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. Author manuscript; available in PMC: 2023 Dec 1.
Published in final edited form as: Dev Growth Differ. 2022 Oct 8;64(9):508–516. doi: 10.1111/dgd.12813

Epigenomic dynamics of early Xenopus embryos

Jeff Jiajing Zhou 1, Ken WY Cho 1,2
PMCID: PMC10550391  NIHMSID: NIHMS1839018  PMID: 36168140

Abstract

Comprehending how the embryonic genome regulates accessibility to transcription factors is one of the major questions in understanding the spatial and temporal dynamics of gene expression during embryogenesis. Epigenomic analyses of embryonic chromatin provide molecular insights into cell-specific gene activities and genomic architectures. In recent years, significant advances have been made to elucidate dynamic changes behind the zygotic genome activation among various model organisms. Here we provide an overview of the current epigenomic studies pertaining to early Xenopus development.

Introduction:

Proper embryonic development requires an extensive reorganization of the zygotic epigenome, which includes a critical transition from transcriptionally silent joined gametes to a transcriptionally active zygote (Eckersley-Maslin et al., 2018). In Xenopus, zygotic genome activation (ZGA) occurs in two phases: minor ZGA and major ZGA. The minor ZGA occurs during the cleavage stage, and results in a low level of transcription. The minor ZGA is required for the major ZGA (Yang et al., 2002; Rosa et al., 2009; Skirkanich et al., 2011), which occurs at the blastula stage (N/F st8). Around the time of ZGA, the embryonic epigenome is established by maternally inherited components stored in the egg. Presumably, maternal transcription factors (TFs) work together with epigenetic modifiers to establish the initial pluripotent state of the embryonic epigenome. The zygotic genome undergoes continuous modifications to activate robust zygotic developmental programs and maintain the integrity of cell fates in response to diverse developmental cues (Schulz and Harrison, 2018). Significant efforts have been made to uncover the dynamic relationship between epigenetic regulation and gene expression during the earliest cell fate decision process in Xenopus (Figure 1). Here we will focus on the initial establishment of embryonic epigenome during germ layer specification in Xenopus.

Figure 1: Characteristics of the epigenome during early Xenopus development.

Figure 1:

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The patterns of different epigenetic signatures emerge around ZGA are shown in a time window from the zygote to the gastrula stage. Blue dashed lines denote minor/major waves of ZGA.

1. Higher-order chromatin structures

The spatial organization of chromatin imposes a critical influence on genome functions. During ZGA, higher-order chromatin structures undergo extensive reorganization. Chromosome conformation capture-based methods reveal the presence of distinct folding configurations within chromatin (Lieberman-Aiden et al., 2009; Pombo and Dillon, 2015). Boundary element binding proteins such as CCCTC-binding factor (CTCF) work together with the cohesin complex to drive the formation of chromatin loops and Topologically Associated Domains (TADs) (Ong and Corces, 2014). TADs are proposed to function as structural scaffolds for regulatory landscape and to facilitate interactions between cis-regulatory modules (CRMs) and their target promoters (Kadauke and Blobel, 2009).

In Xenopus, the notable structural organization of chromatin is not detected at the mid-blastula (st8) stage (Niu et al., 2021). Weak TAD-like structures emerge at the late blastula (st9) stage, suggesting that TAD formation coincides with the occurrence of major ZGA. Chromatin loops between TAD borders and chromatin compartments are continuously consolidated from the gastrula and onward. Inhibition of transcription does not interfere with the formation of TADs (Niu et al., 2021), suggesting that the establishment of early TADs is driven by maternal components. Depletion of CTCF and Rad21, factors involved in TAD regulation and loop structure formation (Heidari et al., 2014), results in compromised TAD formation with weakened TAD borders and attenuated chromatin loop interactions within TADs (Niu et al., 2021). In general, despite strong effects of CTCF and Rad21 knockdowns on TADs formation, the effect on gene expression is moderate (Khoury et al., 2020), indicating that TADs are important for the formation of chromatin architecture, but the expression of genes is strongly dictated by local interactions of nearby CRMs and promoters (Ing-Simmons et al., 2021; Espinola et al., 2021; Kubo et al., 2021; Franke et al., 2021).

