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Epigenetic regulation of nuclear processes in fungal plant pathogens

  • H. Martin Kramer,

    Roles Conceptualization, Funding acquisition, Writing – original draft, Writing – review & editing

    Affiliation Laboratory of Phytopathology, Wageningen University and Research, Wageningen, the Netherlands

  • David E. Cook,

    Roles Conceptualization, Funding acquisition, Writing – review & editing

    Affiliations Laboratory of Phytopathology, Wageningen University and Research, Wageningen, the Netherlands, Department of Plant Pathology, Kansas State University, Manhattan, Kansas, United States of America

  • Michael F. Seidl,

    Roles Conceptualization, Writing – review & editing

    Affiliations Laboratory of Phytopathology, Wageningen University and Research, Wageningen, the Netherlands, Theoretical Biology & Bioinformatics, Department of Biology, Utrecht University, Utrecht, the Netherlands

  • Bart P.H.J. Thomma

    Roles Conceptualization, Funding acquisition, Writing – review & editing

    bthomma@uni-koeln.de

    Affiliations Laboratory of Phytopathology, Wageningen University and Research, Wageningen, the Netherlands, University of Cologne, Institute for Plant Sciences, Cluster of Excellence on Plant Sciences (CEPLAS), Cologne, Germany

Abstract

Through the association of protein complexes to DNA, the eukaryotic nuclear genome is broadly organized into open euchromatin that is accessible for enzymes acting on DNA and condensed heterochromatin that is inaccessible. Chemical and physical alterations to chromatin may impact its organization and functionality and are therefore important regulators of nuclear processes. Studies in various fungal plant pathogens have uncovered an association between chromatin organization and expression of in planta-induced genes that are important for pathogenicity. This review discusses chromatin-based regulation mechanisms as determined in the fungal plant pathogen Verticillium dahliae and relates the importance of epigenetic transcriptional regulation and other nuclear processes more broadly in fungal plant pathogens.

Introduction

Arguably, one of the most important transition towards eukaryotic evolution has been cell compartmentalization, which allowed physical separation of diverse cellular processes [1]. One of the organelles that arose from this compartmentalization is the nucleus, the organelle that harbors most of the DNA of the eukaryotic cell. Nuclear processes include those that are required for short-term cell response, such as transcription, as well as processes that ensure long-term survival and inheritance, such as DNA replication and DNA repair.

Regulation of DNA-templated processes involves the histone code, made up of posttranslational chemical modifications to DNA-interacting histone proteins that help regulate genome functionality [2,3]. Eukaryotes have 4 canonical histone proteins (histone 2A, histone 2B, histone 3, and histone 4) that form globular octameric protein complexes by incorporation of 2 monomers of each histone protein [4]. By binding 145–147 bp of DNA, these protein complexes form nucleosomes that provide the packaging that is required to fit DNA into the confined space of the nucleus [4,5]. Each histone protein carries a flexible N-terminal tail that extends away from the nucleosome complex. These histone tails are enriched for amino acid residues that can undergo chemical modification, such as methylation, acetylation, and phosphorylation [6]. In addition to modifications to canonical histones, eukaryotes can also incorporate histone variants that substitute the core canonical histones in their nucleosomes [7,8]. The combination of histone modifications and variants cannot only locally regulate nuclear processes, but also more globally. Locally, nuclear processes can be regulated through histone modifications or variants that serve as recognition sites for enzymes that act on DNA [2,911], whereas they can affect DNA-accessibility over larger chromosomal regions and shape three-dimensional (3D) genome arrangements within the nucleus, leading global effects [12,13].

In filamentous fungi, the histone code was initially studied in the saprophyte Neurospora crassa [14,15]. In this species, histone modifications affect transcription, DNA replication, DNA repair, and chromosome segregation [14,1621]. The impact of histone modifications have been studied in other filamentous fungi, including saprophytes as well as human pathogens, such as Aspergillus species [22,23], and in various plant pathogens [2428]. One such plant pathogenic fungus is Verticillium dahliae, which causes vascular wilt disease in hundreds of host plants [29]. V. dahliae is an ascomycete, haploid fungus that reproduces predominantly asexually [30]. Genomic analyses on various strains uncovered that V. dahliae harbors a genome that evolved through large-scale chromosomal rearrangements, including duplications that have been followed by reciprocal gene losses [3034]. Eventually, these processes resulted in a genome structure that can be characterized by core genomic regions that are shared by all strains and so-called adaptive genomic regions (AGRs, previously called “lineage-specific,” LS) that show a considerable degree of plasticity among strains [27,31]. Recently, we explored epigenetic features in relation to transcriptional regulation and genome evolution in V. dahliae, resulting in novel insights into the role of epigenetic modifications (Fig 1) [27,3538]. In this review, we discuss epigenetic regulation of nuclear processes in plant pathogenic fungi, based on our recent findings for V. dahliae.

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Fig 1. A model of genome organization and epigenetic modifications in V. dahliae.

Chromosomal regions differentially display distinct chromatin features, associated with differences in 3D genome organization. (A) Representation of the chromatin structure on a linear V. dahliae chromosome. AGRs (green blocks) display a distinct open (uncondensed) chromatin profile, in which the nucleosomes are marked by tri-methylation of histone 3 lysine 27 (H3K27me3, green circles) [27]. H3K27me3-marked regions, consisting of AGRs, as well as particular regions of the core genome, are enriched for differentially expressed genes in vitro and in planta (DEGs, blue blocks) [35]. Centromeres (orange blocks) in V. dahliae, but not in all sister species are specifically associated with the LTR retrotransposon LTRE9 (pink blocks) [37]. The chromatin profile at centromeres consists of tightly packed nucleosomes that are marked by tri-methylation of histone 3 lysine 9 (H3K9me3, red circles) and by DNA-methylation (5mC, red stars) [36,37]. Besides LTRE9 TEs at the centromere, additional inactive TEs in the core genome are marked by H3K9me3 and 5mC, while active TEs in the AGRs are not associated with these marks. (B) Schematic 3D representation of the organization of 3 chromosomes in V. dahliae. Locally, genomic regions form TADs (indicated by dotted circles) that interact more strongly within the domain than with other domains. Intriguingly, TADs within AGRs are less well insulated and interact more freely with neighboring TADs [38]. Centromeres often form single TADs and display strong inter-centromeric interactions [37]. AGR, adaptive genomic region; TAD, topologically associating domain; TE, transposable element.

https://doi.org/10.1371/journal.ppat.1011525.g001

Epigenetic mechanisms affect transcription

To colonize their plant hosts, plant pathogens secrete so-called effector molecules to support host colonization, often through tampering with host immunity [39,40]. The coordinated expression of effectors requires specific transcriptional regulation, likely balancing the need to subvert the host immune system with the potential cost of triggering immunity or other negative consequences of transcription [41,42]. Regulation of transcription in eukaryotes involves binding of transcription factors to promoter regions, followed by recruitment of DNA-dependent RNA polymerases that generate mRNA molecules [43]. In plant pathogenic fungi, only a few transcriptional regulators have been identified that are involved in effector gene expression. For instance, the transcriptional regulator Sge1 of Fusarium oxysporum f. sp. lycopersici is required for expression of particular proteinaceous effector genes during infection, and deletion of Sge1 compromises pathogenicity on tomato [44]. Homologs of Sge1 were shown to also control expression of pathogenicity-related genes in several other plant pathogens [4548]. Similarly, the transcription factor Pf2 regulates expression of some pathogenicity-related genes in various necrotrophic fungal plant pathogens [4951]. The Magnaporthe oryzae regulator of G-protein signaling RGS1 acts as a transcriptional regulator by repressing the expression of numerous effector genes before plant penetration [52]. In these examples, the transcriptional regulators globally coordinate effector gene expression upon infection; however, it is important to note that individual effector genes are often expressed during specific stages of infection [41,42,53]. Therefore, additional transcriptional regulatory mechanisms are required to accurately express effector genes during infection.

The binding and recruitment of transcriptional machinery is influenced by chromatin condensation over broad genomic regions [10,54] and more locally by particular histone modifications that recruit or inhibit local protein binding [55,56]. Thus, epigenetic mechanisms are involved in the regulation of transcription. The chromatin at actively expressed genes is often hyperacetylated, while silent chromatin is hypoacetylated [57]. Therefore, histone acetylation may regulate transcription through activity of histone acetyl transferases (HATs) and histone deacetylase complexes (HDACs) [58,59]. Various fungal plant pathogens utilize HATs during infectious life stages [6062], including the corn smut fungus Ustilago maydis, the banana pathogen Fusarium oxysporum f. sp. cubense, and the wheat pathogen Fusarium graminearum. In these fungi, deletion of genes encoding HAT family members affected fungal virulence as well as lifestyle switches [6062]. In U. maydis, activity of the HDAC member Sir2 is involved in pathogenic development, although it is unclear whether Sir2 is truly active in deacetylation [63]. Recent results in Saccharomyces cerevisiae have raised concerns about the role of histone acetylation in transcriptional regulation, as temporal experiments show that transcriptional activation occurs before histone acetylation, which suggests that even though histone acetylation may be involved in transcriptional regulation during disease development, it may not directly activate transcription [64].

