Advertisement
Research Article

DNA Methylation and Histone Modifications Regulate De Novo Shoot Regeneration in Arabidopsis by Modulating WUSCHEL Expression and Auxin Signaling

  • Wei Li,

    Affiliation: State Key Laboratory of Crop Biology, Shandong Key Laboratory of Crop Biology, College of Life Sciences, Shandong Agricultural University, Taian, China

    X
  • Hui Liu,

    Affiliation: State Key Laboratory of Crop Biology, Shandong Key Laboratory of Crop Biology, College of Life Sciences, Shandong Agricultural University, Taian, China

    X
  • Zhi Juan Cheng,

    Affiliation: State Key Laboratory of Crop Biology, Shandong Key Laboratory of Crop Biology, College of Life Sciences, Shandong Agricultural University, Taian, China

    X
  • Ying Hua Su,

    Affiliation: State Key Laboratory of Crop Biology, Shandong Key Laboratory of Crop Biology, College of Life Sciences, Shandong Agricultural University, Taian, China

    X
  • Hua Nan Han,

    Affiliation: State Key Laboratory of Crop Biology, Shandong Key Laboratory of Crop Biology, College of Life Sciences, Shandong Agricultural University, Taian, China

    X
  • Yan Zhang,

    Affiliation: State Key Laboratory of Crop Biology, Shandong Key Laboratory of Crop Biology, College of Life Sciences, Shandong Agricultural University, Taian, China

    X
  • Xian Sheng Zhang mail

    zhangxs@sdau.edu.cn

    Affiliation: State Key Laboratory of Crop Biology, Shandong Key Laboratory of Crop Biology, College of Life Sciences, Shandong Agricultural University, Taian, China

    X
  • Published: August 18, 2011
  • DOI: 10.1371/journal.pgen.1002243

Abstract

Plants have a profound capacity to regenerate organs from differentiated somatic tissues, based on which propagating plants in vitro was made possible. Beside its use in biotechnology, in vitro shoot regeneration is also an important system to study de novo organogenesis. Phytohormones and transcription factor WUSCHEL (WUS) play critical roles in this process but whether and how epigenetic modifications are involved is unknown. Here, we report that epigenetic marks of DNA methylation and histone modifications regulate de novo shoot regeneration of Arabidopsis through modulating WUS expression and auxin signaling. First, functional loss of key epigenetic genes—including METHYLTRANSFERASE1 (MET1) encoding for DNA methyltransferase, KRYPTONITE (KYP) for the histone 3 lysine 9 (H3K9) methyltransferase, JMJ14 for the histone 3 lysine 4 (H3K4) demethylase, and HAC1 for the histone acetyltransferase—resulted in altered WUS expression and developmental rates of regenerated shoots in vitro. Second, we showed that regulatory regions of WUS were developmentally regulated by both DNA methylation and histone modifications through bisulfite sequencing and chromatin immunoprecipitation. Third, DNA methylation in the regulatory regions of WUS was lost in the met1 mutant, thus leading to increased WUS expression and its localization. Fourth, we did a genome-wide transcriptional analysis and found out that some of differentially expressed genes between wild type and met1 were involved in signal transduction of the phytohormone auxin. We verified that the increased expression of AUXIN RESPONSE FACTOR3 (ARF3) in met1 indeed was due to DNA demethylation, suggesting DNA methylation regulates de novo shoot regeneration by modulating auxin signaling. We propose that DNA methylation and histone modifications regulate de novo shoot regeneration by modulating WUS expression and auxin signaling. The study demonstrates that, although molecular components involved in organogenesis are divergently evolved in plants and animals, epigenetic modifications play an evolutionarily convergent role in this process.

Author Summary

Plants have a strong ability to generate organs from differentiated somatic tissues. Due to this feature, shoot regeneration in vitro has been used as an important way for producing whole plants in agriculture and biotechnology. Phytohormones and the transcription factor WUSCHEL (WUS) are essential for reprogramming during de novo shoot regeneration. Epigenetic modifications are also critical for mammalian cell differentiation and organogenesis. Here, we show that epigenetic modifications mediate the de novo shoot regeneration in Arabidopsis. Mutations of key epigenetic genes resulted in altered WUS expression and developmental rates of regenerated shoots in vitro. Bisulfite sequencing and chromatin immunoprecipitation revealed that the regulatory regions of WUS were developmentally regulated by both DNA methylation and histone modifications. By transcriptome analysis, we identified that some differentially expressed genes between wild type and met1 are involved in signal transduction of the phytohormone auxin. Our results suggest that DNA methylation and histone modifications regulate de novo shoot regeneration by modulating WUS expression and auxin signaling. The study demonstrates that, although molecular components involved in organogenesis are divergently evolved in plants and animals, epigenetic modifications play an evolutionarily convergent role during de novo organogenesis.

Introduction

Differentiated somatic tissues of plants can be reprogrammed to generate various organs, a process called de novo organogenesis. This feature is not only critical for in vitro plant propagation and application of biotechnology, but also provides a good experimental system for understanding regulatory mechanisms underlying organogenesis.

Recent studies have revealed some molecular mechanisms underlying de novo shoot regeneration in Arabidopsis [1][4], in which WUS, a transcription factor, plays a key role [5], [6]. WUS is a master regulator of stem cell fate determination in shoot apical meristem (SAM), on which many signaling pathways converge [7]. It turned out to be also critical for de novo shoot regeneration. During de novo shoot regeneration in Arabidopsis, expression of WUS is sufficient to specify the organizing center, which is required for stem cell induction and subsequent shoot regeneration [5], [6], [8]. WUS induction is also essential for shoot formation during de novo somatic embryogenesis [9]. Induction of the WUS expression during de novo shoot regeneration was regulated by the master phytohormone auxin [2], [5]. Recently, WUS expression in the organizing center of the Arabidopsis plant SAM was shown to be regulated by epigenetic modifications [10].

Epigenetic modifications, including DNA methylation and histone modifications, occur extensively during cellular differentiation and development in mammals [11][13]. In mammals, the patterns of DNA methylation are established by de novo DNA methyltransferase 3 (DNMT3) family and maintained by methyltransferase DNMT1 [14]. DNMT1 plays a vital role in controlling the self-renewal and differentiation of stem cells during hematopoiesis and leukemogenesis and is critical for progenitor maintenance and self-renewal in mammalian somatic tissues [15], [16]. DNA methylation and histone modifications regulate gene expression through changing chromatin structure and transcriptional activities [17][19]. For instance, transcriptional repression is associated with hypermethylation of DNA, histone deacetylation and histone H3K9 methylation, whereas active chromatin is linked with hypomethylation of DNA, histone acetylation and histone H3K4 methylation [17], [20].

