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Epigenetic studies in Developmental Origins of Health and Disease: pitfalls and key considerations for study design and interpretation

Published online by Cambridge University Press:  09 September 2016

L. Yamada  and
S. Chong Open the ORCID record for S. Chong [Opens in a new window]
L. Yamada
Affiliation:
Epigenetics Group, Translational Research Institute, Mater Research Institute – The University of Queensland, Woolloongabba, QLD, Australia
S. Chong*
Affiliation:
Epigenetics Group, Translational Research Institute, Mater Research Institute – The University of Queensland, Woolloongabba, QLD, Australia
*
*Address for correspondence: Dr S. Chong, Epigenetics Group, Mater Research Institute – The University of Queensland, Translational Research Institute, Level 4, 37 Kent Street, Woolloongabba, QLD 4102, Australia. (Email Suyinn.chong@mater.uq.edu.au)
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Abstract

The field of Developmental Origins of Health and Disease (DOHaD) seeks to understand the relationships between early-life environmental exposures and long-term health and disease. Until recently, the molecular mechanisms underlying these phenomena were poorly understood; however, epigenetics has been proposed to bridge the gap between the environment and phenotype. Epigenetics involves the study of heritable changes in gene expression, which occur without changes to the underlying DNA sequence. Different types of epigenetic modifications include DNA methylation, post-translational histone modifications and non-coding RNAs. Increasingly, changes to the epigenome have been associated with early-life exposures in both humans and animal models, offering both an explanation for how the environment may programme long-term health, as well as molecular changes that could be developed as biomarkers of exposure and/or future disease. As such, epigenetic studies in DOHaD hold much promise; however, there are a number of factors which should be considered when designing and interpreting such studies. These include the impact of the genome on the epigenome, the tissue-specificity of epigenetic marks, the stability (or lack thereof) of epigenetic changes over time and the importance of associating epigenetic changes with changes in transcription or translation to demonstrate functional consequences. In this review, we discuss each of these key concepts and provide practical strategies to mitigate some common pitfalls with the aim of providing a useful guide for future epigenetic studies in DOHaD.

Keywords

DNA methylationenvironmental exposureshistone modificationsnon-coding RNAsstudy design
Type
Review
Information
Journal of Developmental Origins of Health and Disease , Volume 8 , Issue 1 , February 2017 , pp. 30 - 43
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This is an Open Access article, distributed under the terms of the Creative Commons Attribution licence (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted re-use, distribution, and reproduction in any medium, provided the original work is properly cited.
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© Cambridge University Press and the International Society for Developmental Origins of Health and Disease 2016

Introduction

Early-life environmental exposures are thought to influence organ development and physiology such that there is an increased risk of disease in later life.Reference Hyatt, Budge and Symonds 1 , Reference Luyckx and Brenner 2 Growing evidence suggests that early-life exposures can also impact the epigenome. Epigenetics has been defined in a number of waysReference Waddington 3 – 5 and the field has yet to reach a generally accepted consensus. For the purposes of this review, we define epigenetics as the study of heritable changes in gene expression, which occur without changes to the underlying DNA sequence.Reference Egger, Liang, Aparicio and Jones 6 Epigenetic marks have the capacity to be stably inherited through successive mitotic cell divisions, providing a possible molecular ‘memory’ of the exposure, and can be associated with altered gene expression, thereby affecting phenotype. As such, epigenetics has the potential to both further our understanding of the mechanisms which underlie the link between early-life exposures and later health outcomes, and to produce novel molecular biomarkers of past exposure and/or future disease.Reference Godfrey, Costello and Lillycrop 7 Here, we cover essential concepts for epigenetic studies in Developmental Origins of Health and Disease (DOHaD), identify potential hazards in study design and interpretation, and highlight strategies for the generation of informative and meaningful results.