During axenic cultivation of numerous plant pathogenic fungi, when effector genes are generally repressed, DNA regions coding effector genes were enriched for H3K9me3 and H3K27me3, and mutants in genes encoding the histone lysine methyltransferases KMT1 and KMT6, respectively, that are responsible for the deposition of these marks displayed de-repressed effector gene expression [2426,28,6568]. As the histone modifications H3K9me3 and H3K27me3 are generally associated with inaccessible heterochromatin, these findings led to the hypothesis that genomic regions containing effector genes are heterochromatic, and therefore, inaccessible to the transcriptional machinery, when the pathogen grows outside of the host plant (Fig 2). Consequently, in order to express the effector genes that facilitate infection, pathogens are generally hypothesized to require chromatin de-condensation at effector gene-containing regions, possibly through depletion of H3K9me3 and H3K27me3 (Fig 2) [25,69,70].

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Fig 2. Hypothesis on epigenetic regulation of effector gene expression in fungal plant pathogens.

(A) Chromosomal regions containing effector genes (yellow blocks) are marked by heterochromatin-associated histone modifications (green circles) and the chromatin is condensed and inaccessible to transcription machinery (colored ellipses) when the pathogen does not require effector gene expression. (B) Upon plant recognition, chromosome regions containing effector genes lose their heterochromatin-associated histone modifications and the chromatin decondenses and becomes accessible to the transcription machinery.

https://doi.org/10.1371/journal.ppat.1011525.g002

To study epigenetic regulation of effector gene expression, analyses are preferably performed in planta, the niche where fungal plant pathogens naturally occur and the context in which they are most studied. However, epigenetic studies of fungal chromatin during plant infection are typically impeded by the usually low pathogen-to-plant biomass, resulting in excessive amounts of plant-derived sequences from interaction samples [69]. Consequently, to our knowledge, no analyses on genome-wide presence of histone modification dynamics during infection have yet been reported. Current reports that discuss epigenetic regulation of transcription in planta rely on association with H3K27me3 in vitro, genetic perturbation altering global H3K27me3 deposition, or the analysis of only a few representative genes in planta [25,28,65,67,68]. Therefore, the full scope of epigenetic regulation and histone modification dynamics that occur during fungal-host infection remains unknown.

In agreement with the notion that effectors are heterochromatically silenced in vitro, V. dahliae genomic regions harboring effector genes and other environmentally regulated genes are enriched for H3K27me3 in vitro, and loss of H3K27me3 in the Set7 deletion mutant, leads to transcription of previously marked genes [27,35]. These results are consistent with the idea that transcriptional regulation of effector gene expression in V. dahliae may also occur in a similar fashion as proposed in other filamentous plant pathogens [27,35]. However, experiments to test the link between histone modification status and transcriptional activity between different axenic conditions reveal a different situation [35]. Here, we find that although H3K27me3 is enriched at effector loci, and it is required for repression, our results do not suggest that depletion of H3K27me3 is the major event leading to transcriptional activation [35]. This is because differential expression of genes located in H3K27me3-marked domains do not require concomitant changes in H3K27me3 status [35], although we cannot fully exclude that H3K27me3 is lost in only a subset of fungal nuclei during effector gene activation and that chromatin immunoprecipitation followed by high-throughput sequencing (ChIP-seq), which is used to determine the presence of this marks, does not have sufficient sensitivity to detect such small differences within the population. Loss of H3K27me3 in the plant pathogenic fungi Fusarium graminearum, Fusarium fujikuroi, and M. oryzae similarly leads to induction of only a subset of previously H3K27me3-associated genes [24,26,65], showing that other factors are involved in transcriptional regulation as well. Interestingly, the lack of concomitant removal of H3K27me3 and transcriptional activation may have to do with the finding in V. dahliae that H3K27me3-marked chromatin is not nearly as condensed as H3K9me3-marked regions during fungal cultivation in vitro (Fig 1A) [27]. Thus, our results in V. dahliae, combined with the results in other fungi, indicate that the previously postulated hypothesis does not fully describe the role of H3K27me3 in regulation of effector gene expression. Rather, H3K27me3 presence may serve to help repress transcription during unwarranted conditions and serve as a binding site or nucleation point to regulate transcriptional activation upon detection of particular environmental signals.

Proteins recognizing and binding to histone modifications are generally known as histone readers and can function in various cellular processes, including transcriptional regulation [71]. For instance, the TAF3 subunit of the basal transcription complex TFIID specifically binds H3K4me3 in a human cell line [72]. In response to genotoxic stress, TAF3 is recruited to H3K4me3-marked genes to stimulate formation of the preinitiation complex and thereby regulates initiation of gene expression [73]. In contrast, binding of H3K4me3 in another human cell line by the histone reader ING2, a subunit of the mSin3a histone deacetylase complex, leads to rapid repression of gene expression [74]. It is also possible for a single histone reader to recognize multiple histone marks and elicit different transcriptional activity depending on the other present histone marks. For instance, the Arabidopsis histone reader EARLY BOLTING IN SHORT DAY (EBS) recognizes both H3K4me3 and H3K27me3 and can switch between binding those histone marks to balance gene expression [75]. As the histone code in plant pathogens has only been partially scrutinized, it is possible that H3K4me3 or another, perhaps yet unidentified, histone modification works in conjunction with H3K27me3 to regulate transcription of effector genes. Although cooperation between histone marks as target of a histone reader is unknown in fungi, it is clear that histone marks themselves do affect each other’s deposition. An interesting example of such interaction between histone modifications occurs between H3K36me3 and H3K27me3 [20,76]. H3K36me3 deposition by the F. fujikuroi histone methyltransferase Ash1 occurs predominantly in the transcriptionally inactive subtelomeric regions, and this presence of H3K36me3 antagonizes deposition of H3K27me3 [76]. In contrast, experimentally induced H3K27me2/3 deposition in N. crassa prevalently occurred at Ash1-deposited H3K36me3 domains [20]. In both fungi, H3K36me3 deposited by another histone methyltransferase, Set2, occurs at transcriptionally active genes [20,76]. These examples indicate that the genomic context, and the accompanying epigenetic context, matters for the functionality of histone marks, and that this may differ between fungal species. Thus, the relatively stable presence of H3K27me3 observed in V. dahliae may lead to different transcriptional outputs depending on the co-occurrence of other histone marks and on specific interactions with opposing histone readers. Interestingly, the recently described H3K27me3 histone readers, EPR-1 and BP1, contribute to transcriptional repression in the filamentous fungi F. graminearum and N. crassa [77,78]. As these histone readers inhibit binding of transcriptional machinery, it is conceivable that dynamic presence of such readers over stable H3K27me3 domains regulates transcription.

Besides potentially functioning as a direct regulator of transcription, H3K27me3 may also affect transcriptional regulation through shaping 3D organization of the genome within the nucleus. In other systems, including animals, plants, and fungi, genomes have been shown to display long-range intra-chromosomal and inter-chromosomal interactions [7982]. At smaller genomic distances, 3D organization results in the formation of topologically associating domains (TADs), which are local genome regions that interact preferentially within themselves, but not with adjacent genomic regions [13,83]. The exact functionality of TADs is still under debate [8487], but TADs have frequently been associated with transcriptional regulation, for instance, by facilitating enhancer–promoter interactions [83,8890]. Connecting the epigenome to the 3D genome, H3K27me3 has been shown to affect both local and global 3D chromosome organization [9193]. Thus, H3K27me3 may be involved in structuring the 3D genome to help regulate transcription, for instance, by providing conducive local 3D-chromatin micro-environments containing components of the transcriptional machinery. Our observations in V. dahliae indicate that genes within the H3K27me3-associated TADs in AGRs are more likely to be transcriptionally co-regulated than genes in core TADs [38]. This suggests that H3K27me3 may be involved in organizing the local 3D genome to help direct the divergent transcriptional profiles seen for TADs in AGRs compared to the core genome. The local 3D genome organization in N. crassa also differs between genomic regions, with transcriptionally silent regions displaying random internal contacts, while organization of active chromatin is more reminiscent of TAD structure [94]. However, this is mostly H3K27me2/3 independent, as H3K27me3 is not present on all the regions that display such random internal contacts [81,92,94]. Recently, such differences in 3D genome organization were investigated and discussed with respect to their impact on genome-wide DNA-templated processes [95].

