Abstract

How developmental gene expression is activated, co-ordinated and maintained is one of the biggest questions in developmental biology. While transcription factors lead the way in directing developmental gene expression, their accessibility to the correct repertoire of genes can depend on other factors such as DNA methylation, the presence of particular histone variants and post-translational modifications of histones. Collectively, factors that modify DNA or affect its packaging and accessibility contribute to a chromatin landscape that helps to control the timely expression of developmental genes. Zebrafish, perhaps better known for their strength as a model of embryology and organogenesis during development, are coming to the fore as a powerful model for interpreting the role played by chromatin in gene expression. Several recent advances have shown that zebrafish exhibit both similarities and differences to other models (and humans) in the way that they employ chromatin mechanisms of gene regulation. Here, I review how chromatin influences developmental transcriptional programmes during early zebrafish development, patterning and organogenesis. Lastly, I briefly highlight the importance of zebrafish chromatin research towards the understanding of human disease and transgenerational inheritance.

Introduction

One of the best-studied problems in biology is the question of how a single-celled zygote gives rise to a multicellular patterned animal. Multiple model systems, including rodents (e.g. Mus musculus, Rattus norvegicus), worms (Caenorhabditis elegans), flies (Drosophila melanogaster), frogs (Xenopus laevis) and zebrafish (Danio rerio), have been instrumental in dissecting the signalling pathways and gene transcriptional programmes that drive development. Developmental pathways converge upon genomic DNA that encodes essentially identical information within multipotent cells, yet intriguingly, such cells do not respond identically to developmental signals. How each cell responds to a given developmental signal is in part controlled by chromatin architecture, which helps to determine which segments of the genome are freely available to be accessed by transcriptional regulators.

Transcription factor (TF) binding to protein-coding and non-coding genes (and their regulatory elements) can be modulated by constraining DNA accessibility in several different ways. Local modulation of the DNA fibre includes DNA methylation on cytosines, post-translational modifications of histone proteins and incorporation of variant histone proteins. Local alterations are integrated with higher order chromatin structure, in which DNA fibres fold three-dimensionally into looped domains and compartments. Local and three-dimensional (3D) chromatin architecture are interdependent in the control of transcriptional accessibility of DNA (reviewed in [1]).

Zebrafish are a popular animal model for developmental biology and for modelling human developmental disorders. A sequenced genome along with optically translucent, externally fertilised embryos make zebrafish ideal for examining the relationship between gene transcription and developmental outcomes. The easy access to large numbers of gametes and embryos at very early stages of development also makes the zebrafish model highly amenable to investigating how chromatin structure influences developmental outcomes from the very start of development [2,3].

This review will cover recent advances on how chromatin architecture controls developmental gene expression in zebrafish.

Early development

The very first steps in embryo development happen during a time of dynamic chromatin organisation and represent a crucial developmental period that has been actively researched in multiple animal models (see [47] for recent reviews). Research in zebrafish has recently shed light on how chromatin structure contributes to early developmental processes.

When a zygote forms, its nascent genome is kept mostly inactive at first. During the first few hours of existence, early zygotic cellular events are regulated solely by maternally inherited molecules. Gradually, the zygotic genome becomes active and is transcribed. This is known as Zygotic Genome Activation (ZGA), which happens during the Maternal to Zygotic Transition (MZT). At this point, all developmental control is transferred to the embryo [8]. Up to the main wave of ZGA (or ‘major’ ZGA), fertilised zebrafish embryos undergo a ‘cleavage’ phase of 10 rapid cell divisions. By 3 hours post-fertilisation (hpf), embryos have 1000 (1k) cells, are transcriptionally active and largely pluripotent [9]. By 4.3 hpf or ‘dome’ stage, the mid-blastula transition (MBT) has occurred, and ZGA is considered to be complete [9].

The earliest transcribed zygotic genes switch on at the 64-cell stage or ‘minor’ ZGA, beginning with transcription of the miR430 genes. Selected protein-coding genes and other non-coding RNAs are later transcribed at the 256- to 512-cell stage [10]. Solely zygotic genes are in the minority (152/592 zygotic transcripts), with 74% of genes being maternally as well as zygotically provided [10]. Products of the miR430 locus are essential for the clearance of a large proportion of maternal RNAs later in development [11].