In plants, pattern changes of DNA methylation and histone modifications leading to changes in chromatin state occur in plant cells undergoing dedifferentiation [21][24]. Furthermore, DNA methylation at some promoters is critical for establishing or maintaining the undifferentiated cell state in plants [25]. However, whether and how epigenetic modifications are involved in cell differentiation during de novo shoot regeneration is unknown. Here we showed that mutations of key epigenetic genes altered WUS expression and developmental rates of regenerated shoots in vitro. In addition, epigenetic marks of DNA methylation and histone modifications in the regions of WUS underwent dynamic changes during de novo shoot regeneration, correlating with dynamic WUS expression levels. Genome-wide transcriptional analysis indicated that some genes involved in auxin signaling and meristem development were methylated within the callus, but were demethylated following an induction treatment. Based on these results, we propose that dynamic DNA methylation and histone modifications mediate de novo shoot regeneration in Arabidopsis through WUS and auxin signaling.

Results/Discussion

Mutations interfering with epigenetic modifications changed developmental rates of de novo shoot regeneration

To find out whether DNA methylation and histone modifications played roles in de novo shoot regeneration, we first compared the capacity and rates of shoot regeneration between wild type and various epigenetic mutants after calli being transferred onto a shoot induction medium (SIM) from a callus induction medium (CIM) [26]. Arabidopsis METHYLTRANSFERASE1 (MET1), KRYPTONITE (KYP), JMJ14 and HISTONE ACETYLTRANSFERASE1 (HAC1), among diverse genes involved in epigenetic modifications, have been well characterized [27][31]. MET1 is an ortholog of DNMT1, which maintains DNA methylation directly at CpG motif and indirectly at non-CG motif [27], [32], [33]. Functional loss of MET1 resulted in delayed transition from vegetative phase to reproductive phase [32]. KYP encodes histone H3K9 methyltransferase, and mutation of which resulted in abnormal number of floral organs [28]. JMJ14 encodes histone H3K4 demethylase that inhibited flowering under long-day condition [29], [34]. HAC1 encodes histone acetyltransferase, regulating flowering time through histone acetylation [31], [35]. We used the final percentage of shoot primordia on SIM to reflect the capacity of de novo shoot regeneration, whereas the timely appearance of shoot primordia to reflect their developmental rates.

Comparable maximal percentages of shoot primordia were reached after 18 days of incubation on SIM for both wild type and all tested mutants, including met1, kyp, jmj14 and hac1 (Figure 1A–1C), indicating that there was no significant difference in the capacities of de novo shoot regeneration. However, it took different induction time for the wild-type calli and the mutant calli to reach half of the maxima (Figure 1A–1C). Specifically, the mutants whose epigenetic changes were associated with more active transcription, such as met1, kyp, jmj14 [27][29], took significantly less time to reach half of the maxima as compared to the wild type (Figure 1A–1C). In contrast, the mutant associated with more repressed transcription such as hac1 took significantly more time to reach half of the maxima (Figure 1C). We obtained similar results indicating precocious or delayed initiation of shoots in these mutants using either pistils or roots as explants (Figure 1A–1C, Figure S1). Interestingly, calli of met1 cultured on SIM develop differently from those of the wild type (Figure 1D). At 4 days on SIM, around 70% met1 calli contained green regions from which the shoots would differentiate, but these green regions could not be identified in the wild-type calli. At 6 to 14 days on SIM, more shoots emerged from the met1 calli than those from the wild-type calli (Figure 1D). At 18 days on SIM, the shoots from the met1 calli were much precocious compared with those from the wild-type calli although the percentages of shoots from both the wild-type and the met1 calli were similar (Figure 1A). We also obtained similar results with roots as explants (Figure S2). Thus, these results indicated that epigenetic modifications, including DNA methylation and histone modifications, played roles in mediating developmental rate of de novo shoot regeneration.

thumbnail

Figure 1. Mutation in key epigenetic genes alters the rate of Arabidopsis shoot regeneration in vitro.

A) Frequency of shoot regeneration from calli of the wild type (Ws) and the mutant met1. B) Frequency of shoot regeneration from calli of the wild type (Ler) and the mutant kyp-2. C) Frequency of shoot regeneration from calli of the wild type (Col) and the mutants jmj14-1 and hac1-3. Calli were induced from pistils on CIM, and were then transferred onto SIM for shoot induction. Data are presented as mean values from three sets of biological replicates. In each replicate, at least 60 calli were examined. D) Calli of the wild type (Ws) and the mutant met1 cultured on SIM for 0 day, 4 days, 6 days, 10 days and 14 days. Scale bars, 1 mm.

doi:10.1371/journal.pgen.1002243.g001

Regulation of WUS expression during de novo shoot regeneration may have resulted from dynamic DNA methylation

It was well established that WUS expression is critical for stem cell formation during de novo shoot regeneration [5], [6]. Here, we showed that induction of wild-type calli on SIM for 4 days (S4) and 6 days (S6) was accompanied by a significant increase of WUS level through qRT-PCR analysis (Figure 2A). In contrast, WUS transcripts were in a low level in wild-type calli on CIM for 16 days (C16) and 20 days (S0, non-induced calli), and similar results were obtained in the prolonged time, such as calli on CIM for 24 days (C24) and 26 days (C26). We further determined the expression patterns of WUS by pWUS::GUS reporter and in situ hybridization, and the results demonstrated that local distribution of WUS transcripts occurred in wild-type calli on SIM (Figure 3, Figure S3). Because it was shown previously that WUS expression was mediated by epigenetic factors [10], we were tempted to hypothesize that the regulation of WUS expression during de novo shoot regeneration might have resulted from reduced DNA methylation.

thumbnail

Figure 2. DNA methylation and histone modifications regulate WUS transcript levels.