Epigenetic mechanisms affect genome evolution

To survive over evolutionary timescales as a species, organisms need to gain, lose, or alter proteins performing particular functions through genome evolution. This may be particularly relevant for plant pathogens, because plants recognize the presence of potential pathogens, or their activity, through intra- and extracellular immune receptors that bind non-self or modified-self ligands, collectively named invasion patterns [40]. Such invasion patterns can be structural components of pathogen cells, also known as microbe-associated molecular patterns (MAMPs), signatures of pathogen-induced plant damage, termed damage-associated molecular patterns (DAMPs), or proteins or metabolites produced by the pathogen during host invasion [40,96]. In order to circumvent recognition by the host plant, successful pathogens evolve to secrete novel proteins that inhibit recognition or evolve to lose or mutate the recognized molecule [40]. In turn, plants evolve novel immune receptors to again restrict pathogen dissemination, leading to a co-evolutionary arms-race between plants and their pathogens, where single gene loss, gain, or mutation can alter the outcome of the interaction from compatible to incompatible or vice versa [39,96,97].

Genetic variation occurs via a combination of mechanisms, including DNA replication errors, external mutagens, chromosomal crossover events, transposable element (TE) activity, partial or whole-genome duplications, chromosome gain or loss, large-scale chromosome rearrangements, etc. [98]. In some cases, the generated genome variation leads to increased cell or organism viability, potentially affecting their frequency in the population, ultimately leading to evolution. Even though genome evolution is considered a stochastic process, it was found that various plant pathogens harbor genomic regions that display increased frequencies of genetic variation, while the majority of the genome, which is typically designated as the core genome, remains evolutionary rather stable [33,99106]. This compartmentalization can generally be described as regions that differ more frequently between strains of a species compared to regions that are more highly conserved, often termed the two-speed genome in pathogenomics [99,100]. The “fast-evolving,” plastic compartments of a two-speed genome are typically TE-rich [100], indicating that TE activity may be one of the more important drivers for evolution. The plastic compartments of V. dahliae are represented by the TE-rich AGRs that evolved through large-scale chromosome rearrangements and segmental duplications, followed by reciprocal gene losses [31,33]. Interestingly, the TEs located in AGRs are relatively young, transcriptionally active, and more frequently polymorphic when compared with TEs in the core genome, suggesting that these TEs actively contribute to shaping of the AGRs [33,34,106]. Additionally, the polymorphic TEs in AGRs are associated with in planta highly expressed pathogenicity-related genes, suggesting that TEs may be involved in transcriptional regulation as well [34,106].

Even though TE activity is beneficial to a certain extent, TE overactivity can be detrimental to genome stability, and, therefore, TEs are generally epigenetically silenced [107110]. In fungi, genomic regions that are enriched for TEs are often epigenetically silenced by H3K9me3 and cytosine methylation (5-methylcytocine, 5mC) [111115]. Similarly, in V. dahliae we found that H3K9me3 and 5mC co-localize on TE-rich genomic regions (Fig 1A) [27,36,37]. However, even though 5mC is generally thought to be involved in transcriptional silencing, we observed that loss of 5mC did not induce TE transcription, whereas loss of H3K9me3, and the accompanying loss of 5mC, leads to the induction of numerous TEs [36]. Thus, 5mC is not strictly necessary for TE silencing. Instead, 5mC may be subject to spontaneous deamination, causing C to T mutations, and thus potentially rendering affected TEs permanently inactive [116]. Cytosine deamination as a driver of genome evolution is well accepted in various taxonomic groups [117119]. For instance, simulations of DNA sequence evolution indicated that mutational pressure by cytosine deamination was vital for the evolution of isochore structures in the mammalian genomes [118]. Additionally, cytosine deamination has been proposed to constitute one of the main evolutionary forces in generating new transcription factor-binding sites in the human genome [120]. However, as cytosine methylation is mainly restricted to genomic regions containing TEs in V. dahliae, spontaneous deamination is likely mainly involved in the inactivation of TEs and less so in genome evolution more broadly. It needs to be noted that there is no evidence for repeat-induced point mutation (RIP) in the presumed asexual fungus V. dahliae, which could alternatively cause C to T mutations instead of spontaneous deamination [27,31,33].

Previous studies in V. dahliae indicated that relatively young and active TEs are associated with the evolution of AGRs [27,33]. Interestingly, the TEs in these AGRs have a lower fraction of C to T mutations (represented by the composite repeat-induced point mutation index, CRI) and display lower association with H3K9me3 and 5mC [27,33,34]. This indicates that C to T mutations happen more frequently in TEs that are marked with 5mC, and thus that spontaneous deamination may be a true end result of DNA methylation, but also that a particular subset of TEs is devoid of H3K9me3 and 5mC. It is interesting to speculate that this contributes to TE activity and helps drive evolution within the AGRs, but further direct experimental evidence is needed.

It remains unclear what dictates the disparate TE silencing observed in V. dahliae, but also other fungi [121,122]. One explanation is simply that of natural selection, where fungal cells with active TEs in the core genome suffer fitness penalty and do not flourish, while cells with active TEs in the AGRs experience less detrimental effects, and thus survive more frequently. Alternatively, the presence of specific epigenetic features of AGRs may constrain the deposition of H3K9me3 and 5mC on TEs in the plastic genome, thereby permitting elevated TE activity within AGRs. Furthermore, it is very intriguing that pairwise clustering of AGRs in the 3D genome corresponds with segmental duplications underlying their evolution [38]. More generally, we observed in V. dahliae that TADs in the H3K27me3-associated AGRs are weaker insulated than TADs in the core genome, meaning that TADs in AGRs are less well separated from their neighboring TADs (Fig 1B) [38]. We speculate that this organization may allow more promiscuous interactions and may promote higher genome instability. It is unclear what makes that AGRs cluster in the 3D genome, nor what the function of these membrane-less nuclear sub-compartments, so-called nuclear bodies, may be [123125]. Various types of nuclear bodies have been described, including nucleoli, Cajal bodies, nuclear speckles, nuclear stress bodies, and polycomb bodies, of which some are formed on chromatin and embed DNA, while others form in the nucleosol and do not contain DNA, but rather interact with chromatin [125,126]. Chromatin-containing nuclear bodies are proposed to form by a process called phase separation through the activity of self-aggregating chromatin-binding molecules or through the activity of individual chromatin bridging factors that cross-link separate chromatin sections, without self-aggregation [126]. Nuclear bodies formed with the non-aggregating bridging factors are usually less stable, as these molecules can more readily disperse into the nucleoplasm, whereas nuclear bodies formed with self-aggregating molecules are more stable, and can exist independent of chromatin [126]. Interestingly, the Drosophila H3K9me3-interacting protein HP1 was shown to aggregate in vitro and to nucleate into foci during early heterochromatin domain formation, suggesting that aggregation of HP1 may drive heterochromatin domain formation [127]. As such, H3K9me3-marked heterochromatin at distal genomic regions may cluster in the nucleus through the presence of HP1, for instance, at centromeres [128]. Similarly, H3K27me3-marked heterochromatin is bound by the polycomb repressive complex 1 (PRC1), of which the CBX2 protein member is capable of assembly through phase separation [129]. PRC1 components are absent from most fungi [130,131], yet other H3K27me3-readers may have an analogous function in fungi. Thus, it is possible that epigenetic differences between the AGR and core segments of V. dahliae [38] function to segregate core and AGR regions to promote the activity of TEs in AGRs in such a way that the core genome remains largely unaffected. In addition to histone modifications influencing TE activity, there is also mounting evidence that histone modifications impact the generation of DNA variation. Two recent studies used mutation accumulation experiments in fungi, subculturing the strains under minimal selection, allowing subsequent identification of de novo genome variation [132,133]. In the plant pathogen Zymoseptoria tritici, the causal agent of Septoria leaf blotch, heterochromatin defined by the presence of H3K9me3 and H3K27me3 impacted the accumulation of DNA variation. The genetic loss of H3K9me3 led to a significantly higher base mutation rate, as well as more frequent loss of specific accessory chromosomes, while genetic loss of H3K27me3 accounted for a small reduction in base accumulation in some genomic regions [132]. In the filamentous fungal saprophyte Neurospora crassa, mutation accumulation was found to be higher in both H3K27me3- and H3K9me3-marked regions [133]. Further research is needed to understand the mechanisms leading to increased variation accumulation rates, but the results clearly indicate that the epigenome is playing an important role in this nuclear process.

