Genome-Wide Negative Feedback Drives Transgenerational DNA Methylation Dynamics in Arabidopsis

Published in the journal: Genome-Wide Negative Feedback Drives Transgenerational DNA Methylation Dynamics in Arabidopsis. PLoS Genet 11(4): e32767. doi:10.1371/journal.pgen.1005154
Category: Research Article
doi: 10.1371/journal.pgen.1005154


DNA methylation is important for controlling activity of transposable elements and genes. An intriguing feature of DNA methylation in plants is that its pattern can be inherited over multiple generations at high fidelity in a Mendelian manner. However, mechanisms controlling the trans-generational DNA methylation dynamics are largely unknown. Arabidopsis mutants of a chromatin remodeler gene DDM1 (Decrease in DNA Methylation 1) show drastic reduction of DNA methylation in transposons and repeats, and also show progressive changes in developmental phenotypes during propagation through self-pollination. We now show using whole genome DNA methylation sequencing that upon repeated selfing, the ddm1 mutation induces an ectopic accumulation of DNA methylation at hundreds of loci. Remarkably, even in the wild type background, the analogous de novo increase of DNA methylation can be induced in trans by chromosomes with reduced DNA methylation. Collectively, our findings support a model to explain the transgenerational DNA methylation redistribution by genome-wide negative feedback, which should be important for balanced differentiation of DNA methylation states within the genome.


Epigenetic variation of gene expression is mediated by chromatin marks, such as modifications of histones and DNA. Importantly, these marks and associated gene expression patterns can be inherited over multiple generations in both animals and plants [1,2]. Transgenerational epigenetic inheritance, especially the one associated with DNA methylation, is widespread in plants, and that could have a significant impact on evolution [35]. The long-term dynamics of DNA methylation has recently been examined genome-wide at single base resolution in the flowering plant Arabidopsis [6,7]; by analysing repeatedly self-pollinated wild type Arabidopsis plants, heritable gain and loss of DNA methylation have been detected, although their frequencies are generally low. A complementary approach to uncover the background mechanisms controlling long-term DNA methylation dynamics is to examine the effects of impaired DNA methylation pattern over multiple generations.

Factors controlling genomic DNA methylation have been studied extensively in Arabidopsis; and many of these factors constitute positive feedback loops to stabilize epigenetic states. Cytosine methylation in the context of dinucleotide CG is maintained by maintenance methyltransferase MET1 [8,9], while cytosine methylation at non-CG site is mediated by chromomethylases (CMTs) [10,11]. The CMTs are recruited to chromatin by methylation of histone H3 lysine 9 (H3K9me), and the H3K9 methylase KYP/SUH4 is also recruited to chromatin with non-CG methylation, generating a self-reinforcing positive feedback loop [1114]. Both H3K9me and non-CG methylation are silent heterochromatin marks normally found in repeats and transposable elements (TEs); and these marks are rarely detectable in transcribed genes. Exclusion of these marks from transcribed genes depends on the H3K9 demethylase IBM1 (Increase in BONSAI Methylation 1) [13,15]. IBM1 removes H3K9me from transcribed genes, generating another positive feedback loop to stabilize active states [13]. In addition, a positive feedback loop is also found in a process called RNA-directed DNA methylation (RdDM). RdDM is a de novo DNA methylation process triggered by double-strand RNA; and factors involved in this process have been extensively studied [1620]. The final step of RdDM is DNA methylation of both CG and non-CG sites by the de novo DNA methyltransferase DRM2 (Domains Rearranged Methylase 2), with the RNAi machinery and small interfering RNA (siRNA) functioning as upstream factors. Interestingly, production of siRNA also depends on DRM2 [21,22], suggesting another positive feedback that stabilizes the silent state. Genome-wide DNA methylation profiles have been determined in mutants of these and other factors controlling DNA methylation [11,23,24], although information for the transgenerational effects of these mutations is limited.

Among the Arabidopsis mutants affecting genomic DNA methylation, ddm1 (decrease in DNA methylation 1) is one of the mutations with the strongest effects. Mutant plants show drastic reduction of DNA methylation at both CG and non-CG sites in heterochromatic repeats and TEs [25,26]. The DDM1 gene encodes a chromatin remodeling factor, which is necessary for DNA methylation in heterochromatic sequences [10,27]. Mutation in its mammalian ortholog Lsh induces loss of DNA methylation, suggesting conserved functions across the animal and plant kingdoms [28,29].

A striking feature of the Arabidopsis ddm1 mutant is the progressive accumulation of the developmental defects; initial generations of the ddm1 mutant grow relatively normally, but many types of developmental abnormalities arise after multiple rounds of self-pollinations [30,31]. Some of the abnormalities are due to DNA sequence changes, such as insertion mutations of de-repressed endogenous TEs [3234] or a rearrangement of repeats [35], but others are due to epigenetic changes in gene expression, which correlate with changes in DNA methylation pattern at the affected loci [36,37].