Epigenetics

DNA methylation

Within mammals, cytosine methylation is the most well-characterized DNA modification. Cytosine methylation occurs most frequently within cytosine phosphate guanine (CpG) dinucleotides, with 70–80% of CpGs within the human genome methylated.Reference Bird 8 Non-CpG cytosine methylation also occurs but is tissue-specific, with higher levels reported in oocytes,Reference Tomizawa, Kobayashi and Watanabe 9 pluripotent cellsReference Ziller, Muller and Liao 10 and various regions of the brain.Reference Xie, Barr and Kim 11 – Reference Guo, Su and Shin 13

Cytosine methylation is catalyzed by the highly conserved DNA methyltransferase (DNMT) family of proteins.Reference Bestor 14 – Reference Goll and Bestor 16 DNMT1 has a higher affinity for hemi-methylated than unmethylated DNA and is responsible for propagating methylation after DNA replication, thus acting to maintain methylation states.Reference Bestor 14 DNMT3A and DNMT3B are essential for the establishment of new, or de novo methylation marks.Reference Okano, Bell, Haber and Li 15 Although DNMT3L lacks catalytic activity,Reference Hata, Okano, Lei and Li 17 it can bind to and stimulate the catalytic activity of DNMT3A and DNMT3B.Reference Suetake, Shinozaki, Miyagawa, Takeshima and Tajima 18 , Reference Gowher, Liebert, Hermann, Xu and Jeltsch 19

Until recently, little was known about how DNA was demethylated. Several mechanisms had been proposed for the active demethylation of DNA, including those involving DNA deamination by methyl-CpG-binding domain protein 4 (MBD4) and glycosylationReference Zhu, Zheng and Angliker 20 , Reference Rai, Huggins and James 21 and cytosine deamination by DNMTs.Reference Metivier, Gallais and Tiffoche 22 , Reference Kangaspeska, Stride and Metivier 23 There is a growing body of literature supporting another mechanism mediated by the Ten-Eleven Translocation family of proteins, which sequentially hydroxylate 5-methylcytosine to 5-hydroxymethylcytosine, 5-formylcytosine and finally to 5-carboxylcytosine.Reference Ito, Shen and Dai 24 – Reference Pfaffeneder, Hackner and Truss 26 Through thymine DNA glycosylation followed by base excision repair, 5-carboxylcytosine is then converted back to the unmodified cytosine.Reference He, Li and Li 25

Passive, replication-dependent DNA demethylation can also occur.Reference Bhutani, Burns and Blau 27 In the early preimplantation mouse embryo, it was observed that chromosome methylation was iteratively lost with each cycle of DNA replication.Reference Rougier, Bourc’his and Gomes 28 This followed an earlier observation that DNMT1 was excluded from the cell nucleus in the very early stages of embryogenesis following fertilization, and only observable in the nucleus from the eight-cell stage.Reference Carlson, Page and Bestor 29 , Reference Howell, Bestor and Ding 30 The absence of a maintenance DNMT from the nucleus would result in a passive reduction in global methylation state with every cell division.

CpG-rich regions, called CpG islands, are often found at the 5' promoter region of genes,Reference Esteller 31 and methylation of these regions is associated with transcriptional silencing.Reference Stein, Razin and Cedar 32 , Reference Bird 33 Promoter methylation is believed to prevent transcriptional initiation.Reference Schubeler, Lorincz and Cimbora 34 In contrast, intragenic CpG methylation in mammalian cells has little effect on transcriptional initiation, instead discouraging transcriptional elongation.Reference Lorincz, Dickerson, Schmitt and Groudine 35 Intragenic methylation can also aid in exon recognition, playing a role in the regulation of alternative splicing.Reference Gelfman, Cohen, Yearim and Ast 36 , Reference Maunakea, Chepelev, Cui and Zhao 37

