Transposons are sequences of DNA with the ability to mobilize and jump from one position to another. In the human genome, the number of transposons far surpasses the number of genes. Furthermore, while transposons have been beneficial for the evolution of the human genome, they can also alter genes and cause cancer and genetic diseases.

The danger posed by transposons has led to numerous mechanisms to keep them under control. In particular, a natural biochemical modification of the DNA molecule called “DNA methylation” plays an important role in keeping transposons inactive or silent. However, during the early development of an embryo, the DNA methylation marks are erased throughout the entire genome. This provides an opportunity for transposons to be active, and it is not clear how the genome manages to control transposons in the absence of this essential protective mark.

Walter et al. have now investigated this process by using a cell type that mimics the loss of DNA methylation that occurs during embryonic development – mouse embryonic stem cells grown in the laboratory. The experiments revealed that when DNA methylation is lost progressively, the transposons are reactivated at first but are later put back into a silent mode by alternative mechanisms. These mechanisms compensate for the disappearance of DNA methylation by encouraging the DNA around transposons to become compacted, which prevents the transposons from moving.

Further analysis revealed that the different families of transposons that exist in the mouse genome can be classified into three groups, and in each group different proteins ensure the transposons remain repressed in the absence of DNA methylation. Together these findings reveal that multiple pathways cooperate to protect the genome against the activity of a variety of transposons. Finally, in mammals, DNA methylation is naturally erased both during the formation of sperm and egg cells and in the early embryo. As such, it will be important to verify whether the mechanisms discovered in the laboratory-grown cells also tame transposons during these critical developmental periods.

Millions of transposable elements reside in mammalian genomes, far surpassing in number the approximately 25000 protein-coding genes (Lander et al., 2001). Most of these elements are retrotransposons, which utilize an RNA intermediate to duplicate and mobilize. Through their activity or their mere presence, transposons can be both beneficial for the evolution of the host genome and deleterious for its integrity. They can modify gene functions through insertional mutagenesis, influence gene transcriptional outputs by acting as promoters or enhancers or induce chromosomal rearrangements through non-allelic recombination (Goodier and Kazazian, 2008). Accordingly, erratic transposon-related events have been linked to congenital diseases, cancer and infertility (Kaer and Speek, 2013).

Successive waves of transposon expansion and decline have shaped mammalian genomes over evolution, leading to a current occupancy rate of approximately half of the genomic space. Reflecting their various evolutionary origin and multiplication success, resident elements are greatly diverse in structures, numbers and functional properties, which define discrete families of transposons. Long Terminal Repeat (LTR) sequences characterize endogenous retroviruses (ERVs, 12% of the mouse genome), which can be further subdivided into three families (ERV1, ERVK and ERVL), according to the infectious retroviruses they derive from (Stocking and Kozak, 2008). Non-LTR elements comprise Long and Short INterspersed Elements (LINEs and SINEs, 20% and 8% of the genome, respectively), and also consist of specific sub-families (Babushok et al., 2007). The majority of transposons have accumulated nullifying mutations and truncations, but around 1–2% of LINEs and ERVs have intact sequences that embed the protein coding information necessary for their mobilization. Notably, ERVK elements show the greatest level of activity, which causes at least 10% of spontaneous mutations in laboratory mice (Maksakova et al., 2006).

To minimize their impact on genome fitness, multiple layers of control antagonize transposons at different steps of their life cycle (Zamudio and Bourc’his, 2010). Notably, restraining mechanisms can differ between cell types. In somatic cells and in the male differentiating germline, DNA methylation is the main transcriptional suppressor of LTR and non-LTR transposons. In these contexts, transposable elements are densely methylated (Rollins et al., 2006; Smith et al., 2012) and DNA hypomethylation leads to their de-repression (Bourc’his and Bestor, 2004; Walsh et al., 1998). In contrast, the early germline and the early embryo manage to globally control their transposon burden without DNA methylation. These cells naturally undergo genome-wide loss of DNA methylation, likely as part of the acquisition of a pluripotent, flexible state (Seisenberger et al., 2013). Moreover, genetic studies have demonstrated that mouse embryonic stem (ES) cells can use DNA methylation-independent mechanisms to silence transposons: knocking-out the three active DNA methyltransferases (Dnmt-tKO) does not yield significant de-repression of transposons, except Intracisternal A Particle (IAP) elements (Karimi et al., 2011b; Matsui et al., 2010)

