Widespread recovery of methylation at gametic imprints in hypomethylated mouse stem cells following rescue with DNMT3A2
© The Author(s) 2016
Received: 9 August 2016
Accepted: 8 November 2016
Published: 22 November 2016
Imprinted loci are paradigms of epigenetic regulation and are associated with a number of genetic disorders in human. A key characteristic of imprints is the presence of a gametic differentially methylated region (gDMR). Previous studies have indicated that DNA methylation lost from gDMRs could not be restored by DNMT1, or the de novo enzymes DNMT3A or 3B in stem cells, indicating that imprinted regions must instead undergo passage through the germline for reprogramming. However, previous studies were non-quantitative, were unclear on the requirement for DNMT3A/B and showed some inconsistencies. In addition, new putative gDMR has recently been described, along with an improved delineation of the existing gDMR locations. We therefore aimed to re-examine the dependence of methylation at gDMRs on the activities of the methyltransferases in mouse embryonic stem cells (ESCs).
We examined the most complete current set of imprinted gDMRs that could be assessed using quantitative pyrosequencing assays in two types of ESCs: those lacking DNMT1 (1KO) and cells lacking a combination of DNMT3A and DNMT3B (3abKO). We further verified results using clonal analysis and combined bisulfite and restriction analysis. Our results showed that loss of methylation was approximately equivalent in both cell types. 1KO cells rescued with a cDNA-expressing DNMT1 could not restore methylation at the imprinted gDMRs, confirming some previous observations. However, nearly all gDMRs were remethylated in 3abKO cells rescued with a DNMT3A2 expression construct (3abKO + 3a2). Transcriptional activity at the H19/Igf2 locus also tracked with the methylation pattern, confirming functional reprogramming in the latter.
These results suggested (1) a vital role for DNMT3A/B in methylation maintenance at imprints, (2) that loss of DNMT1 and DNMT3A/B had equivalent effects, (3) that rescue with DNMT3A2 can restore imprints in these cells. This may provide a useful system in which to explore factors influencing imprint reprogramming.
KeywordsImprinting DNA methylation Reprogramming ESC
In mouse, DNA methylation is found predominantly at cytosine when followed by guanine (CpG) and is associated with various biological functions including the regulation of gene expression, X chromosome inactivation, silencing of retrotransposons and imprinting . Many CpGs are protected from methylation by being clustered into CpG islands (CGI), which are commonly found near the transcriptional start sites of genes and are normally unmethylated, except for CGI on the inactive X or on inactive imprinted alleles. DNMT1, a maintenance methyltransferase , is crucial to ensure the regular propagation of DNA methylation patterns to the daughter strand during replication . This enzyme is predominantly found near replication foci  and preferentially targets hemi-methylated DNA [4–6] suggesting its main functions as a maintenance methyltransferase [7–9]. The addition of methylation to an unmethylated template (de novo) is carried out by DNMT3A and DNMT3B, with the former responsible for most de novo activity in germ cells , while the latter predominates in somatic tissues . However, in addition to their de novo methylation activity, several reports on DNMT3A and DNMT3B indicate a role in methylation maintenance in embryonic stem cells (ESCs), although the extent of their requirement at imprinted loci remains unclear [12, 13].
Once established on a DNA duplex, methylation is stably maintained through most of life [14, 15], but during certain developmental stages undergoes large-scale changes [11–13, 16]. Methylation patterns inherited from the sperm and oocyte are remodelled during pre-implantation development, when the paternal and maternal genomes of the embryo undergo widespread active demethylation involving the TET enzymes as well as passive demethylation via replicative dilution [15, 17]. The blastocyst stage sees methylation reach its nadir, but following implantation, a wave of de novo methylation occurs causing overall global hypermethylation at most non-island CpG in the adult tissues . This de novo activity is present at high levels in ESCs , developing germ cells and early post-implantation embryos  but is present at lower levels in somatic cells [20, 21]. The presence of de novo activity in ESCs makes these cells a suitable model to study the mechanism of de novo methylation in mammals.