DNA methylation appears to be responsive to the environment, with alterations in DNA methylation patterns reported in both humans and animals following a range of adverse early-life exposures, including those of malnutrition, alcohol, choline and arsenic.Reference Tobi, Lumey and Talens 38 – Reference Silver, Kessler and Hennig 44 The DOHaD field has historically focused on promoter DNA methylation; however, the use of unbiased genome-wide screens for DNA methylation has identified associations between early-life exposures and methylation of non-promoter regions such as enhancersReference Tobi, Goeman and Monajemi 45 as well as intergenic regions.Reference Amarasekera, Martino and Ashley 46 , Reference Thompson, Fazzari and Niu 47 Mechanistically, there is some evidence that early-life exposures to alcohol and choline deficiency in rodents can alter the expression of the maintenance methyltransferase Dnmt1.Reference Bekdash, Zhang and Sarkar 41 , Reference Kovacheva, Mellott and Davison 48 , Reference Gardebjer, Anderson, Pantaleon, Wlodek and Moritz 49 One-carbon metabolism is a network of pathways involved in a number of functions, including the synthesis of methionine which can subsequently be adenosylated to S-adenosyl methionine – a major source of methyl groups necessary for DNA methylation.Reference Stover 50 One-carbon metabolism may also mediate environmentally induced changes to DNA methylation as it can be perturbed by early-life exposures to alcoholReference Ngai, Sulistyoningrum and O’Neill 51 and maternal smokingReference Drake, O’Shaughnessy and Bhattacharya 52 as well as by alterations in gestational maternal intake of methyl donors such as choline and folate.Reference Kovacheva, Mellott and Davison 48 , Reference Garro, McBeth, Lima and Lieber 53 – Reference McKay, Waltham, Williams and Mathers 55 Disrupting either Dnmt1 levels or one-carbon metabolism would be expected to impact DNA methylation genome-wide. Although global alterations to DNA methylation have been reported following certain exposures,Reference McKay, Waltham, Williams and Mathers 55 many exposures fail to induce such changes, instead resulting in locus-specific effects.Reference Tobi, Goeman and Monajemi 45 , Reference Kovacheva, Mellott and Davison 48 , Reference Breton, Siegmund and Joubert 56 How perturbations to Dnmt1 expression or one-carbon metabolism could induce locus-specific methylation changes remains unclear, and further study is required to understand the mechanisms by which the environment influences the methylome.

Post-translational histone modifications

In the nucleus, DNA is packaged into chromatin, the individual building blocks of which are nucleosomes. Within the nucleosome, DNA is wrapped around a protein octamer, comprising of two each of histone H2A, histone H2B, histone H3 and histone H4.Reference Cheung, Allis and Sassone-Corsi 57 Although the C-terminal domains of histones are critical for the maintenance of nucleosome structure,Reference Cheung, Allis and Sassone-Corsi 57 the N-terminal tails function to alter the accessibility of the associated DNA.Reference Luger, Mader, Richmond, Sargent and Richmond 58 The N-terminal tail of any histone can, at specific amino acid positions, undergo chemical modifications including acetylation, methylation,Reference Lanzuolo and Orlando 59 phosphorylation,Reference Paulson and Taylor 60 ubiquitinylation,Reference Davie and Murphy 61 carbonylation,Reference Wondrak, Cervantes-Laurean, Jacobson and Jacobson 62 poly(ADP-ribosyl)ationReference Poirier, de Murcia, Jongstra-Bilen, Niedergang and Mandel 63 or sumoylation.Reference Shiio and Eisenman 64 These modifications are thought to alter chromatin structure by affecting electrostatic interactions between the DNA and histones, making them either more, or less tightly packaged and permissive of transcription. In addition, modified histones can be recognized by and directly interact with various proteins which can further modify the histones and/or affect chromatin structure.Reference Bannister and Kouzarides 65 For example, the recognition of histone H3 lysine 4 trimethylation (H3K4me3) by inhibitor of growth 2 results in the recruitment and stabilization of the mSin3a-histone deacetylase 1 (HDAC1) complex at the gene promoter.Reference Shi, Hong and Walter 66 In contrast, SWItch/sucrose non-fermentable is a chromatin remodeling complex which recognizes acetylated histones.Reference Hassan, Prochasson and Neely 67

The histone code hypothesis posits that the combinatorial identity and position of each N-terminal tail modification acts as a code, controlling transcription in a highly specific manner.Reference Strahl and Allis 68 Individually, H3K4me3 at gene promoters is associated with transcriptional activation,Reference Vermeulen, Mulder and Denissov 69 whereas trimethylation of lysine 27 at histone H3 (H3K27me3) is associated with transcriptional repression.Reference Cao, Wang and Wang 70 However, bivalent domains that have both activating (H3K4me3) and repressive (H3K27me3) marks simultaneously also exist.Reference Bernstein, Mikkelsen and Xie 71 Occurring near gene promoters, bivalent domains are thought to poise genes for expression.Reference Bernstein, Mikkelsen and Xie 71