The mechanism controlling TAD formation is context-dependent and regulated differently across species. In fruit flies, frogs, mice, and humans, TADs initially form at or shortly after major ZGA, and are consolidated onward (Sexton et al., 2012; Ogiyama et al., 2018; Niu et al., 2021; Ke et al., 2017; Du et al., 2017; Chen X et al., 2019). In contrast, zebrafish chromatin is highly structured during pre-ZGA, and undergoes a drastic loss in structure at ZGA, then re-establishes the TAD organization post-ZGA (Kaaij et al., 2018). The timing difference in TAD formation between zebrafish and other animals is currently unclear. ZGA is dispensable for early embryonic TAD formation in fruit flies, frogs, and mice (Hug et al., 2017; Niu et al., 2021; Ke et al., 2017; Du et al., 2017). However, ZGA is required for TAD establishment in human embryos (Chen X et al., 2019). These species-specific differences in TAD formation suggest that embryonic 3D genome architecture is controlled by both shared (utilizing CTCF and Rad21) and species-specific mechanisms.

2. Nucleosomes and histone variants

Eukaryotic genome organization requires concise nucleosome positioning, which determines chromatin accessibility for TFs and general transcriptional machinery (Tsompana and Buck, 2014). Accessible DNA regions such as promoters and CRMs for transcriptional regulators are nuclease hypersensitive due to nucleosome destabilization or eviction (Gross and Garrard, 1988;Henikoff, 2008). DNase-seq and ATAC-seq have been used to detect accessibility of DNA regulatory regions of the early Xenopus embryonic genome (Cho et al., 2019, Gentsch et al., 2019, Esmaeili et al., 2020, Bright et al., 2021). Pluripotent factors PouV and Sox3 exhibit a pioneering activity to open chromatin around TF-bound regions during Xenopus ZGA (Gentsch et al., 2019). Overall, embryonic chromatin gradually gains accessibility from late blastula (st9), as revealed by the appearance of co-activator histone acetyltransferase p300 (EP300) binding and active H3K4me3 histone modification (Bright et al., 2021). The sia1 and nodal3.1 genes are some of the earliest genes activated in the dorsal region of gastrulae. Their regulatory regions lose chromatin accessibility, and consequently lose cellular competence toward Wnt signals as gastrulation proceeds (Esmaeili et al., 2020). This finding is consistent with the observation that the expression of organizer expressed sia1 and nodal3.1 occurs shortly after ZGA, but quickly diminishes during gastrulation. The dynamic changes in accessibility of zygotic chromatin observed in Xenopus are generally consistent with the findings in zebrafish and mice (Liu et al., 2018; Lu et al., 2016).

Dynamic incorporation of histone variants into nucleosomes modulates accessibility of underlying DNA to transcriptional regulators (Venkatesh and Workman, 2015). H3.3, the replacement variant of H3, is highly enriched around active transcriptional regions (Ahmad and Henikoff, 2002; Hake et al., 2006), and has been implicated in epigenetic regulation during early development. Mechanistically, phosphorylation of serine 31 in H3.3 promotes acetylation and represses methylation of histone lysine 27 in cis (Sitbon et al., 2020). A nuclear transplant experiment using Xenopus somatic nuclei showed that the epigenetic memory of somatic cells correlates with the association of histone H3.3 (Ng and Gurdon, 2008). The myod promoter maintains its differentiated cell state memory in non-muscle cell lineages of nuclear transplant embryos when it is associated with H3.3, and consequently, myod is abnormally expressed in non-muscle cells. However, the somatic memory is lost when it lacks H3.3 association. Interestingly, the myod promoter eliminates the memory when it is associated with histone H3.3 mutated at K4 lysine residue, indicating a requirement of H3.3 K4 for epigenetic memory. H3.3 is also required for proper gastrulation by activating mesodermal and neuroectodermal genes and maintaining cell viability (Szenker et al., 2012). These observations suggest a role for H3.3 in forming a permissive chromatin state for transcription.

