://ftfd.snu.ac.kr/index.php?a=view) (Park et al., 2008). Our analysis showed that deletion of MoSAS3 does not equally influence expression of different families of TFs. Bromodomain and homeodomain TFs were up‐regulated in the mutant, while the Myb, Zn2Cys6, bHLH and bZIP families of TFs tend to be down‐regulated (Fig. S8, see Supporting Information). Interestingly, many of the knockout mutants for Zn2Cys6 and bZIP TFs were reported to have defects in nutrient utilization and/or hyphal growth (Lu et al., 2014; Tang et al., 2015). This suggests the possibility that at least some genes encoding these TFs are direct targets of MoSAS3‐mediated histone acetylation and that widespread changes in gene expression of the mutant may be a consequence of flow‐on effects resulting from a change in expression of the key genes that control transcription.

Discussion

Sas3 and its homologues (MOZ/MORF, ENOK) are widely conserved across different kingdoms. The roles of MOZ/MORF, ENOK and yeast Sas3 have been well established in human diseases such as leukaemia (Rokudai et al., 2009, 2013; Voss et al., 2009), regulation of global acetylation levels in Drosophila (Scott et al., 2001) and regulation of histone acetylation in yeast (Durant and Pugh, 2006; Reifsnyder et al., 1996; Sterner and Berger, 2000; Takechi and Nakayama, 1999), respectively. However, compared to the GNAT family of HATs, study of the MYST family of HATs is considerably lacking. In this study, we investigated the roles of MoSAS3 as a MYST family HAT in fungal development and pathogenesis in order to bridge this knowledge gap in fungal pathogens.

We first examined cellular localization of MoSAS3 by expressing MoSAS3 protein fused to GFP at its C‐terminal. This showed that MoSAS3 is localized to fungal nuclei as expected for a HAT. However, a faint and diffuse fluorescence signal was also detected in the cytoplasm (Fig. 1B). It should be noted that despite heavy research focus on histones, post‐translation modifications (PTMs) are actually more widespread than previously imagined. Acetylation in particular was found to be a ubiquitous regulatory PTM, showing parallels to phosphorylation (Kouzarides, 2000). Considering that an increasing number of non‐histone targets of previously known HATs are being discovered, our data suggest that MoSAS3 is likely to have cytoplasmic targets as well. In an effort to understand the contribution of MoSAS3 in fungal growth and development, we generated mutant strains where MoSAS3 is deleted in the genome. Absence of MoSAS3 in the fungal genome decreased the global acetylation level of H3K14 by approximately 40–50% (Figs 1C and S2, see Supporting Information). Together with conserved domain architecture and nuclear localization, this result strongly suggests that MoSAS3 is a bona fide histone acetyltransferase in M. oryzae.

Deletion of the MoSAS3 gene significantly impaired vegetative mycelial growth compared to the wild‐type strain. The deletion mutant not only exhibited diminished radial growth, but also exhibited reduced thickness in the mycelial mat compared to the wild‐type strain, pointing towards the direct role of MoSAS3 in regulating fungal vegetative growth. Our RNA‐seq analysis suggests that global down‐regulation of genes involved in primary metabolism is likely to underlie this growth defect. Under conditions where primary metabolism is repressed, we predicted two changes in the mutant transcriptome: an increase in recycle/salvage processes and a decrease in de novo protein production. Contrary to our expectation, however, genes involved in recycling of nitrogen resources such as exo‐peptidases and metallo‐peptidases were not up‐regulated. Autophagy is also an important mechanism for cellular recycling, but none of the autophagy genes were differentially regulated in the mutant. Furthermore, de novo protein synthesis involving tRNA synthetases and ribosome complexes, which is a highly energy demanding process, was not repressed in the mutant. Considering that RNA was extracted from mutant mycelia grown in complete medium, these results suggest that the mutant is probably able to sense the presence of nutrients in the extracellular milieu but it cannot properly express genes implicated in the efficient transport and use of those nutrients due to absence of MoSAS3‐mediated histone acetylation. It is noteworthy that among down‐regulated genes in the mutant are Zn2Cys6 and bZIP TFs, many of which are required for proper growth and asexual sporulation (Lu et al., 2014; Tang et al., 2015). In particular, systematic analysis of Zn2Cys6 TFs by Lu et al. showed that deletion of some Zn2Cys6 TFs leads to defects in growth, conidiation, germination and appressorium formation. Among 33 TFs that are down‐regulated in the MoSAS3 deletion mutant, Zn2Cys6 TFs accounted for 42% (n = 13) in our RNA‐seq data. Such high enrichment of one type of TF suggests the possibility that MoSAS3‐mediated transcriptional regulation preferentially targets Zn2Cys6 TFs, which in turn control the transcription of a multitude of metabolic enzyme genes.

One unexpected observation in our RNA‐seq data was the close resemblance of gene expression profiles between ΔMosas3 and ΔMortt109. In our previous study, we reported that MoRtt109 acetylates H3K56 and is involved in DNA repair and damage responses. In both mutants, DNA repair and damage response genes tended to be up‐regulated, while a diverse array of metabolic genes is down‐regulated. One plausible explanation for this is down‐regulation of MoRtt109 in the ΔMosas3. However, we can rule out this possibility since RNA‐seq data showed marginal up‐regulation of MoRtt109 in the mutant (2.04‐fold, P  < 0.001), indicating that similarity in transcriptomic changes between two mutants is probably based on different mechanisms. Nevertheless, it is tempting to speculate that DNA repair/damage responses and cellular metabolism are linked at the epigenetic level.