All organisms evolved DNA repair mechanisms that correct damage to the genome. Various DNA repair mechanisms exist, including double-strand break repair by homologous recombination or by nonhomologous end joining, and nucleotide and base-excision repair pathways [134,135]. Interestingly, the histone code has been implicated in these DNA repair mechanisms. For instance, histone methylation of lysine 79 on the tail of histone 3, and of lysine 20 on histone 4, as well as phosphorylation and ubiquitination of histone variant H2AX are involved in recruitment of double-strand break machinery [135]. Additionally, budding yeast mutants lacking the N-terminal tails of histones H2A and H3 displayed increased mutation rates due to deficient base-excision repair, indicating that chromatin plays an important role in this DNA repair mechanism [136]. Analysis of CRISPR-Cas–induced DNA double-strand breaks in M. oryzae indicated that multiple DNA repair pathways may function differently across the genome during DNA repair [137]. As such, the creation and repair of DNA lesions may be a significant driver of DNA variation fueling fungal pathogen evolution [138]. In fact, research in V. dahliae identified sequence signatures of double strand-break repair machinery at sites of chromosomal rearrangements, indicating that the evolution of the two-speed genome in V. dahliae involved erroneous double-strand break repair (Faino and colleagues). As these mechanisms are in part regulated on chromatin, there is increasing evidence that epigenetics is important for genome evolution [139].

Epigenetic mechanisms affect cell division

For mitotic and meiotic replication, organisms first require DNA replication, followed by segregation of chromosome pairs, and finally cell division [140]. DNA replication starts with unwinding and separation of DNA strands, resulting in formation of replication forks in which DNA-polymerases attach to the DNA. The formation of replication forks is favored in genomic regions with hyperacetylated euchromatin [141,142], and the timing of replication is further regulated on chromatin [142144]. Heterochromatic regions generally replicate later during the mitotic S-phase and this process involves histone acetylation and methylation, as well as the activity of histone readers recognizing heterochromatin-associated histones [145147]. Interestingly, an exception to late replicating heterochromatic regions are the centromeres in the fungus Schizosaccharomyces pombe, which are heterochromatic yet were shown to replicate relatively early [148].

After DNA replication, the generated chromosome pairs segregate through formation of microtubule spindles that attach to centromeric regions present on each chromosome [149]. Fungal centromeres vary significantly in composition and size between species, ranging from point centromeres of approximately 125 bp in size to regional centromeres of a few kb up till a few hundreds of kb [150,151]. Even though fungal centromeres vary widely, their chromatin is always characterized by presence of nucleosomes carrying the histone H3 variant CenH3 [150,151]. CenH3 is essential for centromere function, as it is the chromatin component that connects chromosomes to the microtubule spindle via the proteinaceous kinetochore complex [149]. Besides CenH3, fungal centromeres are often epigenetically characterized by H3K9me3 and 5mC [151,152], which we also found to be present at V. dahliae centromeres (Fig 1A) [37]. Additionally, in various plants and animals, a large set of histone modifications have been associated with centromeres [153]. The conservation of epigenetic profiles at centromeres in different organisms indicates that the epigenetic landscape likely plays a crucial role in centromere function, and thus in cell division. Such role of the epigenetic landscape, and perhaps especially of H3K9me3, may entail the formation of a nuclear sub-compartment and thereby drive the physical clustering of centromeres. Interestingly, incorporation of the S. pombe CenH3 homolog CENP-A is promoted by nearby heterochromatin, and heterochromatin-bearing minichromosomes in S. pombe localize close to centromeres, suggesting that heterochromatin formation drives the nuclear positioning of centromeres [154]. The crucial role of the epigenetic landscape in centromere functioning is further supported by studies into neocentromere formation, occurring upon centromere defects, showing that neocentromeres often form in H3K9me3-marked genomic regions [155]. However, neocentromeres in the human pathogenic fungus Cryptococcus deuterogattii are formed in genic regions that are not associated with H3K9me3 and 5mC [156,157], suggesting that these heterochromatic features are not essential. Moreover, C. deuterogattii chromosomes lacking their original centromere are unstable and undergo chromosomal fusions, after which the neocentromere loses its function [156,157]. These results suggest that although H3K9me3 and 5mC may not be essential for centromere function and cell viability in short-term, they are important for genome stability over longer evolutionary timescales.

Concluding remarks

In this review, we have highlighted the importance of epigenetics in the regulation of nuclear processes in fungal plant pathogens. Moreover, we describe experimental evidence that genomic regions containing effector genes are characterized by the presence of heterochromatic features, but it remains to be seen if in planta transcriptional activation requires chromatin de-condensation or epigenome remodeling. As we show that differential gene expression in vitro for only a subset of genes located in H3K27me3 domains involves local H3K27me3 depletion, we postulate that gene expression in planta may display local H3K27me3 depletion, but that it is not strictly required. To demonstrate this, future studies will require a functional and reliable procedure to perform in planta chromatin immunoprecipitation assays or other types of nuclear capture or single-cell interrogation.

Histone proteins are not only found in eukaryotes, but are also common in archaea [158,159], indicating that the potential for epigenetic regulation using histones is evolutionary ancient. As such, it is not surprising that nuclear processes have evolved to heavily rely on epigenetic regulation. This evolution resulted in an intricate mechanism, the histone code, in which particular histone modifications may have multiple different functions depending on their genetic localization and the co-occurrence with additional modifications. Therefore, it will be difficult to predict how the function of the genome is affected by specific changes in the histone code. Advances in single-cell sequencing and epigenome analyses [160,161] will deepen the understanding of epigenetics by providing increasingly more fine-grained information about the regulation and output of the epigenome. While the major focus of epigenetic research has been on transcriptional regulation, there is substantial evidence that the epigenome impacts the evolution of genomes, including that of fungal pathogens. This will be an important area to develop a mechanistic understanding in order to predict and intervene on the development of emergent fungal pathogens of plants and animals.

