DNA methylation is a critical epigenetic regulator of development in mammals and social insects, but its significance in development outside these groups is not understood. Here we investigated the genome-wide dynamics of DNA methylation in a mollusc model, the oyster Crassostrea gigas, from the egg to the completion of organogenesis. Large-scale methylation maps reveal that the oyster genome displays a succession of methylated and non methylated regions, which persist throughout development. Differentially methylated regions (DMRs) are strongly regulated during cleavage and metamorphosis. The distribution and levels of methylated DNA within genomic features (exons, introns, promoters, repeats and transposons) show different developmental lansdscapes marked by a strong increase in the methylation of exons against introns after metamorphosis. Kinetics of methylation in gene-bodies correlate to their transcription regulation and to distinct functional gene clusters, and DMRs at cleavage and metamorphosis bear the genes functionally related to these steps, respectively. This study shows that DNA methylome dynamics underlie development through transcription regulation in the oyster, a lophotrochozoan species. To our knowledge, this is the first demonstration of such epigenetic regulation outside vertebrates and ecdysozoan models, bringing new insights into the evolution and the epigenetic regulation of developmental processes.
The methylation of DNA is a prevalent epigenetic mark that is deeply rooted in evolution and found from bacteria to mammals. Despite that metazoan organisms display methylation on cytosines, great variations exist in the amount and distribution of methylcytosines (meCs) across taxa. DNA methylation is an essential feature of mammalian development because meC patterns are associated with a wide range of cell processes whose subtle combination is required for the embryo to develop into a complex adult organism exhibiting differentiated cell types. In mammals, ca. 60 to 80% of CpG cytosines are methylated and exhibit mostly stable patterns across tissues. CpG rich regions (CpG islands) are prevalent at transcription start sites, and the methylation of promoters correlates to gene silencing during development [1]. CpG dinucleotides are overrepresented in promoters of development and housekeeping genes which are protected from methylation by transcription factor binding and subsequent DNA methyltransferase exclusion [2], reflecting poor methylation in the germline over evolutionary time. However, DNA methylation can be highly dynamic at precise locations during development, as illustrated by the demethylation wave observed in parental pronuclei, the epigenetic reprogramming of the germline or the differences between the epigenomes of germ and somatic cells [1,3]. Consistently, DNA methylation shapes cell differentiation (reviewed in [4]) notably through silencing of pluripotency factors [5,6] and of germline specific genes in somatic cells [7] at lineage commitment by de novo methylation. DNA methylation is also implicated in genome defence against transposable element activity [8], maintenance of parental imprints [9,10], and X chromosome inactivation (review in [11]). Developmental processes are not only triggered by DNA methylation, whose causal role remains debated [12,13], but by networks of epigenetic regulators including histone modifiers [14], non coding RNAs [15], transcription factors [16] and DNA methyltransferases [17,18]. DNA methylation stabilizes the chromatin context underlying cell fate decisions that are propagated through cell generations by maintenance of the meC landscapes (review in [4]). In invertebrates, DNA is much less methylated and meCs are not evenly distributed but exhibit mosaic patterns [19,20]. DNA methylation in insect models is rare and mostly confined to gene bodies (gene body methylation, GBM) [20]. In hymenopterans, GBM controls exon selection[21] and governs important developmental outcomes such as caste differentiation in the honeybee [22,23] and in ants [24,25], as well as developmental gene expression in the wasp N. vitripennis [26,27]. However, DNA methylation and its developmental significance seem essentially restricted to a peculiar evolutionary acquisition in hymenopterans. Indeed, in Drosophila, early genes are controlled by cis-regulatory elements, non-coding lnc- and miRNAs, and transcription factors including polycomb and trithorax complexes (review in [28]) but not by DNA methylation. The actual presence and function of meCs in the fruitfly genome have been under discussion, and the nematode C. elegans even lacks conserved DNA methylation machinery. Therefore DNA methylation is considered absent in the