A long-standing question is how early-expressed genes in the embryo are selected, which in turn likely depends on how chromatin patterns are transmitted from the gametes to the zygote and subsequently reprogrammed.

Chromatin structure programs the genome well before most zygotic genes are activated

The genome of newly formed zygotes is reprogrammed by the removal of DNA methylation and chromatin modifications that existed in the gametes, and their replacement or redistribution to a zygote-specific pattern. Locus-specific differences in DNA methylation, particularly in the form of 5-methylcytosine (5mC) at CpG dinucleotides, contribute to cell type identity in subsequent development [12,13]. In zebrafish, a passive and highly localised remodelling of the methylome first occurs just after fertilisation. A further wave of demethylation and re-methylation occurs after gastrulation, during organ specification [14]. The egg and sperm methylomes differ in zebrafish, and following fertilisation, the zygote is reprogrammed to reflect the sperm methylome pattern [15,16]. Surprisingly, paternal DNA is dispensable for this process [15]. Recent studies identified a fascinating interplay between the incorporation of histone variant H2Afv (the zebrafish orthologue of histone variant H2A.Z) and DNA methylation before and during ZGA [17,18].

DNA methylation and H2Avf anti-correlate in the genome of zebrafish embryos, and H2Afv restricts DNA methylation during development [17]. During ZGA, ‘Placeholder’ nucleosomes, comprising H2Afv and H3K4me1, equalise the DNA methylation patterns of the two parental genomes by passively deterring DNA methylation [18]. The authors of this study suggest that the maternal genome does not need to use the paternal genome as a ‘template’ to reprogram the maternal epigenome because reprogramming is driven by TFs that contact placeholder-occupied sites and maintain their identity. These TFs act on both genomes to achieve equivalent methylomes via Placeholders, which deter methylation and mark genes for proper transcriptional regulation [18].

Histone modifications undergo global erasure and re-establishment during early zebrafish development [1821]. Functional post-translational modifications of histones include histone H3 acetylation (associated with chromatin accessibility and active gene expression), histone H3 lysine 4 trimethylation (H3K4me3: associated with active promoters) and histone H3 lysine 27 trimethylation (H3K27me3: associated with transcription repression) [22]. H3K4me3, H3K27me3 and H3K36me3 (associated with transcription elongation) are present in both sperm and eggs of zebrafish [19]. H3K4me3 and H3K27me3 are present in their canonical forms in mature zebrafish gametes, and their genome-wide location differs between eggs and sperm, depending on the identity of genes expressed during gametogenesis and location of their enhancers [19]. After fertilisation and during the parental-to-zygotic transition, histone modifications are removed such that parental-specific enhancers are ‘de-memorised’ [19]. Subsequently, specific histone modifications are added to mark ‘primed’ promoters in the zygotic genome that are activated upon ZGA [19]. Some histone modifications are re-established well before the main wave of ZGA. For example, histone H3 lysine 27 acetylation (H3K27ac: associated with active gene expression and open chromatin) can be detected at the 4-cell stage and anti-correlates with DNA methylation. By the 256-cell stage, H3K27ac almost exclusively co-locates with H2Afv [19]. Intriguingly, DNA methylation is not involved in the positioning of H3K27ac, and it is unclear how this histone mark is specifically located [19].