Though epigenetic studies in DOHaD have primarily focused on DNA methylation, post-translational histone modifications are also subject to the influence of the early-life environment.Reference Bekdash, Zhang and Sarkar 41 , Reference Stevens, Begum and Cook 72 – Reference Zinkhan, Fu and Wang 74 The mechanism by which this occurs; however, is yet to be fully understood. Gestational choline deficiency has been identified to alter the expression of genes involved in the conferral of histone modifications, including the histone lysine methyltransferase Set domain bifurcated 1 (Setdb1) and histone methyltransferase G9a (Kmt1c).Reference Bekdash, Zhang and Sarkar 41 , Reference Davison, Mellott, Kovacheva and Blusztajn 75 One-carbon metabolism may also be involved as S-adenosyl methionine, which in addition to being required for DNA methylation, also contributes to the post-translational methylation of histone tails.Reference Stover 50 Given that early-life exposures can influence one-carbon metabolism,Reference Kovacheva, Mellott and Davison 48 , Reference Ngai, Sulistyoningrum and O’Neill 51 – Reference McKay, Waltham, Williams and Mathers 55 there is potential for these exposures to have consequences on histone as well as DNA methylation but, again, widespread changes might be expected. In support of this, global changes to histone methylation have been reported in rodents following various early-life exposures, including those of gestational choline deficiencyReference Mehedint, Niculescu, Craciunescu and Zeisel 76 and nicotine exposure.Reference Suter, Abramovici and Griffin 77 In contrast, locus-specific effects were observed when H3K4me3 was assayed by chromatin immunoprecipitation and next generation sequencing in the dentate gyrus of inbred C57BL/6 mice following an early-life exposure to arsenic.Reference Tyler, Weber and Labrecque 78 Further, a maternal high-fat diet produced coding region-specific changes in histone H3 lysine 9 trimethylation at the rat Wingless-type MMTV integration site family member 1 (Wnt1) gene in offspring liver,Reference Yang, Cai, Xu and Shi 79 suggesting the presence of mechanisms which allow for conferral of locus- and region-specificity. Therefore, while it is evident that changes in post-translational histone modifications are associated with early-life exposures, further study is required to elucidate both how this occurs and how it impacts offspring health.

Non-coding RNAs

Non-coding RNAs can also affect gene expression, either by transcriptional or post-transcriptional mechanisms. Long non-coding RNAs influence gene expression using a wide array of mechanisms.Reference Wang and Chang 80 For example, the Antisense Igf2r RNA (Air) long non-coding RNA accumulates at the Slc22a3 promoter and recruits the histone H3K9 methyltransferase G9a protein, thereby inducing locus-specific transcriptional repression in the placenta.Reference Nagano, Mitchell and Sanz 81 For a more detailed discussion on the various mechanisms by which long non-coding RNAs influence transcription, we direct the reader to the review by Wang et al.Reference Wang and Chang 80

Small non-coding RNAs can interact with nascent transcripts as well as with single- and double-stranded DNA in a sequence-specific manner.Reference Bonasio, Tu and Reinberg 82 The major categories of small non-coding RNAs include endogenous short interfering RNAs (which are presently poorly characterized in mammalsReference Kim, Han and Siomi 83 ), P-element induced wimpy testis-interacting RNAs (PIWI-interacting RNAs or piRNAs; expressed primarily in germ cellsReference Iwasaki, Siomi and Siomi 84 ), and microRNAs (miRNAs) (relatively well characterized and expressed in many tissuesReference Landgraf, Rusu and Sheridan 85 ).Reference Huang, Zhang and Yu 86 Of these small non-coding RNAs, miRNAs have been the subject of particular interest within the DOHaD field and as such, the remainder of this discussion will focus on miRNAs.

miRNAs are ~22 nucleotides in length, and bind to the 3' untranslated region (UTR) of target messenger RNAs (mRNA) in order to post-transcriptionally regulate their stability and/or translation into protein.Reference Bartel 87 miRNAs regulate target mRNA levels by cleavageReference Yekta, Shih and Bartel 88 or degradation.Reference Bagga, Bracht and Hunter 89 Although some studies have reported exclusive effects of miRNAs on protein translation,Reference Olsen and Ambros 90 , Reference Pillai, Bhattacharyya and Artus 91 others have reported instances whereby miRNAs first inhibit protein translation, and are then subsequently involved in mRNA deadenylation and decay.Reference Djuranovic, Nahvi and Green 92 The extent of base-pair complementarity between the miRNA and target mRNA can influence whether a given miRNAs inhibits or aids translation.Reference Saraiya, Li and Wang 93 Further, Let-7 and a synthetic miRNA (miRcxcr4) were identified to upregulate translation at certain points during the cell cycle, but at other times, the same miRNAs repressed translation of the same target reporter construct.Reference Vasudevan, Tong and Steitz 94

miRNAs were previously estimated to regulate between 20 and 30% of human genes.Reference Lewis, Burge and Bartel 95 , Reference Xie, Lu and Kulbokas 96 However, since the initial estimates were generated, a large number of new miRNAs have been discovered, rendering the figures relatively conservative. Each miRNA has been estimated to target 100–200 mRNAs, with miRNAs likely to act coordinately to aid in the regulation of any given target gene.Reference Brennecke, Stark, Russell and Cohen 97 , Reference Krek, Grun and Poy 98