References

  1. 1. Chen AH, Silver PA. Designing biological compartmentalization. Trends Cell Biol. 2012;22:662–670. pmid:22841504
  2. 2. Dos Santos Á, Toseland CP. Regulation of nuclear mechanics and the impact on DNA damage. Int J Mol Sci. 2021;22:3178. pmid:33804722
  3. 3. Prakash K, Fournier D. Histone code and higher-order chromatin folding: a hypothesis. Genomics Comput Biol. 2017;3:e41. pmid:31245531
  4. 4. Luger K, Mäder AW, Richmond RK, Sargent DF, Richmond TJ. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature. 1997;389:251–260. pmid:9305837
  5. 5. Lowary PT, Widom J. Nucleosome packaging and nucleosome positioning of genomic DNA. Proc Natl Acad Sci U S A. 1997;94:1183–1188. pmid:9037027
  6. 6. Xhemalce B, Dawson MA, Bannister AJ. Histone modifications. In: Meyers RA, editor. Epigenetic regulation and epigenomics. Weinheim: Wiley VCH; 2012. p. 657–703.
  7. 7. Sarma K, Reinberg D. Histone variants meet their match. Nat Rev Mol Cell Biol. 2005;6:139–149. pmid:15688000
  8. 8. Talbert PB, Henikoff S. Histone variants at a glance. J Cell Sci. 2021;134:jcs244749. pmid:33771851
  9. 9. Bannister AJ, Kouzarides T. Regulation of chromatin by histone modifications. Cell Res. 2011;21:381–395. pmid:21321607
  10. 10. Grewal SIS, Jia S. Heterochromatin revisited. Nat Rev Genet. 2007;8:35–46. pmid:17173056
  11. 11. Yun M, Wu J, Workman JL, Li B. Readers of histone modifications. Cell Res. 2011;21:564–578. pmid:21423274
  12. 12. Spielmann M, Lupiáñez DG, Mundlos S. Structural variation in the 3D genome. Nat Rev Genet. 2018;19:453–467. pmid:29692413
  13. 13. Rowley MJ, Corces VG. Organizational principles of 3D genome architecture. Nat Rev Genet. 2018;19:789–800. pmid:30367165
  14. 14. Freitag M, Hickey PC, Khlafallah TK, Read ND, Selker EU. HP1 is essential for DNA methylation in Neurospora. Mol Cell. 2004;13:427–434. pmid:14967149
  15. 15. Tamaru H, Selker EU. A histone H3 methyltransferase controls DNA methylation in Neurospora crassa. Nature. 2001;414:277–283. pmid:11713521
  16. 16. Sasaki T, Lynch KL, Mueller CV, Friedman S, Freitag M, Lewis ZA. Heterochromatin controls γH2A localization in Neurospora crassa. Eukaryot Cell. 2014;13:990–1000.
  17. 17. Basenko EY, Sasaki T, Ji L, Prybol CJ, Burckhardt RM, Schmitz RJ, et al. Genome-wide redistribution of H3K27me3 is linked to genotoxic stress and defective growth. Proc Natl Acad Sci U S A. 2015;112:E6339–E6348. pmid:26578794
  18. 18. Smith KM, Phatale PA, Sullivan CM, Pomraning KR, Freitag M. Heterochromatin is required for normal distribution of Neurospora crassa CenH3. Mol Cell Biol. 2011;31:2528–2542.
  19. 19. Jamieson K, Rountree MR, Lewis ZA, Stajich JE, Selker EU. Regional control of histone H3 lysine 27 methylation in Neurospora. Proc Natl Acad Sci U S A. 2013;110:6027–6032. pmid:23530226
  20. 20. Bicocca VT, Ormsby T, Adhvaryu KK, Honda S, Selker EU. ASH1-catalyzed H3K36 methylation drives gene repression and marks H3K27me2/3-competent chromatin. Elife. 2018;7:e41497. pmid:30468429
  21. 21. Courtney AJ, Kamei M, Ferraro AR, Gai K, He Q, Honda S, et al. The histone variant H2A. Z is required to establish normal patterns of H3K27 methylation in Neurospora crassa. bioRxiv. 2020:2004–2020.
  22. 22. Palmer JM, Perrin RM, Dagenais TRT, Keller NP. H3K9 methylation regulates growth and development in Aspergillus fumigatus. Eukaryot Cell. 2008;7:2052–2060.
  23. 23. Reyes-Dominguez Y, Narendja F, Berger H, Gallmetzer A, Fernandez-Martin R, Garcia I, et al. Nucleosome positioning and histone H3 acetylation are independent processes in the Aspergillus nidulans prnD-prnB bidirectional promoter. Eukaryot Cell. 2008;7:656–663.
  24. 24. Connolly LR, Smith KM, Freitag M. The Fusarium graminearum histone H3 K27 methyltransferase KMT6 regulates development and expression of secondary metabolite gene clusters. PLoS Genet. 2013;9:e1003916. pmid:24204317
  25. 25. Soyer JL, El Ghalid M, Glaser N, Ollivier B, Linglin J, Grandaubert J, et al. Epigenetic control of effector gene expression in the plant pathogenic fungus Leptosphaeria maculans. PLoS Genet. 2014;10:e1004227. pmid:24603691
  26. 26. Zhang W, Huang J, Cook DE. Histone modification dynamics at H3K27 are associated with altered transcription of in planta induced genes in Magnaporthe oryzae. PLoS Genet. 2021;17:e1009376.
  27. 27. Cook DE, Kramer HM, Torres DE, Seidl MF, Thomma BPHJ. A unique chromatin profile defines adaptive genomic regions in a fungal plant pathogen. Elife. 2020;9:e62208. pmid:33337321
  28. 28. Schotanus K, Soyer JL, Connolly LR, Grandaubert J, Happel P, Smith KM, et al. Histone modifications rather than the novel regional centromeres of Zymoseptoria tritici distinguish core and accessory chromosomes. Epigenetics Chromatin. 2015;8:41. pmid:26430472
  29. 29. Fradin EF, Thomma BPHJ. Physiology and molecular aspects of Verticillium wilt diseases caused by V. dahliae and V. albo-atrum. Mol Plant Pathol. 2006:71–86. pmid:20507429
  30. 30. Klosterman SJ, Subbarao KV, Kang S, Veronese P, Gold SE, Thomma BPHJ, et al. Comparative genomics yields insights into niche adaptation of plant vascular wilt pathogens. PLoS Pathog. 2011;7:e1002137. pmid:21829347
  31. 31. de Jonge R, Bolton MD, Kombrink A, van den Berg GCM, Yadeta KA, Thomma BPHJ. Extensive chromosomal reshuffling drives evolution of virulence in an asexual pathogen. Genome Res. 2013;23:1271–1282. pmid:23685541
  32. 32. Faino L, Seidl M, Datema E, van den Berg GCM, Janssen A, Wittenberg AHJ, et al. Single-molecule real-time sequencing combined with optical mapping yields completely finished fungal genome. MBio. 2015;6:e00936–e00915. pmid:26286689
  33. 33. Faino L, Seidl MF, Shi-Kunne X, Pauper M, Van Den Berg GCM, Wittenberg AHJ, et al. Transposons passively and actively contribute to evolution of the two-speed genome of a fungal pathogen. Genome Res. 2016;26:1091–1100. pmid:27325116
  34. 34. Torres DE, Thomma BPHJ, Seidl MF. Transposable elements contribute to genome dynamics and gene expression variation in the fungal plant pathogen Verticillium dahliae. Genome Biol Evol. 2021;13:evab135.
  35. 35. Kramer HM, Seidl MF, Thomma BPHJ, Cook DE. Local rather than global H3K27me3 dynamics associates with differential gene expression in Verticillium dahliae. MBio. 2022;13:e03566–e03521.
  36. 36. Kramer HM, Cook DE, van den Berg GCM, Seidl MF, Thomma BPHJ. Three putative DNA methyltransferases of Verticillium dahliae differentially contribute to DNA methylation that is dispensable for growth, development and virulence. Epigenetics Chromatin. 2021;14:1–15.
  37. 37. Seidl MF, Kramer HM, Cook DE, Lorencini Fiorin G, van den Berg GCM, Faino L, et al. Repetitive elements contribute to the diversity and evolution of centromeres in the fungal genus Verticillium. MBio. 2020;11:e01714–e01720. pmid:32900804
  38. 38. Torres DE, Kramer HM, Tracanna V, Lorencini Fiorin G, Cook DE, Seidl MF, et al. Three-dimensional chromatin organization promotes genome evolution in a fungal plant pathogen. bioRxiv. 2023.
  39. 39. Jones JDG, Dangl JL. The plant immune system. Nature. 2006;444:323–329. pmid:17108957
  40. 40. Cook DE, Mesarich CH, Thomma BPHJ. Understanding plant immunity as a surveillance system to detect invasion. Annu Rev Phytopathol. 2015;53:541–563. pmid:26047564
  41. 41. Lanver D, Müller AN, Happel P, Schweizer G, Haas FB, Franitza M, et al. The biotrophic development of Ustilago maydis studied by RNA-seq analysis. Plant Cell. 2018;30:300–323. pmid:29371439
  42. 42. Gervais J, Plissonneau C, Linglin J, Meyer M, Labadie K, Cruaud C, et al. Different waves of effector genes with contrasted genomic location are expressed by Leptosphaeria maculans during cotyledon and stem colonization of oilseed rape. Mol Plant Pathol. 2017;18:1113–1126. pmid:27474899