Other than vegetative growth, we observed a sharp decline in the ability of the mutant to form conidia, which play a pivotal role as secondary inoculum in the disease cycle of the rice blast. Strikingly, conidia that are produced from the mutant were rarely able to germinate because they were metabolically inactive. To date, only a handful of genes that regulate conidiogenesis have been identified and characterized, and details of morphogenetic and metabolic processes associated with conidiogenesis are largely unknown. Considering misregulation of metabolism genes in the mutant mycelia, we conjecture that MoSAS3 might serve as an important regulator that orchestrates the metabolic changes required during conidiogenesis. Moreover, it appeared that such a defect in initial development has a detrimental effect on down‐stream processes in the mutant, impairing germination, appressorium formation and eventually pathogenesis. However, it should be noted that a number of different factors, including cytological malfunction of cellular organelles or defect in gene expression, may contribute to the inability of the mutant conidia to germinate. Our sheath assay showed that rice cells challenged with ΔMosas3 accumulate brown granules and/or show discolouration. Such a response is typically viewed as being associated with inability of the fungus to suppress plant immune responses and is reminiscent of the ΔModes1 phenotype (Chi et al., 2009), although expression of MoDES1 was not altered in ΔMosas3 compared to the wild‐type. However, it cannot be ruled out that MoDES1 expression is under faulty regulation during the early stage of plant infection. It is noteworthy that a large proportion of genes encoding putative secreted proteins are misregulated in the mutant (Fig. S9, see Supporting Information). Collectively, these observations suggest that many fungal genes involved in interaction with host plants seem to be affected by MoSAS3.

Given that deletion of Sas3 in yeast has little impact on phenotypes (Rosaleny et al., 2007), it is interesting to see large effects of MoSAS3 deletion on development and pathogenicity in M. oryzae. Yeast Sas3 has been shown to function in association with GNAT family HAT, Gcn5 (Rosaleny et al., 2007). In contrast to deletion of the yeast gene pair Gcn5 and Sas3, showing synthetic lethality, deletion of MoGCN5 in ΔMosas3 background did not change the phenotypes of ΔMosas3, suggesting different integration of two evolutionarily conserved components into the epigenetic regulatory system between S. cerevisiae and M. oryzae. We propose that two scenarios, which are not mutually exclusive, can explain such a difference between the two species. First, most if not all MoGCN5 targets could be a subset of MoSAS3 targets. Second, MoSAS3 can have cytoplasmic targets, as evidenced by our MoSAS3 protein localization data, unlike its yeast counterpart. It is of note that MoGCN5 has also been shown to have cytoplasmic targets, one of which is an autophagy protein, MoATG7 (MGG07297) (Zhang et al., 2017). This is in stark contrast to S. cerevisiae, where Esa1, which is a part of the NuA4 complex that is responsible for global H4 acetylation, is the only yeast HAT so far known to have non‐histone substrates (Sapountzi and Cote, 2011). Taken together, these data suggest that there is a degree of functional variation and target protein specificity within each group, and that there is a caveat when generalizing and newly predicting the biological function of acetyltransferases in other species based on previous studies in model organisms.

In summary, our data indicate that MoSAS3 is a conserved epigenetic component responsible for acetylation of H3K14. Although MoSAS3 is likely to have non‐histone substrates as well, such HAT activity seems to be important for growth and pre‐penetration development in M. oryzae. Transcriptome data provide a novel insight into the profound implications of MoSAS3 on DNA damage responses/repair and regulation of cellular metabolisms. Further elucidation of target genes and potential non‐histone substrates would shed light on species‐specific quirks and surprises, and subsequent development of a novel control strategy for fungal diseases of plants.

Experimental Procedures

Fungal isolates and culture conditions

The wild‐type strain M. oryzae KJ201 was procured from the Center for Fungal Genetic Resources (CFGR), South Korea. All strains, including mutants, were grown on oatmeal agar plates (OMA) [5% oatmeal (w/v) and 2% agar (w/v)] at 25 °C under constant white light to promote conidiation. Three‐day‐old vegetative hyphae harvested from the complete medium [CM, 0.6% yeast extract (w/v), 0.6% casamino acid (w/v) and 1% sucrose (w/v); 25 °C with 120 rpm shaking] were used for RNA and DNA extraction. Selection of hygromycin‐resistant transformants was carried out using TB3 agar plates [0.3% yeast extract (w/v), 0.3% casamino acid (w/v), 1% glucose (w/v), 20% sucrose (w/v) and 0.8% agar powder (w/v)] supplemented with 200 ppm hygromycin B.

Test of growth, conidiation and appressorium formation

For measurement of radial mycelial growth rate, conidiation, germination and appressorium formation, strains were cultured on either V8 agar or OMA plates in three replicates for 9 days at 25 °C under continuous light conditions. Conidia were harvested from 9‐day‐old mycelia in 5 mL of sterile water. Conidiation was determined by counting the number of conidia using a haemocytometer under the light microscope. Next, the conidial suspension was adjusted to a concentration of 5 × 104 conidia/mL. Forty microlitres of the conidial suspension was dropped onto plastic coverslips with three replicates and incubated in a humidity box at 25 °C. After 4 h of incubation, the frequency of conidial germination was determined by counting at least 100 conidia per replicate under the bright field microscope. Eight hours after incubation, the frequency of appressorium formation was determined by counting number of conidia that formed appressoria among germinated conidia (at least 100 total conidia in each replicate) under the light microscope. All the assays were performed with three replicates in three independent experiments. Slides were examined under a Leica DM2500 light microscope and imaged with a Leica DFC7000 T digital camera using Leica Application Suite v4.