An increasing number of studies within the DOHaD field are reporting changes in the expression of both long non-coding RNAsReference Laufer, Mantha and Kleiber 42 and miRNAsReference Wang, Zhang and Li 99 – Reference Zhang, Ho, Vega, Burne and Chong 101 following an early-life exposure. A number of studies have also begun to identify circulating miRNAs, in plasma and serum, as potential biomarkers of various early-life exposures.Reference Zhang, Ho, Vega, Burne and Chong 101 – Reference Vrijens, Bollati and Nawrot 104 Interest in non-coding RNAs in the context of DOHaD is relatively recent, and consequently little is known about either the mechanisms by which they are regulated or the downstream functional consequences.

Interactions between epigenetic modalities

In DOHaD studies, the epigenetic modalities of DNA methylation, histone modifications and non-coding RNAs are often considered in isolation, but there is substantial evidence that they regulate gene expression in concert with each other.

DNA methylation and histone modifications

The co-dependent nature of DNA methylation and histone modifications was nicely demonstrated when Zhang et al.Reference Zhang, Fatima and Dufau 105 observed that in order to achieve complete demethylation and activity of the luteinizing hormone receptor promoter in vitro, the addition of both a histone deacetylase inhibitor (trichostatin A) and a DNA demethylating reagent (5-azacytidine) were required. These results built on earlier findings in which the binding of methyl-CpG-binding protein 2 (MeCP2) to methylated DNA was shown to recruit histone deacetylases to impact locally on histone acetylation and chromatin structure.Reference Nan, Ng and Johnson 106 – Reference Eden, Hashimshony, Keshet, Cedar and Thorne 108

Likewise, both histone modifications themselves as well as the proteins responsible for conferring histone modifications can impact DNA methylation. For example, protein arginine methyltransferase 5 (PRMT5) confers symmetric methylation of arginine 3 at histone H4 (H4R3me2s) which then acts as a binding target for DNMT3A.Reference Zhao, Rank and Tan 109 Oocytes deficient in a H3K4 demethylase (KDM1B) exhibited genome-wide DNA hypomethylation, suggesting a critical role of H3K4 demethylation in DNA methylation regulation; however, the exact mechanism for this has not yet been elucidated.Reference Stewart, Veselovska and Kim 110 The histone methyltransferase Enhancer of Zeste homolog 2 (EZH2) directly recruits DNMTs,Reference Vire, Brenner and Deplus 111 and DNMT3L can bind to histone H3 when its lysine 4 is unmethylated, inducing de novo DNA methylation by DNMT3A2.Reference Ooi, Qiu and Bernstein 112

For further information on the complex interplay between DNA methylation and histone modifications, we direct the reader to several reviews.Reference Cedar and Bergman 113 – Reference Du, Johnson, Jacobsen and Patel 115

DNA methylation and non-coding RNAs

DNA methylation can regulate the expression of non-coding RNAs including miRNAs.Reference Han, Witmer, Casey, Valle and Sukumar 116 In turn, DNA methylation itself can also be influenced by various types of non-coding RNAs. DNA methylation can be directed in a sequence-specific manner through the direct interaction of DNMTs with long non-coding RNAs, including Tsix (the antisense transcript of Xist) and numerous promoter-associated non-coding RNAs.Reference Sun, Deaton and Lee 117 – Reference Di Ruscio, Ebralidze and Benoukraf 121 The imprinted H19 non-coding RNA also indirectly regulates the activity of DNMT3B by binding to S-adenosylhomocysteine hydrylase, thereby interfering with the hydrolysis of S-adenyosylhomocysteine – an inhibitor of DNMT3B.Reference Zhou, Yang and Zhong 122