One group of genes that largely escapes global methylation remodelling during somatic development is the imprinted genes [14, 15]. These are a group of genes which exhibit expression from one parental allele only [22, 23]. Regulation of imprinting has biological significance as imprinted genes are important for embryonic development and their dysregulation leads to embryonic death in mouse and to various disease syndromes in human . Initiation of allele-specific gene methylation patterns starts in the male and female germline during gametogenesis . For imprinted genes, one of the parental alleles acquires DNA methylation at certain locations, and these are detected as differentially methylated regions (DMRs) in somatic cells . Due to their origin in the germline, they are known as the gametic differentially methylated regions (gDMRs) , to distinguish them from other types of DMR such as tissue-specific DMR. Some of the gDMR are at cis-acting regulatory regions and are known to control monoallelic expression of more than one linked gene: where this has been proven by experimentation the DMRs are called imprinting control regions (ICRs) [27–31]. Methylation at gDMRs is established in the germline largely by the de novo methyltransferase DNMT3A with the aid of the essential cofactor DNMT3L [32–35]. The gDMRs at imprinted regions exhibit the property of being able to resist the processes of active and passive DNA demethylation during the pre-implantation stages of mammalian development or iPS formation [14, 18, 36, 37].
Loss of imprinting is thought to be irreversible and requires germline passage for its recovery due to the presence of essential factors and de novo methyltransferases needed for imprint establishment there . Previous work has shown that rescuing DNA methyltransferase activity in Dnmt1 −/− (1KO) cells by adding back a cDNA expressing the enzyme failed to restore methylation at paternal and maternal ICRs . Other laboratories confirmed this but reported, however, that the paternally imprinted H19 gDMR regained methylation in Dnmt3a −/−; Dnmt3b −/− double knock-out (3abKO) cells rescued with a DNMT3A2 expression plasmid , suggesting that some imprints could be somatically reprogrammed. As well as these differing results, the early studies were carried out on a very limited number of gDMRs using qualitative approaches, which had limited resolution. Given the important implications somatic resetting could have for imprinted disease syndromes as well as cellular reprogramming generally, we wished to re-examine whether methylation at gDMRs could be established outside of the germline. Recent work has delineated the gDMRs far more sharply since the original studies were carried out, and more quantitative techniques are now available. We aimed to investigate (1) whether deletion of Dnmt3ab gives comparable methylation loss at imprinted loci to Dnmt1 mutated cells; (2) whether imprints can be restored in 3abKO cells, unlike 1KO ESCs; (3) does loss of methylation result in dysregulated expression of imprinted genes; and (4) are there any exceptional imprinted gDMRs that do not regain methylation in rescued cells?
All laboratory experiments were carried out in triplicate with at least one biological repeat, with one or two exceptions as noted. Pyrosequencing, bisulfite sequencing and RT-qPCR data are represented as graphs, where error bars represent standard error of the mean (s.e.m). Statistical analysis was carried out using EXCEL and GraphPad PRISM software; for pyrosequencing data were compared by t test and Kruskal–Wallis, and bisulfite clonal analysis comparison was made using the χ 2 test.
All cell culture media components were purchased from Invitrogen (Paisley, UK). Dnmt1 KO and Dnmt3a/3b double KO cells with matching WT were kind gifts from Dr. Masaki Okano (RIKEN Center for Developmental Biology, Kobe, Japan). ESCs were maintained on Nunc plates (Davidson & Hardy, Belfast, UK) treated with 0.1% gelatin (Sigma-Aldrich, Dorset, UK) and cultured in Knockout DMEM plus 15% knockout serum replacement, 1% ESC-qualified Foetal Bovine Serum, 1× NEAA, 2 mM l-glutamine, 0.1 mM β-mercaptoethanol (Sigma-Aldrich, Dorset, UK) and 1000U/ml LIF (Merck Millipore, Hertfordshire, UK).
