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. 2016 Sep 5;25(22):4870–4880. doi: 10.1093/hmg/ddw311

The FMR1 promoter is selectively hydroxymethylated in primary neurons of fragile X syndrome patients

Rustam Esanov 1, Nadja S Andrade 1, Sarah Bennison 1, Claes Wahlestedt 1, Zane Zeier 1,*
PMCID: PMC6078593  PMID: 28173181

Abstract

Fragile X syndrome (FXS) results from a repeat expansion mutation near the FMR1 gene promoter and is the most common form of heritable intellectual disability and autism. Full mutations larger than 200 CGG repeats trigger FMR1 heterochromatinization and loss of gene expression, which is primarily responsible for the pathological features of FXS . In contrast, smaller pre-mutations of 55–200 CGG are associated with FMR1 overexpression and Fragile X-associated tremor/ataxia syndrome (FXTAS), a late-onset neurodegenerative condition. While the role of 5-methylcytosine (5mC) in FMR1 gene silencing has been studied extensively, the role of 5-hydroxymethylation (5hmC), a newly discovered epigenetic mark produced through active DNA demethylation, has not been previously investigated in FXS neurons. Here, we used two complementary epigenetic assays, 5hmC sensitive restriction digest and ten-eleven translocation-assisted bisulfite pyrosequencing, to quantify FMR1 5mC and 5hmC levels. We observed increased levels of 5hmC at the FMR1 promoter in FXS patient brains with full-mutations relative to pre-mutation carriers and unaffected controls. In addition, we found that 5hmC enrichment at the FMR1 locus in FXS cells is specific to neurons by utilizing a nuclei sorting technique to separate neuronal and glial DNA fractions from post-mortem brain tissues. This FMR1 5hmC enrichment was not present in cellular models of FXS including fibroblasts, lymphocytes and reprogrammed neurons, indicating they do not fully recapitulate this epigenetic feature of disease. Future studies could investigate the potential to leverage this epigenetic pathway to restore FMR1 expression and discern whether levels of 5hmC correlate with phenotypic severity.

Introduction

Fragile X syndrome (FXS) is associated with a spectrum of physical and behavioral characteristics, the most debilitating of which is severe intellectual disability; it is the most common known genetic form of autism and is the most frequent heritable form of mental retardation affecting 1:4000 males and 1:8000 females (1,2). Typically, FXS results from a trinucleotide CGG repeat expansion mutation within the 5′ untranslated region of the FMR1 gene that leads to transcriptional repression via epigenetic restructuring (3–5). The FMR1 gene product is an RNA binding protein, called FMRP, which is necessary for proper RNA metabolism and synaptic plasticity (6,7). Both transcriptional repression and reduced translational efficiency of expanded mRNAs conspire to halt the production of FMRP, which is primarily responsible for the pathological features of FXS (8). Occurring early in embryogenesis (9), premutated FMR1 alleles (55–200 CGG repeats) of the maternal lineage expand into larger full mutations (>200 repeats) in an anticipatory manner where larger premutations are more likely to transmit (10,11). While premutated FMR1 alleles do not undergo heterochromatinization, they are not without consequence, as the over-production of mutant FMR1 mRNA results in Fragile X-associated tremor/ataxia syndrome (FXTAS) at a rate of 1 in 250 male and 350 female adults (2,12). Remarkably, full FMR1 mutations compromise the chromatin superstructure so profoundly that a portion of the X chromosome becomes precariously displaced and appears ‘fragile’ when prepared for karyotype analysis (13). The FMR1 expansion was identified in 1991 (5) placing it among the first to be discovered in a rapidly growing family of repeat expansion disorders that has now reached nearly 40 members (14,15).

Expanded FMR1 alleles are depleted of active chromatin marks and are instead adorned with repressive epigenetic marks. Specifically, the level of acetylation at lysine residues of histones 3 and 4—permissive chromatin markers—are reduced whereas repressive markers (H3K9me3, H4K20me3 and H3K27me3) are enriched (16–21). In addition to histone tail modifications, methylation of cytosine residues (5-methylcytosine, 5mC), primarily in the context of CpG dinucleotides, is enriched within the promoter and repeat sequence (4,22,23), which is highly predictive of phenotypic severity and is diagnostic of FXS (24). It has been proposed that 5mC is restricted to a region upstream of the FMR1 promoter due to a DNA methylation boundary, but spreads downstream into the FMR1 promoter and CGG repeat region in FXS cells where this boundary is lost (25).