A) Transcript levels of WUS in calli of the wild type (Ws) and the mutant met1. B) Transcript levels of WUS in calli of the wild type (Ler) and the mutant kyp-2. C) Transcript levels of WUS in calli of the wild type (Col) and the mutants, hac1-3, hac1-5, jmj14-1 and jmj14-2. Total RNAs were isolated from calli of wild type (Ws, Ler and Col) and various mutants (met1, kyp-2, jmj14-1, jmj14-2, hac1-3 and hac1-5) cultured on SIM at the indicated time points, respectively. WUS transcript levels were quantified by qRT-PCR. The results are shown as mean values of three biological replicates with standard errors. The relative expression level of WUS gene, corresponding to the expression level of TUBULIN2, was calculated using the comparative C(T) method. After incubating on CIM for 20 days (S0), some of the calli were transferred onto SIM for further induction for 4 days (S4) and 6 days (S6), other calli were still cultured on CIM as controls (C24, C26). C16, C24, C26 indicated that pistils as explants were cultured on CIM for 16 days, 24 days and 26 days, respectively.

doi:10.1371/journal.pgen.1002243.g002
thumbnail

Figure 3. Regulation of WUS expression in met1 mutant.

A) By roots as explants, pWUS::GUS transgenic calli in the wild type transferred onto SIM for 6 days, 8 days, 10 days and 14 days, and pWUS::GUS transgenic calli in the met1 mutant transferred onto SIM for 6 days, 8 days, 10 days and 14 days. Arrowheads indicate pWUS::GUS signals. Scale bars, 1 mm. B) Longitudinal sections of pWUS::GUS transgenic calli in both the wild type and the met1 mutant transferred onto SIM for 6 days, 8 days, 10 days and 14 days, respectively. Scale bars, 50 µm.

doi:10.1371/journal.pgen.1002243.g003

To test this possibility, we first compared DNA methylation of the ~10 kb WUS genomic sequences between the calli of wild type on CIM (C16 and S0) and those on SIM (S6) by bisulfite genomic sequencing. Three regions within the WUS genomic sequences were hyper-methylated in S0 calli but substantially decreased in S6 calli (Figure 4A and 4B). Among the three regions, region I was previously proposed to regulate WUS expression [36]. Both CpG dinucleotide motifs and non-CG motifs in the three regions of the WUS genomic sequences showed induced demethylation upon induction on SIM (Figure 4B). These results showed that de novo shoot regeneration was accompanied with demethylation on methylated WUS genomic sequences. That could partially contribute to the regulation of WUS expression during de novo shoot regeneration.

thumbnail

Figure 4. Analysis of WUS methylation through bisulfite genomic sequencing.

A) A diagram of WUS structure, with +1 as the transcription start site and rectangles representing the methylated region I, II and III. B) Cytosine methylation at region I, II and III of WUS was determined by bisulfite genomic sequencing. Genomic DNA methylation status of WUS is shown in calli of the wild type on CIM for 16 days (WT, C16) and for 20 days (WT, S0), and on SIM for 6 days (WT, S6). Calli of met1 are incubated on CIM for 16 days (met1, C16) and for 20 days (met1, S0), and on SIM for 6 days (met1, S6). H represents A, T or C.

doi:10.1371/journal.pgen.1002243.g004

Demethylation and regulation of WUS expression in met1 mutant

Because DNA methylation was significantly reduced in met1 mutant [27], we wondered whether DNA methylation in the WUS genomic sequences would be affected in met1 mutant. To find out, we used two approaches. First, we compared the expression patterns of WUS in wild-type calli and met1 calli at different induction points. Indeed, the met1 mutant showed much higher WUS level than that in the wild type at each time point by qRT-PCR (Figure 2A). Then, in situ hybridization analysis demonstrated that localization of WUS in the met1 calli on SIM was earlier than that in the wild-type calli on SIM (Figure S3AS3F, Table S1). GUS staining confirmed that the pattern of WUS expression is similar to that in situ hybridization (Figure 3), and the number of GUS signal distribution in both the met1 calli and the wild-type calli on SIM is consistent to percentages of shoot primordia on SIM at different induction points (Figure 3, Figure S3, Table S2). Thus, the results indicated that WUS expression and corresponding developmental rate of de novo shoot regeneration were mediated by reduced DNA methylation.

Next, we tested whether MET1 loss of function affected the methylation status of WUS genomic region by bisulfite genomic sequencing. We found that the calli of met1 mutant on CIM (C16 and S0) and on SIM (S6) showed much lower level of DNA methylation in the WUS genomic region than those of wild type under the same condition (Figure 4B). WUS expression was detected in met1 calli earlier than in wild type based on in situ hybridization and GUS reporter analysis (Figure 3 and Figure S3). In addition, met1 contained more WUS-expressing regions than wild type, indicating that increased WUS expression level contributed to elevated the number of organizing centers (Figure 3 and Figure S3). These results suggested that the regulation of WUS expression in met1 mutant during de novo shoot regeneration could at least partially be contributed by DNA demethylation on methylated WUS genomic sequences.

Dynamic changes of histone modifications at the genomic regions of WUS during de novo shoot regeneration

Higher WUS level in the met1 mutant suggested the involvement of MET1-mediated DNA methylation in the regulation of WUS expression. However, the expression of WUS still responded to the induction by incubation on SIM in met1 mutant (Figure 2A), indicating additional pathways that regulated the dynamic expression of WUS. Because we showed that histone modifications were also important for de novo shoot regeneration (Figure 2B and 2C), we next tested whether histone modifications played a role in mediating WUS expression during de novo shoot regeneration.

We analyzed several histone modifications for the WUS genomic sequences using chromatin immunoprecipitation at two developmental stages: S0 and S6. Methylation at histone H3 at lysine 4 (H3K4me3) was shown to occur in euchromatin undergoing active transcription [37]. Whereas methylation at histone H3 at lysine 9 (H3K9me2) was shown to inhibit transcription [38]. Additionally, acetylation at histone H3 at lysine 9 (H3K9ac) is one of the most characterized epigenetic marks invariably associated with active transcription in all species investigated so far [18]. It also plays a crucial role in plant development [39].

Our results showed that these three histone modifications were dynamically regulated at the WUS genomic sequences during de novo shoot regeneration. Compared with S0, S6 showed an increase in the levels of H3K4me3 at region a and d, but not at b and c (Figure 5A and 5B). H3K4me3 occurred in euchromatin undergoing active transcription [37], therefore increased H3K4me3 levels were consistent with WUS induction during de novo shoot regeneration (Figure 1C, Figure 2C). A mark for chromatin acetylation, H3K9ac, also showed increased levels at all four regions during induction (Figure 5C). In contrast to these epigenetic marks associated with active transcription, H3K9me2, which is associated with transcription suppression [37] were reduced during de novo shoot regeneration in all four regions (Figure 5B). The changes at these epigenetic marks around WUS genomic region explained the active state of WUS chromatin structure, and might well contribute to the regulation of WUS expression during de novo shoot regeneration.

thumbnail

Figure 5. ChIP assays of calli of wild type on SIM at the WUS locus.