Chromatin structure and the onset of the zygotic gene expression programme

By the time of the main wave of ZGA, the ‘primed’ promoters are divided into two groups; those that replace H3K27ac with H3K27me3 and correspond to silenced developmental genes, and those that retain H3K27ac and acquire H3K36me3 in gene bodies, indicating gene activation [19]. Promoters marked by H3K27ac at the 256-cell stage, therefore resolve into activated or repressed promoters by the dome stage, when the majority of zygotic genes are activated [19]. The H3K27ac mark is likely to be ‘permissive’ for promoter activation [19] and corresponds to accessible regions of the genome [23]. Thus, distinctive chromatin marks that appear well before most zygotic genes are activated to predict subsequent developmental programmes. From the onset of the main wave of ZGA, transcription-specific histone modifications appear [20,24,25], and nucleosomes become strongly positioned at promoters by the ‘dome’ stage [21]. Cap analysis of gene expression (CAGE) together with H3K4me3 distribution showed that two different promoter classes are evident during ZGA: ‘sharp’ promoters that have a single transcription start site (TSS), often associated with a TATA box, and ‘broad’ promoters that often overlap a CpG island and contain multiple TSSs. A transition from the ‘sharp’ to the ‘broad’ promoter class occurs commensurate with the main wave of genome activation and indicates that genes can differentially interpret maternal and zygotic regulatory environments [26].

What triggers the transcription of early-expressed genes? In zebrafish, early-activated genes rely on the maternally supplied (and also zygotically transcribed) pluripotency factors, sox19a, nanog and pou5f3 [19,27,28]. These genes encode TFs with ‘pioneer’ activity, which promotes transcription of the majority of early zygotic genes (reviewed in [8]). Nanog, for example, is essential for the transcription of particular patterning genes [29]. However, a direct, cell-autonomous role for Nanog in ZGA remains contentious; analysis of maternal only and maternal-zygotic (see Abbreviations) nanog mutants showed that its primary role is in the formation of the extra-embryonic yolk syncytial layer [30]. Published [19] and preliminary [31] evidence suggests that histone acetyl transferase (HAT) activity is essential for gene activation at the 1k-cell stage. HAT activity is maternally supplied [19,31], including as mRNA that needs to be translated [31]. The timing of maternal mRNA translation is also important for activation of the genome, including temporal translation of HAT transcripts [31,32]. Translation timing, in turn, may be controlled by mRNA polyadenylation [32]; furthermore, RNA metabolism and location are also controlled by particular RNA-binding proteins [33,34]. Maternal RNAs also need to be cleared before the majority of zygotic genes can be activated [8], and interestingly, clearance of a large proportion of maternal mRNA is mediated by N6-methyladenosine (m6A) modification [35] in addition to the degradation by the miR430 gene products [11].

Conversely, what prevents premature activation of early genes? Joseph et al. showed that histone proteins loaded into the zygotic nucleus in excess, by sheer abundance, can compete with transcriptional machinery to prevent access to early-expressed genes [36]. In support of this concept, a recent study provides evidence that Nanog and Pou5f3 non-specifically compete with histones to reduce nucleosome occupancy on DNA, particularly at nucleosome-enriched regions [37]. Another recent study used reflected light-sheet microscopy to track fluorescently labelled TFs (including Sox19b) to show that the chromatin-bound fraction of TFs increases over successive cell divisions. The authors propose that decreasing nuclear size with each cell division could physically concentrate TFs onto chromatin [38]. This is consistent with the idea that a gradual increase in nuclear to cytoplasmic ratio, which happens with the successive cell divisions that take place in the cleavage phase, plays a role in ZGA [39].

As embryos transition through early development to full ZGA, global changes in chromatin organisation and accessibility occur [7]. In zebrafish, ATAC-seq (assay for transposase-accessible chromatin using sequencing) was used to profile chromatin accessibility from the 64-cell stage through to ‘dome’ stage. At the 64-cell stage, very few loci are accessible, and accessible loci include the early-expressed miR-430 cluster [23,40]. A burst of accessible chromatin appears during the main wave of ZGA at the 1k-cell stage, 63% of which is located at the promoters [23]. By the ‘dome’ stage, accessible chromatin at promoters decreases to 29% [23]. These dynamic changes in chromatin accessibility indicate that a dramatic nuclear reorganisation takes place during early development.

Three-dimensional genome organisation in early development

The spatial organisation of the nucleus changes as cells commit to developmental fates (reviewed in [41]). One obvious example of this is the organisation of chromosomes into topologically associated domains, known as TADs (topologically associated domains) [1]. TAD borders are highly populated by architectural proteins in the nucleus such as CCCTC-binding factor (CTCF) and cohesin [42,43]. These 3D domains work in tandem with ‘compartments’ to segregate active and inactive regions of the genome [1]. Spatial organisation within TADs can facilitate transcription of developmental loci, such as Hox genes [4447], thereby determining cell fate and driving embryo development.