Here we analyze the transgenerational effects of the ddm1 mutation genome-wide, by comparing DNA methylation of the ddm1 mutants before and after the repeated self-pollinations. This analysis revealed ectopic accumulation of non-CG methylation at hundreds of loci; and unexpectedly, this hypermethylation could only be seen after repeated self-pollinations. Furthermore, when ddm1-derived chromosomes with disrupted heterochromatin were introduced into a DDM1 wild type background, de novo accumulation of non-CG methylation was induced in trans. These results lead us to propose a model in which loss of heterochromatin is progressively compensated for through a negative feedback mechanism that leads to heterochromatin redistribution across the genome.


Early and late generations of ddm1 mutants show distinct genomic DNA methylation patterns

To understand the changes in DNA methylation patterns during self-pollinations of ddm1 mutant genome-wide, we compared DNA methylation before and after the self-pollination of the mutant. We examined DNA methylation in four individuals of ddm1 homozygous mutants segregated in progeny of a heterozygote (hereafter called 1G for the 1st Generation) and also four lines of ddm1 plants independently self-pollinated eight times (hereafter called 9G) (S1 Fig). In 1G, the ddm1 mutation already induced reduction of DNA methylation in heterochromatic regions [10,25,26]. Methylation in repetitive sequences, such as transposable elements (TEs) (Fig 1D1F), was much more severely affected than that in low copy sequences, such as genes (Fig 1A1C). The reduction was found for both CG sites (Fig 1A and 1D) and non-CG sites. In non-CG sites, both CHG sites (Fig 1B and 1E) and CHH sites (Fig 1C and 1F) were affected (H can be A, T, or C). When we compared average DNA methylation of 9G to 1G, two features were noted for both genes and TEs: further decrease of CG methylation and an increased methylation at non-CG sites (Fig 1).

DNA methylation in <i>ddm1</i> mutants before and after repeated self-pollination.
Fig. 1. DNA methylation in ddm1 mutants before and after repeated self-pollination.
DNA methylation patterns of WT, 1G ddm1, and 9G ddm1 mutants for cellular genes (A-C) or genes within transposable elements (transposable element genes, or TEGs, D-F). “WT” is a DDM1/DDM1 plant segregating as a sibling of the 1G ddm1/ddm1 plants. The black bars in the bottom represent transcribed regions. A chromosome-wide view of DNA methylation is also shown in S2 Fig.

Progressive reduction of CG methylation in the self-pollinated ddm1 lines

Although the ddm1 mutation immediately induces a drastic loss of DNA methylation in repeats, further reduction of methylation in later generations has been reported for a few CG sites [30]. Our genome-wide analysis revealed that many loci behave in a similar manner (Fig 2A). The progressive reduction of DNA methylation can have significant phenotypic effects; for example, the promoter of the imprinted gene FWA remains methylated in the 1G ddm1 but the methylation is lost stochastically in 9G ddm1 (Fig 2B), generating heritable epialleles that cause late-flowering phenotype [31,36,38]. The progressive reduction is seen genome-wide for both genes and TEs (Fig 1A and 1D).

Change of CG methylation during self-pollination of <i>ddm1</i> mutants.
Fig. 2. Change of CG methylation during self-pollination of ddm1 mutants.
(A) CG methylation level compared for each transcription unit. Each dot represents the DNA methylation level in a gene (black dot) or a transposable element gene (TEG, red dot). The top half shows effects in four different 1G ddm1 plants, while the bottom half shows effects in four different 9G ddm1 plants. Each of the 9G plants was originated from independent self-pollinations. Comparison of the 9G ddm1 plants to independently self-pollinated 9G DDM1 plants (S1 Fig) is shown in S3 Fig. “WT” is a DDM1/DDM1 plant segregating as a sibling of the 1G ddm1/ddm1 plants. (B, C) Genome browser views of loci with CG methylation reduced in 9G ddm1 using the Integrated Genome Browser [74]. FWA locus (B) and AT2G04350 locus (C) are shown. The FWA gene has dense CG methylation around the 5’ end, which is lost during self-pollination of the ddm1 mutant. (D) H3K9me2 level of differently hypo-methylated regions (hypo-DMRs) in CG context. Left (1G - WT): Distribution of 119,883 DMRs between WT and 1G ddm1 mutant. Center (9G - specific): Distribution of 25,861 DMRs between WT and 9G ddm1, excluding DMRs between WT and 1G ddm1. Distribution of 100,000 randomly chosen 100 bp regions is also shown as a control (right). H3K9me2 level of wild type is shown by reads per million (RPM) in ChIP-seq data obtained from GEO (GSE28398 [72]). (E) Change in CG methylation level in 1G ddm1 (left) and 9G ddm1 (right) from wild type, plotted against enrichment of H3K9me2 in wild type (data from Inagaki et al. 2010).

To compare the features of the regions hypomethylated immediately or gradually, we defined differentially methylated regions (DMRs; details in Materials and Methods). The regions ddm1 affects immediately (1G-WT DMRs) were enriched in dimethylation of histone H3 lysine 9 (H3K9me2) (Fig 2D left and 2E). H3K9me2 is a mark of silent heterochromatin, and these results are consistent with previous reports [10,26]. In marked contrast, however, regions affected slowly (9G-specific DMRs) have much lower level of H3K9me2 in wild type (Fig 2D middle). DDM1 gene function is necessary for CG methylation in heterochromatin, but in the long-term DDM1 also has significant effects on CG methylation in less heterochromatic regions including gene bodies (Fig 2C).