A) A diagram of WUS structure, with +1 as the transcription start site, and bars representing the regions examined by ChIP. B) ChIP analysis using antibodies against trimethyl H3K4 and dimethyl H3K9 at 5′ and 3′ regions of WUS in calli of wild type for 20 days on CIM (S0) and 6 days on SIM (S6). C) ChIP analysis using antibodies against H3 acetyl Lys 9 at 5′ and 3′ regions of WUS in calli of wild type (S0, S6). ACTIN was used as a control. The input was chromatin before immunoprecipitation. ‘No AB’ corresponds to chromatin treated with normal mouse IgG as the negative control. Three biological replicates were analyzed and each was tested by three technical replicates, and similar results were obtained. Representative data were shown.

doi:10.1371/journal.pgen.1002243.g005

WUS expression was changed in mutants defective in histone modifications

Dynamic histone modifications at the genomic regions of WUS during de novo shoot regeneration indicated that histone modifications contributed to regulation of WUS expression during de novo shoot regeneration. To provide further evidence that histone modifications regulated WUS expression in this process, we examined transcript level of WUS in mutants that were defective in histone modifications by qRT-PCR. As stated before, KYP, JMJ14 and HAC1 encoded enzymes for histone modification, mutations of which affected the developmental rate of de novo shoot regeneration (Figure 1B and 1C, Figure S1). Comparing with the wild-type calli, levels of WUS expression in the calli of the mutant kyp-2 were significantly enhanced compared to those of wild type for 6 days on SIM (Figure 2B). Similar results were obtained for the mutants jmj14-1 and jmj14-2 (Figure 2C). Contrast to the mutants kyp and jmj14, the levels of WUS transcripts in two different allelic hac1 mutants were reduced compared to that of wild type (Figure 2C).

Then, we used kyp-2 calli on SIM (S0, S4, and S6) to do in situ hybridization analysis. The results showed that localization of WUS signals in kyp-2 calli on SIM occurred early comparing to that in wild-type calli on SIM (Figure S3GS3L). Also, the number of localized WUS signals in kyp-2 calli on SIM (S4 and S6) was more than that in wild-type calli at the same time points (Table S1). Similar to the case of met1, expression of WUS appeared earlier in kyp-2 calli than in wild type (Figure S3). Thus, changes of WUS expression in these mutants correlated with their different developmental rates of de novo shoot regeneration, suggesting that WUS expression was regulated by histone modifications.

SIM-induced as well as MET1-dependent transcriptional changes during de novo shoot regeneration

Our results showed that DNA methylation and histone modifications regulated WUS expression during de novo shoot regeneration. To get a whole picture of epigenetic modifications during this process, we decided to do a genome-wide expression profiling using the Affymetrix ATH1 full genome array. We analyzed the transcriptomes of wild-type calli being transferred to CIM for 20 days (S0) and to SIM for 6 days (S6). Because met1 calli showed significantly different developmental rate from wild-type calli, we also analyzed transcriptomes of met1 calli being transferred to CIM for 20 days (M0) for comparison. Significance Analysis of Microarrays software package analysis was conducted for three biological samples replicates between the Ws and met1. The q value≤0.05 and fold change ≥2 were used as the threshold for candidate gene selection (Figure 6A). This criterion gave 1334 upregulated genes, and 501 downregulated genes by induction on SIM (S6 versus S0) (Table S3). 768 candidate genes showed over 2 fold difference between M0 and S0, suggesting that they might be regulated by MET1-dependent DNA methylation (Table S4). 308 candidate genes showed over 2 fold difference both between S6 versus S0 and between M0 versus S0, suggesting that they might be induced on SIM and be regulated by MET1-dependent DNA methylation (Table S5). By qRT-PCR analysis, we confirmed the microarray data (Figure S4).

thumbnail

Figure 6. Identification of the candidate genes regulated by DNA methylation.

A) The overlap between differentially-expressed genes of S6 versus S0 (Table S3) and M0 versus S0 (Table S4) were identified as candidate genes, and were listed in Table S5. A two-fold difference in the expression level of genes with a q value≤0.05 between S6 versus S0 and M0 versus S0 was set as the threshold for the selection of differentially-expressed genes. B)–E) Cytosine methylation levels of ARF3, ARF4, IAA18 and BLH7 genes in calli of wild type (S0, S6), and calli of met1 (M0) were determined by bisulfite genomic sequencing. H represents A, T or C.

doi:10.1371/journal.pgen.1002243.g006

Because auxin and cytokinin are essential for de novo shoot regeneration [2], [5], we selected genes involved in cytokinin and auxin signaling for bisulfite sequencing analysis. Indeed, some displayed differential methylation patterns during de novo shoot regeneration, such as AUXIN RESPONSE FACTOR3 (ARF3), AUXIN RESPONSE FACTOR4 (ARF4), INDOLE-3-ACETIC ACID INDUCIBLE18 (IAA18) and BELL1-LIKE HOMEODOMAIN7 (BLH7) (Figure 6B–6E). A loss of DNA methylation occurred in these genes, along with increased levels of their transcription in induced wild-type calli (Figure S4). Their expression levels were also higher in met1 than those in the wild type, suggesting that the expression of these genes might be regulated by a MET1-dependent dynamic DNA methylation during shoot regeneration. On the other hand, some candidate genes selected from SIM-induced and MET1-dependent pathways displayed no methylation, such as ASMMETRIC LEAVES1 (AS1), ARABIDOPSIS RESPONSE REGULATOR15 (ARR15), CYTOKININ OXIDASE/DEHYDROGENASE1 (CKX1), INDOLE-3-ACETIC ACID27 (IAA27) and PINOID2 (PID2), but they displayed great changes in their transcriptional levels upon SIM-induction, implying that those genes might not be directly regulated by MET1 (Table S5).