Several studies have shown that 3D chromatin structure is dynamically rearranged in early embryo development. Similar to somatic cells, the sperm of mice have TADs and chromatin loops, while in contrast, oocytes do not appear to have an organised nucleus (reviewed in [4,5]). In most studied organisms, fertilised zygotes do not have a defined 3D chromatin structure, and TADs gradually emerge commensurate with the onset of zygotic transcription [5]. In contrast with other studied animals to date, TADs appear to be present pre-ZGA in zebrafish (∼128-cell stage); remarkably, they almost completely disappear by 4 hpf (just post-ZGA) and re-emerge by 8 hpf (gastrulation) [48]. It is not clear at exactly what point TADs are removed; or whether they are also present much earlier in development, before the 64-cell stage, at which most chromatin modifications predictive of gene activation have already been established [18,19]. It is probably significant that the dissolution and re-emergence of TADs coincide with the period of maximum chromatin accessibility of gene promoters [23]. The importance of TADs to genome activation is not yet known, but in Drosophila, TAD formation during ZGA does not depend on transcription [49].

The cohesin complex, along with CTCF, is crucial for TAD formation and 3D genome organisation (reviewed in [50,51]). Cohesin loss completely eliminates TADs [52]. In zebrafish, morpholino depletion of the Rad21 subunit of cohesin delays the onset of ZGA and results in nucleolar fragmentation and the dispersal of RNA Polymerase II (Pol II) foci at the ‘dome’ stage [40]. Overall, cohesin is essential for normal nuclear structure in zebrafish. Interestingly, between the 256-cell and ‘dome’ stages, there is a genome-wide shift of Rad21/cohesin binding location from repetitive regions of the genome (primarily pericentromeric satellite DNA) to gene-rich regions [40]. The timing of this shift coincides with the dis-establishment and re-establishment of TADs [48] and dynamic changes in chromatin accessibility [23]. Spatially structured, physical compartments appear to be involved in early transcription events in zebrafish. A recent study used live imaging to show that pri-miR430 and several zinc-finger gene transcripts, all encoded by genes on chromosome 4, cluster in a compartment with Pol II that represents a focus of detectable transcription before the MBT [53]. This DNA region is one of the very few locations enriched for cohesin binding ahead of the main wave of ZGA, suggesting a possible role for cohesin in the formation of these pre-MBT transcription compartments. By the post-ZGA ‘dome’ stage, when miR430 is down-regulated, cohesin binding disappears from this locus [40].

Figure 1 illustrates how chromatin structure changes as fertilised zygotes progress to the ‘dome’ stage, at which the zygotic genome is fully activated.

Summary of chromatin changes from fertilisation up to the ‘dome’ stage of zebrafish development.

Figure 1.
Summary of chromatin changes from fertilisation up to the ‘dome’ stage of zebrafish development.

The flow diagram depicts the order of appearance of histone variants, histone modifications, DNA methylation, TADs, chromatin accessibility, cohesin location and transcription factor activity. Maternal mRNA metabolism is also indicated. Green zones indicate the onset of transcription of zygotic genes.

Figure 1.
Summary of chromatin changes from fertilisation up to the ‘dome’ stage of zebrafish development.

The flow diagram depicts the order of appearance of histone variants, histone modifications, DNA methylation, TADs, chromatin accessibility, cohesin location and transcription factor activity. Maternal mRNA metabolism is also indicated. Green zones indicate the onset of transcription of zygotic genes.

Patterning

Once the zygotic genome has been activated, gastrulation begins, and cells shift from pluripotent transcriptional states to tissue-specific gene regulation. Chromatin structure plays an essential role in cell type specification.