Pathogenicity assay

For pathogenicity assay, we either sprayed spore suspensions (c.105/mL) directly onto the rice seedlings or injected into the leaves of rice seedlings (Oryza sativa ‘Nakdongbyeo’) of three‐ to four‐leaf stages through wound sites. The inoculated plants were placed in a humidity box for 24 h under dark conditions at 25 °C, then transferred to an incubator at 25 °C with a 16‐h photoperiod with fluorescent lights. Disease lesions were assessed and photographed at 7 days post‐inoculation. Plants grown in three pots were used for pathogenicity assay of individual strains in each of three independent experiments.

Targeted gene deletion and complementation

Knockout constructs were generated using the strategy of double‐joint PCR (Yu et al., 2004). For this, about 1.5 kb of flanking sequences of each gene were amplified and fused on either side of the hygromycin resistance gene (HPH) cassette. Next, the knockout constructs were transformed into the wild‐type protoplasts to generate transformants. Candidate transformants were screened using a PCR‐assisted screening method using specific primer pairs for the genes. Positive candidate knockout mutants were confirmed by Southern blot analysis using one of the flanking sequences as a probe. Southern blot DNA hybridization was performed with the selected transformants to ensure correct gene replacement events and absence of ectopic integration. Genomic DNA was digested with EcoR1, and blots were probed with 1‐kb 5′‐flanking or 3′‐flanking sequences (Fig. S1). Southern blot DNA hybridization was performed using a standard method. Complementation experiments on knockout mutants were carried out by amplifying the target gene and flanking sequences (~1 kb on each side) from the wild‐type and introducing the resulting PCR products along with pII99 vector harbouring the geneticin resistance gene marker into the mutant protoplasts. Geneticin‐resistant colonies were selected and subsequently screened by PCR for the presence of the target gene.

Western blot analysis and histone acetyltransferase assay for H3K14 acetylation

Total protein extracted from mycelia was separated by 12% SDS‐PAGE. Blots were probed with histone H3 acetyl‐lysine K14 antibody (1:2000, Cat. No. 39599, Active Motif, CA, USA) and H3 antibody as loading control (1:2000, Cat. No. 39164, Active Motif, CA, USA). Pierce Fast Western Blot Kit, ECL Substrate (Pierce Biotechnology, Rockford, IL, USA) was used and signals were detected on X‐ray films. Independent measurement of global H3K14 acetylation levels was performed using EpiQuik Global Acetyl Histone H3‐K14 Quantification Kit (colourimetric) according to the manufacturer's instruction (Epigentek, Farmingdale, NY, USA).

RNA sequencing and real‐time PCR analysis

Three‐day‐old mycelia harvested from the complete medium were used for total RNA extraction using an Easy‐spin total RNA extraction kit (iNtRON Biotechnology, Seoul, Korea) for all the strains. In RNA‐seq experiment, paired‐end sequencing of mRNAs from three biological replicates was carried out on the Illumina platform. TopHat was used to map the reads to the annotated M. oryzae transcripts, followed by analysis of transcript abundance and differential expression between the strains using Cufflinks. Expression levels were estimated as the number of fragments per kilobase per million reads (FPKM). Differentially expressed genes between the wild‐type and mutant strain were detected at the 0.01 significance level and with two‐fold change criteria. The FDR was used to maintain the significance level at 0.01. Fold changes of expression levels were calculated after adding one to each FPKM value to avoid division by zero. Gene ontology was carried out using the topGO package in R programming (Alexa and Rahnenfuhrer, 2019). Enrichment of genes in particular GO terms was analysed using Fisher's exact test (P < 0.01). For qRT‐PCR, 1 μg of total RNA was reverse transcribed into first‐strand cDNA with ImProm‐II Reverse Transcription System (Promega, Madison, WI, USA). Real‐time PCR was performed in a 20 μL volume containing 2 μL of cDNA (25 ng of input RNA), 1 μM of primer mix (forward and reverse) and 10 μL of 2 × Power SYBR Green PCR Master Mix (Applied Biosystems, Warrington, UK). Reactions were run on the Applied Biosystems 7500 Real Time PCR System (Applied Biosystems, Foster City, CA, USA) for 40 cycles of 15 s at 95 °C, 30 s at 60 °C and 30 s at 72 °C with three technical replicates. Resulting values of mean threshold cycles (Ct) were normalized as previously described (Livak and Schmittgen, 2001). The results of real‐time PCR were confirmed through three independent experiments.

Supporting information

Fig. S1 Targeted gene replacement of MoSAS3 (here shown as MoHAT10). Flanking sequences of MoSAS3 were amplified and fused to hygromycin cassette using double‐joint PCR (A). This PCR product was directly used for transformation of wild‐type protoplasts. Resulting transformants were screened by PCR (B) and correct gene replacement event was confirmed by Southern blot analysis (C). Location of primers used in PCR‐based screening (B) are designated in (A) as blue arrows. Primer pairs are supposed to give c.3 kb products only in the deletion mutant, and bands in the lower position in the gel appear to be non‐specific products.