Epigenetic modifications: evolutionary recurring themes for reprogramming

DNA methylation and histone modifications are critical epigenetic processes that control chromatin structure and gene expression during development and differentiation [17], [18], and there are likely complicated interactions between these processes [20], [40]. In human, a crosstalk between DNA methylation and histone modifications has been proposed to regulate gene transcription in tumors [20]. Similarly, DNA methylation controls histone H3K9 methylation and further affect heterochromatin assembly in Arabidopsis [41]. Recent study has indicated that chromatin status facilitates the accessibility of transcription factor to FLOWERING LOCUS T (FT) in Arabidopsis, and distant regulatory regions are required for FT transcription [42]. WUS transcription is regulated through a fairly complicated chromatin remodeling mechanism in the SAM of the Arabidopsis plant [43]. It was shown that WUS expression was positively correlated with FASCIATA1 (FAS1)/FAS2, subunits of ASSEMBLY FACTOR-1 (CAF-1), and BRUSHY1 (BRU1), both of which regulate post-replicative stabilization of chromatin structure [44], [45]. Another study showed that the chromatin remodeling factor SPLAYED (SYD) directly regulated WUS to maintain proper WUS transcript levels in its spatial expression domain [46]. It has been demonstrated that at least 3.5 kb fragment upstream of WUS is required for its spatiotemporal expression during plant development [36]. Here, we showed that the 5′ and 3′ regions of WUS were regulated by SIM-induced changes of DNA methylation and histone modifications. Because the met1-3 kyp-7 double mutant displayed more severe phenotypes than each single mutant [19], we propose that regulation of WUS by DNA methylation and histone modifications may function in a partially redundant manner during de novo shoot regeneration. To understand mechanism of the in vitro organogeneis mediated by the factors involved in both DNA methylation and histone modifications, knocking out both DNA methylation and histone modifications remains to be investigated in the future.

It has long been thought that animal cells, once committed to a specific lineage, can no longer change their fate. However, recent studies suggested that differentiated animal cells do maintain plasticity and can be induced to undergo reprogramming [47], [48]. Further studies have shown that differentiated cells in mouse can be reprogrammed to pluripotent stem cells by introducing four transcription factors [49]. Plant cells can easily regenerate organs from the differentiated tissues under proper cultured conditions [1]. Previously, we used Arabidopsis ptstils as explants on CIM to obtain the callus, a mass of pluripotent cells [26], and by transferring calli onto SIM, the expression of WUS was induced in a group of cells termed the organizing center as a self-renewing source of stem cells within calli. The induced organizing center and stem cells were responsible for subsequent shoot regeneration. Here, we showed that expression of many genes was induced by SIM-induction (Figure 6A). Those genes were divided into either MET1-dependent or MET1-independent. Among MET1-dependent genes, WUS is a key transcription factor to regulate shoot regeneration [1]. ARF3 was required for shoot induction (Cheng et al., unpublished data). Previous study showed that ARF3 and ARF4 act redundantly to establish the abaxial cell fate of the Arabidopsis leaves [50]. Thus, ARF3 and ARF4 may function on de novo meristem formation mediated by epigenetic modifications. MET1-independent genes might also be involved in the process of shoot induction. Our results suggested that pluripotent cells of the callus can be reprogrammed to stem cells and subsequent, shoot formation through the regulation of both MET1-dependent genes, such as WUS and ARFs, and some MET1-independent genes.

In conclusion, our results indicate that dynamic DNA methylation and histone modifications contribute to the control of stem-cell formation and subsequent shoot regeneration. These epigenetic modifications regulate WUS and probably hormone-related genes, whose spatiotemporal expression was critical for de novo shoot regeneration. In mammals, epigenetic modifications of transcription factors and of components in hormone signaling pathways also play crucial roles in cell differentiation and organogenesis [51], [52]. Our results thus provide an interesting scenario in which epigenetic modifications were adopted as recurring themes during evolution for de novo organogenesis.

Materials and Methods

Plant materials

The met1 mutant in the Wassilewskija (Ws) background was a kind gift from Dr. J. Bender (The MCB Department of Brown University) [27]. The kyp-2 [28] mutant in the Landsberg (Ler) background, jmj14-1, jmj14-2 [29], hac1-3, and hac1-5 [31] mutants in the Columbia (Col) background were generously provided by Dr. Xiaofeng Cao (Institute of Genetics and Developmental Biology, Chinese Academy of Sciences).

Plant growth and shoot regeneration

Plants were grown as previously described [9]. Arabidopsis seeds were surface sterilized and plated on germination medium [53]. After cold treatment for 2 days at 4°C in the dark, they were transferred to sterile conditions or the growth chamber at 22°C in a 16 h light/8 h dark cycle. Shoot regeneration procedures used in this study were based on the previously described protocols [26], [54]. Pistils were excised from sterile Arabidopsis plants and transferred onto callus induction medium (CIM, MS medium [53] with 0.5 mg/L 2, 4-dichlorophenoxyacetic acid (2, 4-D) and 1.0 mg/L 6-benzylaminopurine (6-BA)). The explants were incubated for 20 days on CIM to induce callus production, and calli were then transferred onto shoot induction medium (SIM, MS medium with 0.01 mg/L indole-3-acetic acid (IAA) and 2 mg/L zeatin (ZT)). Root explants of 5–10 mm length were excised from 7-day-sterile seedlings, then transferred onto callus induction medium (CIM, Gamborg's B5 medium [55] with 0.5 g/L MES, 2% glucose, 0.2 µmol/L kinetin, and 2.2 µmol/L 2,4-dichlorophenoxyacetic acid (2,4-D), 0.8% agar), and incubated for 6 days in continuous light. Finally, explants were transferred onto shoot-inducing medium (SIM, Gamborg's B5 medium with 0.5 g/L MES, 2% glucose, 0.9 µmol/L 3-indoleacetic acid, 0.5 µmol/L 2-isopentenyladenine) and incubated in continuous light.

The morphology of calli was examined and photographed with an Olympus microscope. We defined the number of regenerated shoots as the number of at least 2 mm long shoots on each callus.

In situ hybridization

Probes were labeled using digoxigenin RNA labeling kit (Boehringer Mannheim). An antisense probe from a full-length WUS cDNA clone was generated using T7 RNA polymerase, and a sense probe was synthesized using SP6 RNA polymerase. The detailed protocol was carried out as described previously [56]. Primer sequences used for probes amplification are summarized in Table S6.

β-glucuronidase (GUS) assay

Plant tissues were incubated in GUS assay solution (50 mmol/L Na2HPO4, 50 mmol/L KH2PO4, pH 7.2, 10 mmol/L Na2EDTA, 0.5 mmol/L K3Fe(CN)6, 0.5 mmol/L K4Fe(CN)6, 1% Triton X-100 and 2 mmol/L X-Gluc (Bio. Basic Inc., Canada)) at 37°C for 12 h. To further investigate WUS expression pattern, some GUS-stained tissues were embedded in paraffin (Sigma) and sectioned. To display the outline of cells clearly, ruthenium red (200 mg/L) was used to stain cell walls.