Accumulation of non-CG methylation in ddm1 lines after propagation by self-pollination

More counter-intuitively, our genome-wide analysis revealed a large number of genes and TEs ectopically hypermethylated at non-CG sites in the self-pollinated ddm1 lines (Figs 3A, 3B, 4A and 5A5E). The regions CHG hypermethylated also showed hypermethylation at CHH sites (Figs 3D, 5A5D, and S6A Fig). In addition, although genic CG methylation tends to decrease progressively from 1G to 9G on average (Figs 1 and 2), non-CG hypermethylated regions show an increase in CG methylation (Fig 3D). The CG and non-CG hypermethylation was found reproducibly at specific loci (S8 Fig). The affected loci include BONSAI and other sequences we have reported previously [37,39], but the majority of the affected loci could only be detected by whole-genome bisulfite sequencing (WGBS), because that can detect increased non-CG methylation with high sensitivity even at loci already CG methylated. In addition to genes, a large number of TEs showed increase in non-CG methylation (Figs 3A,3B, 4E, and S9S11 Figs).

Change of non-CG methylation during self-pollination of <i>ddm1</i> mutants.
Fig. 3. Change of non-CG methylation during self-pollination of ddm1 mutants.
(A, B) Effects of 1G and 9G ddm1 mutation on CHG methylation (A) and CHH methylation (B). The format is as shown for CG sites in Fig 2A. Comparison of the 9G ddm1 plants to independently self-pollinated 9G DDM1 plants is shown in S3 Fig. (C) The number of genes that gained non-CG methylation in ddm1 mutant (methylation level < 0.1 in WT and ≥ 0.1 in ddm1). Results for the four 1G and four 9G of ddm1 mutants are shown for CHG and CHH sites. (D) Coordinated hypermethylation of CG, CHG and CHH sites. “Genes CHG-hypermethylated in 9G ddm1” are those with methylation level < 0.1 in 1G ddm1 and ≥ 0.1 in 9G ddm1. DNA methylation levels for three contexts are shown for WT, 1G ddm1, and 9G ddm1. On the right, total genes are shown as controls. Although CHG hypermethylated genes tend to have more CG methylation in wild type, the body methylation is not an absolute requirement; even genes without CG methylation occasionally non-CG hypermethylated in 9G ddm1 (S4 Fig). Pattern of CG methylation change from 1G ddm1 to 9G ddm1 is further characterized in S5 Fig.

<i>BONSAI</i> hypermethylation in self-pollinated <i>ddm1</i> mutants is associated with H3K9 methylation.
Fig. 4. BONSAI hypermethylation in self-pollinated ddm1 mutants is associated with H3K9 methylation.
(A) Genome browser views of CHH and CHG methylation in AT1G73177 (BONSAI) locus. (B) H2K9me detected by chromatin immunoprecipitation (IP). “input” is the sample before IP; “mock” denotes samples after IP procedure without antibody. H3mK9me1 and H3K9me2 are samples after IP with the respective antibodies. Amplified regions around the BONSAI locus are indicated in (A). LINE and ACT2 are used as positive and negative controls, respectively. 11G plants are the ddm1 mutants in the 11th generation. 11G #1 and 11G #3 samples are prepared from progenies of direct sibling of 9G ddm1 #1 and #3 plants (shown in A), respectively. Results for other loci are shown in S6 and S7 Figs. Although the BONSAI locus accumulated both CHG and CHH methylation, some of the CHG hypermethylated loci have less CHH methylation than others (S6A Fig). In our preliminary analyses, H3K9me1 is more prevalent in those loci than H3K9me2 (S6 and S7 Figs).

Spread of non-CG methylation in self-pollinated <i>ddm1</i> mutants.
Fig. 5. Spread of non-CG methylation in self-pollinated ddm1 mutants.
(A-E) Genome browser views of loci with non-CG methylation in the 9G ddm1 plants. AT5G16880 (A-B), AT3G06480 (C-D), and AT4G07518 loci (E) are shown for CHG (A, C) and CHH (B, D, E) contexts. Direction of transcription is shown by an arrow in A-D. (F) Histogram of correlation coefficient between the CHG methylation level and the relative centroid position of CHG methylation within the DMR. The centroid position was determined by averaging relative position of the methylated cytosine weighed with the methylation level for each residue. The coefficient was calculated among the four 9G ddm1 plants in each conDMR for CHG methylation between 9G and 1G ddm1 (details in Materials and Methods section) overlapping with genes. The coefficient becomes negative when the centroid moves to the 5’ regions as the average level of CHG methylation in the conDMR increase. A large proportion of the contiguous DMRs with the coefficient near -1 reflects spread of CHG methylation from 3’ to 5’ regions as the CHG methylation levels increase.