Click here for additional data file. (5.1MB, pdf)

Fig. S2 Histone acetyltransferase activity of MoSAS3 and MoGCN5. (A) Western blot analysis of global H3K14 acetylation level in the ΔMosas3 strain compared to wild‐type (KJ201) and complementation strain (MoSAS3 c). Total protein extracted was separated on a 15% polyacrylamide gel and probed with antibodies against H3K14ac. (B) Independent measurement of H3K14 acetylation levels in ΔMosas3 and ΔMogcn5 compared to the wild‐type (EpiQuik Global Acetyl Histone H3‐K14 Quantification Kit, Epigentek).

Click here for additional data file. (158.7KB, pdf)

Fig. S3 Phenotypes of ΔMogcn5. Germination and appressorium formation of ΔMogcn5 (A, B and C). Reduced pathogenicity of ΔMogcn5 on host plants (D and E). Asterisks indicate statistically significant differences (P < 0.001, Tukey HSD test).

Click here for additional data file. (592.7KB, pdf)

Fig. S4 Validation of RNA‐seq data via qRT‐PCR. Twelve (six up‐ and six down‐regulated) differentially expressed genes in our RNA‐seq data were randomly selected and their transcript abundance in the mutant relative to the wild‐type was examined using qRT‐PCR. Mean Ct values were obtained from three technical replicates. The consistent pattern of expression observed here was confirmed via two independent experiments. Correlation between qRT‐PCR and RNA‐seq data was calculated using Spearman's rank‐order in R programming language.

Click here for additional data file. (78KB, pdf)

Fig. S5 Summary of GO enrichment analysis (biological process). GO terms that are enriched with up‐regulated genes are shown (Fisher's exact test, P < 0.01). The area of the circle represents the number of genes assigned to the particular GO term. The colour of the circle indicates the proportion of genes assigned to the GO term in our data set among the total number of genes having that GO term in the genome.

Click here for additional data file. (876.9KB, pdf)

Fig. S6 Summary of GO enrichment analysis (biological process). GO terms that are enriched with down‐regulated genes are shown (Fisher's exact test, P < 0.01). The area of the circle represents the number of genes assigned to the particular GO term. The colour of the circle indicates the proportion of genes assigned to the GO term in our data set among the total number of genes having that GO term in the genome.

Click here for additional data file. (2.1MB, pdf)

Fig. S7 Venn diagram summarizing the overlap of differentially expressed genes between ΔMosas3 and ΔMortt109.

Click here for additional data file. (142.1KB, pdf)

Fig. S8 Genes encoding transcription factors (TF) that are differentially expressed in ΔMosas3. Percentage (y‐axis) of up‐ or down‐regulated (red and green bars, respectively) TF genes are shown relative to the total number of differentially expressed TFs (blue bars) across different families of TFs (x‐axis).

Click here for additional data file. (101.9KB, pdf)

Fig. S9 Number of differentially expressed genes among putative secreted protein‐coding genes. A whole list of putative secreted proteins was retrieved from the Fungal Secretome Database (http://fsd.snu.ac.kr/). SP and SP3 indicate secreted proteins predicted by SignalP and three other signal peptide prediction programs, respectively.

Click here for additional data file. (63.8KB, pdf)

Table S1 List of up‐regulated genes in ΔMosas3.

Click here for additional data file. (47KB, csv)

Table S2 List of down‐regulated genes in ΔMosas3.

Click here for additional data file. (57.9KB, csv)

Table S3 RNA‐seq analysis on genes involved in glycolysis.

Click here for additional data file. (2KB, csv)

Table S4 RNA‐seq analysis on genes involved in TCA cycle.

Click here for additional data file. (1.5KB, csv)

Table S5 Changes in expression of nitrogen metabolism genes in Mosas3 relative to the wild‐type.

Click here for additional data file. (13.5KB, docx)

Acknowledgements

This work was supported by grants from National Research Foundation of Korea (NRF‐2017R1A2A1A17069504, NRF‐2015M3A9B8028679, NRF‐2018R1A5A1023599, and NRF‐2017R1D1A3B03033932) and Korea Institute of Planning and Evaluation for Technology in Food, Agriculture, and Forestry through Agricultural Microbiome Program (918017‐04‐1‐CG000). SK is grateful for a graduate fellowship through the Brain Korea 21 Plus Program.

Contributor Information

Yong‐Hwan Lee, Email: yonglee@snu.ac.kr.

Junhyun Jeon, Email: jjeon@yu.ac.kr.