Genomic bisulfite sequencing

DNA methylation assays were performed by bisulfite sequencing as previously described [57]. PCR products were cloned into the pMD19-T Simple Vector (Takara), and 12 clones were sequenced to determine the methylation status of a locus in each genotype. Primer sequences are shown in Table S6. Bisulfite sequencing data were analyzed by the CyMATE software [58]. The results returned by CyMATE were input into SigmaPlot 10.0 to illustrate DNA methylation frequencies at CG, CHG and CHH (where H = A, C or T) at the various cultured stages of each genotype.

Chromatin immunoprecipitation assay

The Arabidopsis calli grown on CIM for 20 days (S0) and on SIM for 6 days (S6) were vacuum-infiltrated with formaldehyde crosslinking solution. Chromatin immunoprecipitation was performed according to manufactures' instructions (Epigentek Group Inc. USA, Catalogno. P-2014). Chromatin samples were immunoprecipitated with antibodies against a negative control normal mouse IgG and H3 dimethyl Lys 9 (both included in EpiQuik™ Plant ChIP Kit), or with antibodies against H3 trimethyl Lys 4 (Abcam USA, Catalogno. ab1012) and H3 acetyl Lys 9 (Abcam USA, Catalogno. ab10812). PCR amplification was performed in 25 µL volumes for 32 to 37 cycles to determine the appropriate conditions for the PCR products of each region. Primer sequences are shown in Table S6. The PCR products were electrophoresed in a 2% agarose gel. Three biological replicates were analyzed and each was tested by three technical replicates.

Total RNA isolation and quantitative real-time PCR analysis

Total RNAs were isolated from callus tissues 2 to 3 mm deep from the surface. Quantitative real-time PCRs (qRT-PCRs) were performed as described previously [9]. To check the specificity of amplification, the melting curve of the PCR products was detected. The expression levels of specific genes were standardized to the housekeeping gene TUBULIN2. Each reaction was carried out in three biological replicates. The relative expression level of each gene, corresponding to the expression level of TUBULIN2, was calculated using the comparative CT method [59]. Primer sequences used for qRT-PCR are summarized in Table S6.

DNA microarray analysis

RNA of three plant samples was prepared from each of the following tissue types: the wild-type calli cultured on CIM for 20 days (S0), and on SIM for 6 days (S6); the met1 mutant calli cultured on CIM for 20 days (M0). RNA purification, probe labeling, chip hybridization, probe array scanning and data pre-processing normalization were performed by the Affymetrix custom service (CapitalBio, Beijing, China). Significance Analysis of Microarrays software package analysis was conducted for three biological samples replicates between the Ws and met1. When all replicates clustered together, further analysis was performed based on mean values. A two-fold change in the gene expression levels between one versus another samples with a q value≤0.05 was set as the threshold for altered gene expression. Microarray data are available in the ArrayExpress database (www.ebi.ac.uk/arrayexpress) under accession number E-MEXP-3120.

Supporting Information

Figure S1.

Frequency of shoot regeneration of met1 mutant and the mutants defective in histone modifications. Frequency of shoot regeneration of the wild type (Col) and the mutants jmj14-2 and hac1-5 was shown, using pistils as explants. Frequency of shoot regeneration of the wild type (Ws, Ler and Col) and the mutants met1, kyp-2, jmj14-1, jmj14-2, hac1-3 and hac1-5 was shown, using roots as explants. Standard errors were calculated from three sets of biological replicates. In each replicate, at least 60 calli were examined.

doi:10.1371/journal.pgen.1002243.s001

(TIF)

Figure S2.

MET1 mutation promotes shoot regeneration in Arabidopsis using roots as explants. Calli of the wild type (Ws) and the met1 mutant were cultured on SIM for 6 to 18 days. Scale bars, 1 mm.

doi:10.1371/journal.pgen.1002243.s002

(TIF)

Figure S3.

Expression patterns of WUS were changed in met1 and kyp-2 mutants. In situ hybridization of WUS expression in calli of the wild type (Ws) cultured on SIM for A) 0 day, B) 4 days and C) 6 days, and that of met1 mutant cultured on SIM for D) 0 day, E) 4 days and F) 6 days. In situ hybridization of WUS expression in calli of the wild type (Ler) cultured on SIM for G) 0 day, H) 4 days and I) 6 days, and that of kyp-2 mutant cultured on SIM for J) 0 day, K) 4 days and L) 6 days. Scale bars, 50 µm.

doi:10.1371/journal.pgen.1002243.s003

(TIF)

Figure S4.

Expression patterns of candidate genes validated by qRT-PCR. Total RNAs were isolated from calli of wild type and met1 cultured on SIM at the indicated time points, and the transcripts of genes ARF3, ARF4, IAA18, BLH7, ANT, AS1, CKX1, and ARR15 were measured by qRT-PCR. Three independent RNA preparations were analyzed for each time point. Mean values were calculated from triplicate qRT-PCR analysis with standard errors. The relative expression level of each gene, corresponding to the expression level of TUBULIN2, was calculated using the comparative C(T) method.

doi:10.1371/journal.pgen.1002243.s004

(TIF)

Table S1.

The percentage of the calli with WUS expressing signals detected by in situ hybridization.

doi:10.1371/journal.pgen.1002243.s005

(DOC)

Table S2.

The number of pWUS::GUS signal distribution detected in each callus.

doi:10.1371/journal.pgen.1002243.s006

(DOC)

Table S3.

List of 1334 up-regulated genes and 501 down-regulated genes in S6 as compared to S0.

doi:10.1371/journal.pgen.1002243.s007

(XLS)

Table S4.

List of 768 genes showing more than two-fold difference between M0 and S0.

doi:10.1371/journal.pgen.1002243.s008

(XLS)

Table S5.

List of 308 genes showing more than two fold difference both between S6 and S0 and between M0 and S0.

doi:10.1371/journal.pgen.1002243.s009

(XLS)

Table S6.

Sequences of primers used in this study.

doi:10.1371/journal.pgen.1002243.s010

(XLS)

Acknowledgments

The authors thank Dr. J. Bender (The MCB Department of Brown University) for her generously providing the mutant met1 and Dr. Xiaofeng Cao (Institute of Genetics and Developmental Biology, Chinese Academy of Sciences) for her kindly providing the mutants kyp-2, jmj14-1, jmj14-2, hac1-3, and hac1-5.