  43. 43. Kadonaga JT. Eukaryotic transcription: an interlaced network of transcription factors and chromatin-modifying machines. Cell. 1998;92:307–313. pmid:9476891
  44. 44. Michielse CB, Van Wijk R, Reijnen L, Manders EMM, Boas S, Olivain C, et al. The nuclear protein Sge1 of Fusarium oxysporum is required for parasitic growth. PLoS Pathog. 2009;5:e1000637. pmid:19851506
  45. 45. Michielse CB, Becker M, Heller J, Moraga J, Collado IG, Tudzynski P. The Botrytis cinerea Reg1 protein, a putative transcriptional regulator, is required for pathogenicity, conidiogenesis, and the production of secondary metabolites. Mol Plant Microbe Interact. 2011;24:1074–1085. pmid:21635139
  46. 46. Jonkers W, Dong Y, Broz K, Kistler HC. The Wor1-like protein Fgp1 regulates pathogenicity, toxin synthesis and reproduction in the phytopathogenic fungus Fusarium graminearum. PLoS Pathog. 2012;8:1–18. pmid:22693448
  47. 47. Brown DW, Busman M, Proctor RH. Fusarium verticillioides SGE1 is required for full virulence and regulates expression of protein effector and secondary metabolite biosynthetic genes. Mol Plant Microbe Interact. 2014;27:809–823. pmid:24742071
  48. 48. Santhanam P, Thomma BPHJ. Verticillium dahliae Sge1 differentially regulates expression of candidate effector genes. Mol Plant Microbe Interact. 2013;26:249–256. pmid:22970788
  49. 49. Cho Y, Ohm RA, Grigoriev IV, Srivastava A. Fungal-specific transcription factor AbPf2 activates pathogenicity in Alternaria brassicicola. Plant J. 2013;75:498–514. pmid:23617599
  50. 50. Rybak K, See PT, Phan HTT, Syme RA, Moffat CS, Oliver RP, et al. A functionally conserved Zn2Cys6 binuclear cluster transcription factor class regulates necrotrophic effector gene expression and host-specific virulence of two major Pleosporales fungal pathogens of wheat. Mol Plant Pathol. 2017;18:420–434. pmid:27860150
  51. 51. Jones DAB, John E, Rybak K, Phan HTT, Singh KB, Lin SY, et al. A specific fungal transcription factor controls effector gene expression and orchestrates the establishment of the necrotrophic pathogen lifestyle on wheat. Sci Rep. 2019;9:15884. pmid:31685928
  52. 52. Tang B, Yan X, Ryder LS, Bautista MJA, Cruz-Mireles N, Soanes DM, et al. Rgs1 is a regulator of effector gene expression during plant infection by the rice blast fungus Magnaporthe oryzae. Proc Natl Acad Sci U S A. 2023;120:e2301358120. pmid:36913579
  53. 53. Yan X, Tang B, Ryder LS, MacLean D, Were VM, Eseola AB, et al. The transcriptional landscape of plant infection by the rice blast fungus Magnaporthe oryzae reveals distinct families of temporally co-regulated and structurally conserved effectors. Plant Cell. 2023:koad036. pmid:36808541
  54. 54. Lee DY, Hayes JJ, Pruss D, Wolffe AP. A positive role for histone acetylation in transcription factor access to nucleosomal DNA. Cell. 1993;72:73–84. pmid:8422685
  55. 55. Chen Y, Jørgensen M, Kolde R, Zhao X, Parker B, Valen E, et al. Prediction of RNA Polymerase II recruitment, elongation and stalling from histone modification data. BMC Genomics. 2011;12:1–16. pmid:22047616
  56. 56. Angelov D, Molla A, Perche P-Y, Hans F, Côté J, Khochbin S, et al. The histone variant macroH2A interferes with transcription factor binding and SWI/SNF nucleosome remodeling. Mol Cell. 2003;11:1033–1041. pmid:12718888
  57. 57. Kurdistani SK, Tavazoie S, Grunstein M. Mapping global histone acetylation patterns to gene expression. Cell. 2004;117:721–733. pmid:15186774
  58. 58. Lee KK, Workman JL. Histone acetyltransferase complexes: one size doesn’t fit all. Nat Rev Mol Cell Biol. 2007;8:284–295. pmid:17380162
  59. 59. Yang X-J, Seto E. The Rpd3/Hda1 family of lysine deacetylases: from bacteria and yeast to mice and men. Nat Rev Mol Cell Biol. 2008;9:206–218. pmid:18292778
  60. 60. Kong X, van Diepeningen AD, van der Lee TAJ, Waalwijk C, Xu J, Xu J, et al. The Fusarium graminearum histone acetyltransferases are important for morphogenesis, DON biosynthesis, and pathogenicity. Front Microbiol. 2018;9:654.
  61. 61. Liu J, An B, Luo H, He C, Wang Q. The histone acetyltransferase FocGCN5 regulates growth, conidiation, and pathogenicity of the banana wilt disease causal agent Fusarium oxysporum f. sp. cubense tropical race 4. Res Microbiol. 2022;173:103902.
  62. 62. González-Prieto JM, Rosas-Quijano R, Domínguez A, Ruiz-Herrera J. The UmGcn5 gene encoding histone acetyltransferase from Ustilago maydis is involved in dimorphism and virulence. Fungal Genet Biol. 2014;71:86–95.
  63. 63. Navarrete B, Ibeas JI, Barrales RR. Systematic characterization of Ustilago maydis sirtuins shows Sir2 as a modulator of pathogenic gene expression. Front Microbiol. 2023:14.
  64. 64. Martin BJE, Brind’Amour J, Kuzmin A, Jensen KN, Liu ZC, Lorincz M, et al. Transcription shapes genome-wide histone acetylation patterns. Nat Commun. 2021;12:210. pmid:33431884
  65. 65. Studt L, Rösler SM, Burkhardt I, Arndt B, Freitag M, Humpf HU, et al. Knock-down of the methyltransferase Kmt6 relieves H3K27me3 and results in induction of cryptic and otherwise silent secondary metabolite gene clusters in Fusarium fujikuroi. Environ Microbiol. 2016;18:4037–4054. pmid:27348741
  66. 66. Möller M, Schotanus K, Soyer JL, Haueisen J, Happ K, Stralucke M, et al. Destabilization of chromosome structure by histone H3 lysine 27 methylation. PLoS Genet. 2019;15:e1008093. pmid:31009462
  67. 67. Carlier F, Li M, Maroc L, Debuchy R, Souaid C, Noordermeer D, et al. Loss of EZH2-like or SU (VAR) 3–9-like proteins causes simultaneous perturbations in H3K27 and H3K9 tri-methylation and associated developmental defects in the fungus Podospora anserina. Epigenetics Chromatin. 2021;14:1–28.
  68. 68. Lukito Y, Lee K, Noorifar N, Green KA, Winter DJ, Ram A, et al. Regulation of host-infection ability in the grass-symbiotic fungus Epichloë festucae by histone H3K9 and H3K36 methyltransferases. Environ Microbiol. 2021;23:2116–2131.
  69. 69. Soyer JL, Möller M, Schotanus K, Connolly LR, Galazka JM, Freitag M, et al. Chromatin analyses of Zymoseptoria tritici: methods for chromatin immunoprecipitation followed by high-throughput sequencing (ChIP-seq). Fungal Genet Biol. 2015;79:63–70.
  70. 70. Chujo T, Scott B. Histone H3K9 and H3K27 methylation regulates fungal alkaloid biosynthesis in a fungal endophyte-plant symbiosis. Mol Microbiol. 2014;92:413–434. pmid:24571357
  71. 71. Musselman CA, Lalonde M-E, Côté J, Kutateladze TG. Perceiving the epigenetic landscape through histone readers. Nat Struct Mol Biol. 2012;19:1218–1227. pmid:23211769
  72. 72. Vermeulen M, Mulder KW, Denissov S, Pijnappel WWMP, van Schaik FMA, Varier RA, et al. Selective anchoring of TFIID to nucleosomes by trimethylation of histone H3 lysine 4. Cell. 2007;131:58–69. pmid:17884155
  73. 73. Lauberth SM, Nakayama T, Wu X, Ferris AL, Tang Z, Hughes SH, et al. H3K4me3 interactions with TAF3 regulate preinitiation complex assembly and selective gene activation. Cell. 2013;152:1021–1036. pmid:23452851
  74. 74. Shi X, Hong T, Walter KL, Ewalt M, Michishita E, Hung T, et al. ING2 PHD domain links histone H3 lysine 4 methylation to active gene repression. Nature. 2006;442:96–99. pmid:16728974
  75. 75. Yang Z, Qian S, Scheid RN, Lu L, Chen X, Liu R, et al. EBS is a bivalent histone reader that regulates floral phase transition in Arabidopsis. Nat Genet. 2018;50:1247–1253. pmid:30082787
  76. 76. Janevska S, Baumann L, Sieber CMK, Münsterkötter M, Ulrich J, Kämper J, et al. Elucidation of the two H3K36me3 histone methyltransferases Set2 and Ash1 in Fusarium fujikuroi unravels their different chromosomal targets and a major impact of Ash1 on genome stability. Genetics. 2018;208:153–171. pmid:29146582