References

  1. Alexa, A. , Rahnenfuhrer, J. (2019) topGO: enrichment analysis. for gene ontology. R package version 2.36.0.
  2. Allfrey, V.G. and Mirsky, A.E. (1964) Structural modifications of histones and their possible role in the regulation of RNA synthesis. Science, 144, 559. [DOI] [PubMed] [Google Scholar]
  3. Allfrey, V.G. , Faulkner, R. and Mirsky, A.E. (1964) Acetylation and methylation of histones and their possible role in the regulation of RNA synthesis. Proc. Natl. Acad. Sci. USA. 51, 786–794. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Allis, C.D. , Berger, S.L. , Cote, J. , Dent, S. , Jenuwien, T. , Kouzarides, T. , Pillus, L. , Reinberg, D. , Shi, Y. , Shiekhattar, R. , Shilatifard, A. , Workman, J. and Zhang, Y . (2007) New nomenclature for chromatin‐modifying enzymes. Cell, 131, 633–636. [DOI] [PubMed] [Google Scholar]
  5. Altschul, S.F. , Gish, W. , Miller, W. , Myers, E.W. and Lipman, D.J. (1990) Basic local alignment search tool. J. Mol. Biol. 215, 403–410. [DOI] [PubMed] [Google Scholar]
  6. Canovas, D. , Marcos, A.T. , Gacek, A. , Ramos, M.S. , Gutierrez, G. , Reyes‐Dominguez, Y. and Strauss, J. (2014) The histone acetyltransferase GcnE (GCN5) plays a central role in the regulation of Aspergillus asexual development. Genetics, 197, 1175–1214. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Chi, M.H. , Park, S.Y. , Kim, S. and Lee, Y.H. (2009) A novel pathogenicity gene is required in the rice blast fungus to suppress the basal defenses of the host. PLoS Pathog. 5, e1000401. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Choi, J. , Kim, K.T. , Huh, A. , Kwon, S. , Hong, C. , Asiegbu, F.O. , Jeon, J. and Lee, Y‐H. (2015) dbHiMo: a web‐based epigenomics platform for histone‐modifying enzymes. Database, 2015, bav052. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Clarke, A.S. , Lowell, J.E. , Jacobson, S.J. and Pillus, L. (1999) Esa1p is an essential histone acetyltransferase required for cell cycle progression. Mol. Cell Biol. 19, 2515–2526. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Couch, B.C. and Kohn, L.M. (2002) A multilocus gene genealogy concordant with host preference indicates segregation of a new species, Magnaporthe oryzae, from M. grisea . Mycologia, 94, 683–693. [DOI] [PubMed] [Google Scholar]
  11. Couch, B.C. , Fudal, I. , Lebrun, M.H. , Tharreau, D. , Valent, B. , van Kim, P. , Nottéghem, J.L. and Kohn, L.M. (2005) Origins of host‐specific populations of the blast pathogen Magnaporthe oryzae in crop domestication with subsequent expansion of pandemic clones on rice and weeds of rice. Genetics, 170, 613–630. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Dean, R.A. (1997) Signal pathways and appressorium morphogenesis. Annu. Rev. Phytopathol. 35, 211–234. [DOI] [PubMed] [Google Scholar]
  13. Dean, R. , Van Kan, J.A. , Pretorius, Z.A. , Hammond‐Kosack, K.E. , Di Pietro, A. , Spanu, P.D. , Rudd, J.J , Dickman, M. , Kahmann, R. , Ellis, J. and Foster, G.D . (2012) The top 10 fungal pathogens in molecular plant pathology. Mol. Plant Pathol. 13, 414–430. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Ding, S.L. , Liu, W. , Iliuk, A. , Ribot, C. , Vallet, J. , Tao, A. , Wang, Y. , Lebrun, M.H. and Xu, J.R. (2010) The tig1 histone deacetylase complex regulates infectious growth in the rice blast fungus Magnaporthe oryzae . Plant Cell, 22, 2495–2508. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Dubey, A. and Jeon, J. (2017) Epigenetic regulation of development and pathogenesis in fungal plant pathogens. Mol. Plant Pathol. 18, 887–898. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Durant, M. and Pugh, B.F. (2006) Genome‐wide relationships between TAF1 and histone acetyltransferases in Saccharomyces cerevisiae . Mol. Cell Biol. 26, 2791–2802. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Essary, B.D. and Marshall, P.A. (2009) Assessment of FUN‐1 vital dye staining: yeast with a block in the vacuolar sorting pathway have impaired ability to form CIVS when stained with FUN‐1 fluorescent dye. J. Microbiol. Meth. 78, 208–212. [DOI] [PubMed] [Google Scholar]
  18. Feller, C. , Forne, I. , Imhof, A. and Becker, P.B. (2015) Global and specific responses of the histone acetylome to systematic perturbation. Mol. Cell, 57, 559–571. [DOI] [PubMed] [Google Scholar]
  19. Finn, R.D. , Attwood, T.K. , Babbitt, P.C. , Bateman, A. , Bork, P. , Bridge, A.J. , Chang, H.‐Y. , Dosztányi, Z. , El‐Gebali, S. , Fraser, M. , Gough, J. , Haft, D. , Holliday, G.L. , Huang, H. , Huang, X. , Letunic, I. , Lopez, R. , Lu, S. , Marchler‐Bauer, A. , Mi, H. , Mistry, J. , Natale, D.A. , Necci, M. , Nuka, G. , Orengo, C.A. , Park, Y. , Pesseat, S. , Piovesan, D. , Potter, S.C. , Rawlings, N.D. , Redaschi, N. , Richardson, L. , Rivoire, C. , Sangrador‐Vegas, A. , Sigrist, C. , Sillitoe, I. , Smithers, B. , Squizzato, S. , Sutton, G. , Thanki, N. , Thomas, P.D. , Tosatto, S.C.E , Wu, C.H. , Xenarios, I. , Yeh, L.‐S. , Young, S.‐Y. and Mitchell, A.L. (2017) InterPro in 2017‐beyond protein family and domain annotations. Nucleic Acids Res. 45, D190–D199. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Georgakopoulos, T. and Thireos, G. (1992) Two distinct yeast transcriptional activators require the function of the GCN5 protein to promote normal levels of transcription. EMBO J. 11, 4145–4152. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Gomez‐Rodriguez, E.Y. , Uresti‐Rivera, E.E. , Patron‐Soberano, O.A. , Islas‐Osuna, M.A. , Flores‐Martinez, A. , Riego‐Ruiz, L. , Rosales‐Saavedra, M.T. and Casas‐Flores, S . (2018) Histone acetyltransferase TGF‐1 regulates Trichoderma atroviride secondary metabolism and mycoparasitism. PLoS One, 13, e0193872. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Gonzalez‐Prieto, J.M. , Rosas‐Quijano, R. , Dominguez, A. and Ruiz‐Herrera, J. (2014) The UmGcn5 gene encoding histone acetyltransferase from Ustilago maydis is involved in dimorphism and virulence. Fung. Genet. Biol. 71, 86–95. [DOI] [PubMed] [Google Scholar]
