Genome-wide analysis validates aberrant methylation in fragile X syndrome is specific to the FMR1locus
- Reid S Alisch†2, 5,
- Tao Wang†1, 2,
- Pankaj Chopra2,
- Jeannie Visootsak2, 4,
- Karen N Conneely2 and
- Stephen T Warren2, 3, 4Email author
© Alisch et al.; licensee BioMed Central Ltd. 2013
Received: 23 October 2012
Accepted: 25 January 2013
Published: 29 January 2013
Fragile X syndrome (FXS) is a common form of inherited intellectual disability caused by an expansion of CGG repeats located in the 5′ untranslated region (UTR) of the FMR1 gene, which leads to hypermethylation and silencing of this locus. Although a dramatic increase in DNA methylation of the FMR1 full mutation allele is well documented, the extent to which these changes affect DNA methylation throughout the rest of the genome has gone unexplored.
Here we examined genome-wide methylation in both peripheral blood (N = 62) and induced pluripotent stem cells (iPSCs; N = 10) from FXS individuals and controls.
We not only found the expected significant DNA methylation differences in the FMR1 promoter and 5′ UTR, we also saw that these changes inverse in the FMR1 gene body. Importantly, we found no other differentially methylated loci throughout the remainder of the genome, indicating the aberrant methylation of FMR1 in FXS is locus-specific.
This study provides a comprehensive methylation profile of FXS and helps refine our understanding of the mechanisms behind FMR1 silencing.
KeywordsEpigenetics DNA methylation Fragile X syndrome
Individuals with fragile X syndrome (FXS) exhibit a broad range of phenotypes, including varying degrees of intellectual disability, social impairment, macroorchidism, and an elongated face with large, everted ears. The most common mutation that causes FXS is an expansion of the CGG trinucleotide repeat located within the 5′ untranslated region (UTR) of the FMR1 gene [1–3]. When this expansion is greater than 200 repeats (known as the full mutation), the FMR1 promoter becomes hypermethylated, which prevents the expression of FMR1. Deletions and sequence variants within FMR1 result in a very small fraction of FXS cases [4–6], arguing that the loss of FMR1 function is the cause of FXS.
An association between the hypermethylation of the FMR1 trinucleotide repeats and FXS was first recognized over two decades ago , which sparked intensive study of the DNA methylation dynamics within the FMR1 locus. Research into the developmental timing of FMR1 silencing in chorionic villi (CV) samples from FXS patients has revealed that the FXS full mutation alleles are still expressed during early embryogenesis (i.e. during gastrulation), indicating that epigenetic repression is established at a later developmental time point . Furthermore, human embryonic stem cells (hESCs) derived from FXS patient embryos also express FMR1 in undifferentiated cells, until cellular differentiation triggers the recruitment of specific histone modifications, followed by DNA methylation and subsequent silencing of FMR1 transcription . In contrast, the FMR1 locus remains hypermethylated in induced pluripotent stem (iPS) cell lines derived from FXS patients, suggesting that, once the methylation marks are established at this locus, they are stable and resistant to current reprograming methodologies .
A long-standing question in the field is whether the full mutation triggers DNA methylation elsewhere in the genome or only at the FMR1 locus. Resolving this question could modify theories of how an expanded CGG repeat triggers aberrant DNA hypermethylation. For example, RNA-induced transcriptional silencing (RITS) has been proposed as a mechanism to explain the silencing of FMR1. RITS is a form of gene silencing triggered by small interfering RNA (siRNA) and generally causes the transcriptional downregulation of a genomic region . This model is attractive in that the unmethylated full mutation allele is known to be expressed in early development, presumably producing a transcript with a long riboCGG tract, and this riboCGG tract is cleaved in vitro by Dicer, producing small siRNA-like fragments of the riboCGG tract. Thus, it may be that small CGG RNAs could target chromatin-modifying activities back to the FMR1 locus. If true, there could be other CGG tracts in the genome that also are modified by this mechanism. To test this hypothesis, we examined DNA methylation levels at nearly half a million sites throughout the genome in the peripheral blood and fibroblast iPS cells of FXS patients using a highly sensitive genome-wide assay that quantitates methylation level at single CpG dinucleotide resolution; our results show that the hypermethylation of the FMR1 locus in FXS is indeed locus-specific.
The study protocol and consent form used in this investigation wasreviewed and approved by the Emory Internal Review Board on August 3, 2012and given the approval number CR8_IRB00001764.
Derivation of iPS cells
Human normal fibroblasts CRL2097 were obtained from ATCC, and GM0011 (normal), GM05848, and GM07730 (fragile X patients) were obtained from the Coriell Cell Repositories. The fibroblasts were cultured in DMEM containing 10% FBS, 1× glutamine, 1× Non-Essential amino acids, and 1× Pen/Strep.