Short non-coding RNAs can also influence DNA methylation. When MitoPLD, a protein involved in primary piRNA synthesis was mutated in mice, the de novo DNA methylation of the RAS protein-specific guanine nucleotide-releasing factor 1 (Rasgrf1) differentially methylated region (DMR) in spermatogonia was impaired, suggesting a role for piRNAs in de novo DNA methylation.Reference Watanabe, Tomizawa and Mitsuya 123 Similarly, the PIWI proteins MILI and MIWI2, which interact with piRNAs, were essential for the establishment of de novo methylation of retrotransposons in male fetal germ cells.Reference Kuramochi-Miyagawa, Watanabe and Gotoh 124 A number of miRNAs, which include miR-148a and miR-152, are able to directly target the expression of Dnmt1.Reference Braconi, Huang and Patel 125 , Reference Huang, Wang, Guo and Sun 126

Histone modifications and non-coding RNAs

Although histone modifications can regulate the expression of non-coding RNAsReference Barski, Jothi and Cuddapah 127 , Reference Wu, Kallin and Zhang 128 , non-coding RNAs themselves are also capable of directing histone modifications.Reference Guil and Esteller 129 , Reference Peschansky and Wahlestedt 130 The long non-coding RNA, HOTAIR, facilitates the conferral of histone modifications to the Homeobox D cluster by acting as a scaffold for both the polycomb repressive complex 2 (PRC2) and the Lysine-specific demethylase 1 (LSD1)/coRepressor element-1 silencing transcription factor (coREST)/RE1-silencing transcription factor (REST) complex, which in turn recruit enzymes to trimethylate histone H3 lysine 27 and demethylate histone H3 lysine 4, respectively.Reference Rinn, Kertesz and Wang 131 , Reference Tsai, Manor and Wan 132 A number of histone modifying enzymes, including HDAC1Reference Noonan, Place and Pookot 133 and EZH2,Reference Wong and Tellam 134 have also been identified to be direct targets of miRNAs.

For further information regarding the interaction between non-coding RNAs and the other epigenetic modalities, we direct the reader to other reviews.Reference Guil and Esteller 129 , Reference Peschansky and Wahlestedt 130

Epigenetic reprogramming in mammals

There are two major developmental periods, preimplantation development and gametogenesis, when the epigenome is erased and reset genome-wide in a process called epigenetic reprogramming.Reference Messerschmidt, Knowles and Solter 135 – Reference Morgan, Santos, Green, Dean and Reik 138 It has been proposed that the epigenome is most susceptible to environmental exposures during these periods of epigenetic reprogramming.Reference Kaminen-Ahola, Ahola and Maga 39

In preimplantation development, DNA methylation changes include an initial global demethylation event post-fertilization,Reference Reik, Dean and Walter 139 in which paternally derived DNA undergoes active demethylation,Reference Oswald, Engemann and Lane 140 whereas maternally derived DNA undergoes replication-dependent passive demethylation.Reference Rougier, Bourc’his and Gomes 28 Methylation is then re-established de novo from implantation onwards (approximately gestational day 4.5 in the mouse), with somatic tissues becoming increasingly methylated.Reference Monk, Boubelik and Lehnert 141 This reprogramming of the epigenome in the preimplantation embryo is necessary to allow cells of the early embryo to achieve a state of pluripotency,Reference Bhutani, Brady and Damian 142 and to then set up distinct patterns of gene expression that are associated with differentiation and cell fate determination.

Imprinted genes, which are resistant to preimplantation epigenetic reprogramming, have been the subject of a number of studies in the DOHaD field.Reference Reik, Dean and Walter 139 Although most genes are expressed from both the maternally and paternally derived alleles (biallelic expression), imprinted genes are expressed monoallelically – that is, exclusively from either the maternally or paternally derived allele.Reference Reik, Dean and Walter 139 Imprinting is a known epigenetic process; many of these genes have well-characterized DMRs that are associated with and thought to control monoallelic expression. The finding that imprinted gene DMR methylation in somatic tissues can be altered by gestational environmental exposuresReference Liu, Balaraman, Wang, Nephew and Zhou 143 , Reference Chen, Ganguly and Rubbi 144 appears at odds with their resistance to preimplantation epigenetic reprogramming; however, it is possible that the exposure compromises this resistance. Interestingly, other studies have identified no changes in imprinted gene DMR methylation in response to gestational environmental perturbations, despite identifying changes in expression,Reference Ivanova, Chen, Segonds-Pichon, Ozanne and Kelsey 145 , Reference Radford, Isganaitis and Jimenez-Chillaron 146 leading to speculation that these expression changes are due to transcription-factor-mediated mechanisms rather than epigenetic mechanisms.Reference Radford, Isganaitis and Jimenez-Chillaron 146 Indeed, the idea that imprinted regions are of no greater importance than any other genomic region in the epigenetic response to early-life exposures has been extensively discussed in a recent review.Reference Lecomte, Youngson, Maloney and Morris 147