ecdysozoan common ancestor, in line with animal genomes evolving towards an overall loss of DNA methylation in protostomes [20] such as insects, compared to deuterostomes [29] such as mammals. As a consequence of this basic divergence between ‘methylated vertebrates’ and ‘unmethylated invertebrates’, and in spite of the tremendous variability of organisms and life traits within protostomes, DNA methylation is largely neglected outside insects. However, recent studies in lophotrochozoans (that include molluscs and annelids), the sister group of ecdysozoans (that include insects and nematodes), suggest a more complex situation. Although meCs similarly exhibit a mosaic distribution, mollusc genomes are far more methylated than insects’, where methylated genomes display ca. 0. 15% of meCs [30], whereas this value reaches ca. 2% in the snail Biomphalaria glabrata [31] and in the gills [32] and mantle [33] of the oyster Crassostrea gigas. In this bivalve of greatest ecological and economical importance, GBM is predominantand associated to mRNA content [32,34]. Surprisingly, exposure to a DNMT inhibitor disrupts the oyster embryogenesis [35], and meCs are present in the promoter of some development genes with a direct influence on their expression [36]. These data point to developmental significance for DNA methylation in a lophotrochozoan species [37], challenging the current view on the evolution of epigenetic regulation of developmental processes. Here, to shed light on this point, we provide the first characterisation, to our knowledge, of genome-wide DNA methylation dynamics covering the development of a lophotrochozoan species. Using a development stage-wise MeDIP-seq approach, we characterized the methylome dynamics from the egg to the completion of organogenesis in the oyster C. gigas. Epigenetic landscapes were analysed at both a global, physical and more local, feature-related scales, together with mRNA expression and functional annotation, and indicate a dynamic regulation of DNA methylation at critical developmental steps.
The methylated DNA immunoprecipitation followed by high throughput sequencing (MeDIP-seq) approach enabled genome-wide assessment of methylation and its variations during oyster development. Large-scale genome-wide methylome dynamics were investigated by analyses of differentially methylated regions (DMRs) and physical maps. DMRs highlighted 4 main developmental phases (oocytes, 2–8 cells, mid-larval, spat) separated by 3 main developmental steps: cleavage (C step), gastrulation and organogenesis (I step, intermediate) and metamorphosis (M step), respectively (S1A Fig). The morula, blastula, gastrula, trochophore and D-larva stages were grouped into an intermediate mid-larval stage, because DMRs and individual feature methylation profiles (see below) showed only minor differences. Physical methylation maps of genomic scaffolds confirm the mosaic characteristic of oyster DNA methylomes, which display a succession of methylated and non methylated regions bearing gene clusters of variable length with no obvious organisation or relationship to CpG content (Fig 1). Developmental methylation dynamics mostly affect regions that were already methylated in oocytes, with hypermethylation prevailing during development (Fig 1). Regions not methylated in oocytes mostly remain unmethylated and only little de novo methylation of previously unmethylated regions is observed that lie almost exclusively between two adjacent previously methylated regions or at their direct proximity (Fig 1). Indeed, 98. 7% of the genes that are methylated at the spat stage were already methylated in oocytes. The mean distance from genomic features to the nearest DMR was drastically shorter at the M step (ca. 5 kb for CDS to 20 kb for TE) than at other steps (ca. 100 to 200 kb at C and I), indicating that DNA methylation is more evenly regulated throughout the genome at the M step than in the C step (S1B Fig). Although DMR length did not exhibit marked variations, they were not equally distributed regarding genome features amongdevelopment steps. Many more DMRs were found at the C (n = 1043) and M (n = 2230) steps than at the I step (n = 14), with methylation being preferentially regulated in exons (CDS), repeats (REP) and transposable elements (TEs) (S1C Fig), and in class I TEs (i. e. retrotransposons) compared to class II TEs (i. e. DNA transposons) (Pearson’s χ2: p<0. 0001***; C step: 122 DMRs in class I TEs vs. 81 in class II (60. 1% vs. 39. 9%); M step: 121 in class I TEs vs. 63 in class II (65. 7% vs. 34. 2%); genome, 64150 class I TEs vs. 21263 class II (75. 1 vs. 24. 