Chromatin and cell fate specification during development

After genome activation (‘dome’ stage), chromatin modifications resolve to mark genes that are expressed, as well as those that are not [20]. Inactive genes can be marked with H3K4me3 together with H3K27me3 (i.e. containing transcription activating and repressing histone modifications; known as ‘bivalent’ [54]) or marked with K3K27me3 alone (transcription repressing) [20]. These histone signatures are similar to those observed in embryonic stem cells and could mean that non-expressed genes thus marked may be ‘poised’ for activation at a later developmental time [24]. Changes in the genome-wide distribution of H3K4me1 and H3K27ac revealed that a dramatic shift in the location of H3K27ac in particular accompanies the activation of enhancers for developmental genes [55].

DNA methylation profiling has been used to identify developmental stage-specific regions of methylation (dsDMRs) during zebrafish development. Six developmental stages incorporating sperm through pre-gastrulation (‘shield’ stage) and post-gastrulation (24 hpf) were surveyed for global methylation using MeDIP and MRE-seq (see Abbreviations) [56]. Many dsDMRs are marked with enhancer-associated histone modifications and TF binding motifs that correspond with developmental genes [56]. The dsDMR enhancer methylation pattern is strongly correlated with the expression of predicted gene targets [56]. Unusually, highly active enhancers are hyper-methylated in zebrafish, whereas those that are primed or ‘poised’ are hypo-methylated and expressed later in development [57]. This is opposite to what has been described in many other model systems. Inactive, hypo-methylated enhancers are marked prior to ZGA with H3K4me1, and subsequently gain H3K4me2/3, and sometimes H3K27me3 [57]. Interestingly, selected hypo-methylated enhancers tested by 4C-seq (see Abbreviations) remain stably associated with their target genes regardless of whether or not those genes are transcribed [57]. Formation of looped structures may coincide with TAD formation, which happens progressively after the main wave of ZGA [48]. Later in development, enhancer methylation patterns are more consistent with other species; widespread demethylation of enhancers occurs during zebrafish organogenesis [14].

DNA replication timing correlates with chromatin conformation and can also be used to predict changes in chromatin organisation during development. Expressed regions, including putative enhancers, replicate earlier in S phase than non-expressed regions [58]. Early-expressed genes, including those that are expressed in minor or major ZGA, replicate early in S phase even if these genes are yet to be activated [58]. Notably, the long arm of chromosome 4, which contains the miR430 locus, switches from early to late replication during gastrulation. Interestingly, cohesin binding spreads from the gene-poor long arm of chromosome 4 to the gene-rich short arm at ‘dome’ stage [40]. Considering cohesin is also required during DNA replication for sister chromatid cohesion from S phase until G2 in the cell cycle [59], it is possible that cohesin may play a role in developmentally regulated replication switches.

Developmental enhancers

Developmental signalling pathways converge on transcription networks to confer cell identity. During differentiation, specific TFs contact enhancers to activate the expression of cell type-specific genes. Large enhancer domains that are capable of recruiting multiple TFs and putatively regulate multiple genes are known as ‘super-enhancers’ (SEs) [60]. Genome-wide distribution of H3K27ac was used to confirm that zebrafish also have SEs and that SEs occupy locations near to important developmental genes that are conserved through evolution [61]. However, in contrast with mammalian SEs, zebrafish SEs were not especially enriched at gene TSSs or intronic regions and were more inclined to be intergenic [61]. Zebrafish SEs are also not conserved at the DNA sequence level, making it difficult to determine if SEs have a common evolutionary origin [61]. Using chromatin dynamics as a guide, recent research has made progress on identifying which enhancers are active in which tissues.

In differentiated cells, it can be technically difficult to find chromatin signatures associated with tissue-specific enhancer activity. This is because cells of interest must be isolated in enough numbers and in a pure enough population in order for experiments, such as ChIP-seq (chromatin immunoprecipitation followed by sequencing), to identify enhancers of interest. To overcome these challenges, one study applied fluorescent-activated nuclei sorting (FANS) followed by ATAC-seq to identify tissue-specific enhancers, in this case, in endothelial cells. Nuclei from endothelial cells marked with GFP were sorted and analysed with ATAC-seq to identify areas of open chromatin that could correspond to tissue-specific enhancers. Selected identified enhancers tested did indeed drive GFP expression in endothelial cells [62]. A similar study used an enhancer for Smarcd3 that is active in early mouse mesoderm to drive GFP expression in zebrafish early cardiac progenitors [63]. GFP expression was used to purify cardiac progenitors, and ATAC-seq was used to identify regions of open chromatin. Open regions were overlapped with H3K4me3, H3K4me1 and H3K27ac profiles, and compared with conserved non-coding element databases to predict enhancer locations [63].