Tissues of interest were derived from outbred TO mice (Harlan, Huntingdon, UK). Sperm collection was carried out as previously described .
RNA extraction, cDNA synthesis and RT-qPCR
RNA was extracted using the RNeasy kit (Qiagen, Crawley, UK), according to the manufacturer’s instructions. For cDNA synthesis, 300–500 ng RNA was used in combination with 0.5 μg random primers (Roche, West Sussex, UK), 40 U RNaseOUT 0.5 μM dNTPs (Invitrogen, Paisley, UK) 1× RT Buffer (Fermentas, Cambridge, UK) and RevertAid reverse transcriptase (Fermentas, Cambridge, UK) made up to a final volume of 20 μl using RNase-free water (Qiagen, Crawley, UK). Reactions were carried out in a thermocycler with conditions—25 °C for 10 min, 42 °C for 60 min and 70 °C for 10 min. One microlitre cDNA per well on a 96-well plate (Roche) was used for RT-qPCR with SYBR Green reagent and remaining cDNA stored at −80 °C. RT-qPCRs were performed using a LightCycler 480 Instrument II (Roche, West Sussex, UK). Gene expression was normalised to Hprt and relative expression calculated by the ΔΔC T method . Each RT-qPCR contained 1× buffer, 0.4 mM dNTPs, 50 μM primers (Additional file 1: Table S1), 0.01 U Taq DNA polymerase (Invitrogen, Paisley, UK) and nuclease-free water (Qiagen, Crawley, UK). Four primer sets for Dnmt1, Dnmt3a, Dnmt3b  and Hprt  were used. The general thermocycler conditions are as follows—94 °C for 3 min, followed by 30 cycles of 94 °C for 30 s, 63 °C for 1 min, 72 °C for 1 min with a final elongation step of 72 °C for 4 min.
Protein was extracted from cells growing in log phase using protein extraction buffer (50 mM Tris–HCl, 150 mM NaCl, 1% Triton-X, 10% glycerol, 5 mM EDTA; all Sigma-Aldrich) and 0.5 µl protease inhibitor mix (Sigma-Aldrich, Dorset, UK). For Western blotting, 30 μg protein was denatured in the presence of 5 μl 4× LDS sample buffer (Invitrogen, Paisley, UK) and 2 μl 10× reducing agent (Invitrogen) in a total volume of 20 μl nuclease-free water (Qiagen, Crawley, UK) via incubation at 70 °C. Proteins were fractionated on a 4–12% SDS-PAGE gel, then electroblotted onto a nitrocellulose membrane (Invitrogen, Paisley, UK) and blocked in 5% non-fat milk for 1 h at room temperature (RT). Membranes were incubated with anti-DNMT1 (ab87654, Abcam), anti-DNMT3A (clone 64B1446, Novus Biologicals, Abingdon, UK), anti-GAPDH (clone 14C10, Cell Signalling Technologies, Leiden, Netherlands) or anti-β-actin (clone AC-15, Sigma-Aldrich) overnight at 4 °C, followed by incubation with the relevant HRP-conjugated secondary antibody (Sigma-Aldrich, Dorset, UK) for 1 h at RT and then visualised using ECL (ThermoFisher Scientific, Loughborough, UK).
DNA isolation and bisulfite conversion
DNA extraction from sperm and tissues was as previously described . All ESCs were pelleted and incubated overnight at 55 °C in lysis buffer (50 mM Tris pH 8, 0.1 M EDTA, 0.5% SDS (all from Sigma-Aldrich, Dorset, UK), 0.2 mg/ml proteinase K (Roche, West Sussex, UK) with rotation. DNA was extracted next day using the phenol/chloroform/isoamylalcohol (25:24:1, pH 8.0; Sigma-Aldrich, Dorset, UK) extraction method. The integrity of the DNA was checked on a 1% agarose gel (Eurogentec, Southampton, UK) and quality and quantity checked using a NanoDrop UV spectrophotometer (Labtech International, Ringmer, UK). For bisulfite conversion, 500 ng of DNA was processed with the EpiTect Bisulfite Kit (Qiagen, Crawley, UK) or EZ DNA Methylation Kit (Zymo, Cambridge, UK) according to manufacturer’s instructions.