Demethylation of 5mC has an important role in neurodevelopment and gene regulation that has only recently been realized (26,27). The first step of active DNA demethylation is conversion of 5mC to 5-hydroxymethylcytosine (5hmC), a process catalyzed by the ten-eleven translocation (TET) family proteins (27,28). Apart from being a stable intermediate of DNA demethylation, 5hmC serves as an epigenetic mark often referred as ‘sixth’ base. Now a well-substantiated epigenetic mark, 5hmC, has been implicated in transcriptional activation of genes, in part by changing the affinity of DNA-binding proteins (29,30). 5hmC levels are enriched at regions upstream of transcription start site (TSS) including CpG islands and are highest in mammalian embryonic stem cells and brain tissue (29,31–33). Aberrant 5hmC levels have been implicated in impaired learning, short-term memory, fear extinction and long-term depression (34–37).

Previous investigations of FMR1 hypermethylation in FXS utilized the standard bisulfite conversion technique, which is incapable of distinguishing 5mC from 5hmC (38). In light of this, we sought to investigate the potential role of DNA demethylation in FMR1 silencing. We employed improved methods to quantify 5mC and 5hmC at the FMR1 locus in FXS and FXTAS patient brains as well as cellular models of FXS including primary fibroblasts, immortalized lymphocytes, induced pluripotent stem cell derived neurons (iPSC-Neurons) and embryonic stem cell derived neurons (ES-Neurons). Demethylation of the silenced FMR1 locus could have therapeutic implications, given the opposing effects of 5mC and 5hmC on transcription rates and the newly discovered importance of 5hmC in neuronal gene expression.

Results

Characterization of the FMR1 locus in post-mortem brain samples

A total of 18 post-mortem brain tissue samples were evaluated in this study. Based on genetic assessment of the FMR1 repeat size, we divided samples into three groups: full-mutation carriers (FM), pre-mutation carriers (PM), and unaffected individuals with no detectable expansion mutation (CTL); each group consisted of samples from six individuals (Table 1). To assess the presence and size of the FMR1 repeat expansion mutation in our study groups, we performed a CGG repeat-primed PCR followed by capillary electrophoresis (PCR/CE) assay for all brain samples used in the study (Supplementary Material, Fig. S1). Data output from one representative sample for each experimental group is shown in Figure 1. A detailed description and data interpretation of this method have been reported elsewhere (39). To confirm that the FMR1 repeat size corresponded to levels of FMR1 expression in our samples, we assessed FMR1 transcription levels by QPCR. As expected, we found FMR1 expression to be significantly reduced in the FM group and a trend of decreased FMR1 antisense transcript levels compared to the CTL group (Fig. 2). Individuals with pre-mutations had higher expression levels of both FMR1 sense and antisense transcripts as compared to unaffected individuals, which is consistent with the premutation causing FMR1 overexpression in FXTAS (12). Our data show that all samples corresponded with clinical diagnosis, with three exceptions: (1) Patient 1031-09 LZ, despite possessing an expansion beyond 200 repeats (Supplementary Material, Fig. S1), showed normal expression of FMR1 (Fig. 2). (2) While clinically classified as FXTAS, our PCR/CE analyses indicate the presence of both a PM and FM for patient 5006 (Supplementary Material, Fig. S1). (3) While patient 4751 was initially diagnosed with FXS, the clinical report indicates this patient possessed only 88 CGG repeats (as determined by Southern blot) and our own analysis using PCR/CE shows the presence of a PM (Fig. 1 andSupplementary Material, Fig. S1). Thus, genetic analyses indicate that patient 4751 is more similar to FXTAS individuals while the clinical presentation was consistent with FXS. Since somatic repeat instability, FMR1 methylation mosaicism and X chromosome inactivation in females can lead to differences between clinical diagnosis and the FMR1 genotype in a particular tissue sample, we have stratified the cohort according the repeat size, as determined by repeat primed PCR analysis of DNA extracted from post mortem brain tissues. In all cases, our repeat primed PCR assessment confirmed the repeat size as determined by Southern blot analysis (Supplementary Material, Table S1).

Table 1.

Demographic data for human post-mortem brain tissue samples

Sample ID Group Age (years) PMI (h) Sex Brain region Source
1790 CTL 13 18 Male OC NICHD
8265 CTL 52 18 Male PFC HBB
13574 CTL 53 21 Male PFC HBB
13295 CTL 56 22 Male PFC HBB
10028 CTL 36 22 Male PFC HBB
8288 CTL 57 22 Male PFC HBB
5319 FM 71 17 Male OC NICHD
1061-09 JB FM 64 30 Female OC MIND
1031-08 GP FM 58 20 Male OC MIND
1033-08 WS FM 79 18 Male OC MIND
1031-09 LZ FM 65 12 Male OC MIND
1018-10 RH FM 60 54 Male OC MIND
4751 PM 21 5 Male OC NICHD
4555 PM 80 12 Male OC NICHD
4664 PM 71 3 Male OC NICHD
5006 PM 85 5 Male PC NICHD
5212 PM 80 12 Male PC NICHD
1005-06 CB PM 55 NO INFO Male OC MIND
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PMI: post mortem interval; CTL: healthy control; FM: full-mutation; PM: pre-mutation; OC: occipital cortex; PFC: prefrontal cortex; PC: parietal cortex; NICHD: National Institute of Child Health and Human Development; HBB: Harvard Brain Bank; MIND: Medical Investigation of Neurodevelopmental Disorders Institute.