The location of particular histone variants can also be used to identify enhancers that are active in distinct tissues and processes. For example, by expressing a tagged histone H3.3 in cardiomyocytes in zebrafish, Goldman et al. [64] were able to identify enhancers that are active following cardiac injury and during regeneration. In this experiment, a biotin-tagged histone H3.3 replaces histone H3 during nucleosome turnover at activated enhancers, allowing a streptavidin-based enrichment and sequencing approach to profile enhancer-mediated gene regulatory changes [64].

A less technical method involves predicting where enhancers might be based on phenotypic observations in a mutant or knock-down embryo. For example, a nonsense mutation in rad21, which encodes a subunit of cohesin, causes loss of expression of the runx1 gene in the posterior lateral mesoderm (PLM), but not in Rohon–Beard neurons, of bud-stage embryos [65]. This observation implies that different enhancers control runx1 expression in haematopoietic zones of the PLM versus neuronal tissue. The authors reasoned that enhancers involved in the tissue-specific expression of runx1 likely reside in proximity to Rad21/cohesin binding sites near the runx1 gene. Using a ChIP-PCR approach based on predicted cohesin/CTCF binding sites, they identified several regulatory elements that reside in the intron between runx1’s two promoters, P1 and P2 [66]. The location of these regulatory elements are broadly conserved in mice, and in turn, several regulatory elements surrounding the mouse Runx1 gene behave as active enhancers in zebrafish haematopoietic tissues [67].

An illustration of chromatin changes at enhancers and promoters that occur during cell type specification is shown in Figure 2.

Chromatin transitions upon cell fate specification during zebrafish embryogenesis.

Figure 2.
Chromatin transitions upon cell fate specification during zebrafish embryogenesis.

1. Soon after fertilisation, H2Afv and H3K4me1 marks appear. Some H2Afv colocalises with H3K27ac. The H2Afv Placeholder nucleosomes deter passive DNA methylation. 2. As embryos transition through ZGA, enhancers resolve to have H3K4me1 with H3K27ac, or H3K4me1 with H3K27me3. Active promoters (not shown) gain H3K4me3 and H3K27ac, while ‘poised’ promoters can have H3K4me3 and H3K27me3. 3. Upon cell specification, developmental enhancers are activated and recruit H3K27ac. The formation of 3D structure (TADs and compartments), partly due to cohesin activity, enables developmental enhancers to contact their target genes. Promoters have H3K4me3 and acquire H3K27ac upon activation.

Figure 2.
Chromatin transitions upon cell fate specification during zebrafish embryogenesis.

1. Soon after fertilisation, H2Afv and H3K4me1 marks appear. Some H2Afv colocalises with H3K27ac. The H2Afv Placeholder nucleosomes deter passive DNA methylation. 2. As embryos transition through ZGA, enhancers resolve to have H3K4me1 with H3K27ac, or H3K4me1 with H3K27me3. Active promoters (not shown) gain H3K4me3 and H3K27ac, while ‘poised’ promoters can have H3K4me3 and H3K27me3. 3. Upon cell specification, developmental enhancers are activated and recruit H3K27ac. The formation of 3D structure (TADs and compartments), partly due to cohesin activity, enables developmental enhancers to contact their target genes. Promoters have H3K4me3 and acquire H3K27ac upon activation.

Chromatin and organogenesis

Chromatin factors are emerging as key, and surprisingly specific, players in the development and patterning of distinct tissues in zebrafish (for recent diverse examples, see [6874]). The development of particular organs appears to be especially sensitive to perturbations in chromatin components and their regulators. For example, correct patterning (or regeneration) of the heart requires wild type levels of the histone lysine methyltransferase Smyd4 [68], chromatin remodellers Brg1 [69] and Smarce1 [73] and cohesin subunit Rad21 [74].