Bisulfite-converted DNA was PCR amplified in a reaction containing 1 μM primers, 1× buffer and 0.4 mM dNTPs, with MgCl2 at a concentration specific to the primer set and 0.01U Taq DNA polymerase (all reagents from Invitrogen, Paisley, UK). Combined bisulfite restriction analysis (COBRA) on genes was carried out as previously described  using TaqαI enzyme for H19 and KvDMR and BstUI for Snrpn (both New England Biolabs, Hitchin, UK). Clonal analysis of bisulfite-converted PCR-amplified products in pJET1.2 vector (Fermentas, Cambridge, UK) was carried out using a PRISM 3130 Genetic Analyzer (Applied Biosystems, Paisley, UK). All pyrosequencing assays (Additional File 2: Table S2) were designed in-house using PyroMark (V2.0) assay design software (Qiagen, Crawley, UK). The PyroMark PCR Kit (Qiagen, Crawley, UK) was used to amplify genes using a thermocycler (Techne, Stone, UK) with conditions: 95 °C, 15 min; followed by 45 cycles of 94 °C for 30 s, 56 °C for 30 s and 72 °C for 30 s; with final elongation at 72 °C for 10 min. Subsequent pyrosequencing was carried out using Pyromark reagents as per manufacturer’s recommendations (Qiagen, Crawley, UK); 2 M NaOH was from Sigma-Aldrich (Dorset, UK) and Sepharose beads from GE Healthcare (Chalfont St. Giles, UK).
The Luminometric Methylation Assay (LUMA) using 300 ng/μl of genomic DNA from the respective cell lines was carried out exactly as described previously [13, 49]. HCT116 WT DNA (hypermethylated) and DKO DNA (hypomethylated) samples were used as a control (data not shown).
Optimising primer alignment with galaxy user-defined tracks
Wang et al.  provided chromosomal coordinates for numerous known and putative germline imprints as part of their supplemental material. The coordinates delineated for each imprint were used as a tool to define the minimal gDMR regions, from which the respective genomic sequence was extracted by visualising these regions on UCSC genome browser. The extracted genomic sequence was used to promote specificity in the design of pyrosequencing assays. BED files were generated using these chromosomal coordinates and uploaded through the Galaxy interface  as user-defined tracks visible on UCSC genome browser. The genomic sequence of interest generated from each respective imprint primer set created was matched using the BLAT tool at UCSC against the user-defined track to confirm the positions of the assays (Additional file 1: Table S1).
Initial gDMR examined and regions assayed
Methylation at imprinted gDMRs in WT ESCs is similar to that in normal tissues
Using pyrosequencing assays designed to match the known gDMR, we found that the H19 gDMR (paternally imprinted) was hypermethylated (84.2%) in sperm samples, while all maternally imprinted gDMRs displayed very low methylation (Fig. 1b), as expected. All the gDMRs assayed also showed methylation around 50% (range normally observed 40–60% ) in somatic tissues and WT ESCs, although the level of Igf2r methylation was reproducibly higher in WT ESC. Further, we confirmed a normal level of methylation for H19 and Snrpn gDMRs in WT ESCs by clonal analysis (Fig. 1c), which at 56.3 and 45.8%, respectively, was very comparable to that seen by pyrosequencing (58.5 and 42%) (Fig. 1b). These data: (1) indicated that the regions assayed by pyrosequencing showed the expected levels of methylation in somatic tissue, validating these assays, and (2) that the parental ESCs from which all the subsequent knockouts were derived had relatively normal levels of methylation at the gDMR.