Figure 1.

Figure 1.

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Assessment of the FMR1 expansion mutation. Representative electropherograms of CGG repeat-primed PCR products. (Top trace) A large 320 bp peak indicates the presence of an unexpanded allele. (Middle trace) Premutation alleles produce a saw-tooth amplification pattern with the largest PCR products ranging from 320–820 bp. (Bottom trace) Amplification of fully expanded FMR1 alleles yields large PCR products represented by a peak beyond 820 bp.

Figure 2.

Figure 2.

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Expression of FMR1 in clinical samples. Levels of FMR1 sense (A) and antisense (B) mRNAs in post-mortem brain samples of individuals with full-mutation (FM), pre-mutation (PM) and healthy controls (CTL) were assessed by qPCR. Relative quantification (RQ) of FMR1 normalized to the GAPDH endogenous control is shown; error bars are SEM (*P < 0.05). The data point representing an outlier FM sample ‘LZ’, found to express FMR1 at normal levels, is indicated.

Site-specific hydroxymethylcytosine assessment of the FMR1 promoter in post-mortem brain tissues reveals enrichment in full-mutation samples

Enrichment of 5mC within the FMR1 promoter, relative to unaffected controls, is a characteristic feature in FXS, but not FXTAS (40). However, previous studies used a standard bisulfite conversion method to examine levels of 5mC, which cannot disambiguate 5mC from 5hmC (38). Given the recent discovery of 5hmC as a demethylation intermediate associated with active transcription (29,30), it has become necessary to distinguish between these two epigenetic marks using improved methodologies. Here, we used 5mC and 5hmC sensitive restriction digest followed by PCR amplification to quantify 5mC and 5hmC levels at the FMR1 locus (see Materials and Methods for experimental details). A total of six MspI/HpaII restriction sites located both upstream and downstream of the FMR1 TSS and CGG repeat were interrogated. The genomic location of each site, and distance from the FMR1 TSS in base pairs (bp) is illustrated in Figure 3. We observed significantly higher 5mC levels across all six restriction sites in FM brain tissues when compared to the PM and CTL samples (Fig. 4A and Supplementary Material, Fig. S2A). On the other hand, 5hmC levels were significantly elevated at the −431, −344, −44 and +667 bp restriction sites, but not those spanning the CGG repeat expansion, +6 and +359 bp (Fig. 4B and Supplementary Material, Fig. S2B). Mean 5mC and 5hmC abundances across all six restriction sites for each clinical subgroup are plotted in Fig. 4C. We found elevated levels of 5hmC in all FM samples except one coded 1061-09 JB (Supplementary Material, Fig. S2B). Since this outlier is the only female patient in our study, and therefore the only subject with two FMR1 alleles, we suggest that such low levels of 5hmC could result from X-inactivation as discussed below.

Figure 3.

Figure 3.

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The FMR1 locus. Filled circles represent CpG dinucleotides within the FMR1 promoter region plotted to scale as the distance in base pairs from the TSS + 1. The MspI/HpaII restriction sites located at –431, –344, −44, +6, +359 and +667 bp where methyl and hydroxymethylcytosine assessment was performed are indicated with open circles. A grey shaded region within the FMR1 promoter indicates the location of 22 CpG dinucleotides interrogated by methyl and hydroxymethylcytosine sensitive pyrosequencing analysis. The location of the FMR1 CGG repeat sequence from downstream of the TSS is also indicated.

Figure 4.

Figure 4.

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Locus-specific FMR1 DNA methylation and hydroxymethylation assessment in clinical samples. The MspI/HpaII restriction digest assay was used to assess 5mC (A) and 5hmC (B) at six sites within the FMR1 promoter across three experimental groups. Grey dashed line indicates SEM, (*P < 0.05, **P < 0.005, ***P < 0.00005). Mean abundances across all six restriction sites for FM, PM and CTL patient brain samples are shown as relative percentages (C).