  23. Grimaldi, B. , Coiro, P. , Filetici, P. , Berge, E. , Dobosy, J.R. , Freitag, M. , Selker, E.U. and Ballario, P. (2006) The Neurospora crassa white collar‐1 dependent blue light response requires acetylation of histone H3 lysine 14 by NGF‐1. Mol. Biol. Cell. 17, 4576–4583. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Horton, P. , Park, K.J. , Obayashi, T. , Fujita, N. , Harada, H. , Adams‐Collier, C.J. and Nakai, K. (2007) WoLF PSORT: protein localization predictor. Nucleic Acids Res. 35, W585–587. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Howe, L. , Auston, D. , Grant, P. , John, S. , Cook, R.G. , Workman, J.L. and Pillus, L. (2001) Histone H3 specific acetyltransferases are essential for cell cycle progression. Genes Dev. 15, 3144–3154. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Huang, F. , Paulson, A. , Dutta, A. , Venkatesh, S. , Smolle, M. , Abmayr, S.M. and Workman, J.L. (2014) Histone acetyltransferase Enok regulates oocyte polarization by promoting expression of the actin nucleation factor spire. Genes Dev. 28, 2750–2763. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Huang, F. , Abmayr, S.M. and Workman, J.L. (2016) Regulation of KAT6 acetyltransferases and their roles in cell cycle progression, stem cell maintenance, and human disease. Mol. Cell Biol. 36, 1900–1907. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Jeon, J. , Kwon, S. and Lee, Y.H. (2014) Histone acetylation in fungal pathogens of plants. Plant Pathol. J. 30, 1–9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. John, S. , Howe, L. , Tafrov, S.T. , Grant, P.A. , Sternglanz, R. and Workman, J.L. (2000) The something about silencing protein, Sas3, is the catalytic subunit of NuA3, a yTAF(II)30‐containing HAT complex that interacts with the Spt16 subunit of the yeast CP (Cdc68/Pob3)‐FACT complex. Genes Dev. 14, 1196–1208. [PMC free article] [PubMed] [Google Scholar]
  30. Kouzarides, T. (2000) Acetylation: a regulatory modification to rival phosphorylation? EMBO J. 19, 1176–1179. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Kouzarides, T. (2007) Chromatin modifications and their function. Cell, 128, 693–705. [DOI] [PubMed] [Google Scholar]
  32. Kwon, S. , Lee, J. , Jeon, J. , Kim, S. , Park, S.Y. , Jeon, J. and Lee, Y.‐H . (2018) Role of the histone acetyltransferase Rtt109 in development and pathogenicity of the rice blast fungus. Mol. Plant–Microbe Interact. 31, 1200–1210. [DOI] [PubMed] [Google Scholar]
  33. Lafon, A. , Chang, C.S. , Scott, E.M. , Jacobson, S.J. and Pillus, L. (2007) MYST opportunities for growth control: yeast genes illuminate human cancer gene functions. Oncogene, 26, 5373–5384. [DOI] [PubMed] [Google Scholar]
  34. Li, F. , Zheng, L.D. , Chen, X. , Zhao, X.L. , Briggs, S.D. and Du, H.N. (2017) Gcn5‐mediated Rph1 acetylation regulates its autophagic degradation under DNA damage stress. Nucleic Acids Res. 45, 5183–5197. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Livak, K.J. , Schmittgen, T.D. (2001) Analysis of relative gene expression data using real‐time quantitative PCR and the 2−ΔΔCT method. Methods. 25, 402–408. [DOI] [PubMed] [Google Scholar]
  36. Lu, J. , Cao, H. , Zhang, L. , Huang, P. and Lin, F. (2014) Systematic analysis of Zn2Cys6 transcription factors required for development and pathogenicity by high‐throughput gene knockout in the rice blast fungus. PLoS Pathog. 10, e1004432. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Maciel, J.L.N. , Ceresini, P.C. , Castroagudin, V.L. , Zala, M. , Kema, G.H.J. and McDonald, B.A. (2014) Population structure and pathotype diversity of the wheat blast pathogen Magnaporthe oryzae 25 years after its emergence in Brazil. Phytopathology, 104, 95–107. [DOI] [PubMed] [Google Scholar]
  38. Martin‐Urdiroz, M. , Oses‐Ruiz, M. , Ryder, L.S. and Talbot, N.J. (2016) Investigating the biology of plant infection by the rice blast fungus Magnaporthe oryzae . Fung. Genet. Biol. 90, 61–68. [DOI] [PubMed] [Google Scholar]
  39. McCullough, C.E. and Marmorstein, R. (2016) In vitro activity assays for MYST histone acetyltransferases and adaptation for high‐throughput inhibitor screening. Methods Enzymol. 573, 139–160. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Millard, P.J. , Roth, B.L. , Thi, H.P. , Yue, S.T. and Haugland, R.P. (1997) Development of the FUN‐1 family of fluorescent probes for vacuole labeling and viability testing of yeasts. Appl. Environ. Microbiol. 63, 2897–2905. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Molloy, S. (2010) Magnaporthe effectors on the move. Nat. Rev. Microbiol. 8, 466. [DOI] [PubMed] [Google Scholar]