Comparable demethylation at imprinted loci in cells lacking DNMT3A/B or DNMT1
Methylation can be restored following loss of DNMT3A/B, but not DNMT1
As methylation at gDMR was not completely restored to WT levels in the 3abKO + 3a2, we compared whole-genome methylation levels using LUMA , a bisulfite-independent quantitative assay that uses methylation-sensitive and methylation-insensitive enzymes to estimate total genomic methylation levels. LUMA analysis of 1KO and 3abKO cells shows clear loss of methylation compared to WT (set to 100%). While both 1KO + 1 and 3abKO + 3a2 cells showed gain in methylation, neither fully recovered to WT levels (Fig. 3b). HCT116 WT and DKO were used to confirm the LUMA was working: as expected, HCT116WT was found to be hypermethylated and methylation drops significantly for DKO cells lacking DNMT1 and DNMT3B (data not shown). To further confirm our results with respect to the imprinted gDMR, we carried out COBRA on Snrpn and H19 gDMRs, which indicated clear losses of methylation in both KO cell types, no recovery in 1KO + 1 and almost complete methylation restoration in 3abKO + 3a2 samples (Fig. 3c). We also used clonal analysis for these gDMRs, which gave similar results (Fig. 3d, e).
Transcriptional activity tracks with methylation at the H19 locus
To test whether the loss of these methyltransferases and their recovery are associated with abnormal expression, we carried out RT-qPCR on a number of imprinted loci. While most imprinted genes tested were not transcribed at significant levels in these ESC, precluding assessment of response, we did find that the expression level of H19 was significantly higher in 1KO and 3abKO cells as compared to WT, consistent with biallelic expression of H19 in those cell lines (Fig. 3f). Rescuing 1KO cells with DNMT1 did not restore repression but rescuing 3abKO cells with DNMT3A significantly reduced (p < 0.05) levels of H19 mRNA (Fig. 3f). In keeping with the regulatory mechanism in place at this locus [Fig. 1a(i)], Igf2 showed the opposite pattern, with repression in 1KO and 3abKO cells and greatest recovery in 3abKO + 3a2 cells (Fig. 3f), though this did not reach significance because of the lower transcription levels. In addition, the Peg1 mRNA, which is mildly depressed in the 3abKO line, showed a significant repressive effect of adding back in the DNMT3A2 enzyme (Fig. 3f).
Failure to restore methylation at KvDMR
Loss of methylation of remaining known and putative gDMR in Dnmt3ab KO cells
Assays designed to cover four novel putative gDMRs described in Wang et al.  (Neurog3 upstream, FR149454 promoter, Pvt1 promoter and 6330408a02Rik 3′ end) also validated in a range of mouse somatic tissue (Fig. 5b), with average values for the assays falling within the threshold criteria as per Fig. 5a. Notably, there was greater variation in the methylation values for these loci than for the known gDMR. There was less deviation from the median methylation value in liver than heart tissue, which has consistently exhibited a wider spread of results (Fig. 5a, b). The results for the known and putative imprints in mouse sperm samples are shown in Fig. 5c, where they displayed low (22.8% or below) methylation for maternally methylated gDMRs, or conversely hypermethylation (78%) at the respective paternally methylated regions.
With our validated assays, we next examined these nine additional known or putative gDMRs in the mouse ESCs. All of these loci showed decreases in methylation in 3abKO cells when compared to WT (Fig. 5d), though differences at some (Innp5f, Dlk1-Gtl2 IG and Pvt1 promoter) did not reach statistical significance.