Targeted hydroxymethycytosine pyrosequencing of the FMR1 promoter confirms enrichment in full-mutation samples

To confirm our findings, we utilized a second method of 5hmC and 5mC assessment to interrogate 22 CpG dinucleotides from –294 to –155 bp, relative to the FMR1 TSS, within the FMR1 promoter region (Fig. 3). Genomic DNA from 5 FM, 1 PM and 1 CTL post-mortem brain tissue samples was subjected to TET-assisted bisulfite pyrosequencing (TAB-seq) to quantify levels of 5mC and 5hmC. Our results confirmed that 5hmC is enriched at the FMR1 promoter in FM samples (Fig. 5). Interestingly, we found that while 5mC levels were uniform across the CpG sites of FM samples, 5hmC levels were relatively more varied among CpGs (Supplementary Material, Fig. S3).

Figure 5.

Figure 5

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Locus-specific pyrosequencing of the FMR1 promoter in FXS clinical samples. (A) 5mC and (B) 5hmC levels were quantified by TET-assisted bisulfite pyrosequencing (TAB-seq) at 22 CpG dinucleotides within the FMR1 promoter, plotted relative to their position (X axis) in base pairs from the FMR1 TSS. The relative abundance in FM patient samples (n = 5) is shown as mean values (Y axis), where grey dashed lines indicate SEM. For reference, one PM and one healthy control sample (CTL) was included in the analysis. (C) Mean abundance of 5hmC, 5mC and 5C across all 22 CpGs are plotted as relative percentages for each sample.

Cell-type specific 5-hydroxymethylcytosine assessment of the FMR1 promoter reveals neuronal enrichment in FXS

In light of previous reports showing genome-wide enrichment of 5hmC in neurons when compared to glia (41), we next explored the cell-type specificity of 5mC and 5hmC distribution levels at the FMR1 promoter in a representative FM post-mortem brain tissue sample (FXS patient 5319). In order to isolate neuronal and non-neuronal DNA, we collected nuclei from frozen post-mortem tissues then labeled the neuronal nuclei with an anti NeuN antibody conjugated to a fluorescent dye. Genomic DNA was then isolated following fluorescence-activated cell sorting (FACS). Our data indicate that the sorting process enriches nuclei to nearly 100% purity (Supplementary Material, Fig. S4). Due to the large amount of tissue required for the nuclei sorting technique (∼700 mg), it was not possible to evaluate the entire patient group. We performed site-specific 5mC/5hmC assessment using the MspI/HpaII restriction PCR method to interrogate, unsorted brain, NeuN+ and NeuN− genomic DNA fractions (Fig. 6A). The neuronal marker NeuN used for sorting is a general neuronal marker localized to the nuclear envelope of most classes of neurons, including nearly all cortical neurons used in the present study, with few exceptions (42,43). While NeuN-nuclei could arise from alternative cell types including blood or connective tissue, they are presumed to be glia due to their overwhelming majority of glia relative to these other cell types in brain tissue. Our epigenetic analysis was focused on two Msp1/HpaII restriction sites located at −431 and −344 bp due to the elevated 5hmC levels observed in patient brain samples at these sites and their location within the FMR1 promoter, which is hypothesized to confer a dominant role in transcriptional regulation. We found higher levels of 5hmC and lower levels of 5mC at the FMR1 promoter of neurons when compared to glia at the −431 and −344 bp MspI/HpaII restriction sites (Fig. 6B and C, respectively). These results indicate 5hmC enrichment at the FM FMR1 promoter of FXS patients is specific to neurons, which may reflect the more pronounced DNA demethylation capability of these cells.

Figure 6.

Figure 6

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Cell-type specific DNA methylation and hydroxymethylation assessment of the FMR1 promoter in post-mortem brain tissues. (A) Illustration of the nuclei sorting strategy used to isolate glial (NeuN−) and neuronal (NeuN+) DNA fractions from a full-mutation FXS post-mortem brain tissue sample (5319). Mean abundance of 5hmC, 5mC and 5C at two sites located −431 bp (B) and −344 bp (C) from the FMR1 TSS are shown.