Unlike H3.3, the linker histone H1 is not assembled into the core histone octamer. The linker histone (H1 and its variants) binds to the DNA entry/exit point of the nucleosome core through its globular domain at linker DNA regions (Fyodorov et al., 2017). Maternally supplied linker histone variant B4 (also known as H1M) shares 30% identity with H1 (Smith et al., 1988), and functions like H1 (Dimitrov et al., 1993). The expression level of zygotic H1 gradually increases to reach the level of maternal B4 by the blastula stage, and vastly exceeds by the neurula stage, whereas the level of maternal B4 drastically diminishes after the blastula stage. Histone B4 is less basic than H1, and has a reduced binding affinity to nucleosomes than H1, thus promoting a permissive chromatin state for transcription (Ura et al., 1996). These studies show that differential incorporation of histone variants during early embryogenesis shapes chromatin states of the embryonic genome.

3. Post-translational modifications of histones

Approximately 25–30% of the mass of individual histones is derived from the histone tails, which protrude from the surface of the chromatin polymer (Wolffe and Hayes, 1999). The histone tails frequently gain post-translational modifications including acetylation, methylation, phosphorylation and ubiquitination. The patterns of covalent histone modifications, referred to as the ‘histone code’, significantly impact overall chromatin structures, and are interpreted by ‘reader’ proteins to carry out diverse cellular functions (Strahl and Allis, 2000). In Xenopus embryos, the major epigenetic regulations via histone modifications occur around ZGA and affect both temporal and spatial transcriptomes (Akkers et al., 2009; Gupta et al., 2014; van Heeringen et al., 2014; Hontelez et al., 2015).

3.1. Histone acetylation

Histone acetylation occurs on the ε-amino group of the lysine residues within N-terminal tails of all four core histones H2A, H2B, H3 and H4 (Inoue and Fujimoto, 1969; Seto and Yoshida, 2014). Histone acetylation is often indicative of permissive chromatin states (thus associated with gene activation) because the acyl groups neutralize the positive charge on lysine residues, thereby reducing the affinity of histones to DNA (Wang et al., 2000; Anderson et al., 2001). The acetylation status of histone is determined by equilibrated activities between histone acetyltransferases (HATs, e.g. Ep300) (Lee and Workman, 2007) and histone deacetylases (HDACs, e.g. Hdac1) (Seto and Yoshida, 2014). Current genomic analyses of Xenopus blastula stage embryos show that the embryonic genome is devoid of major histone acetylation marks such as H3K27ac (Akkers et al., 2009). However, H3K27ac modification appears during the late blastula stage and is widespread in the genome at gastrula stages (Gupta et al., 2014). This finding is also supported by mass spectrometry analysis of histone modifications present in Xenopus embryos (Schneider et al., 2011). Interestingly, ChIP-qPCR analysis shows that H3K9/14ac modifications are detected around the promoters of sia1 and nodal3.1 during the early blastula stage (Blythe et al., 2010), suggesting a possibility that selected promoters are subject to specific histone acetylation modifications around or prior to the blastula stage. Despite the fact that some genes are marked by H3K9/14ac, histone acetylation levels in pluripotent blastula cells are low, which is essential for developmental plasticity and competence (Rao and LaBonne, 2018). Consistent with the notion, histone acetyltransferase Ep300 progressively binds to zygotic genomic loci from blastula and onward (Hontelez et al., 2015). Similarly, histone deacetylases such as Hdac1 also occupy the zygotic genome around the same time, and establish germ-layer specific histone acetylomes during gastrulation (Zhou et al., 2022). These findings illustrate that the early Xenopus embryonic genome is naive, and that the establishment of histone acetylation occurs during blastula and onward, coinciding with the onset of major ZGA. Region-specific histone acetylation patterns are then generated via dynamic interactions between HATs and HDACs.