Methylation can be restored following loss of DNMT3A/B at the majority of imprinted gDMR in DNMT3A2-rescued ES cells
Summary of findings with regard to methylation at gametic differentially methylated regions at imprinted loci
Chromosomal regiona delineated by pyro assay
Sperm meth %
Somatic meth %d
Location within gene
Gain in 3abKO + 3a2 ESCs
Maintenance methylation is a vital process as it is responsible for the stable inheritance of this epigenetic signature from mother to daughter cells during the process of mitosis. At one time, DNMT1 was thought to be the only enzyme associated with maintenance of methylation due to its preferential binding to hemi-methylated DNA and its presence at the replication foci [4, 5]. Chen et al.  showed, however, that DNMT3A and DNMT3B were also important for maintenance methylation at some repeats and, using a qualitative technique, at certain imprinted loci. In a previous study, we confirmed that DNMT3A and 3B were needed at a few selected imprinted loci using a more quantitative approach and extended this observation to transiently imprinted genes, which also require DNMT3A/B for maintenance in ESCs . Here we looked in greater depth at all the known gametic DMR as well as some newly identified imprinted gDMR and confirm their reliance (with 1–2 exceptions such as Dlk1-Gtl2 IG) on the DNMT3A/B enzymes for maintenance of methylation. Interestingly, the decrease in methylation at these gDMRs was found to be approximately the same in 1KO and 3abKO cells, suggesting an equal contribution by DNMT3A/B and DNMT1 in maintenance of methylation at imprinted gDMRs in ESCs.
Overexpression of DNMT1 in 1KO cells resulted in a global increase in methylation as reported previously; similar global increases in methylation were observed here in DNMT3A/B rescued cells using LUMA, although this increase does not bring the methylation level to the normal WT level globally. This could be due to a number of reasons: (1) it may indicate the presence of some sequences which are refractory to remethylation in ESCs; (2) the expression of the cDNA in the rescued cells may not be as high as the endogenous levels of DNMT3A2; and/or (3) some sequences may require both DNMT3B and DNMT3A2 to fully recover methylation to WT levels .
We showed here that in the 1KO + 1 cells there was no gain in methylation seen at any of the gDMR examined, confirming earlier results from a number of groups. In contrast, 3abKO cells rescued with DNMT3A2 showed clear and reproducible gains in methylation at the majority of imprinted gDMR. These results were confirmed using up to three techniques per locus-pyrosequencing, clonal analysis and COBRA. Additionally, the transcriptional status of H19 and Igf2 responded appropriately to the loss and regain of methylation, confirming that functional imprinting was being affected, at least at these loci (other loci showed transcription levels which were too low to reliably quantitate in these cells). While some previous studies have found that none  or only one  imprinted locus showed any gains in methylation on rescue, these were based on more qualitative techniques and in many cases could not examine the locus except at a low level of resolution using techniques such as Southern blotting. Here we show gain in methylation of greater than 10% at 11/14 gDMR, with substantially greater gains at most. Average gain was 28%, which in the context of an incomplete overall rescue as indicated from the global methylation levels (above) represents a corrected gain of close to 50%.
There were a few loci (3/14-KvDMR, Grb10 and 6330408a02Rik 3′ end), which showed no gain in methylation, and in fact displayed evidence for further hypomethylation relative to the 3abKO cells. This latter is not unexpected since we have shown that ESC rely on DNMT3A/B for maintenance methylation as well as de novo activity, so if these three loci are refractory to the action of DNMT3A2 in the rescued line, they would be expected to continue to lose methylation. Our examination of ENCODE data and of the current literature has found so far no common denominator for these three loci. Nevertheless, these results show that for the majority of imprinted loci, methylation at the gDMR, and in some cases functional imprinting, can be restored in a somatic cell type without germline passage.