Enrichment of 5-hydroxymethylcytosine at the FMR1 promoter is absent in cellular models of FXS

To determine if FMR1 5hmC enrichment that we observed in post-mortem FXS brain samples is recapitulated in cellular models of disease, we interrogated genomic DNA isolated from various patient derived cell types using the site-specific 5mC/5hmC MspI/HpaII restriction PCR method. We confirmed the presence of full mutations and FMR1 transcriptional silencing in all FXS cell lines using repeat-primed PCR/EC (Supplementary Material, Fig. S1) and QPCR (Supplementary Material, Fig. S5), respectively. Somatic patient tissue derived cells included two FXS fibroblast (GM09497 and GM05848) and one immortalized FXS lymphocyte (GM09237) cell lines. While 5mC levels were elevated in fibroblast and lymphocyte FXS patient cells when compared to healthy controls, we did not observe 5hmC enrichment at the FMR1 promoter (Fig. 8A and B). To examine more disease-relevant cellular models of FXS, we generated induced pluripotent stem cells (iPSCs) using cellular reprogramming of fibroblast and lymphocyte cell lines then differentiated them into neurons. In addition, we derived neurons from a well-characterized FXS human embryonic stem cell (hESC) line (WCMC-37). To confirm neuronal maturity, we used immunolabeling of multiple neuronal markers including the vesicular glutamate transporter 1, beta-III tubulin, microtubule-associated protein 2, synapsin 1, neurofilament heavy and NeuN (Fig. 7). Neither FXS iPSC nor hESC derived neurons showed 5hmC enrichment at the FMR1 promoter when compared to controls (Fig. 8C and D). To eliminate the possibility that cell-cycle progression prevents 5hmC enrichment at the FMR1 promoter, we used thymidine to block the cell cycle. Thymidine treatment had no effect on 5hmC abundance at the FMR1 promoter in FXS fibroblasts, lymphocytes or iPSC-derived neurons (Supplementary Material, Fig. S6A and B). To eliminate the possibility that incomplete neuronal differentiation could explain the lack of FMR1 5hmC enrichment in iPSC and ES-derived FXS neurons, we performed a time-course study where DNA was extracted after 2, 4, 6 and 8 weeks of terminal neuronal differentiation (Supplementary Material, Fig. S6C and D). Our results demonstrate that even mature neurons, terminally differentiated for 8 weeks do not recapitulate FMR1 5hmC enrichment that we observed in FM FXS post-mortem brain tissues. Lastly, to determine if a reduced TET expression could explain the lack of FMR1 5hmC promoter enrichment in neuronal cell lines, we performed quantitative PCR (qPCR) for three TET genes in FXS iPSC and ES derived neurons. We compared TET expression to levels expressed in FM, PM and CTL patient brain samples (Supplementary Material, Fig. S7). We found no significant difference in TET expression between primary FM neurons and stem cell derived neurons. Interestingly, we observed elevated levels of TET expression in PM patient samples as compared to FM and unaffected controls. Global 5hmC disturbances in a mouse model of FXTAS was reported previously (44) and further studies are warranted to investigate a potential role of TET dysregulation in FXTAS patients.

Figure 8.

Figure 8

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Site-specific FMR1 DNA methylation and hydroxymethylation assessment in cellular models of FXS. Relative percentage of 5C, 5mC and 5hmC at two restriction sites located −431 (A and C) and −344 (B and D) bp from the FMR1 transcriptional start site. Somatic patient derived cell lines (A and B) include control fibroblasts (CTL FB), two FXS fibroblast lines (FXS FB 1, 2) and FXS lymphocytes. Neuronal lines were derived from three FXS induced pluripotent stem cells (FXS iPSC N 1–3), one FXS embryonic stem cell line (FXS ES N) and a control iPSC cell line (C and D).

Figure 7.

Figure 7

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Characterization of FXS neuronal cell lines. Immunostaining of FXS iPSC-neurons for the neuronal markers VGLUT1, TUJ1, MAP2, SYN1, NFH and NeuN (A–F), scale bar = 20 μm. Cells were counterstained with DAPI to reveal nuclei (blue).

Discussion

Using two independent methods, we show that 5hmC is enriched at the FMR1 promoter in primary FXS neurons with FMR1 full mutations but not stem cell derived neurons or non-neuronal cell lines. Levels of FMR1 5hmC were more variable than 5mC across CpG dinucleotides within the FMR1 promoter. In addition, we found 5hmC both upstream and downstream of the FMR1 TSS, but not at sites proximal to the CGG repeat sequence itself. These findings indicate the repeat expansion may impede TET-mediated demethylation at the FMR1 locus. In a previous study, we found similar 5hmC enrichment of the C9ORF72 promoter, a gene that harbors a repeat expansion mutation responsible for one genetic form of amyotrophic lateral sclerosis (45). Others have found 5hmC enrichment near the GAA repeat expansion mutation within the Frataxin gene, which is responsible for Friedreich’s ataxia (46). Notably, disruption of DNA methylation has been demonstrated for all three of these expansion mutations (22,47,48). Thus there is evidence to suggest that local perturbations of DNA methylation caused by repeat expansion mutations involve DNA demethylation as a common epigenetic feature as well.