  77. 77. Tang G, Yuan J, Wang J, Zhang Y-Z, Xie S-S, Wang H, et al. Fusarium BP1 is a reader of H3K27 methylation. Nucleic Acids Res. 2021;49:10448–10464. pmid:34570240
  78. 78. Wiles ET, McNaught KJ, Kaur G, Selker JML, Ormsby T, Aravind L, et al. Evolutionarily ancient BAH–PHD protein mediates Polycomb silencing. Proc Natl Acad Sci U S A. 2020;117:11614–11623. pmid:32393638
  79. 79. Dekker J, Rippe K, Dekker M, Kleckner N, Woodcock CL, Dimitrov S, et al. Capturing chromosome conformation. Science. (80-). 2002;295:1306–1311. pmid:11847345
  80. 80. Dong P, Tu X, Chu PY, Lü P, Zhu N, Grierson D, et al. 3D chromatin architecture of large plant genomes determined by local A/B compartments. Mol Plant. 2017;10:1497–1509. pmid:29175436
  81. 81. Klocko AD, Ormsby T, Galazka JM, Leggett NA, Uesaka M, Honda S, et al. Normal chromosome conformation depends on subtelomeric facultative heterochromatin in Neurospora crassa. Proc Natl Acad Sci U S A. 2016;113:15048–15053. pmid:27856763
  82. 82. Torosin NS, Anand A, Golla TR, Cao W, Ellison CE. 3D genome evolution and reorganization in the Drosophila melanogaster species group. PLoS Genet. 2020;16:e1009229.
  83. 83. Gonzalez-Sandoval A, Gasser SM. On TADs and LADs: spatial control over gene expression. Trends Genet. 2016;32:485–495. pmid:27312344
  84. 84. Beagan JA, Phillips-Cremins JE. On the existence and functionality of topologically associating domains. Nat Genet. 2020;52:8–16. pmid:31925403
  85. 85. Cavalheiro GR, Pollex T, Furlong EEM. To loop or not to loop: what is the role of TADs in enhancer function and gene regulation? Curr Opin Genet Dev. 2021;67:119–129. pmid:33497970
  86. 86. Arzate-Mejía RG, Cerecedo-Castillo AJ, Guerrero G, Furlan-Magaril M, Recillas-Targa F. In situ dissection of domain boundaries affect genome topology and gene transcription in Drosophila. Nat Commun. 2020;11:1–16.
  87. 87. Ghavi-Helm Y. Functional consequences of chromosomal rearrangements on gene expression: not so deleterious after all? J Mol Biol. 2020;432:665–675. pmid:31626801
  88. 88. Rao SSP, Huntley MH, Durand NC, Stamenova EK, Bochkov ID, Robinson JT, et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell. 2014;159:1665–1680. pmid:25497547
  89. 89. Dixon JR, Selvaraj S, Yue F, Kim A, Li Y, Shen Y, et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature. 2012;485:376–380. pmid:22495300
  90. 90. Kim J, Dean A. Enhancers navigate the three-dimensional genome to direct cell fate decisions. Curr Opin Struct Biol. 2021;71:101–109. pmid:34280668
  91. 91. Donaldson-Collier MC, Sungalee S, Zufferey M, Tavernari D, Katanayeva N, Battistello E, et al. EZH2 oncogenic mutations drive epigenetic, transcriptional, and structural changes within chromatin domains. Nat Genet. 2019;51:517–528. pmid:30692681
  92. 92. Galazka JM, Klocko AD, Uesaka M, Honda S, Selker EU, Freitag M. Neurospora chromosomes are organized by blocks of importin alpha-dependent heterochromatin that are largely independent of H3K9me3. Genome Res. 2016;26:1069–1080.
  93. 93. Cheutin T, Cavalli G. Polycomb silencing: from linear chromatin domains to 3D chromosome folding. Curr Opin Genet Dev. 2014;25:30–37. pmid:24434548
  94. 94. Rodriguez S, Ward A, Reckard AT, Shtanko Y, Hull-Crew C, Klocko AD. The genome organization of Neurospora crassa at high resolution uncovers principles of fungal chromosome topology. G3. 2022;12:jkac053.
  95. 95. Torres DE, Reckard AT, Klocko AD, Seidl MF. Nuclear genome organization in fungi: From gene folding to Rabl chromosomes. FEMS Microbiol Rev. 2023:fuad021. pmid:37197899
  96. 96. Boller T, Felix G. A renaissance of elicitors: perception of microbe-associated molecular patterns and danger signals by pattern-recognition receptors. Annu Rev Plant Biol. 2009;60:379–406. pmid:19400727
  97. 97. Cesari S. Multiple strategies for pathogen perception by plant immune receptors. New Phytol. 2018;219:17–24. pmid:29131341
  98. 98. Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P. How genomes evolve. Molecular biology of the cell. 4th edition. Garland Science; 2002.
  99. 99. Raffaele S, Farrer RA, Cano LM, Studholme DJ, MacLean D, Thines M, et al. Genome evolution following host jumps in the irish potato famine pathogen lineage. Science (80-). 2010;330:1540–1543. pmid:21148391
  100. 100. Dong S, Raffaele S, Kamoun S. The two-speed genomes of filamentous pathogens: Waltz with plants. Curr Opin Genet Dev. 2015:57–65. pmid:26451981
  101. 101. Haas BJ, Kamoun S, Zody MC, Jiang RHY, Handsaker RE, Cano LM, et al. Genome sequence and analysis of the Irish potato famine pathogen Phytophthora infestans. Nature. 2009;461:393–398. pmid:19741609
  102. 102. Torres DE, Oggenfuss U, Croll D, Seidl MF. Genome evolution in fungal plant pathogens: looking beyond the two-speed genome model. Fungal Biol Rev. 2020;34:136–143.
  103. 103. Raffaele S, Kamoun S. Genome evolution in filamentous plant pathogens: why bigger can be better. Nat Rev Microbiol. 2012;10:417–430. pmid:22565130
  104. 104. Frantzeskakis L, Kusch S, Panstruga R. The need for speed: compartmentalized genome evolution in filamentous phytopathogens. Mol Plant Pathol. 2019;20:3. pmid:30557450
  105. 105. Dutheil JY, Mannhaupt G, Schweizer G, Sieber MK, Münsterkötter M, Güldener U, et al. A tale of genome compartmentalization: the evolution of virulence clusters in smut fungi. Genome Biol Evol. 2016;8:681–704. pmid:26872771
  106. 106. Seidl MF, Thomma BPHJ. Transposable elements direct the coevolution between plants and microbes. Trends Genet. 2017;33:842–851. pmid:28800915
  107. 107. Grandaubert J, Balesdent MH, Rouxel T. Evolutionary and adaptive role of transposable elements in fungal genomes. Adv Bot Res. 2014;70:79–105.
  108. 108. Rebollo R, Horard B, Hubert B, Vieira C. Jumping genes and epigenetics: towards new species. Gene. 2010;454:1–7. pmid:20102733
  109. 109. Fedoroff N V. Transposable elements, epigenetics, and genome evolution. Science (80-). 2012;338:758–767.
  110. 110. Fouché S, Oggenfuss U, Chanclud E, Croll D. A devil’s bargain with transposable elements in plant pathogens. Trends Genet. 2021. pmid:34489138
  111. 111. Kouzminova E, Selker EU. Dim-2 encodes a DNA methyltransferase responsible for all known cytosine methylation in Neurospora. EMBO J. 2001;20:4309–4323. pmid:11483533
  112. 112. Zhang X, Liu X, Zhao Y, Cheng J, Xie J, Fu Y, et al. Histone H3 lysine 9 methyltransferase DIM5 is required for the development and virulence of Botrytis cinerea. Front Microbiol. 2016. pmid:27597848
  113. 113. Möller M, Habig M, Lorrain C, Feurtey A, Haueisen J, Fagundes WC, et al. Recent loss of the Dim2 DNA methyltransferase decreases mutation rate in repeats and changes evolutionary trajectory in a fungal pathogen. PLoS Genet. 2021;17:e1009448. pmid:33750960
  114. 114. He C, Zhang Z, Li B, Tian S. The pattern and function of DNA methylation in fungal plant pathogens. Microorganisms. 2020;8:227. pmid:32046339
  115. 115. Bewick AJ, Hofmeister BT, Powers RA, Mondo SJ, Grigoriev IV, James TY, et al. Diversity of cytosine methylation across the fungal tree of life. Nat Ecol Evol. 2019;3:479–490. pmid:30778188
  116. 116. Holliday R, Grigg GW. DNA methylation and mutation. Mutat Res—Fundam Mol Mech Mutagen. 1993;285:61–67. pmid:7678134
  117. 117. Lynch M, Sung W, Morris K, Coffey N, Landry CR, Dopman EB, et al. A genome-wide view of the spectrum of spontaneous mutations in yeast. Proc Natl Acad Sci U S A. 2008;105:9272–9277. pmid:18583475