A very unexpected feature revealed by WGBS is that non-CG hypermethylation of genes is almost undetectable in the first generation of ddm1 but is specifically and reproducibly seen in the repeatedly self-pollinated ddm1 lines. In Fig 3A and 3B, many black dots can be seen along the vertical axis in the panels for 9G but not for 1G. The non-CG hypermethylation of genes is not a simple extension of the effect seen in the first generation. This feature can only be detected in later generations (Fig 3C). In order to further understand the transgenerational dynamics, we examined four independently self-pollinated 2G ddm1 plants. If the hypermethylation proceeds equally at each self-pollination, the increase from 1G to 2G would be 1/8 or more of the increase from 1G to 9G, provided that the methylation level should saturate at specific level (the methylation level can not exceed 100%). Interestingly, although hypermethylation proceeded in 2G, the difference between 1G and 2G was much less than 1/8 of that between 1G and 9G, suggesting that the increase is slow initially but accelerated in later generations (S12 and S13 Figs).

Spread of H3K9me and non-CG methylation in ddm1 mutants

How is this non-CG hypermethylation induced? Our genome-wide bisulfite analyses revealed that the genes non-CG hypermethylated in the self-pollinated ddm1 tend to have low levels of non-CG methylation already in wild type plants (Fig 3D), suggesting that preexisting small heterochromatin domains may function as seed for further heterochromatin formation. Interestingly, distribution of H3K9me2 around the DMR is asymmetric; it is enriched in the 3’ of the DMRs (S14 Fig). We have previously shown that the BONSAI gene is flanked by insertion of a heterochromatic LINE in the 3’ region [37] (Fig 4A and S13A Fig). The BONSAI hypermethylation in ddm1 is induced in a strain with the LINE insertion but not found in a strain without the LINE insertion [37]. The DNA methylation spreads from the 3’ LINE to the BONSAI region during repeated self-pollination of ddm1 mutants [37]. Spread of non-CG methylation from 3’ to 5’ regions was also noted at other loci (Fig 5A5D). When the methylation level differs among the four 9G ddm1 plants, plants with stronger signals tended to show relative centroid positions more upstream than plants with weaker signals, suggesting that the signal spreads from 3’ to 5’ (Fig 5F). These observations suggest that common mechanisms may operate in BONSAI and many, even if not all, affected loci.

We have previously shown that the de novo non-CG methylation in the self-pollinated ddm1 does not require components of the RdDM machinery, such as RDR2, DCL3, and DRM2 [39]. On the other hand, the non-CG methylase CMT3 and H3K9 methylase KYP are necessary for the de novo methylation, suggesting that the ectopic methylation occurs by mechanisms mediated by the heterochromatin marks H3K9me and non-CG methylation [39]. Indeed, the non-CG hypermethylation at the BONSAI locus is associated with ectopic H3K9me (Fig 4B).

The self-reinforcing loop of non-CG methylase and H3K9 methylase activities could be the basis for the acceleration of hypermethylation as the generation proceeds (S13B Fig). As the two processes enhance each other, the positive feedback would accelerate the spread of the heterochromatin in later generations [12, 13].

Properties of loci hypermethylated in the self-pollinated ddm1

Increased non-CG methylation has been reported in mutants of the CG methyltransferase gene MET1 [4042], which results at least in part from a reduction of full-length IBM1 transcript [43]. The IBM1 gene encodes a demethylase for histone H3K9; and mutation in this gene induces accumulation of H3K9me2 and non-CG methylation in gene bodies. Interestingly, developmental phenotypes of the ibm1 mutation also become progressively stronger during self-pollinations [15]. We compared the regions of non-CG hypermethylation in the ibm1 and self-pollinated ddm1. Although an overlap can be detected, the majority of the DMRs in ddm1 mutants before and after the self-pollinations were distinct from the DMRs of ibm1 mutants (Fig 6B and S16 Fig). Just as progressive loss of CG methylation in the ddm1 mutant, ibm1 mutant shows progressive accumulation of non-CG methylation in later generations (Fig 6A, S15 and S16 Figs). This is consistent with a recent report [44] and likely accounts for the progressive developmental defects in the ibm1 mutant.

Hypermethylated regions in <i>ddm1</i> and <i>ibm1</i> mutants.
Fig. 6. Hypermethylated regions in ddm1 and ibm1 mutants.
(A) Increase of CHG methylation in 1G and 3G ibm1 mutants. Genes hypermethylated in 1G ibm1 (CHG methylation level < 0.1 in WT and ≥ 0.1 in 1G ibm1) are shown (right) with total genes (left). Profiles for multiple 1G and 3G ibm1 mutant plants are shown in S15 Fig. (B) Comparison of regions CHG hypermethylated in ibm1 and 9G ddm1. DMRs between 9G and 1G ddm1 (blue), between 1G ibm1 and WT (orange), and between 3G ibm1 and WT (red) are shown. Heat map of CHG methylation for these DMRs are shown in S16B Fig. (C) DNA methylation profile for the genes CHG hyper-methylated in 9G ddm1 (shown in Fig 3D). The top and bottom half represent genes and TEGs, respectively. In these regions, CHH methylation also increased in 9G ddm1.