While it is well established that epigenetic silencing of FMR1 FM alleles leads to FXS, the mechanism remains incompletely defined and cannot fully explain rare and variable phenotypes that are observed in patient populations. For example, males with FXS typically possess FM with hypermethylated FMR1 promoters (+CpGme) that are transcriptionally silent (−mRNA) (+FXS, +FM, +CpGme, −mRNA). However, exceptions to this rule exist: (1) In about 15% of individuals—often with a mild phenotype—both PM and FMs are detectable, rendering them mosaic for expansion size (+/− FXS, ±/−FM, +/−CpGme, +/−mRNA) (49–51). In the current study, we identified one individual (5006) that fit this scenario, and showed mixed representation of PM and FM FMR1 expansions. (2) In about half of male FXS individuals, FMR1 mRNA is detected despite the presence of hypermethylated FMs (+FXS, +FM, ±CpGme, ±mRNA) (1). In this case, it has been suggested that CpGme and mRNA production are uncoupled due to unknown differences in the pattern of CpGme between cells, or ‘cryptic inter-cell mosaicism’ (52). We saw that one individual 1031-09 LZ fit with this scenario demonstrating FMR1 transcriptional activity despite being methylated and having a FM. Notably, this individual had the highest levels of 5hmC and lowest levels of 5mC at the FMR1 promoter amongst FXS patient group (Fig. 5C and Supplementary Material, Fig. S3). (3) Extremely rare individuals have been identified who possess unmethylated transcriptionally active FMs and present with a mild or normal phenotype (−FXS, +FM, −CpGme, +mRNA) (17,25,53). While we did not identify individuals with such condition, future studies could examine whether such uncoupling of FMs from CpGme, could be due to active DNA demethylation mediated by TET enzymes. One potential mechanism could be that 5hmC enrichment may recruitment proteins that increase transcriptional permissiveness (54). Alternatively, the enrichment may reduce the affinity of repressive methylated DNA binding proteins that maintain FMR1 heterochromatinization (55,56). Finally, it is tempting to speculate that 5hmC enrichment could alter the binding affinity of CTCF, a critical DNA binding protein implicated in FMR1 silencing (57,58). Interestingly, 5hmC levels of a female FXS patient (1061-09 JB) were lowest in the group and we hypothesize it is a consequence of random inactivation of X chromosomes in females. Since X-inactivation is known to cause global DNA methylation at the X chromosome (59), it is likely that the full-mutation FMR1 allele is inactivated in a portion of neurons within the tissue sample and is therefore not accessible for TET-mediated DNA demethylation. Additionally, 5mC levels were highest in this patient amongst FM group further supporting our reasoning. These findings advance our knowledge of the composite epigenetic organization of expanded FMR1 alleles in FXS leading us to postulate active DNA demethylation at the FMR1 promoter occurs in FXS patients in vivo and that this process could modulate FMR1 expression rates and contribute to the observed variability of clinical phenotype.

For one clinical FXS post-mortem FM brain sample, of which there was sufficient material, we compared isogenic neurons and glia in order to examine neuron-specific aspects of FMR1 DNA methylation and hydroxymethylation. An investigation of the FMR1 epigenetic status specifically in neurons is warranted for multiple reasons. First, the primary affected cell type in FXS is neurons where FMRP deficiency alters synaptic function and morphology (6). Second, 5hmC levels are enriched in neurons (26,31), where TET expression is higher than other cell types including glia (60). Thus, neurons may be uniquely capable of reversing the hypermethylation of FMR1 that is associated with full mutations. Our data indicate that for one FXS patient, 5hmC enrichment was unique to neurons. One explanation is that DNA replication and subsequent reacquisition of DNA methylation, which occurs in glial cells—but not terminally differentiated, non-proliferative neurons—is known to affect epigenetic status of FMR1 gene (61–63). As far as we are aware, this is the first study to perform cell-type specific epigenetic assessment using FXS patient post-mortem brain tissue. Future studies are warranted in order to determine the applicability of this finding to broader clinical populations.