Chromatin components and their regulators are also well-known contributors to the development of the haematopoietic system, and mutations in genes encoding these factors frequently occur in leukaemia (reviewed in [75,76]). Zebrafish studies have contributed a great deal of information on how haematopoietic development can be influenced by chromatin. An in situ hybridisation-based reverse genetics screen identified 44 chromatin factors that affect primitive and definitive blood development in zebrafish [77]. The function of 425 zebrafish orthologues of human chromatin factors was blocked using morpholino oligonucleotides, and targeted embryos were screened using riboprobes detecting markers of blood development. The screen uncovered chromatin remodellers, histone modifying enzymes and members of the Polycomb repressor complex as regulators of haematopoiesis [77].

In addition to the activity of chromatin factors, DNA methylation status is also important for haematopoietic stem cell (HSC) emergence. The Tet2 and Tet3 enzymes, which convert 5-methylcytosine to 5-hydroxymethylcytosine (5hmC) to facilitate DNA demethylation, have overlapping functions upstream of Notch signalling to regulate HSC production [78]. Conversely, the DNA methyl transferase, Dnmt3bb, acts downstream of Notch and Runx1 to specifically methylate a CpG island in intron 1 of the c-myb gene. Interestingly, increased methylation at this location promotes c-myb expression, which in turn supports the maintenance of HSC fate [79]. It was not determined whether the methylated CpG island in intron 1 of c-myb corresponds to an enhancer for this gene. Finally, in addition to its role in maternal mRNA clearance, m6A mRNA methylation also appears to have an important role in HSC specification [80]. In 28 hpf zebrafish embryos, m6A-mediated clearance of mRNA encoded by arterial endothelial genes notch1a and rhoca dampens Notch signalling to allow HSCs to emerge from the haemogenic endothelium [80].

Chromatin factors and DNA methylation essential for blood development likely contribute to the selection and activity of enhancers that specify haematopoietic cell fate. Some chromatin regulators, for example, Dnmt3bb [79] and Rad21/cohesin [65], appear to have extraordinarily specific roles in controlling the expression of specific genes; however, it is far from clear how this specificity is achieved.

Zebrafish models for chromatin mechanisms

Zebrafish oddities

While in many cases zebrafish have equivalency with mammalian systems, at the same time many chromatin-based mechanisms that regulate early developmental processes in zebrafish appear to be unique to that system. Erasure and re-establishment of DNA methylation following fertilisation is not as clear cut in fish as in mammals [23,56]; instead, the DNA methylation pattern of the newly formed zebrafish zygote quickly resolves to reflect the methylation pattern of sperm within a couple of hours post-fertilisation [15,16]. 3D genome structure gradually appears the following fertilisation in mammalian and Drosophila zygotes and is established at ZGA [5], whereas in zebrafish, the 3D structure is already present by 2.25 hpf and disappears during the main wave of ZGA [48]. Nevertheless, early selective action of DNA and chromatin modifications later sustains tightly regulated developmental programmes in zebrafish [14,19,56,57]. Therefore, early plasticity in the zebrafish chromatin landscape conforms to the ‘hourglass’ model of development, where early diversity in regulatory mechanisms converge to conserved and constricted bottlenecks of gene regulation [81].

Modelling epigenetic developmental disorders

High-throughput sequencing technologies to understand the molecular basis of developmental disorders and cancer are now common in use, and identification of disease-associated mutations in chromatin factors is on the rise. One major challenge is to understand how mutations in genes encoding chromatin or ‘epigenetic’ factors, which are seemingly non-specific and usually ubiquitously expressed, could give rise to such specific phenotypes. In both development and cancer, chromatin mutations interact with, and influence the outcomes of, signalling pathways. Understanding how chromatin mutations operate in a biologically relevant context relies on valuable information from tractable model systems. The zebrafish model is proving its value in this work, which reinforces the importance of zebrafish chromatin research to the field. For example, recent work using zebrafish was instrumental in uncovering precise function and regulation of the TET enzymes that demethylate DNA [82]. Cornerstone experiments providing functional data to explain the effects of human mutations (or variants) in gene regulatory components are often provided by zebrafish (see [8386] for recent examples).