We examined DNA methylation patterns of the genes and TEs hypermethylated in the self-pollinated ddm1 lines (Fig 6C). Compared to the ibm1 mutant, the peak in the ddm1 was shifted toward 3’ end. Interestingly, the shift of the peak in the hypermethylation was also found for CG methylation (S5D Fig). Although CG methylation of gene body in wild type peaks around the center (S5C Fig), increase of genic CG methylation in 9G ddm1 was not proportional to the methylation of wild type; instead, the increase of CG methylation was shifted toward 3’ regions (S5D Fig). Together with the observation that CHG-hypermethylated genes tend to show CG-hypermethylation (Fig 3D), these results suggest a link between the ectopic CG methylation and non-CG methylation, as we discussed previously [39].

The bias of the hypermethylation signal toward the 3’ region in 9G ddm1 is especially evident in the hypermethylated TEs; the peak was often located outside of the transcription unit for both CHG and CHH methylations (Fig 6C, bottom half). When different families of TEs are compared, the peak in the downstream region was especially evident in the GYPSY-like LTR retrotransposons (S10 Fig). Generally, these TEs lost DNA methylation in 1G ddm1, but regained methylation during the self-pollinations (S5A and S9S11 Figs).

The hypomethylated chromosomes from a ddm1 mutant could induce hypermethylation in trans even in a DDM1 wild type background

The ddm1 mutation can induce increased DNA methylation at hundreds of genes and TEs. The hypermethylation can be a direct consequence of impaired DDM1 function, or alternatively, an indirect effect of disruption of heterochromatin in the mutants. To test these possibilities, we examined the effect of chromosomes introduced from ddm1 into wild type DDM1 background. Chromosomes losing DNA methylation in the ddm1 mutants remain unmethylated even after introduction into wild type DDM1 background [25,45]. We examined DNA methylation data of epigenetic recombinant inbred lines (epiRILs) [46]. In the epiRILs, a ddm1 mutant plant was crossed to wild type plant twice to segregate DDM1/DDM1 lines with around one quarter of chromosome segments derived from ddm1. Although remethylation can be induced in regions associated with small RNA, hundreds of DMRs remain unmethylated in the wild type DDM1 background [46,47]. Each of these segregating lines have been self-pollinated seven times, which makes most of the genomic regions fixed in ddm1-derived haplotype or wild-type derived haplotype [46].

We examined if the loci exhibiting hypermethylation in the self-pollinated ddm1 lines also showed hypermethylation in some of the epiRILs. We utilized DNA methylation data for the 123 epiRILs, which are based on immunoprecipitation (IP) of genomic DNA by anti-methylcytosine antibody. As the context of methylation cannot be distinguished, we examined seven loci that show increased methylation in 9G ddm1 but a relatively low level of methylation at CG sites in wild-type. In six out of the seven loci examined, we could detect hypermethylation in multiple epiRILs, suggesting that the hypermethylation can be induced or maintained in the DDM1 background (Figs 7A, 7C, 7E and S17 Fig). In all of them, the hypermethylation showed a strong positive correlation with the amount of disrupted heterochromatin in each of these lines (Fig 7, S17 Fig and S1 Table), suggesting that the hypermethylation was induced or maintained in the background of disrupted heterochromatin in other genomic regions.

Effects of disrupted heterochromatin in the <i>DDM1</i> wild type background examined by IP.
Fig. 7. Effects of disrupted heterochromatin in the DDM1 wild type background examined by IP.
(A, C, E) Changes in local DNA methylation plotted against the global level of DNA hypomethylation in 123 epigenetic recombinant inbred lines (epiRILs). Each dot represents the value for one line. Three loci, AT1G73177 (BONSAI) (A), AT5G52480 (C), and AT5G35510 (E) are shown. Results with four other loci are shown in S17 Fig, with values for the F0 ddm1 and WT parents. (B, D, F) WT (light green) / ddm1 (dark blue) haplotype for epiRILs that showed increase of cytosine methylation for each locus (numbered 1–6 for each locus, the line names can be different among the panels). In each panel, the chromosome including the target locus (arrowhead) is shown. Haplotypes of all five chromosomes are shown in S18S23 Figs. The filled circles indicate centromere positions. The haplotypes are predicted from stably hypomethylated markers [46]. The regions not covered by any markers are indicated in gray. Names of epiRILs numbered 1–6 in each panel are in Materials and Methods. Data of epiRILs were obtained from GEO (GSE37284 [46]).

The hypermethylation could be induced de novo or alternatively maintained from the parental ddm1. The parental ddm1 plant originally used for making epiRILs was already self-pollinated three times (4G) and that plant also show low level of ectopic methylation at some loci (S17 Fig), which may have the potential to be maintained in DDM1 background [37]. Very importantly, however, the hypermethylation was found even in chromosome segments originated from wild type DDM1 (Figs 7B, 7D, 7F and S18S23 Figs), demonstrating that the hypermethylation could be induced de novo after the initial crosses and subsequent repeated self-pollinations in the background of functional DDM1.