Large GC-rich sequences, such as the FMR1 repeat expansion, are difficult to model using standard molecular genetic approaches. Therefore, most studies have relied on post-mortem tissues or patient-derived cell lines such as primary fibroblasts or immortalized lymphocytes to investigate FMR1 repeats in their native genetic environment. Recently, more relevant neuronal model systems derived from hESCs (64–66) and iPSCs (67,68) have been employed and offer substantial advantages over other in vitro model systems. While certain FXS phenotypes are mirrored in iPSC-derived neurons, the epigenetic features of the FMR1 repeat in this model has been a source of controversy (67,68). In the current study, we sought to determine whether cellular models of FXS recapitulate 5mC and 5hmC levels found in primary FXS neurons isolated from clinical post-mortem brain tissue. We determined that FXS patient primary fibroblasts, immortalized lymphocytes, ES-neurons, and iPSC-neurons do not recapitulate 5hmC enrichment at the FMR1 promoter. Neither cell-cycle progression, time of neuronal differentiation nor expression of TET enzymes could explain this lack of 5hmC enrichment. These findings suggest that cellular models of FXS are not representative with regard to DNA demethylation of the FMR1 promoter. This is particularly important for future therapeutic strategies targeting epigenetic pathways to reactivate the locus. While reactivation of the locus has been achieved using the nucleoside analog 5-aza 2′-deoxycitidine (69,70), future studies could examine the potential role of 5hmC in FMR1 reactivation as well as potentially impeding FMR1 silencing. In summary, we have established a novel epigenetic perturbation associated with the FMR1 FM, its neuronal-specific abundance, and absence in widely used FXS cellular models.

Materials and Methods

Clinical samples and cell lines

FXS and FXTAS patient post-mortem brain tissues were obtained from the UC-Davis MIND Institute and University of Maryland Brain and Tissue Bank; healthy control (CTL) post-mortem brain tissues were provided by the Harvard Brain Tissue Resource Center at McLean Hospital. Primary FXS fibroblasts (GM09497, GM05848) and lymphocytes (GM09237) were ordered from Coriell Institute Biorepository. The FXS embryonic stem cell line (WCMC-37, NIH registration #0211) was obtained from the Joan & Sanford I. Weill Medical College of Cornell University with approval from the Embryonic Stem Cell Research Oversight Committee (ESCRO) at the University of Miami. Transfer of all tissues and cell lines was performed under IRB approved protocols at the University of Miami. DNA was extracted from 30 mg of brain tissue or 5 × 106 cells in culture using QIAamp DNA mini kit (Qiagen) as per manufacturer instructions.

Quantitative real-time PCR

Isolation of RNA was performed using Trizol® reagent for cell lysis, followed by addition of chloroform to facilitate phase separation. RNAs within the aqueous phase were then purified using RNeasy® columns (Qiagen) with on-column DNase treatment according to the manufacturer’s protocol. Reverse transcription was performed using the qSCRIPT kit (Quanta Biosciences) with random priming. The cDNA was amplified to quantify TET1, TET2, TET3, FMR1 sense and antisense RNAs using TaqMan® master mix (Life Technologies) with primer/probe sets: TET1 (Hs00286756_m1), TET2 (Hs00325999_m1), TET3 (Hs00379125_m1), FMR1 (Hs00924547_m1) and FMR1-AS (Hs03680973_g1) transcripts. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) expression was used as an endogenous control and relative expression levels were calculated using the ΔΔCt method.

AmplideX FMR1 repeat primed PCR and capillary electrophoresis

Repeat size was assessed using a repeat primed PCR followed by capillary electrophoresis (CE) as reported previously (39). Briefly, 40 ng of genomic DNA was amplified using the Asuragen Amplidex® PCR/CE kit containing FMR1-FAM labeled primers, according to manufacturer’s manual. The amplified product was then subjected to denaturation and addition of Hi-Di formamide and ROX-1007 ladder. The denatured PCR products were visualized using ABI3730 DNA Analyzer and GeneMapper software. Genotyping followed the American College of Medical Genetics guidelines (71) as follows: <45 CGG repeats were categorized as unexpanded controls, while 55–200 CGG repeats were considered pre-mutation and >200 CGG repeats as full mutation alleles.

DNA methylation assessment of FMR1 promoter

The EpiMark® 5-hmC and 5-mC analysis kit (New England Biolabs Inc., #E3317) was used for relative quantification of 5hmC and 5mC as we previously reported (45). Briefly, the kit distinguishes 5mC from 5hmC by the glycosylation of 5hmC residues using T4 β-glucosyltransferase, which blocks MspI endonuclease activity at CCGG sequences but not HpaII activity. Following the digest, qPCR was performed to amplify the region containing the MspI/HpaII restriction site and relative quantification values were used to calculate the abundance of 5hmC and 5mC. Genomic DNA (2 µg) was initially glycosylated and digested with MspI or HpaII endonucleases for 4–6 h at 37 °C. QPCR was performed using two primer sets spanning each MspI/HpaII restriction sites located at −431, −344, −44, +6, +359 and +667 bp relative to the FMR1 transcriptional start site. Primer sequences were as follows: −431 bp (F/R: 5′CC TAT TCTCGCCTTCCACTC/5′GGTCTGGTTTGGTTTGGTTTG); −344 bp (F/R: 5′AGAGGCCGAACTGGGATAA/5′GCCA GAACGCCCATTTCT); −44 bp (F/R: 5′GGAGGGAACAGCGTTGA/5′CTGA CTGAGG CCGA ACC); +6 bp (F/R: 5′GTGACGTGGTTTCAGTGTTTAC/5′CTCC ACC GG AAGTGAAACC); +359 bp (F/R: 5′CCCTTCCTTCCCTCCCTTT/5′CCAA GTCCAGTCCTTCCCT); +667 bp (F/R: 5′CCGAA ATCGG CG C TAAGT/5′CAACTACCCACACGACAGG). Relative 5hmC and 5mC levels were calculated according to manufacturer’s protocol.