Author Contributions

Conceived and designed the experiments: XSZ WL. Performed the experiments: XSZ WL HL ZJC YHS HNH. Analyzed the data: XSZ WL HL. Contributed reagents/materials/analysis tools: XSZ WL HL ZJC YZ. Wrote the paper: XSZ YZ.

References

  1. 1. Che P, Lall S, Nettleton D, Howell SH (2006) Gene expression programs during shoot, root, and callus development in Arabidopsis tissue culture. Plant Physiol 141: 620–637.
  2. 2. Atta R, Laurens L, Boucheron-Dubuisson E, Guivarc'h A, Carnero E, et al. (2009) Pluripotency of Arabidopsis xylem pericycle underlies shoot regeneration from root and hypocotyl explants grown in vitro. Plant J 57: 626–644.
  3. 3. Pernisová M, Klíma P, Horák J, Válková M, Malbeck J, et al. (2009) Cytokinins modulate auxin-induced organogenesis in plants via regulation of the auxin efflux. Proc Natl Acad Sci USA 106: 3609–3614.
  4. 4. Che P, Lall S, Howell SH (2007) Developmental steps in acquiring competence for shoot development in Arabidopsis tissue culture. Planta 226: 1183–1194.
  5. 5. Gordon SP, Heisler MG, Reddy GV, Ohno C, Das P, et al. (2007) Pattern formation during de novo assembly of the Arabidopsis shoot meristem. Development 134: 3539–3548.
  6. 6. Gallois JL, Nora FR, Mizukami Y, Sablowski R (2004) WUSCHEL induces shoot stem cell activity and developmental plasticity in the root meristem. Genes Dev 18: 375–380.
  7. 7. Dodsworth S (2009) A diverse and intricate signalling network regulates stem cell fate in the shoot apical meristem. Dev Biol 336: 1–9.
  8. 8. Schoof H, Lenhard M, Haecker A, Mayer KFX, Jürgens G, et al. (2000) The stem cell population of Arabidopsis shoot meristems is maintained by a regulatory loop between the CLAVATA and WUSCHEL genes. Cell 100: 635–644.
  9. 9. Su YH, Zhao XY, Liu YB, Zhang CL, O'Neill SD, et al. (2009) Auxin-induced WUS expression is essential for embryonic stem cell renewal during somatic embryogenesis in Arabidopsis. Plant J 59: 448–460.
  10. 10. Shen WH, Xu L (2009) Chromatin remodeling in stem cell maintenance in Arabidopsis thaliana. Mol Plant 2: 600–609.
  11. 11. Meissner A, Mikkelsen TS, Gu H, Wernig M, Hanna J, et al. (2008) Genome-scale DNA methylation maps of pluripotent and differentiated cells. Nature 454: 766–770.
  12. 12. Wang K, Chen Y, Chang EA, Knott JG, Cibelli JB (2009) Dynamic epigenetic regulation of the Oct4 and Nanog regulatory regions during neural differentiation in rhesus nuclear transfer embryonic stem cells. Cloning Stem Cells 11: 483–496.
  13. 13. Shukla V, Vaissière T, Herceg Z (2008) Histone acetylation and chromatin signature in stem cell identity and cancer. Mutat Res 637: 1–15.
  14. 14. Goll MG, Bestor TH (2005) Eukaryotic cytosine methyltransferases. Annu Rev Biochem 74: 481–514.
  15. 15. Bröske AM, Vockentanz L, Kharazi S, Huska MR, Mancini E, et al. (2009) DNA methylation protects hematopoietic stem cell multipotency from myeloerythroid restriction. Nat Genet 41: 1207–1215.
  16. 16. Sen GL, Reuter JA, Webster DE, Zhu L, Khavari PA (2010) DNMT1 maintains progenitor function in self-renewing somatic tissue. Nature 463: 563–567.
  17. 17. Vaillant I, Paszkowski J (2007) Role of histone and DNA methylation in gene regulation. Curr Opin Plant Biol 10: 528–533.
  18. 18. Kouzarides T (2007) Chromatin modifications and their function. Cell 128: 693–705.
  19. 19. Mathieu O, Reinders J, Čaikovski M, Smathajitt C, Paszkowski J (2007) Transgenerational stability of the Arabidopsis epigenome is coordinated by CG methylation. Cell 130: 851–862.
  20. 20. Vaissière T, Sawan C, Herceg Z (2008) Epigenetic interplay between histone modifications and DNA methylation in gene silencing. Mutat Res 659: 40–48.
  21. 21. Grafi G (2004) How cells dedifferentiate: a lesson from plants. Dev Biol 268: 1–6.
  22. 22. Koukalova B, Fojtova M, Lim KY, Fulnecek J, Leitch AR, et al. (2005) Dedifferentiation of Tobacco cells is associated with ribosomal RNA gene hypomethylation, increased transcription, and chromatin alterations. Plant Physiol 139: 275–286.
  23. 23. Grafi G, Ben-Meir H, Avivi Y, Moshe M, Dahan Y, et al. (2007) Histone methylation controls telomerase-independent telomere lengthening in cells undergoing dedifferentiation. Dev Biol 306: 838–846.
  24. 24. Grafi G, Avivi Y (2004) Stem cells: a lesson from dedifferentiation. Trends in Biotechnology 22: 388–389.
  25. 25. Berdasco M, Alcázar R, García-Ortiz MV, Ballestar E, Fernández AF, et al. (2008) Promoter DNA hypermethylation and gene repression in undifferentiated Arabidopsis cells. PLoS ONE 3: e3306. doi:10.1371/journal.pone.0003306.
  26. 26. Cheng ZJ, Zhu SS, Gao XQ, Zhang XS (2010) Cytokinin and auxin regulates WUS induction and inflorescence regeneration in vitro in Arabidopsis. Plant Cell Reports.
  27. 27. Bartee L, Bender J (2001) Two Arabidopsis methylation-deficiency mutations confer only partial effects on a methylated endogenous gene family. Nucleic Acids Res 29: 2127–2134.
  28. 28. Jackson JP, Lindroth AM, Cao X, Jacobsen SE (2002) Control of CpNpG DNA methylation by the KRYPTONITE histone H3 methyltransferase. Nature 416: 556–560.
  29. 29. Lu F, Cui X, Zhang S, Liu C, Cao X (2010) JMJ14 is an H3K4 demethylase regulating flowering time in Arabidopsis. Cell Res 20: 387–390.