For human iPSC reprogramming, 1 × 105 fibroblasts were seeded in a well of a 6-well plate. The next day, concentrated pMXs-hOCT4, hSOX2, hKLF4, and c-hMYC retrovirus were added to cells in the presence of 6 μg/ml polybrene. A second round of transduction was repeated the following day. On day 7 after initial transduction, the cells were reseeded in 10-cm dishes with irradiated MEF feeders. The hiPSC colonies were picked between days 18–25. iPSCs were maintained in hiPSC standard medium (DMEM/F12, 20% KnockOut Serum Replacement, 1× MEM Non-Essential Amino Acids, 1× glutamine, 0.11 mM 2-mercaptoethanol, 10 ng/ml bFGF) on irradiated MEF feeders. The established iPSC cell lines were subsequently confirmed with AP staining, and pluripotent markers by immunofluorescence staining and the ability to differentiate into 3 germ layers. Before isolating genomic DNA, iPS cells were subcultured in mTeSR1 feeder free system (STEMCELL Technologies) for at least 3 passages to reduce potential contamination.
DNA methylation profiling
Five hundred nanograms of human genomic DNA was sodium bisulfite–treated for cytosine (C) to thymine (T) conversion using the EZ DNA Methylation-Gold kit (Zymo Research). The converted DNA was purified and prepped for analysis on the Illumina HumanMethylation450 BeadChips following the manufacturer’s guidelines. Briefly, converted DNA was amplified, fragmented, and hybridized to the HumanMethylation450 pool of allele-differentiating oligonucleotides. After a series of extension, ligation, and cleanup reactions, the DNA was labeled with a fluorescent dye. The labeled DNA was then scanned using an Illumina BeadArray Reader or iScan. Image analysis and signal determination were performed using the GenomeStudio software, Methylation Module (Illumina).
Interpretation and QC of DNA methylation data
CpG DNA methylation data were interpreted using GenomeStudio to quantify methylated (M) and unmethylated (U) signal intensities for genomic DNA. The signals were quantile normalized separately, and overall methylation levels (β) were calculated as the ratio of methylated to total signal [i.e. β = M/(M + U + 100)], where β ranges from 0 (unmethylated) to 1 (methylated). Quality control of data resulted in removal of samples with aberrantly low signal intensity (mean <2000) or with fewer than 90% of CpG loci detected, where a given locus was deemed not detected if the detection P-value was >0.01 (detection P-value provided by GenomeStudio and calculated relative to background signal). Any probe having more than 25% detected P-values >0.01 was discarded from the analysis. Missing data were imputed using the “impute.knn” function from the “impute” package in R (Cran). Assay controls were inspected to remove samples with poor bisulfite conversion, staining, extension (single nucleotide extension assay), hybridization, or specificity. Furthermore, outliers identified by hierarchical clustering and/or dissimilarity matrices were removed. Additionally, one control DNA replicate was run on each BeadChip to assess overall assay reproducibility. Methylation profiles of the control DNA correlated well, with an average Pearson correlation coefficient (R) of 0.990 between replicates.
Analysis of FXS-associated CpG loci
To analyze DNA methylation differences associated with FXS, we fit a separate regression for each CpG site. Although samples were randomly distributed across BeadChips and experiments with respect to disease, BeadChip was also included as a random effect covariate in all analyses to account for potential batch effects. The package “nlme” in R (Cran) was used for the mixed effect model. Fixed effects included in the model were intensity, position, and age (for blood). To test several FMR1-related and -unrelated hypotheses, we filtered the data to include only those probes that reside in the following genomic regions: 1500 base pairs of a transcription start site; 200 base pairs of a transcription start site; the first exon of a gene; the “3′UTR” of a gene; the “5′UTR” of a gene; a CpG Island; the “N_Shore” or “S_Shore” of a CpG island; or near a CGG trinucleotide repeat containing at least 8 consecutive repeats. We also filtered the data to include only those probes annotated to genes found to play a role in recurrent genomic abnormalities. To correct for multiple hypothesis testing, we applied a Benjamini-Hochberg False Discovery Rate (FDR) correction using the R function “p.adjust,” but to avoid false positives due to the small sample size, we used conservative Bonferroni adjustment for our ultimate determination of significance.
All permutation analyses were conducted in R using the same linear model as the actual analysis, where BeadChip was treated as a mixed-effects covariate, but in each permutation the disease status of the subjects was randomly reassigned. In total, 1000 permutations were conducted for both the peripheral blood and iPS cell groups independently. Permutation P-values for each CpG locus were calculated by assessing the number of times each locus was more significantly associated with FXS in the 1000 permuted data sets than the actual association (Additional file 1: Figure S3; Additional file 2: Table S1 and Additional file 3: Table S2).
We have submitted the data generated from the 9 FXS samples and the 53 controls for this study to the Gene Expression Omnibus (GEO), which can be found under the Gene Series: GSE41273.