9%) ). In parallel to genome-scale investigations, methylation landscapes were examined at the level of individual genomic features (i. e. exons (CDS), introns (INT), promoters (PRO), repeats (REP) and transposable elements (TE) ). Most of the reads (81. 5 ± 0. 95%) mapped to the considered features and a great majority (ca. 90%) of methylation was found within gene bodies (CDS and INT, S4A Fig). Overall, the distribution of methylation depends on the development stage (Pearson’s χ2: p<2. 10−16***, S2A Fig). The relative methylation of CDS is strongly increased after metamorphosis at the expense of the other features, especially introns (correlation between methylation in CDS and INT: Pearson R = -0. 985, p<0. 0001***), but not TEs (S2A Fig). This is because the intermediately methylated genes (ca. 2 to 6 log counts per million (CPM) ) have their CDS markedly hypermethylated at the spat stage (Fig 2). The individual methylation level of an important number of TEs increases in 2–8 cell embryos, and the stability observed thereafter is due to compensation between individual TE hypermethylation and hypomethylation in spats (Fig 2). The biological coefficient of variation (BCV) analyses of feature methylation between development stages clearly discriminate the 2–8 cells and spat stages from one another and apart from the other stages, which are grouped and may display a gradual distribution regarding embryogenesis chronology (S4B Fig). The methylation profiles depend on the feature considered (S2C Fig) thereby confirming both the feature- and development stage-specificity of the dynamics of oyster DNA methylomes, especially marked at the 2–8 cells (cleavage) and spat (post metamorphosis) stages. DMR proximity is associated to gene expression variability, and whether the DMR lies upstream or downstream has no influence (S3A Fig). Consistently, DMR-associated genes have their expression level more regulated than genes not associated to a DMR at each developmental step, although DMR and mRNA level variations were not correlated (S3B Fig). Compared to moderate changes, extreme methylation variations tend to hinder mRNA level regulation (S3B Fig). At a finer scale, most genes display a detectable methylation (20704 methylated genes vs. 7197 non methylated genes) during oyster development. The non methylated genes are mostly silent whereas the methylated genes are dramatically more expressed (Fig 3A). These genes have their mRNA level positively associated to their CDS methylation level, with a slight drop for genes within the 10th expression decile. Conversely, the methylation level decreases with expression variability (Fig 3A). These results indicate that methylation marks highly and stably expressed genes. Although the exact localisation of methylcytosines is hampered by the resolution of MeDIPseq (ca. 250 bp), gene expression decreases with the hypermethylation of the INT or CDS feature over the other (ANOVA p<0. 001), and the expression variability is correlated to the methylation pattern variability (p<2. 10−16***) suggesting an optimal in-gene methylation pattern for maximum transcription (S4 Fig). A large set (26%) of oyster genes exhibits a dynamic CDS methylation during development (ANOVA across stages p<0. 01**), and gene clusters can be discriminated based on their distinct CDS methylation kinetics (Fig 3B). However, methylation and mRNA level kinetics are correlated for only ca. 10% of these genes (r2>0, p<0. 05*) (Fig 3B). All together, these results indicate that during oyster development, unmethylated DNA is associated to transcription repression whereas methylated DNA corresponds to gene expression, with the dynamics of gene-body methylation being associated with transcription regulation. There was no association between PRO methylation and gene expression. The methylation level or coefficient of variation of gene features (PRO, CDS or INT) was not found to be correlated to the number of transcript variants during oyster development. Gene ontology (GO) annotation of DMRs depends on the development step considered. Cleavage DMRs bear genes with functional annotation (Biological Process ontology) related to egg vitellogenic resource consumption, mRNA metabolism and nuclear genome processes, whereas metamorphosis DMR genes are enriched in terms related to transport within the cell and protein degradation (Table 1). The gene clusters based on methylation kinetics exhibit specific GO term distribution in ontologies (Biological Process, Molecular Function and Cell Component) and little GO terms in common (S5A Fig). Conversely, selected Biological Process ontology terms display specific methylation level and developmental dynamics (S5B Fig), indicating that methylation dynamics correspond to the distinct functional pathways related to specific steps of oyster development.