Can epigenetic marks be trans-generationally inherited and how do they respond to the environment?

A future challenge is understanding not only how chromatin influences development, but how chromatin modifiers respond to the environment, and if functional modifications can be transmitted between generations (reviewed in [87]). Zebrafish are emerging as a potent model to explore these questions. Interfering with chromatin structure during development certainly has lasting effects in zebrafish, for example, inhibition of DNA methylation using 5-aza-2′-deoxycytidine (5-AC) during early development feminised zebrafish and permanently altered the adult female gonadal transcriptome [88]. While altering chromatin status can affect the life course of the animal, the evidence is also emerging that forced alterations to chromatin can be inherited by subsequent zebrafish generations. Chemically induced DNA methylation changes (using 5-AC and MEHP — see Abbreviations) at two specific loci were apparent in first and second generation larvae of treated parents [89]. Finally, zebrafish can be used to show that commonly used therapeutics can elicit intergenerational effects. For example, Vera-Chang et al. showed that the commonly used SSRI (selective serotonin reuptake inhibitor), fluoxetine, suppresses cortisol levels for at least three zebrafish generations after F0 fish were exposed to physiologically relevant levels during the first 6 days of life, with behavioural consequences [90]. Genes encoding DNA methyltransferases and histone deacetylases were among those genes that were differentially expressed between parental and affected generations, implying an epigenetic mechanism of inheritance.

Perspectives
  • Zebrafish have recently emerged as a powerful model to understand the contribution of chromatin to genetic control of early development, patterning and environmental response.

  • There are similarities and differences in chromatin-controlled transcriptional mechanisms between zebrafish, mammalian models and humans. The diversity of mechanisms can teach us just as much about the governing principles of development as can the similarities, highlighting the potential of zebrafish models to further explore this growing field.

  • Going forward, the emergence of new technologies, such as single-cell sequencing [91], and resources, such as the DanioCode database (https://danio-code.zfin.org/), will enhance the contribution of zebrafish as a model to understand how chromatin structure influences development.

Abbreviations

     
  • 3D

    three-dimensional

  •  
  • 4C-seq

    circular chromosome conformation capture followed by sequencing

  •  
  • 5-AC

    5-aza-2′-deoxycytidine

  •  
  • ATAC-seq

    assay for transposase-accessible chromatin using sequencing

  •  
  • ChIP-seq

    chromatin immunoprecipitation followed by sequencing

  •  
  • DMR

    differentially methylated region

  •  
  • FANS

    fluorescent activated nuclei sorting

  •  
  • HAT

    histone acetyl transferase

  •  
  • HSC

    haematopoietic stem cell

  •  
  • Maternal-zygotic mutant

    product of a cross where the germline giving rise to the embryo is mutant for a given gene, as well as the zygotic genome of that embryo. This means that no maternal product from the mutant gene can be transmitted to the zygote.

  •  
  • MBT

    mid-blastula transition

  •  
  • MeDIP

    methylation-dependent DNA immunoprecipitation followed by sequencing

  •  
  • MEHP

    mono(2-ethylhexyl) phthalate

  •  
  • MRE-seq

    methyl-sensitive restriction enzyme digestion followed by sequencing

  •  
  • MZT

    maternal-to-zygotic transition

  •  
  • PLM

    posterior lateral mesoderm

  •  
  • SE

    super enhancer

  •  
  • TAD

    topologically associated domain

  •  
  • ZGA

    zygotic genome activation

Acknowledgements

The author is grateful to Drs Judith Marsman and Jisha Antony for comments on the manuscript and to the Royal Society of New Zealand Marsden Fund [#16-UoO-072], the Health Research Council of New Zealand [#15/229 and #15/623] and the Maurice Wilkins Centre for Molecular Biodiscovery for research funding.

Competing Interests

The Author declares that there are no competing interests associated with this manuscript.

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