We next tested several FXS-related and -unrelated hypotheses, including recurrent genomic abnormalities associated with intellectual delay and annotated genomic structures (e.g. CpG islands; Methods). Although this approach reduces our multiple testing burden and effectively increases our power to find subtle yet significant changes in DNA methylation, as before the only probes identified as differentially methylated were probes annotated to the FMR1 gene (data not shown). Note that even for the strict Bonferroni criteria employed for the genome-wide analysis, 75% of the CpG sites had >80% power to detect β – value differences of 0.10 or greater, so the lack of observed widespread methylation differences in genes other than FMR1 cannot be explained by low power. Finally, to gain insight into the potential mechanism(s) behind FXS-associated DNA methylation changes, we annotated the genome for all CGG trinucleotide repeats containing at least eight consecutive repeats (N = 136 tracts; N = 452 probes; see Methods) and found two juxtaposed probes annotated to ZFHX3 that reached significance (Additional file 6: Table S3). In contrast to the FMR1 locus, the ZFHX3 locus had no distinct FXS-associated DNA methylation or gene expression differences (Additional file 7: Figure S4; data not shown). These findings imply that the significant DNA methylation differences observed at the two probes do not have a functional consequence on ZFHX3 expression. Together, these data suggest that the FXS-associated hypermethylation of the FMR1 promoter is locus-specific and does not alter DNA methylation elsewhere in the genome.
To corroborate these findings, we derived a total of ten iPS cell lines from fibroblasts of two FXS patients (FXS-iPS) and two control individuals (iPS). DNA was extracted from 12 FXS-iPS cell lines and 11 iPS cell lines and epityped using Infinium HumanMethylation450 BeadChips. Limiting the FXS-associated DNA methylation analysis in this group to only those loci that satisfied an FDR <0.05 in the peripheral blood analysis (N = 1183 probes; Figure 1A; see Methods) yielded results similar to those found in peripheral blood: eight FXS-methylated loci and one FXS-demethylated locus; all nine differentially methylated loci are annotated to the FMR1 promoter (Bonferroni <0.05) (Figure 1C; Additional file 8: Table S4). Since iPS cells show significant reprogramming variability, we also excluded the hotspots of aberrant reprogramming regions reported by Lister et al.  and still found FXS-associated DNA methylation changes only at the FMR1 locus (data not shown). Subsequent hypothesis- and mechanism-driven analyses also failed to uncover any non-FMR1 annotated FXS-associated DNA methylation changes, including at the ZFHX3 locus. Therefore, FXS-derived iPS cells exhibit similar genome-wide methylation profiles as terminally differentiated cells of blood, a FMR1-specific epigenetic disruption.
This study provides a sensitive and comprehensive quantitative analysis of genome-wide DNA methylation levels in a group of FXS and control individuals. We found that FXS-associated hypermethylation is profound throughout the CpG island encompassing the FMR1 5′ UTR, revealing CpG dinucleotides whose distinct FXS methylation profile could improve current diagnostic methods. For example, there are four CpG loci in FMR1 that show a clear distinction between all FXS and control samples with no overlap (Figure 1B), suggesting that interrogation of these loci for methylation would be diagnostic. The finding that FXS-associated methylation is significantly decreased at one CpG in the FMR1 gene body is consistent with previous reports indicating that gene body hypermethylation is associated with active gene expression . It would be interesting to explore whether this trend persists throughout the remainder of the gene.
Our examination reported here shows that CGG repeats elsewhere in the genome do not appear abnormally methylated in trans with the full mutation. Thus, either the hypothesis of a RITS role in silencing of FMR1 is false, or there may be a threshold of length for a CGG repeat tract to be susceptible to silencing, since there are no known CGG tracts in the reference genome that even approach the size of a premutation, let alone a full mutation allele. Indeed, the expression of the normal FMR1 allele opposite the full mutation in FXS females would be consistent with a threshold model.
When FMR1 was first identified, the question posed in this study was unanswerable. Today, our knowledge of the human genome sequence allows genome-wide examination of DNA methylation differences. Here we report that only probes located in the FMR1 promoter or gene body exhibit FXS-associated DNA methylation differences in DNA from peripheral blood and iPS cells of FXS individuals. Thus, while this study does not determine the mechanism behind the aberrant methylation in the expanded FMR1 repeat, it does help refine our mechanistic picture of FMR1 silencing in fragile X syndrome. Since we did not find any non-FMR1-associated differentially methylated loci, we have made a significant stride toward finally put a long-standing question in FXS research to rest.
Fragile X syndrome
Human embryonic stem cells
Induced pluripotent stem
Pearson correlation coefficient
RNA-induced transcriptional silencing
Small interfering RNA.
The authors would like to thank Julie Mowrey and Brian Lynch for technical assistance and Cheryl Strauss for editorial comments. This work was supported in part by a Simons Foundation (SFARI) award and NIH grants MH089606 and HD24064, all to STW. We are grateful to all the families collected from the Department’s Fragile X Syndrome Clinic and those participating at the SFARI Simplex Collection (SSC) sites, as well as the principal investigators (A. Beaudet, R. Bernier, J. Constantino, E. Cook, E. Fombonne, D. Geschwind, D. Grice, A. Klin, D. Ledbetter, C. Lord, C. Martin, D. Martin, R. Maxim, J. Miles, O. Ousley, B. Peterson, J. Piggot, C. Saulnier, M. State, W. Stone, J. Sutcliffe, C. Walsh, and E. Wijsman). We also appreciate the access to phenotypic data on SFARI Base. Approved researchers can obtain the SSC population dataset described in this study by applying at https://base.sfari.org. Emory University’s research IT service center and its high performance computer cluster also supported this research.
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