In order to confirm and extend this observation, we used WGBS for an epiRIL with genome-wide reduction of heterochromatic DNA methylation. The epiRIL98, which contains large amount of chromosomes with reduced DNA methylation, showed CHG hypermethylation in many genes (Fig 8A), which include BONSAI gene (S24A Fig) and genes with body methylation (S24BS24C Fig). In the CHG hypermethylated genes, the CHG methylation level was generally much higher than that of the parental 4G ddm1 plant (Fig 8B), suggesting that the hypermethylation was amplified or induced de novo in the background of functional DDM1. A large number of CHG hypermethylated genes were found in chromosome regions of wild type haplotype (Fig 8C and S25 Fig), again suggesting that they can be induced de novo. In control epiRILs with much lower levels of disrupted chromatin, the hypermethylation was undetectable, confirming that the disrupted heterochromatin was responsible (Fig 8A). Taken together, these results indicate that the hypermethylation can be induced de novo by trans-acting effects of disrupted heterochromatin.

Effects of disrupted heterochromatin in the <i>DDM1</i> wild type background examined at single base resolution.
Fig. 8. Effects of disrupted heterochromatin in the DDM1 wild type background examined at single base resolution.
(A) Methylation level was compared for each transcription unit in CG, CHG, and CHH contexts. The format is as shown in Fig 2A. A globally hypomethylated epiRIL (epiRIL98: plant #3 in Fig 7A and 7B and plant #2 in Fig 7E and 7F) and two epiRILs with lower level of hypomethylation (epiRIL260 and epiRIL480) are shown. Global hypomethylation indexes of epiRIL98, epiRIL260, and epiRIL480 are 0.38, 0.04, and 0.09, respectively. “WT” data are from the parental wild-type Col plant used to generate the epiRILs. (B) CHG methylation levels in the genes that were not methylated in WT but methylated in epiRIL98 (methylation level < 0.1 in WT and ≥ 0.1 in epiRIL98: n = 232). For these transcription units, distributions of the methylation levels were compared among the parental WT, the parental 4G ddm1 plant, and the epiRIL98. (C-D) Ectopic CHG methylation in epiRIL98 compared to wild type. Each gene was assigned to the inferred haplotypes in epiRIL98: WT-like (C) or ddm1-like (D). The ectopic methylation could be detected in genes of the WT-like haplotype. Examples of such genes are shown in S25 Fig.


Local spread of heterochromatin by positive feedback loop

Here we report short- and long-term effects of the ddm1 mutation. The mutation immediately induces a drastic loss of DNA methylation in heterochromatic regions in the first generation when it becomes homozygous. In later generations, the ddm1 mutation reproducibly induces ectopic accumulation of DNA methylation in hundreds of genes and TEs. This work and previous work [39] suggest that the ectopic methylation occurs by spread of heterochromatin marks mediated by the non-CG methylase CMT3 and H3K9 methylase KYP. Interestingly, this effect was slow in the initial generations but accelerated in later generations, suggesting strong positive cooperativity for the heterochromatin accumulation. That could be explained by the self-reinforcing positive feedback of H3K9me and non-CG methylation [12, 13].

Global negative feedback for heterochromatin redistribution

In addition to the local positive feedback, global negative feedback seems important for the DNA methylation dynamics. The ectopic DNA methylation seems to reflect negative feedback of disrupted heterochromatin in other genomic regions, because the ectopic methylation could also be induced in DDM1 wild type background when the genome contains large amount of chromosomal segments with disrupted heterochromatin (Figs 7 and 8). How does the negative feedback work? One possible explanation is that disruption of heterochromatin in the ddm1 mutant results in release of heterochromatin-forming factors such as CMTs and H3K9 methylases, which then become available in other regions. As these factors are normally recruited to heterochromatin, disruption of a large proportion of heterochromatin in the genome would result in increased level of these factors in released conditions, which would induce spread of heterochromatin into normally euchromatic regions and its amplification by the self-reinforcing loop of H3K9me and non-CG methylation (Fig 9).

A model for the transgenerational heterochromatin redistribution.
Fig. 9. A model for the transgenerational heterochromatin redistribution.
The cylinder indicates a nucleosome. Red dots above the nucleosome indicate methylation of H3K9. Red and blue lines indicate DNA with and without non-CG methylation, respectively. The CMTs are non-CG methylases, such as CMT3 and CMT2 [10,11]. SUVHs are H3K9 methylases, such as SUVH4/KYP, SUVH5 and SUVH6 [75]. In both WT and ddm1 mutant plants, the histone demethylase IBM1 removes H3K9me from transcribed genes.

In the model we proposed, global reduction of heterochromatin induces ectopic non-CG methylation (Fig 9). That would account for the correlation between the global reduction of methylation and ectopic methylation in epiRILs (Fig 7A, 7C, 7E and S17 Fig). An alternative mechanism would be that ddm1 induces change in a specific locus, such as transcriptional de-repression or repression of a specific gene, and the change is inherited in the DDM1 wild type background and induces the ectopic methylation. For example, ROS1 gene expression is reduced in mutants with reduced DNA methylaiton [48], which would lead to hypermethylation at specific loci. However, although ROS1 gene expression is reduced in ddm1, it is expressed almost normally in epiRIL98, which show strong non-CG hypermethylation (S26A Fig). In addition, DMRs hypermethylated in 9G ddm1 and ros1-dml2-dml3 triple mutant do not overlap much, further suggesting that the hypermethylation in 9G ddm1 is not due to reduced ROS1 expression (S26B Fig). More generally, we could not find a locus consistently derived from ddm1 parent in all of the plants showing the high level of ectopic hypermethylation in the six loci (S18S23 Figs). Although we cannot exclude the possibility that two or more specific loci redundantly mediate the ectopic methylation, a more parsimonious explanation derived from available data would be that the trans-interaction is mediated by global homeostasis.