  118. 118. Fryxell KJ, Zuckerkandl E. Cytosine deamination plays a primary role in the evolution of mammalian isochores. Mol Biol Evol. 2000;17:1371–1383. pmid:10958853
  119. 119. Lu Z, Cui J, Wang L, Teng N, Zhang S, Lam H-M, et al. Genome-wide DNA mutations in Arabidopsis plants after multigenerational exposure to high temperatures. Genome Biol. 2021;22:1–27.
  120. 120. Żemojtel T, Arndt PF, Behrens S, Bourque G, Vingron M. CpG deamination creates transcription factor–binding sites with high efficiency. Genome Biol Evol. 2011;3:1304–1311. pmid:22016335
  121. 121. Gupta YK, Marcelino-Guimarães FC, Lorrain C, Farmer A, Haridas S, Ferreira EGC, et al. Major proliferation of transposable elements shaped the genome of the soybean rust pathogen Phakopsora pachyrhizi. Nat Commun. 2023;14:1–16.
  122. 122. Montanini B, Chen P-Y, Morselli M, Jaroszewicz A, Lopez D, Martin F, et al. Non-exhaustive DNA methylation-mediated transposon silencing in the black truffle genome, a complex fungal genome with massive repeat element content. Genome Biol. 2014;15:411. pmid:25091826
  123. 123. Morimoto M, Boerkoel CF. The role of nuclear bodies in gene expression and disease. Biology (Basel). 2013;2:976–1033. pmid:24040563
  124. 124. Sawyer IA, Dundr M. Nuclear bodies: Built to boost. J Cell Biol. 2016;213:509–511. pmid:27241912
  125. 125. Mao YS, Zhang B, Spector DL. Biogenesis and function of nuclear bodies. Trends Genet. 2011;27:295–306. pmid:21680045
  126. 126. Erdel F, Rippe K. Formation of chromatin subcompartments by phase separation. Biophys J. 2018;114:2262–2270.
  127. 127. Larson AG, Elnatan D, Keenen MM, Trnka MJ, Johnston JB, Burlingame AL, et al. Liquid droplet formation by HP1α suggests a role for phase separation in heterochromatin. Nature. 2017;547:236–240.
  128. 128. Singh PB, Newman AG. On the relations of phase separation and Hi-C maps to epigenetics. R Soc Open Sci. 2020;7:191976. pmid:32257349
  129. 129. Tatavosian R, Kent S, Brown K, Yao T, Duc HN, Huynh TN, et al. Nuclear condensates of the Polycomb protein chromobox 2 (CBX2) assemble through phase separation. J Biol Chem. 2019;294:1451–1463. pmid:30514760
  130. 130. Lewis ZA. Polycomb group systems in fungi: new models for understanding polycomb repressive complex 2. Trends Genet. 2017;33:220–231. pmid:28196760
  131. 131. de Potter B, Raas MWD, Seidl MF, Snel B, Verrijzer CP. Uncoupled evolution of the Polycomb system and deep origin of non-canonical PRC1. bioRxiv. 2023:2004–2023.
  132. 132. Habig M, Lorrain C, Feurtey A, Komluski J, Stukenbrock EH. Epigenetic modifications affect the rate of spontaneous mutations in a pathogenic fungus. Nat Commun. 2021;12:5869. pmid:34620872
  133. 133. de la Peña MV, Summanen PAM, Liukkonen M, Kronholm I. Chromatin structure influences rate and spectrum of spontaneous mutations in Neurospora crassa. Genome Res. 2023;33:599–611.
  134. 134. Branzei D, Foiani M. Regulation of DNA repair throughout the cell cycle. Nat Rev Mol Cell Biol. 2008;9:297–308. pmid:18285803
  135. 135. Huertas D, Sendra R, Muñoz P. Chromatin dynamics coupled to DNA repair. Epigenetics. 2009;4:31–42. pmid:19218832
  136. 136. Meas R, Smerdon MJ, Wyrick JJ. The amino-terminal tails of histones H2A and H3 coordinate efficient base excision repair, DNA damage signaling and postreplication repair in Saccharomyces cerevisiae. Nucleic Acids Res. 2015;43:4990–5001.
  137. 137. Huang J, Rowe D, Subedi P, Zhang W, Suelter T, Valent B, et al. CRISPR-Cas12a induced DNA double-strand breaks are repaired by multiple pathways with different mutation profiles in Magnaporthe oryzae. Nat Commun. 2022;13:7168. pmid:36418866
  138. 138. Huang J, Cook DE. The contribution of DNA repair pathways to genome editing and evolution in filamentous pathogens. FEMS Microbiol Rev. 2022;46:fuac035. pmid:35810003
  139. 139. Seidl MF, Cook DE, Thomma BPHJ. Chromatin biology impacts adaptive evolution of filamentous plant pathogens. PLoS Pathog. 2016;12:e1005920.
  140. 140. Ohkura H. Meiosis: an overview of key differences from mitosis. Cold Spring Harb Perspect Biol. 2015;7:a015859. pmid:25605710
  141. 141. Aggarwal BD, Calvi BR. Chromatin regulates origin activity in Drosophila follicle cells. Nature. 2004;430:372–376.
  142. 142. Sansam CG, Pietrzak K, Majchrzycka B, Kerlin MA, Chen J, Rankin S, et al. A mechanism for epigenetic control of DNA replication. Genes Dev. 2018;32:224–229. pmid:29483155
  143. 143. Gindin Y, Valenzuela MS, Aladjem MI, Meltzer PS, Bilke S. A chromatin structure-based model accurately predicts DNA replication timing in human cells. Mol Syst Biol. 2014;10:722. pmid:24682507
  144. 144. MacAlpine DM, Almouzni G. Chromatin and DNA replication. Cold Spring Harb Perspect Biol. 2013;5:a010207. pmid:23751185
  145. 145. Casas-Delucchi CS, van Bemmel JG, Haase S, Herce HD, Nowak D, Meilinger D, et al. Histone hypoacetylation is required to maintain late replication timing of constitutive heterochromatin. Nucleic Acids Res. 2012;40:159–169. pmid:21908399
  146. 146. Schwaiger M, Kohler H, Oakeley EJ, Stadler MB, Schübeler D. Heterochromatin protein 1 (HP1) modulates replication timing of the Drosophila genome. Genome Res. 2010;20:771–780.
  147. 147. Brustel J, Kirstein N, Izard F, Grimaud C, Prorok P, Cayrou C, et al. Histone H4K20 tri-methylation at late-firing origins ensures timely heterochromatin replication. EMBO J. 2017;36:2726–2741. pmid:28778956
  148. 148. Kim S-M, Dubey DD, Huberman JA. Early-replicating heterochromatin. Genes Dev. 2003;17:330–335. pmid:12569122
  149. 149. Freitag M. The kinetochore interaction network (KIN) of ascomycetes. Mycologia. 2016;108:485–505. pmid:26908646
  150. 150. Yadav V, Sreekumar L, Guin K, Sanyal K. Five pillars of centromeric chromatin in fungal pathogens. PLoS Pathog. 2018;14:e1007150. pmid:30138484
  151. 151. Smith KM, Galazka JM, Phatale PA, Connolly LR, Freitag M. Centromeres of filamentous fungi. Chromosome Res. 2012;20:635–656. pmid:22752455
  152. 152. Roy B, Sanyal K. Diversity in requirement of genetic and epigenetic factors for centromere function in fungi. Eukaryot Cell. 2011;10:1384–1395. pmid:21908596
  153. 153. Achrem M, Szućko I, Kalinka A. The epigenetic regulation of centromeres and telomeres in plants and animals. Comp Cytogenet. 2020;14:265. pmid:32733650
  154. 154. Wu W, McHugh T, Kelly DA, Pidoux AL, Allshire RC. Establishment of centromere identity is dependent on nuclear spatial organization. Curr Biol. 2022;32:3121–3136. pmid:35830853
  155. 155. Nishimura K, Komiya M, Hori T, Itoh T, Fukagawa T. 3D genomic architecture reveals that neocentromeres associate with heterochromatin regions. J Cell Biol. 2019;218:134–149. pmid:30396998
  156. 156. Schotanus K, Heitman J. Centromere deletion in Cryptococcus deuterogattii leads to neocentromere formation and chromosome fusions. Elife. 2020;9:e56026.
  157. 157. Schotanus K, Yadav V, Heitman J. Epigenetic dynamics of centromeres and neocentromeres in Cryptococcus deuterogattii. PLoS Genet. 2021;17:e1009743.
  158. 158. Henneman B, Van Emmerik C, van Ingen H, Dame RT. Structure and function of archaeal histones. PLoS Genet. 2018;14:e1007582. pmid:30212449
  159. 159. Stevens KM, Swadling JB, Hocher A, Bang C, Gribaldo S, Schmitz RA, et al. Histone variants in archaea and the evolution of combinatorial chromatin complexity. Proc Natl Acad Sci U S A. 2020;117:33384–33395. pmid:33288720
  160. 160. Harada A, Kimura H, Ohkawa Y. Recent advance in single-cell epigenomics. Curr Opin Struct Biol. 2021;71:116–122.
  161. 161. Agbleke AA, Amitai A, Buenrostro JD, Chakrabarti A, Chu L, Hansen AS, et al. Advances in chromatin and chromosome research: perspectives from multiple fields. Mol Cell. 2020;76:881–901. pmid:32768408