  30. 30. Searle IR, Pontes O, Melnyk CW, Smith LM, Baulcombe DC (2010) JMJ14, a JmjC domain protein, is required for RNA silencing and cell-to-cell movement of an RNA silencing signal in Arabidopsis. Genes Dev 24: 986–991.
  31. 31. Deng W, Liu C, Pei Y, Deng X, Niu L, et al. (2007) Involvement of the histone acetyltransferase AtHAC1 in the regulation of flowering time via repression of FLOWERING LOCUS C in Arabidopsis. Plant Physiol 143: 1660–1668.
  32. 32. Kankel MW, Ramsey DE, Stokes TL, Flowers SK, Haag JR, et al. (2003) Arabidopsis MET1 cytosine methyltransferase mutants. Genetics 163: 1109–1122.
  33. 33. Lister R, O'Malley RC, Tonti-Filippini J, Gregory BD, Berry CC, et al. (2008) Highly integrated single-base resolution maps of the epigenome in Arabidopsis. Cell 133: 523–536.
  34. 34. Yang W, Jiang D, Jiang J, He Y (2010) A plant-specific histone H3 lysine 4 demethylase represses the floral transition in Arabidopsis. Plant J 62: 663–673.
  35. 35. Earley KW, Shook MS, Brower-Toland B, Hicks L, Pikaard CS (2007) In vitro specificities of Arabidopsis co-activator histone acetyltransferases: implications for histone hyperacetylation in gene activation. Plant J 52: 615–626.
  36. 36. Bäeurle I, Laux T (2005) Regulation of WUSCHEL transcription in the stem cell niche of the Arabidopsis shoot meristem. Plant Cell 17: 2271–2280.
  37. 37. Barski A, Cuddapah S, Cui K, Roh TY, Schones DE, et al. (2007) High-resolution profiling of histone methylations in the human genome. Cell 129: 823–837.
  38. 38. Jackson JP, Johnson L, Jasencakova Z, Zhang X, PerezBurgos L, et al. (2004) Dimethylation of histone H3 lysine 9 is a critical mark for DNA methylation and gene silencing in Arabidopsis thaliana. Chromosoma 112: 308–315.
  39. 39. Zhou J, Wang X, He K, Charron JB, Elling AA, et al. (2010) Genome-wide profiling of histone H3 lysine 9 acetylation and dimethylation in Arabidopsis reveals correlation between multiple histone marks and gene expression. Plant Mol Biol 72: 585–595.
  40. 40. Deleris A, Greenberg MV, Ausin I, Law RW, Moissiard G, et al. (2010) Involvement of a Jumonji-C domain-containing histone demethylase in DRM2-mediated maintenance of DNA methylation. EMBO Rep 11: 950–955.
  41. 41. Soppe WJ, Jasencakova Z, Houben A, Kakutani T, Meister A, et al. (2002) DNA methylation controls histone H3 lysine 9 methylation and heterochromatin assembly in Arabidopsis. EMBO J 21: 6549–6559.
  42. 42. Adrian J, Farrona S, Reimer JJ, Albani MC, Coupland G, et al. (2010) cis-Regulatory elements and chromatin state coordinately control temporal and spatial expression of FLOWERING LOCUS T in Arabidopsis. Plant Cell 22: 1425–1440.
  43. 43. Williams L, Fletcher JC (2005) Stem cell regulation in the Arabidopsis shoot apical meristem. Curr Opin Plant Biol 8: 582–586.
  44. 44. Kaya H, Shibahara K-i, Taoka K-i, Iwabuchi M, Stillman B, et al. (2001) FASCIATA genes for Chromatin Assembly Factor-1 in Arabidopsis maintain the cellular organization of apical meristems. Cell 104: 131–142.
  45. 45. Takeda S, Tadele Z, Hofmann I, Probst AV, Angelis KJ, et al. (2004) BRU1, a novel link between responses to DNA damage and epigenetic gene silencing in Arabidopsis. Genes Dev 18: 782–793.
  46. 46. Kwon CS, Chen C, Wagner D (2005) WUSCHEL is a primary target for transcriptional regulation by SPLAYED in dynamic control of stem cell fate in Arabidopsis. Genes Dev 19: 992–1003.
  47. 47. Graf T (2002) Differentiation plasticity of hematopoietic cells. Blood 99: 3089–3101.
  48. 48. Liu Y, Rao MS (2003) Transdifferentiation-fact or artifact. J Cell Biochem 88: 29–40.
  49. 49. Takahashi K, Yamanaka S (2006) Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126: 663–676.
  50. 50. Pekker I, Alvarez JP, Eshed Y (2005) Auxin response factors mediate Arabidopsis organ asymmetry via modulation of KANADI activity. Plant Cell 17: 2899–2910.
  51. 51. Birnbaum KD, Alvarado AS (2008) Slicing across kingdoms: regeneration in plants and animals. Cell 132: 697–710.
  52. 52. Liu J, Casaccia P (2010) Epigenetic regulation of oligodendrocyte identity. Trends in Neurosciences 33: 193–201.
  53. 53. Murashige T, Skoog F (1962) A revised medium for rapid growth and bioassays with Tobacco tissue cultures. Physiologia Plantarum 15: 473–497.
  54. 54. Buechel S, Leibfried A, To JP, Zhao Z, Andersen SU, et al. (2010) Role of A-type ARABIDOPSIS RESPONSE REGULATORS in meristem maintenance and regeneration. Eur J Cell Biol 89: 279–284.
  55. 55. Gamborg OL, Miller RA, Ojima K (1968) Nutrient requirements of suspension cultures of soybean root cells. Exp Cell Res 50: 151–158.
  56. 56. Zhao XY, Cheng ZJ, Zhang XS (2006) Overexpression of TaMADS1, a SEPALLATA-like gene in wheat, causes early flowering and the abnormal development of floral organs in Arabidopsis. Planta 223: 698–707.
  57. 57. Jacobsen SE, Sakai H, Finnegan EJ, Cao X, Meyerowitz EM (2000) Ectopic hypermethylation of flower-specific genes in Arabidopsis. Curr Biol 10: 179–186.
  58. 58. Hetzl J, Foerster AM, Raidl G, Scheid OM (2007) CyMATE: a new tool for methylation analysis of plant genomic DNA after bisulphite sequencing. Plant J 51: 526–536.
  59. 59. Schmittgen TD, Livak KJ (2008) Analyzing real-time PCR data by the comparative CT method. Nat Protocols 3: 1101–1108.