The de novo methylation in the epiRILs might also be related to mechanisms such as paramutation [49,50], or transchromosomal methylation (TCM) [51]. In these phenomena, methylated sequences induce methylation in related sequences. However, the ectopic hypermethylation in the epiRILs is generally much higher than that of the parental ddm1 (Fig 8B), suggesting that even if paramutation-like or TCM-like mechanisms are involved, the effect should be much amplified during self-pollinations of epiRILs; and the degree of the amplification correlates with global disruption of heterochromatin (Fig 7 and S17 Fig), which is due to the ddm1-derived chromosomes.

This trans-acting negative feedback could also be understood as a hypersensitive reaction to the challenge by active and proliferating TEs. Our genome-wide analyses revealed that many of the TEs can be targets of the negative feedback (Fig 3A and 3B and S9S11 Figs). Active TEs often keep parts of heterochromatin, which can function as seeds of the self-reinforcing heterochromatin formation.

An increase in non-CG methylation is also seen in mutants of the histone demethylase gene IBM1. However, targets of IBM1 are generally euchromatic and they do not overlap much with regions hypermethylated in the self-pollinated ddm1 lines (Fig 6B and S16 Fig). An increase in non-CG methylation is also found in the maintenance CG methylase gene MET1 [4042]. As a mechanism for the met1-induced increase in non-CG methylation, loss of IBM1 function is suggested, as IBM1 transcripts become truncated in the met1 mutant [43]. On the other hand, it has been reported that the main targets of the met1-induced accumulation of H3K9me2 are genes with H3K27me3, another modification for silent chromatin [52]. The negative feedback of heterochromatin marks comparable to that seen in the self-pollinated ddm1 lines may also operate in met1 mutants. In our analyses, although regions affected by met1, ibm1, and self-pollinated ddm1 all differ, significant overlaps are noted (S27 Fig). For these mutants, the local triggers for heterochromatin accumulation appear to be distinct, despite the possible overlap in the downstream mechanisms, including the self-reinforcing loop of non-CG methylation and H3K9me.


Heterochromatin homeostasis mechanisms analogous to those we have uncovered in Arabidopsis may also be operating in other eukaryotes. Mice with a disruption of its DDM1 homolog Lsh show global reduction of genomic DNA methylation, but interestingly it is also associated with increased DNA methylation at specific regions [29]. In human cancer, hypomethylation of repeats and TEs is often associated with local hypermethylation of genes, such as tumor suppressor genes [53,54]. In Drosophila, an increase in the amount of heterochromatic Y chromosome can results in a release of silencing at multiple loci in trans [55], suggesting a negative feedback similar to that discussed here. Furthermore, Drosophila modifiers of position effect variegation often function in dosage-dependent manners [56,57], consistent with the pathway proposed in Fig 9. Positive feedback loops would stabilize and enhance silent and active states [12,13,21,58], but they carry the risk of going out of control to excess. A global negative feedback mechanism, together with the local positive feedback, would ensure a robust and balanced chromatin differentiation within the genome, as has been discussed for pattern formation during development [59,60].

In the context of evolution in plants, a large variation in the amount of repetitive sequences is often noted between related species or even within a species [6163]. On such occasions, fine-tuning of the amount of trans-acting heterochromatin factors would be especially important, as an imbalance would not only immediately affect gene expression level but also influence the epigenotype in a transgenerational manner.

Materials and Methods

Plant materials and annotations

Isolation of the ddm1-1 and ibm1-4 mutants has been described previously [15,25]. Self-pollination of ddm1 lines was described previously [30]. In order to remove heritable effects of the ddm1 mutation, the original ddm1 mutant was backcrossed six times in the heterozygous state. The heterozygous plants were propagated by self-pollination. 1G ddm1 mutant plants were selected from self-pollinated progeny of the heterozygote. 9G ddm1 plants were generated by independently self-pollinating different ddm1 segregants eight times (S1 Fig). Generation of epiRILs has been described previously [64].

The annotations of genes and TEs are based on The Arabidopsis Information Resource ( TAIR8 was used for analyzing ChIP chip data (Fig 2E), TEG (TE gene) data, and epiRILs data. TAIR10 was used for other analyses. The details of the annotation of TEGs were described in a document in TAIR web (

DNA methylation analyses

For the 1G and 9G ddm1 plants and their controls, genomic DNA was isolated from rosette leaves using the Illustra Nucleon Phytopure genomic DNA extraction kit, and genome-wide bisulfite sequencing was performed as described previously [65]. Raw sequence data were deposited in the DDBJ (DNA Data Bank of Japan) Sequence Read Archive (DRA; accession nos. DRA002545, DRA002546, DRA002548, DRA002549, DRA002551, DRA002554, DRA002555, DRA003018, DRA003019 and DRA003020). The adaptor sequences were clipped out using the FASTX-toolkit ( Reads were trimmed to 90 nucleotide length (45 nucleotide for the data obtained from GEO—GSE39901) and mapped to reference genomes (Release 10 of the Arabidopsis Information Resources) using the Bowtie alignment algorithm [66] with the following parameters, "-X 500-e 90-l 20-n 1". Only uniquely mapped reads were used. Clonal reads were removed except one with the best quality. Any read with three consecutive methylated CHH sites were eliminated. The level of methylation of cytosine in a genomic region was calculated using the ratio of the number of methylated cytosine to that of total cytosine. For the three epiRILs and two parental lines, whole-genome bisulfite sequencing was described previously [46] and the data are in GEO (GSE62206).