An increase in histone acetylation is a conserved feature of ZGA during animal development. Histone acetylation in fruit flies appears on a few zygotic genes during the minor wave of ZGA, but increases dramatically at the major wave (Li et al., 2014). In zebrafish, a small number of CRMs are H3K27ac marked by the blastula stage, and the number of H3K27ac marked CRMs increases significantly during gastrulation (Bogdanovic et al., 2012; Chan et al., 2019). In mice, H3K27ac mark increases genome-wide around the 2 cell-stage, correlating with the major wave of ZGA (Dahl et al., 2016). The correlation between the deposition of histone acetylation and the timing of ZGA begs the question of which enzymes regulating histone acetylation are recruited to selected zygotic genes, and form spatially and temporally restricted activities of histone acetylation during embryogenesis. We speculate that localized expression of TFs coordinates the selective recruitment of HATs and HDACs to target sites in the genome. Proteomic analyses of Ep300 and Hdac1 associated complexes as well as quantitative acetylome studies will be essential to address this question.

3.2. H3K4 methylation

Unlike histone acetylation, which primarily promotes gene activation, histone methylation can activate or repress transcription. CRMs marked with both H3K4me3 and H3K27ac are associated with active chromatin states, whereas CRMs decorated with H3K4me3 and H3K27me3 are associated with repressive chromatin states (Skvortsova et al., 2018). H3K4me3 catalysis is controlled by lysine methyltransferases KMT2G/F (also known as SETD1B/1A) and lysine demethylases KDM5A-D (also known as JARID1A-D) (Beacon et al., 2021). In Xenopus, deposition of H3K4me3 is observed at sia1 and nodal3.1 promoter regions before the onset of their transcription (Blythe et al., 2010). H3K4me3 is progressively deposited to the embryonic epigenome after ZGA (Hontelez et al., 2015). These observations suggest that the appearance of promoter H3K4me3 is tightly associated with the expression of zygotic genes. In Drosophila, genome-wide H3K4me3 marks appear shortly after ZGA (Chen et al., 2013; Li et al., 2014), whereas in zebrafish, H3K4me3 is present before the onset of ZGA (Vastenhouw et al., 2010; Zhang et al., 2014). In mice, the broad domains of H3K4me3 are detected in oocytes, but disappear in most of the regions in fertilized embryos except promoter regions of active genes at the 2-cell stage (Dahl et al., 2016; Zhang et al., 2016). This mechanism poises the selected promoters during ZGA. In humans, the oocyte genome is marked with H3K4me3 at gene promoters, and weak de novo deposition of H3K4me3 is observed in pre-ZGA embryos (Xia et al., 2019). These findings suggest that H3K4me3 deposition, while the onset may vary among different animal species, is a functionally conserved feature associated with major ZGA.

H3K4me1 is enriched at active and primed CRMs to fine-tune CRM activities and functions (Dorighi et al., 2017; Local et al., 2017; Rickels et al., 2017). The primary enzymes regulating H3K4me1 catalysis are KMT2C/D (also known as MLL3/4) and KDM1A/B (Herz et al., 2012; Beacon et al., 2021). The presence of H3K4me1, coupled with the absence of H3K4me3, distinguishes CRMs from proximal promoters (Heintzman et al., 2009). The coexistence of H3K27ac and H3K4me1 demarcates active CRMs from primed CRMs (Creyghton et al., 2010; Rada-Iglesias et al., 2011). Genome-wide detection of H3K4me1 occurs around the onset of major ZGA in Xenopus (Hontelez et al., 2015). H3K4me1 deposition on CRMs is controlled by both maternally and zygotically expressed TFs (Paraiso et al., 2021). Recent studies suggested that dense clusters of CRMs with high H3K4me1 signals act as super-enhancers (SEs), which underlie robust tissue-specific gene expression (Lovén et al., 2013; Whyte et al., 2013). Ectoderm, mesoderm and endoderm germ-layer specific SE formation occurs by the early gastrula stage (Paraiso et al., 2019, Paraiso et al., 2021). The formation of endodermal SEs was shown to be dependent on the presence of vegetally localized maternal TFs. In Drosophila, H3K4me1 signals are widely spread in the zygotic genome after ZGA, which flanks regions bound by TF Zelda (Li et al., 2014; Moshe and Kaplan, 2017). CRMs marked by H3K4me1 are poised for further activation or repression during tissue patterning (Koenecke et al., 2017). In zebrafish, H3K4me1 is detected in a few regions at blastula, but expands greatly afterward (Bogdanovic et al., 2012). These data are consistent with the view that H3K4me1 primes CRMs during ZGA for robust zygotic transcription. However, further work is needed to determine key factors in establishing and interpreting H3K4me1 patterns on CRMs.