In fact, transposon control in ES cells seems to rely primarily on post-translational histone methylation, notably at lysine 9 of histone H3 (H3K9). H3K9 dimethylation (H3K9me2), which is deposited by the EHMT2/G9a and EHMT1/GLP lysine methyltransferases, directly and specifically represses class L ERVs (Maksakova et al., 2013). H3K9 trimethylation (H3K9me3) can be catalyzed by the SETDB1 (also known as ESET) or the SUV39H enzymes. The SUV39H system targets H3K9me3 at evolutionary young LTR and non-LTR transposons, but Suv39h mutant ES cells principally up-regulate LINE1 elements (Bulut-Karslioglu et al., 2014). In parallel, SETDB1, together with its associated co-repressor, the Krüppel-associated box domain (KRAB)-Associated Protein 1 (TRIM28, also known as KAP1), mainly control H3K9me3-dependent suppression of ERVK transposons- a family to which IAP elements belong (Karimi et al., 2011b; Matsui et al., 2010; Rowe et al., 2010). TRIM28 is recruited to specific genomic sites via direct interactions with KRAB-zinc finger proteins (Friedman et al., 1996), which are a large family of DNA binding factors that co-evolved with ERVs (Emerson and Thomas, 2009). Therefore, different H3K9 methylation-based mechanisms are utilized to silence different transposons families in ES cells. In contrast, the repressive spectrum of polycomb-mediated H3 lysine 27 trimethylation (H3K27me3) is limited: only Murine Leukemia Virus (MuLV) elements are reactivated upon H3K27me3 deficiency (Leeb et al., 2010).

However, the prevailing view that H3K9 methylation acts as the main transposon controller in ES cells may be biased by two confounding factors. First, conclusions are based on analyses of chromatin modifier mutants, which still harbor high DNA methylation levels. Second, proper transposon repression in Dnmt-tKO ES cells may reflect a long-term adaptation to a DNA methylation-free state rather than a lack of significant role of DNA methylation per se. In fact, how the ES cell genome transitions from a DNA methylation-dependent to -independent mode of transposon control has never been investigated.

To study the dynamics of transposon regulation upon DNA methylation loss, we modulated the ES cell methylome by using interconvertible culture systems, which do not modify pluripotency potential. ES cells grown in standard serum-based conditions have heavily methylated genomes (~75% of CpG methylation)(Stadler et al., 2011), which is linked to the expression of de novo DNA methyltransferases. ES cells grown in presence of two small kinase inhibitors (2i) down-regulate these enzymes, and have reduced DNA methylation levels (Leitch et al., 2013; Ying et al., 2008). Upon transfer from serum to 2i medium, demethylation occurs with a slow kinetics: several weeks are required to reach 20–30% of CpG methylation. Notably, imprinted genes, major satellite repeats and IAP elements maintain persistent DNA methylation after 2i adaptation (Ficz et al., 2013; Habibi et al., 2013). Addition of vitamin C (vitC) can also lower the ES cell methylome. This compound promotes active demethylation by stimulating the TET (Ten Eleven Translocation) enzymes, which oxidize 5-methylcytosines to 5-hydroxymethylcytosines that are potential intermediates towards unmethylated cytosines (Blaschke et al., 2013).

Here, by switching ES cells directly from a serum-based to a 2i+vitC medium, we were able to induce rapid and extensive demethylation genome-wide, mimicking a situation occurring in the early embryo. By combining DNA methylation, chromatin and transcriptional profiling of transposons along with genetic analyses, we found that DNA methylation represses multiple families of transposons in ES cells, but an epigenetic switch towards histone-based control is progressively implemented as DNA methylation disappears. Importantly, we reveal for the first time the specific and overlapping roles of H3K9 and H3K27 trimethylation in controlling distinct transposon families upon DNA demethylation. These findings have important implications for understanding the molecular underpinning of transposon control in the pluripotent cells of the early mammalian embryo.

Our study provides unprecedented insight into the dynamic adaptation of the pluripotent genome to a loss of DNA methylation-based control of transposons. This was achieved through detailed kinetic assessment of transcription and chromatin states during conversion of WT ES cells from serum to 2i+vitC media, as a way to reproduce the DNA methylation erasure that occurs during embryogenesis. Despite their heterogeneous origins and structures, we found that various transposon families residing in the mouse genome adopted a common regulatory fate: after an initial transcriptional burst, repression was re-established in a DNA methylation-independent manner. Distinct combinations of H3K9me2/3 and H3K27me3 were observed among transposon families, defining three functional categories of chromatin-based responses to DNA methylation loss: joint H3K9me3 and H3K27m3 (A), H3K9me3-exclusive (B), and H3K27me3-exclusive (C) (Figure 8). Importantly, Dnmt-tKO cells, which have endured long-term adaptation to a DNA methylation-free state, displayed similar transposon-specific chromatin patterns when grown in serum, which excludes a medium-related effect. In conclusion, our work revises the previous assumption that DNA methylation is dispensable for transposon silencing in ES cells; rather, we reveal here that various histone-based repression strategies are implemented upon DNA methylation loss, thereby safeguarding pluripotent cells against a multitude of heterogeneous transposon entities.