DMRs (differentially methylated regions) were defined by comparing the methylation level of 100-bp windows throughout the genome between two genotypes. The windows with at least 20 sequenced cytosines were used for the comparison. The level of methylation was calculated using the weighed methylation level of each genotype [67]. The windows were selected as DMRs when difference of methylation level was 0.5 or more at CG site or 0.3 or more at CHG sites. For defining contiguous DMR (conDMR), multiple DMRs were merged if they were adjacent to each other or there was only one gap of the 100-bp window. The centroid of cytosine methylation in conDMR was calculated using the relative position within that region weighed by methylation level of each cytosine. In Fig 5F, we used conDMR of 500 bp or longer and overlapping with genes. Each contiguous DMR was aligned according to the orientation of the corresponding gene. The correlation coefficient between the level and the relative centroid position of DNA methylation was calculated among the four 9G ddm1 plants in each conDMR. To plot DNA methylation patterns over genes or TEGs in ddm1 mutants, #1 samples of each genotype (Figs 2A, 3A and 3B) in 1G ddm1 and 9G ddm1 were used. To draw the heatmap of methylation of cytosine, cluster 3.0 [68] and Java Treeview [69] were used.

Chromatin IP analysis

15-day-old seedlings were fixed with formaldehyde and ChIP was performed as described previously [70], using antibody against H3K9me1 (CMA316) and H3K9me2 (CMA307) [71]. To assure the equal amount of chromatin in each line, input DNA were quantified by quantitative PCR using TaKaRa Dice_Real Time System TP800 and ACT7 primers. Then, input DNA and each sample were diluted according to the estimated input DNA concentrations. Input DNA, mock (without antibody), and ChIP samples were analyzed by PCR. The PCR conditions were as follows: pre-incubation for 2 min at 94°C, 27 cycles at 94°C for 30 sec, 58°C for 20 sec, 72°C for 45 sec and a final extension at 72°C for 4 min. Primers used for the ChIP are listed in S2 Table. In addition to the BONSAI locus, we examined six loci with CHG methylation increased more than 0.3 from 1G ddm1 to 9G ddm1. Three of them were selected for relatively high level of ectopic CHH methylation (H1, H2, H3) and three with relatively low CHH methylation (L1, L2, L3). The increase of CHH methylation from 1G ddm1 to 9G ddm1 is more than 0.2 for the three H loci, and it is less than 0.02 for the three L loci. The lengths of amplicons for the six loci are between 250 bp and 300 bp.

Processing ChIP-seq data

ChIP-seq data of various histone modifications [72] in GEO (GSE28398) were used for our analysis. The coordinates were remapped onto TAIR10 annotation using a script in TAIR [73]. Enrichment of histone modification in a DMR was calculated by the density of ChIP-seq reads, and normalized by the mean and the standard deviation of the density of reads in 100,000 windows randomly chosen across the genome.

Processing MeDIP-chip data of epiRILs

The MeDIP-chip data of 123 epigenetic recombinant inbred lines (epiRILs), ddm1 and WT are in GEO (GSE37284). The regions that were methylated (M) in WT and unmethylated (U) in ddm1 were selected as targets of ddm1 mutation using the values for HMM (hidden Markov model) status (M (methylated) or I (Intermediate) or U (Unmethylated)) [46]. Global hypo-methylation index of an epiRIL was calculated as the genome-wide average of the values for HMM status of probes on the chip (M = 0, I = 0.5, U = 1) in the target regions of ddm1 mutation. The data of inference of inherited haplotypes were shown in the previous study [46]. Following are names of lines numbered 1–6 in Fig 7 and S18S23 Figs. (Fig 7A and 7B and S18 Fig) epiRIL208 epiRIL122 epiRIL98 epiRIL232 epiRIL70 epiRIL114; (Fig 7C and 7D and S19 Fig) epiRIL122 epiRIL208 epiRIL114 epiRIL258 epiRIL438 epiRIL508; (Fig 7E and 7F and S20 Fig) epiRIL208 epiRIL98 epiRIL438 epiRIL508 epiRIL122 epiRIL114; (S21 Fig) epiRIL208 epiRIL73 epiRIL71 epiRIL394 epiRIL98 epiRIL438; (S22 Fig) epiRIL508 epiRIL114 epiRIL122 epiRIL438 epiRIL208 epiRIL93; (S23 Fig) epiRIL208 epiRIL114 epiRIL556 epiRIL71 epiRIL244 epiRIL98.

Supporting Information































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