3.3. Heterochromatic histone methylation

The formation of heterochromatin, a repressive chromatin state, requires distinct histone methylation marks. Histone H3K9 methylation, which is mediated by the interplay between KMT1 family proteins (also known as Su(var)39 family) and KDM3/4/7 (also known as JHDM2/JHDM3/PHF8), is essential for the formation of constitutive heterochromatin through the recruitment of Heterochromatin Protein 1 (HP1) (Hyun et al., 2017; Allshire and Madhani, 2017). In Xenopus, H3K9me2/3 and H4K20me3 marks are widespread in the genome from late-blastula and onward (Hontelez et al., 2015). Generally, H3K9 methylation marks repetitive DNA sequences to safeguard genome stability such as transposon silencing (Karimi et al. 2011). Indeed, retrotransposons are marked by H3K9me2/3 with H4K20me3, and a subset of the DNA transposons lose these histone methylation modifications during gastrulation (van Kruijsbergen et al., 2017). However, whether H3K9 methylation plays a role in developmental gene repression during Xenopus ZGA remains elusive.

Tri-methylation of histone H3K27 (H3K27me3) is catalyzed by Polycomb Repressive Complex 2 (PRC2) and removed by KDM6A/B (Pan et al., 2018). Clusters of genomic regions enriched with H3K27me3 can function as silencers by forming repressive intra-chromatin loops (Kundu et al., 2017; Cai et al., 2021). In Xenopus, the early landscape of H3K27me3 marks is instructed primarily by maternal factors (Hontelez et al., 2015). H3K27me3 enrichment on CRMs increases from blastula and onward, which coincides with the developmental period of increasing tissue complexity (Akkers et al., 2009). Presumably, the dynamic appearance of spatially and temporally defined H3K27me3 patterns are critical for generating defined spatial gene expression patterns.

The regulation of repressive H3K27me3 modification seems to diverge among different animal species. In nematode worms and fruit flies, H3K27me3 is maternally transmitted from oocytes to embryos, modulating the balance between H3K27ac and H3K27me3 around CRMs (Gaydos et al., 2014; Zenk et al., 2017). In zebrafish, CRMs are frequently marked by both H3K27me3 and H3K4me3 (Vastenhouw et al., 2010; Lindeman et al., 2011), similar to the bivalent chromatin domains identified in mouse embryonic stem cells and epiblast (Bernstein et al., 2006; Rugg-Gunn et al., 2010). While the function of bivalent domains has not been fully established, these bivalently marked domains tend to coincide with genes at low expression levels. However, such bivalent domains are largely absent in Drosophila and Xenopus embryos at ZGA (Li et al., 2014; Akkers et al., 2009). This discrepancy between mouse and zebrafish embryos vs. Drosophila and Xenopus embryos could be due to species-specific modes of gene regulation or caused by experimental differences in embryo collection stages. In mouse embryos, maternally inherited H3K27me3 marks persist at distal CRMs until blastocysts, whereas H3K27me3 marks are erased from paternal alleles upon fertilization and later redeposited after post-implantation (Zheng et al., 2016). In human embryos, repressive H3K27me3 marks are absent at ZGA due to the global erasure of H3K27me3 at both parental alleles (Xia et al., 2019). Taken together, H3K27me3 marks are dynamically regulated among different species.