The present work constitutes, to our knowledge, the first description of genome-wide methylome dynamics in a lophotrochozoan model. Dynamics of DNA methylation in gene bodies are associated with transcriptional regulation, and the control of transposable elements may imply DNA methylation. The shifts in methyl DNA profiles and their functional outcomes are prevalent at cleavage and metamorphosis, and suggest the importance of inherited methylomes. These results demonstrate that DNA methylation dynamics underlie Crassostrea gigas development. The developmental significance of gene body methylation in the oyster brings new insights into the epigenetic regulation of developmental processes and its evolution.
Oyster embryos were obtained as previously described [50]. Wild individuals were collected in Marennes- Oléron, France in August 2008 then transferred in mesh bags in February 2009 to Paimpol (northern Brittany, France, 48°48’ 24. 49”N, 3° 0’ 22. 84”W) until February 2010 and then to the Ifremer grow-out farm located at Aber-Benoît (northern Brittany, France, 48° 34’ 29. 976”N, 4°36’ 18. 378”W). These animals were exposed to disease during the spring of 2009 and suffered ca. 75% mortality. In April 2010 (Experiment 1 and 2) and February 2011 (Experiment 3), 60 individuals were transferred to the Ifremer marine station located at Argenton (Brittany, France, 48° 31’ 16. 320”N, 4°46’ 01. 998”W) for broodstock conditioning (6 weeks in 500 L flow-through tanks with UV-treated and 1 μm filtered seawater (TSW) at 19°C, enriched with a 1: 1 in dry weight mixture of Isochrysis affinis galbana and Chaetoceros gracilis corresponding to a daily diet of a ration equivalent to 6% of the oyster dry weight). Diploidy of oysters was confirmed by flow cytometry of gill cells from randomly sampled animals as previously described [51]. Gametes from mature specimen (13♂ and 27♀, 10♂ and 24♀, 13♂ and 21♀, Experiment 1,2 and 3 respectively) were obtained by stripping and filtered on a 100 μm mesh for the removal of large debris. For females, oocytes were harvested as the remaining fraction on a 30 μm mesh; for males, spermatozoa were harvested as the passing fraction through a 30 μm mesh. Oocytes were pre-incubated in TSW then mixed in a 5 L jar at 50–100 spermatozoids per oocyte (22 November 2010,5 January 2011 and 12 April 2011 for experiment 1,2 and 3 respectively). The embryonic development was completed in TSW in oxygenated 150 L tanks at 21°C for 48 h. The D-larvae were then collected and reared in flow- through rearing systems at 25°C. At the end of the pelagic phase (16 d), competent larvae were collected on a 225 μm sieve and allowed to settle on cultch. Post-larvae were maintained in downwelling systems where they were continuously supplied with enriched seawater. After 10 d, the spat were collected on 400 μm mesh. In the larval and post-larval stages, the oysters were fed the same diet as the broodstock. Throughout this time, the oysters were free of any abnormal mortality and OsHV-1 virus. Embryos were left unattended until sampling, i. e. before fertilization for control oocytes, and ca. 1 hour post-fertilization (hpf) for 2–8 cells stage, ca. 3 hpf for morulae, ca. 6 hpf for blastulae, ca. 9 hpf for gastrulae, ca. 16 hpf for trochophore larvae, and ca. 24 hpf for D larvae. Spat was collected at 26 days, after settlement and metamorphosis. Developmental stages were assayed by microscopic observation based on morphological and motility criteria before and after fixation using 70% ethanol. Samples were split in aliquots of 2 million larvae, stored dry at -80°C and thawed only once before use. Each development stage was sampled from three distinct fertilization experiments (experiments 1,2 and 3). Genomic DNA from ca. 2 million larvae per sample was purified by affinity chromatography (Macherey Nagel) following the manufacturer’s instructions. Degradation of contaminating RNA was realized using RNAse. DNA purity and concentration were assayed by spectrometry (Nanodrop, Thermo) and