Figure 8

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Upon 2i+vitC-mediated DNA methylation loss, the repertoire of repressive histone marks is profoundly remodeled (Figure 8A): H3K9me2 enrichment decreases, while H3K27me3 is enhanced and H3K9me3 levels are globally constant. Interestingly, repressive chromatin reorganization has also been cytologically observed in primordial germ cells (PGCs) undergoing genome-wide demethylation (Hajkova et al., 2008; Seki et al., 2007). Moreover, the relocalization of H3K27me3 at transposons and its co-occurrence with H3K9me3 was also reported in hypomethylated PGCs (Liu et al., 2014). Our cellular system could therefore represent an adequate model to study in vivo events of chromatin reprogramming occurring at transposons upon DNA methylation loss.

The persistence of H3K9me3 upon loss of DNA methylation highlights that DNA methylation does not exert significant control over H3K9me3-targeting of transposons in ES cells. Interestingly, we consistently observed that regions of persistent DNA methylation (RMRs) coincide with high H3K9me3 enrichment on transposon sequences in fully 2i+vitC-converted cells, e.g. on the 5’ end of LINE1 elements and throughout the length of ERVK elements. This supports previous evidence that H3K9me3 can confer protection against DNA demethylation (Leung et al., 2014). Inversely, the rapid disappearance of H3K9me2 upon serum to 2i+vitC conversion could reflect a direct role of DNA methylation in the maintenance of these marks. Accordingly, H3K9me2 reduction was also observed in DNA methylation-free Dnmt-tKO ES cells grown in serum (data not shown). Coupled losses of DNA methylation and H3K9me2 have also been previously reported in vivo, during normal primordial germ cell development (Hajkova et al., 2008; Seki et al., 2005) and in DNA methylation-deficient spermatocytes (Zamudio et al., 2015). A mechanism of H3K9me2 methyltransferase recruitment via DNA methylation has been resolved in plants (Du et al., 2015); the evolution of an analogous mechanism in mammals should be explored.

Of particular importance to this study is our observation of an epigenetic switch occurring between DNA methylation- and H3K27me3-based control. H3K27me3 is barely detectable at transposons in DNA hypermethylated WT ES cells grown in serum; in contrast, transposons accumulate H3K27me3 in both 2i+vitC-converted cells and in serum-grown Dnmt-tKO cells. This is in line with the prevailing notion that DNA methylation and H3K27me3 are mutually exclusive genome-wide and that DNA methylation antagonizes H3K27me3 deposition (Brinkman et al., 2012; Jermann et al., 2014; Tanay et al., 2007). Saliently, this raises the question as to how transposons acquire H3K27me3 upon DNA methylation loss. In mammalian genomes, polycomb is typically targeted to unmethylated GC-rich gene promoters (Jermann et al., 2014; Mendenhall et al., 2010). Notably, transposon sequences have a GC content superior to the genome average (Figure 5—figure supplement 2B): this signature could be sufficient to attract polycomb-mediated H3K27me3 deposition in the absence of DNA methylation. Intermediate methyl-sensitive DNA binding proteins may be involved: the BEND3 protein was recently identified as a sensor of DNA methylation states at pericentromeric repeats, recruiting polycomb-dependent H3K27me3 marks in Dnmt-tKO ES cells (Saksouk et al., 2014). Interestingly, we also observed H3K27me3 relocalization towards pericentromeric repeats in hypomethylated 2i+vitC ES cells. Similarly, the H3K36 demethylase KDM2B, which targets unmethylated CpG-rich sequences, was shown to recruit PRC1, potentially leading to H3K27me3 deposition through PRC2 recruitment (Blackledge et al., 2014; Farcas et al., 2012). Comparable mechanisms might be at play for the recruitment of H3K27me3 at hypomethylated transposons, involving BEND3, KDM2B and/or other methyl-sensitive DNA binding proteins.

Thus, based on previous observations, we posit that H3K27me3 invades the transposon space left unmarked by DNA methylation upon 2i+vitC conversion. Moreover, we provide evidence that the three possible chromatin configurations that the different transposon families adopt are further determined by H3K9me3 occupancy (Figure 8B). Mutual exclusion between H3K9me3 and H3K27me3 marks has been previously documented at gene promoters and pericentromeric repeats (Mikkelsen et al., 2007; Peters et al., 2003), but not at transposons. We found that category B transposons, which constantly maintain H3K9me3 marks throughout their entire length, do not acquire H3K27me3-based chromatin even though they lose DNA methylation. In contrast, category C transposons, exemplified by MERVL elements, become strongly enriched for H3K27me3 as H3K9me2/3 depletes during medium conversion. Finally, category A elements provide a striking illustration of the physical segregation of H3K9me3 and H3K27me3: as H3K9me3 constitutively marks the 5’ end of this transposon category, only their 3’ end is accessible to H3K27me3 deposition upon DNA methylation loss. The presence of H3K27me3 at the 3’ end of transcription units has not been described before and its functional significance was unknown. Our transcriptome analysis of 2i+vitC-converted Eed-KO cells reveals that 3’ end enrichment of H3K27me3 does not confer transcriptional repression of transposons of the A category: it may rather represent a passive response to the lack of DNA methylation and H3K9me3 at this position.