TET-assisted bisulfite pyrosequencing (TAB-seq)

To distinguish 5mC and 5hmC nucleotides, we used TAB-seq, which uses glucose protection of 5hmC residues, TET-mediated conversion of 5mC residues and bisulfite conversion (72). Genomic DNA (2 µg) was subjected to glycosylation reaction followed by TET enzyme treatment. Afterwards, bisulfite conversion and pyrosequencing was performed by EpigenDx Inc. to assess FMR1 promoter methylation levels. The analysis was performed using the PSQTM96HS system and custom designed primers provided by EpigenDx (ADS1451FS1 and ADS1451FS2). The assays cover 22 CpG dinucleotides within the promoter from −294 to +155 relative to the transcriptional start site of the human genomic sequence.

Nuclei sorting

Neuronal nuclei were extracted according to fluorescence-activated sorting protocol adapted from Jiang et al. (73). Nuclei were dounced in hypotonic lysis solution and purified by 2.5-h centrifugation at 107 000 rcf at 4 °C. The nuclei pellet was then resuspended in 300 µl PBS, 100 µl of blocking mix (containing 0.10% normal goat serum and 0.5% BSA), and 1.2 µl of an Alexa 488 conjugated anti-NeuN antibody (Millipore MAB377X). Samples were incubated for 45 min with rotation in the dark at 4 °C. Nuclei were then separated using a BD FACS Aria-IIu flow cytometric cell sorter. The NeuN positive (neuronal) and negative (non-neuronal) nuclei fractions were pelleted by centrifugation at 1800 rcf for 15 min at 4 °C and resuspended in douncing buffer (10 mM Tris, pH 7.5; 4 mM MgCl2; and 1 mM CaCl2).

Neuronal differentiation

Induced pluripotent stem cells were derived from primary patient fibroblast, or immortalized lymphocytes using previously published protocol (45,74). The iPSCs and hESCs were directed towards neural stem cells by culturing in ultra-low attachment flasks with Neurobasal media (Life technologies) supplemented with 2% B27, 100 ng/ml βFGF, 100 ng/ml EGF and 5 µg/ml heparin. For neuronal differentiation, neural precursor spheres were plated onto poly-l-ornithine/laminin coated plates in DMEM/F12 (Life Technologies) with 2% B27, 1% N2, 10 ng/ml BDNF, 10 ng/ml GDNF, 1 µM cAMP and 200 ng/ml ascorbic acid for a minimum of 30 days.

Immunocytochemistry

Neuronal cultures were fixed in 4% paraformaldehyde at room temperature for 10 min, blocked in antibody buffer with 20% goat serum, permeabilized with 0.2% Triton X-100TM and incubated in the presence of primary antibodies against beta-III-tubulin (TUJ1, Sigma Aldrich T8578), neurofilament heavy chain (NFH, Abcam ab4680), microtubule-associated protein 2 (MAP2, Abcam 5392), synapsin 1 (SYN1, Abcam ab8), NeuN (Millipore MAB77X) and vesicular glutamate transporter 1 (VGLUT1, Synaptic Systems 135302) overnight at 4 °C. The next day, samples were washed 3 times with PBS and incubated with secondary antibody (1:500) for 2 h. DAPI counterstaining was performed and images were acquired using confocal fluorescent microscopy.

Cell cycle arrest

Cells were arrested in S phase of the cell cycle using a standard single thymidine block, according to standard practice (74). Specifically, cells were treated with 2 mM of thymidine for 18 h prior to DNA extraction and DNA hydroxymethylation assessment.

Supplementary Material

Supplementary Material is available at HMG online.

Supplementary Material

Supplementary Data
Click here for additional data file. (4.2MB, zip)

Acknowledgements

We would like to thank Dr Flora Tassone form UC Davis MIND Institute for providing FXS brain samples.

Conflict of Interest statement. None declared.

Funding

This study was supported by the National Institute of Mental Health (grant R01 MH084880 to C.W.).

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Supplementary Materials

Supplementary Data
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