  42. Musselman, C.A. and Kutateladze, T.G. (2011) Handpicking epigenetic marks with PHD fingers. Nucleic Acids Res. 39, 9061–9071. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Park, J. , Park, J. , Jang, S. , Kim, S. , Kong, S. , Choi, J. , Ahn, K. , Kim, J. , Lee, S. , Kim, S. , Park, B. , Jung, K. , Kim, S. , Kang, S. and Lee, Y‐H . (2008) FTFD: an informatics pipeline supporting phylogenomic analysis of fungal transcription factors. Bioinformatics, 24, 1024–1025. [DOI] [PubMed] [Google Scholar]
  44. Reifsnyder, C. , Lowell, J. , Clarke, A. and Pillus, L. (1996) Yeast SAS silencing genes and human genes associated with AML and HIV‐1 Tat interactions are homologous with acetyltransferases. Nat. Genet. 14, 42–49. [DOI] [PubMed] [Google Scholar]
  45. Rokudai, S. , Aikawa, Y. , Tagata, Y. , Tsuchida, N. , Taya, Y. and Kitabayashi, I. (2009) Monocytic Leukemia Zinc Finger (MOZ) interacts with p53 to induce p21 expression and cell‐cycle arrest. J. Biol. Chem. 284, 237–244. [DOI] [PubMed] [Google Scholar]
  46. Rokudai, S. , Laptenko, O. , Arnal, S.M. , Taya, Y. , Kitabayashi, I. and Prives, C. (2013) MOZ increases p53 acetylation and premature senescence through its complex formation with PML. Proc. Natl. Acad. Sci. USA. 110, 3895–3900. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Rosaleny, L.E. , Ruiz‐Garcia, A.B. , Garcia‐Martinez, J. , Perez‐Ortin, J.E. and Tordera, V. (2007) The Sas3p and Gcn5p histone acetyltransferases are recruited to similar genes. Genome Biol. 8, R119. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Sapountzi, V. and Cote, J. (2011) MYST‐family histone acetyltransferases: beyond chromatin. Cell Mol. Life Sci. 68, 1147–1156. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Scott, E.K. , Lee, T. and Luo, L. (2001) enok encodes a Drosophila putative histone acetyltransferase required for mushroom body neuroblast proliferation. Curr. Biol. 11, 99–104. [DOI] [PubMed] [Google Scholar]
  50. Smith, E.R. , Eisen, A. , Gu, W.G. , Sattah, M. , Pannuti, A. , Zhou, J.X. , Cook, R.G , Lucchesi, J.C and Allis, C.D . (1998) ESA1 is a histone acetyltransferase that is essential for growth in yeast. Proc. Natl. Acad. Sci. USA. 95, 3561–3565. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Sterner, D.E. and Berger, S.L. (2000) Acetylation of histones and transcription‐related factors. Microbiol. Mol. Biol. Rev. 64, 435–459. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Takechi, S. and Nakayama, T. (1999) Sas3 is a histone acetyltransferase and requires a zinc finger motif. Biochem. Bioph. Res. Co. 266, 405–410. [DOI] [PubMed] [Google Scholar]
  53. Talbot, N.J. (2003) On the trail of a cereal killer: exploring the biology of Magnaporthe grisea . Annu. Rev. Microbiol. 57, 177–202. [DOI] [PubMed] [Google Scholar]
  54. Tang, W. , Ru, Y.Y. , Hong, L. , Zhu, Q. , Zuo, R.F. , Guo, X.X. , Wang, J. , Zhang, H. , Zheng, X. , Wang, P. and Zhang, Z. (2015) System‐wide characterization of bZIP transcription factor proteins involved in infection‐related morphogenesis of Magnaporthe oryzae . Environ. Microbiol. 17, 1377–1396. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Vicente‐Munoz, S. , Romero, P. , Magraner‐Pardo, L. , Martinez‐Jimenez, C.P. , Tordera, V. and Pamblanco, M. (2014) Comprehensive analysis of interacting proteins and genome‐wide location studies of the Sas3‐dependent NuA3 histone acetyltransferase complex. FEBS Open Bio. 4, 996–1006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Voss, A.K. , Collin, C. , Dixon, M.P. and Thomas, T. (2009) Moz and retinoic acid coordinately regulate H3K9 acetylation, Hox gene expression, and segment identity. Dev. Cell, 17, 674–686. [DOI] [PubMed] [Google Scholar]
  57. Wilson, R.A. and Talbot, N.J. (2009) Under pressure: investigating the biology of plant infection by Magnaporthe oryzae . Nat. Rev. Microbiol. 7, 185–195. [DOI] [PubMed] [Google Scholar]
  58. Yan, X. and Talbot, N.J. (2016) Investigating the cell biology of plant infection by the rice blast fungus Magnaporthe oryzae . Curr. Opin. Microbiol. 34, 147–153. [DOI] [PubMed] [Google Scholar]
  59. Yan, Y. , Barlev, N.A. , Haley, R.H. , Berger, S.L. and Marmorstein, R. (2000) Crystal structure of yeast Esa1 suggests a unified mechanism for catalysis and substrate binding by histone acetyltransferases. Mol. Cell, 6, 1195–1205. [DOI] [PubMed] [Google Scholar]
  60. Yu, J.H. , Hamari, Z. , Han, K.H. , Seo, J.A. , Reyes‐Dominguez, Y. and Scazzocchio, C. (2004) Double‐joint PCR: a PCR‐based molecular tool for gene manipulations in filamentous fungi. Fung. Genet. Biol. 41, 973–981. [DOI] [PubMed] [Google Scholar]
  61. Zhang, N. , Cai, G. , Bhattacharya, D. , Luo, J. and Qiu, H. (2016) Progress on genomic analysis and taxonomy of the rice blast fungus (Pyricularia oryzae) and other Magnaporthales species. Phytopathology, 106, 29–29.26368515 [Google Scholar]
  62. Zhang, S. , Liang, M. , Naqvi, N.I. , Lin, C. , Qian, W. , Zhang, L.H. and Deng, Y.Z. (2017) Phototrophy and starvation‐based induction of autophagy upon removal of Gcn5‐catalyzed acetylation of Atg7 in Magnaporthe oryzae . Autophagy, 13, 1318–1330. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Zhang, Q. , Akhberdi, O. , Wei, D. , Chen, L. , Liu, H. , Wang, D. , Hao, X. and Zhu, X . (2018) A MYST histone acetyltransferase modulates conidia development and secondary metabolism in Pestalotiopsis microspora, a taxol producer. Sci. Rep. 8, 8199. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