on-chip gel electrophoresis (Tape Station 2200, Agilent). DNA was sheared in ca. 250 bp fragments using a Covaris S2 sonicator (duty cycle: 10%, intensity: 4, cycles: 200, time: 80s). Twenty samples (n = 2 to 3 biological replicates per development stages) were processed for MeDIP-seq library preparation following the protocol of Taiwo et al. [52]. Briefly, 5μg DNA from each sample were used for DNA end-repair and dA-tailing (NEBNext reagents, New England Biolabs). Immunoprecipitation of methylated DNA (MeDIP) was realised on 1 μg DNA after end-repair, dA-tailing and purification using the MagMeDIP kit (Diagenode) using the manufacturer’s instructions. All DNA purifications were carried out using Ampure XP magnetic beads (Beckman Coulter) according to the recommended procedure. Ten randomly chosen samples were assayed following the manufacturer’s recommendations for MeDIP specificity with a mean value of 96. 9%. Immunoprecipitated DNA was then amplified using Phusion DNA polymerase (New England Biolabs) and purified. After size selection and quality control, libraries were submitted to 2x76 bp paired-end sequencing using a GAIIx sequence analyzer (Illumina). This strategy produced ca. 80 million paired end reads i. e. 3. 6±0. 25x106 reads per sample among which 80. 7±0. 1% were aligned to the genome, giving a 24. 8±3. 3-fold mean coverage. Data were analysed using a using a combination of dedicated R (bioconductor. org) and bash script as well as in-house R, bash, TiCL and PERL scripts. Data source files (NCBI project PRJNA324546) and scripts used for analyses are publicly available (github. com/BOREA-UNICAEN/MeDIPSeq-Dev-Gigas). Primary analysis was performed with RTA (Illumina) with default parameters and reads were demultiplexed using CASAVA v. 1. 8. Bases with a QC>30 were retained for further analyses. Paired-end reads were mapped to the oyster genome (assembly v. 9) using BWA with default parameters and pair-sorted. Paired reads mapping to the following genomic features: exons (CDS), introns (INT), promoters (PRO), repeats (REP) and transposable elements (TE) [34] were counted using HTseq-count [53]. Only promoter sequences longer than 100 bp were retained for further analyses and ambiguous read pairs were discarded.
Elucidating the mechanisms which govern the development of multicellular animals and their evolution is a fundamental task. Epigenetic mechanisms like DNA methylation have recently emerged as critical regulators of mammalian development through the control of genes that determine the identity of cells and the transmission of parental imprints. In invertebrates however, DNA is mostly unmethylated and does not play a role in development except in the peculiar case of social insects. Therefore the significance of DNA methylation in development is thought to be restricted to vertebrates, and thereby considered a recent evolutionary acquisition, and the situation in more distant organisms is unknown. Here we investigated the dynamics of genome-wide DNA methylation patterns in a mollusc, the oyster C. gigas, throughout its development. We found that the dynamics of DNA methylation correspond to the expression dynamics of distinct functional gene clusters that control two critical development steps, cleavage and metamorphosis, and we provide insights into the underlying molecular mechanisms in a non-vertebrate species. These findings challenge the present considerations on the evolution of developmental processes and their epigenetic regulation, and open a new area of research in molecular and developmental biology in invertebrates.
invertebrates
gene regulation
animals
invertebrate genomics
dna transcription
developmental biology
epigenetics
dna
dna methylation
chromatin
chromosome biology
gene expression
oysters
molluscs
chromatin modification
dna modification
animal genomics
biochemistry
bivalves
cell biology
nucleic acids
genetics
biology and life sciences
metamorphosis
genomics
organisms
Dynamics of DNA methylomes underlie oyster development