What mechanism is associated with imprint recovery in 3abKO + 3a2 and not 1KO + 1 cells? This is particularly puzzling since the two rescued cell types both have all three enzymes present. Two possibilities are that (1) loss of DNMT3A/B proteins could alter histone marks on chromatin, which then act to attract de novo methylation by DNMT3A2 on rescue or (2) loss of DNMT1 protein causes a change in histone marks, which mean that even after rescue, the DNA cannot be remethylated. Notably, triple KO cell lines lacking all three enzymes also fail to show imprint restoration when rescued , suggesting that it is the loss of DNMT1 which leads to an irreversible change in epigenetic potential, precluding rescue with DNMT3A2. It has been reported that the loss of DNMT1 results in loss of H3K9me3 in ESC . One possibility is that loss of H3K9me3 occurs in 1KO cells, but not 3abKO cells, and that the presence of this mark facilitates remethylation by DNMT3A2 in the latter. It has also been reported that the PWWP domain of DNMT3A is linked with targeting of chromatin carrying H3K36me3 . Loss and gain of methylation marks on imprinted gDMRs could be due to the presence and absence of such interactions between methyltransferases and histone marks associated with chromatin, which require further experimental exploration in this system.
We clearly identified three gDMR, including KvDMR, where methylation once lost cannot be recovered. This supports other evidence, suggesting that mechanism of imprinting and response to methylation loss and recovery can vary among imprinted genes . In future, it will be interesting to compare the histone marks associated with KvDMR and with those associated with gDMRs that recover methylation in rescued cells. The Dlk1-Gtl2 IG was also interesting in that it showed overmethylation in our experiments, gaining almost 40% methylation in 3abKO + 3a2. The tendency of this locus to become hypermethylated in human ES and iPS cells has been noted before [53, 54] and may reflect some fundamental mechanistic feature of imprinting at this locus, which in practise could act as a barrier to somatic reprogramming efforts.
During the course of writing, a paper from the Wong group investigating the behaviour of UHRF1 rescue cells found that a number of imprinted genes showed gains in methylation in that system too . Methylation gain was only seen at some of the imprinted loci, and there was no clear link to the location of the gDMR, the presence of antisense transcripts or the type of imprint. Furthermore, they investigated common histone marks and found no relationship between any specific mark and the ability of the locus to gain methylation in the rescues. They did not, however, investigate transcriptional changes at the loci in their cells. Their data, taken together with the findings we present here, show that gametically acquired methylation at imprinted loci can be reset somatically in certain circumstances.
We have shown that (1) both DNMT1 and DNMT3A/B loss generate similar methylation changes at imprinted gDMRs in ESCs; (2) recovery of imprints in 1KO + 1 cell lines is not possible but imprints can be recovered in DNMT3A2-rescued 3abKO cells; and (3) there are some exceptional gDMRs where imprints, once lost, cannot be re-established. Our findings highlight important differences between the two cell systems and indicate that it may be possible to restore imprints somatically under certain circumstances, an observation of clear relevance for imprinting disorders. This may provide a useful model system in which to further explore reprogramming.
AT made the initial observations, carried out wet laboratory and statistical analyses of the first five loci, assembled figures and wrote the first draft. SJM carried out bioinformatics and wet laboratory analyses of all other loci, did statistical analysis, assembled figures and contributed to writing. REI carried out all westerns. From a paper ran RT-qPCR and assembled figures. GP carried out RT-PCR and RT-qPCR. KON carried out some cell culture, supervised laboratory work and contributed to design and writing. CPW conceived and designed the study, interpreted results and wrote the MS. All authors read and approved the final manuscript.
We are very grateful to Masaki Okano for the gift of cells. We thank Philip Logue, Rhonda Black, Bernie McKay and Keith Thomas for technical support and members of the Genomic Medicine Research Group for comments.
The authors declare that they have no competing interests.
Availability of data and material
All data generated or analysed during this study are included in this published article (and its supplementary information files).
Ethics approval and consent
Animals were housed and kept in accordance with UK Home Office regulations and relevant University Institutional guidance. Ethical approval was obtained from the University of Ulster Ethics Committee.
Work in the C.P.W. laboratory was supported by a Grant from the Medical Research Council (MR/J007773/1). AT was the recipient of a Vice Chancellor’s Research Studentship from Ulster University.
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