4. DNA methylation

Methylation of the 5th carbon on cytosine is one of the most studied and conserved epigenetic modifications among fungi, plants, and animals (Feng et al., 2010). Transfer of DNA methylation patterns to the progeny cells ensures the integrity of cell states during development (Bird, 2002). DNA methylation patterns are often unique among different cell types, and their changes influence the expression of cell type specific genes (Schübeler, 2015). The deposition and maintenance of methylation on symmetrical CpG DNA are primarily controlled by DNA methyltransferases (Dnmts) (Li et al., 1992; Okano et al., 1999; Ramsahoye et al., 2000). Dnmts can directly interact with histone-modifying enzymes to confer a repressive state in the regulatory regions of genes (Moore et al., 2013), and affect the binding of TFs to DNA (Yin et al., 2017), suggesting that remodeling of DNA methylation patterns is critical in cellular differentiation and ZGA.

Genome-wide DNA methylome survey in early Xenopus embryos showed that promoters and gene bodies of actively transcribed genes are hypermethylated, but H3K4me3 enriched transcriptional start sites are hypomethylated (Veenstra and Wolffe, 2001; Bogdanovic et al., 2011). An early biochemical study demonstrated that in vitro methylated DNA is transcriptionally inactive when microinjected into Xenopus oocytes (Vardimon et al., 1982), suggesting a functional link between DNA methylation and gene repression. However, genome-wide DNA methylome patterns do not support the repressive role of DNA methylation during blastula and gastrula stages. The repressive H3K27me3 histone mark is found around the regions of the genome lacking DNA methylation, and is minimally present until stage 9 (after ZGA) (Akkers et al., 2009; Gupta et al., 2014; van Heeringen et al., 2014). In light of this finding, it is interesting to note that Xenopus embryos depleted of Dnmt1 express zygotic genes approximately two cell cycles earlier than wild type embryos (Stancheva and Meehan, 2000). This is consistent with the view that Dnmt1 functions as a repressor. Surprisingly, the ectopic expression of catalytically inactive human DNMT1 in Dnmt1 deficient Xenopus embryos abolishes the premature activation of ZGA (Dunican et al., 2008). This demonstrates that Dnmt1 catalytic activity is not essential for repression, implying that non-catalytic function of Dnmt1 controls the timing of ZGA and thus, the transcriptional competence of embryos. Further studies are needed to understand how DNA methylation influences transcriptional activities of the early embryonic genome.

Conclusions:

The rich history of Xenopus research, combined with the ability to obtain a large collection of synchronized embryos, to manipulate embryos for experimental procedures, and to microinject macromolecules into embryos for over- and under-expression studies are obvious experimental advantages for using Xenopus to uncover epigenetic mechanisms regulating Xenopus ZGA. Along with these advantages, modern single embryo proteomic approaches (Lombard-Banek et al., 2019; Saha-Shah et al., 2019) to determine TF and epigenetic modifier protein interactions, nascent transcriptomic methods (Chen H et al., 2019; Gentsch et al., 2019; Wissink et al., 2019; Chen and Good, 2022) to examine the onset of ZGA, chromatin conformation capture and more advanced chromatin capture methods (McCord et al., 2020; Niu et al., 2021) to identify DNA looping interactions will provide much needed tools to gather high resolution genomic data. Additional tools that will be useful in the future are high resolution imaging techniques such as expansion microscopy and super resolution microscopy (Wassie et al., 2019), to examine 3D architecture of chromatin. While the current work has uncovered major spatial and temporal events required for the proper assembly of chromatin signatures surrounding both active and inactive CRMs, there are still many unanswered questions. For instance, most of the enzymatic components involved in epigenetic modifications are maternally expressed, but their biological activities are stalled until later developmental stages. How is the assembly of functional histone methyl and acetyl transferases controlled? How do these enzymes target specific genomic regions and confer proper epigenetic modifications? Are maternal TFs required for the recruitment of enzyme complexes to the genome? If so, what maternal TFs are major players in the process? How does the change in chromatin architecture impact zygotic developmental programs? The Xenopus system will remain instrumental to address these fundamentally important biological questions.

Acknowledgments:

We thank current Cho lab member Clark L. Hendrickson for critical comments. We apologize for any oversights or omissions. This work is supported by NIH R01GM126395, R35GM139617 and NSF 1755214 to K.W.Y.C.

Footnotes

Declaration of interests:

The authors declare no competing interests.

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