Fig. S1 Targeted gene replacement of MoSAS3 (here shown as MoHAT10). Flanking sequences of MoSAS3 were amplified and fused to hygromycin cassette using double‐joint PCR (A). This PCR product was directly used for transformation of wild‐type protoplasts. Resulting transformants were screened by PCR (B) and correct gene replacement event was confirmed by Southern blot analysis (C). Location of primers used in PCR‐based screening (B) are designated in (A) as blue arrows. Primer pairs are supposed to give c.3 kb products only in the deletion mutant, and bands in the lower position in the gel appear to be non‐specific products.

Click here for additional data file. (5.1MB, pdf)

Fig. S2 Histone acetyltransferase activity of MoSAS3 and MoGCN5. (A) Western blot analysis of global H3K14 acetylation level in the ΔMosas3 strain compared to wild‐type (KJ201) and complementation strain (MoSAS3 c). Total protein extracted was separated on a 15% polyacrylamide gel and probed with antibodies against H3K14ac. (B) Independent measurement of H3K14 acetylation levels in ΔMosas3 and ΔMogcn5 compared to the wild‐type (EpiQuik Global Acetyl Histone H3‐K14 Quantification Kit, Epigentek).

Click here for additional data file. (158.7KB, pdf)

Fig. S3 Phenotypes of ΔMogcn5. Germination and appressorium formation of ΔMogcn5 (A, B and C). Reduced pathogenicity of ΔMogcn5 on host plants (D and E). Asterisks indicate statistically significant differences (P < 0.001, Tukey HSD test).

Click here for additional data file. (592.7KB, pdf)

Fig. S4 Validation of RNA‐seq data via qRT‐PCR. Twelve (six up‐ and six down‐regulated) differentially expressed genes in our RNA‐seq data were randomly selected and their transcript abundance in the mutant relative to the wild‐type was examined using qRT‐PCR. Mean Ct values were obtained from three technical replicates. The consistent pattern of expression observed here was confirmed via two independent experiments. Correlation between qRT‐PCR and RNA‐seq data was calculated using Spearman's rank‐order in R programming language.

Click here for additional data file. (78KB, pdf)

Fig. S5 Summary of GO enrichment analysis (biological process). GO terms that are enriched with up‐regulated genes are shown (Fisher's exact test, P < 0.01). The area of the circle represents the number of genes assigned to the particular GO term. The colour of the circle indicates the proportion of genes assigned to the GO term in our data set among the total number of genes having that GO term in the genome.

Click here for additional data file. (876.9KB, pdf)

Fig. S6 Summary of GO enrichment analysis (biological process). GO terms that are enriched with down‐regulated genes are shown (Fisher's exact test, P < 0.01). The area of the circle represents the number of genes assigned to the particular GO term. The colour of the circle indicates the proportion of genes assigned to the GO term in our data set among the total number of genes having that GO term in the genome.

Click here for additional data file. (2.1MB, pdf)

Fig. S7 Venn diagram summarizing the overlap of differentially expressed genes between ΔMosas3 and ΔMortt109.

Click here for additional data file. (142.1KB, pdf)

Fig. S8 Genes encoding transcription factors (TF) that are differentially expressed in ΔMosas3. Percentage (y‐axis) of up‐ or down‐regulated (red and green bars, respectively) TF genes are shown relative to the total number of differentially expressed TFs (blue bars) across different families of TFs (x‐axis).

Click here for additional data file. (101.9KB, pdf)

Fig. S9 Number of differentially expressed genes among putative secreted protein‐coding genes. A whole list of putative secreted proteins was retrieved from the Fungal Secretome Database (http://fsd.snu.ac.kr/). SP and SP3 indicate secreted proteins predicted by SignalP and three other signal peptide prediction programs, respectively.

Click here for additional data file. (63.8KB, pdf)

Table S1 List of up‐regulated genes in ΔMosas3.

Click here for additional data file. (47KB, csv)

Table S2 List of down‐regulated genes in ΔMosas3.

Click here for additional data file. (57.9KB, csv)

Table S3 RNA‐seq analysis on genes involved in glycolysis.

Click here for additional data file. (2KB, csv)

Table S4 RNA‐seq analysis on genes involved in TCA cycle.

Click here for additional data file. (1.5KB, csv)

Table S5 Changes in expression of nitrogen metabolism genes in Mosas3 relative to the wild‐type.

Click here for additional data file. (13.5KB, docx)

Articles from Molecular Plant Pathology are provided here courtesy of Wiley

RESOURCES