The main message conveyed by our work is that compensatory histone-based mechanisms ensure transposon silencing when DNA methylation-based control is alleviated in ES cells. We cannot rule out that other mechanisms-such as small RNA-based post-transcriptional repression-could also participate to transposon control. Importantly, genetic analyses globally confirmed the functionality of the chromatin patterns that we identified. In particular, H3K27me3-dependency was very well illustrated by the failure to repress several transposon families – in particular MERVL and LINE1– in Eed-KO ES cells undergoing medium-based DNA methylation loss. However, the transposon category B (IAPEz), which remains enriched for H3K9me3 throughout media conversion, gave complex, disparate phenotypes in the mutants of the different H3K9me3 pathways. While these elements failed to be repressed in Setdb1 and Trim28-deficient ES cells, the complete suppression of IAPEz reactivation in Suv39-dko cells was unexpected. We suspect that alternative repressive processes likely obscure IAPEz transcriptional responses to DNA methylation loss in this mutant. This is akin to Dnmt-tKO cells, which also exhibit global transposon repression. Thus, our analyses highlight the possible unexplained phenotypes of mutant cells that have adapted to long-term chromatin-based deficiencies.

Finally, one important point to raise is that the epigenetic switch from a DNA methylation-dependent to -independent mode of transposon silencing is not perfectly synchronized: ES cells experience an acute burst of transposon expression at D6 of medium conversion. At this time point, we showed that DNA methylation has been mostly erased but H3K27me3 patterns have not been established yet. Interestingly, the stability of H3K9me3 marks at category A and B transposons is not sufficient to ensure their continuous silencing upon conversion. This may imply that H3K9me3 readers are transiently deficient in this system. The lag between DNA methylation loss and subsequent implementation of histone-based repression could create an opportunistic window for transposon reactivation, provided that adequate transcription factors are available. Several studies have previously pointed out that transposons are enriched in pluripotency transcription factor binding motifs (Kunarso et al., 2010; Wang et al., 2014), in particular for NANOG and OCT4, and that upregulation of these transcription factors was sufficient to promote transposon expression (Grow et al., 2015).

We propose that the simultaneous disappearance of DNA methylation marks and increased availability of pluripotency activators create favorable conditions to transposon expression at D6 of serum to 2i+vitC conversion (Figure 8B). After a brief silencing release, functional repressive chromatin is recovered, in an H3K9me3 and/or H3K27me3-dependent manner. Notably, we repeatedly observed a peak of massive cell death of H3K27me3-deficient Eed-KO ES cells between D6 and D10 of medium conversion, when DNA methylation has mostly disappeared. This phenotype was even exacerbated in Suv39h-Eed-tKO ES cells, which did not survive after a week of medium conversion. These observations support the critical role for H3K27me3 in supplementing DNA methylation-based control in ES cells.

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Author details

1. Marius Walter

   1. Department of Genetics and Developmental Biology, Institut Curie, Paris, France
   2. Paris Science Lettres Research University
   3. UMR3215, CNRS, Paris, France
   4. U934, Inserm, Paris, France

   Contribution

   MW, Performed all the experiments, at the exception of the immunofluorescence studies, Conducted bioinformatics analyses, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

2. Aurélie Teissandier

   1. Department of Genetics and Developmental Biology, Institut Curie, Paris, France
   2. UMR3215, CNRS, Paris, France
   3. U934, Inserm, Paris, France
   4. Paris Science Lettres Research University
   5. Bioinformatics, Biostatistics, Epidemiology and Computational Systems Biology of Cancer, Institut Curie, Paris, France
   6. Mines Paris Tech, Paris, France
   7. U900, Inserm, Paris, France

   Contribution

   AT, Conducted bioinformatics analyses, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

3. Raquel Pérez-Palacios

   1. Department of Genetics and Developmental Biology, Institut Curie, Paris, France
   2. UMR3215, CNRS, Paris, France
   3. U934, Inserm, Paris, France
   4. Paris Science Lettres Research University

   Contribution

   RPP, Performed the immunofluorescence studies, Acquisition of data

   Competing interests

   The authors declare that no competing interests exist.

4. Déborah Bourc'his

   1. Department of Genetics and Developmental Biology, Institut Curie, Paris, France
   2. UMR3215, CNRS, Paris, France
   3. U934, Inserm, Paris, France
   4. Paris Science Lettres Research University

   Contribution

   DB, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   deborah.bourchis@curie.fr

   Competing interests

   The authors declare that no competing interests exist.

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