Later in development, a second major epigenetic reprogramming event occurs during gametogenesis.Reference Reik, Dean and Walter 139 This reprogramming event involves the removal of parent-of-origin DNA methylation from imprinted loci, allowing for the establishment of new sex-specific methylation patterns, such that the alleles are imprinted with either a maternal pattern (in oocytes) or a paternal pattern (in sperm). This erasure of parental imprints begins during the migration of primordial germ cells to the genital ridge (from approximately gestational day 9.5 to 11.5 in the mouse).Reference Kawasaki, Lee and Matsuzawa 148 In murine male germ cells, DNA remethylation occurs when these primordial germ cells become prospermatogonia (from approximately gestational day 13 in the mouse) and is completed by birth.Reference Sasaki and Matsui 149 In contrast, DNA remethylation in female germ cells does not commence until after birth, occurring during preovulatory oocyte growth and maturation.Reference Messerschmidt, Knowles and Solter 135 , Reference Lucifero, Mann, Bartolomei and Trasler 150 Given that the timing of DNA remethylation is sexually dimorphic in nature, sensitivity to environmental exposures may also differ between sperm and oocytes. Furthermore, gametogenesis may be a period during which imprinted genes may be most sensitive to exposures. Specifically, differences in imprinting may be most likely to occur in offspring derived from males exposed during late gestation or offspring derived from females exposed in the preconceptional period (during oocyte maturation).

Given the difficulty of obtaining germ cell and/or preimplantation embryo samples from humans, much of the knowledge pertaining to epigenetic reprogramming has been obtained using murine models. Nonetheless, a small number of studies utilizing human samples have found that many of these processes appear to be conserved between species.Reference Geuns, De Rycke, Van Steirteghem and Liebaers 151 , Reference Okae, Chiba and Hiura 152

Mammalian epigenetic reprogramming is a complex phenomenon, and while a very brief overview is provided here, we direct the reader to more comprehensive reviews for further information.Reference Messerschmidt, Knowles and Solter 135 – Reference Morgan, Santos, Green, Dean and Reik 138

Considerations for epigenetic studies in DOHaD

As much promise as epigenetics has in unraveling the molecular mechanisms underlying adverse health outcomes following early-life environmental exposures, there are a number of challenges when conducting these studies. In this section, we discuss the central challenges in epigenetic studies in both humans and animal models, how they may impact the interpretation of results and highlight strategies to mitigate some of these issues (summarized in Tables 1 and 2).

Table 1 Strategies to address common challenges in epigenetic studies

Table 2 A selection of recent DOHaD publications that have addressed two or more of the challenges highlighted in this review. Black boxes denote the challenge addressed by the study

The epigenome is influenced by the genome

In addition to the extrinsic influence of the environment, the epigenetic landscape is also shaped intrinsically by the underlying DNA sequence. For example, the comparison of adolescent and middle-aged monozygotic and dizygotic twins suggested a greater contribution of genetics than environmental factors to DNA methylation at the imprinted Insulin-like growth factor 2 (IGF2) DMR.Reference Heijmans, Kremer, Tobi, Boomsma and Slagboom 153 Genetic influences on DNA methylation have also been reported genome-wide. When two well-characterized inbred mouse strains, C57BL/6 and BALB/c, with several hundred differentially methylated loci were mated, the F1 hybrid (C57BL/6xBALB/c) offspring exhibited strain-specific methylation patterns on each allele, likely driven by the local genomic context in cis.Reference Schilling, El Chartouni and Rehli 154

In humans, widespread associations between single nucleotide polymorphisms and DNA methylation have been reported in methylation quantitative trait loci studies.Reference Zhi, Aslibekyan and Irvin 155 – Reference McClay, Shabalin and Dozmorov 157 The underlying mechanisms as to how these genetic variants influence methylation remain poorly understood; however it has been proposed that the creation or disruption of CpG sites,Reference Zhi, Aslibekyan and Irvin 155 or perturbations in transcription factor bindingReference Banovich, Lan and McVicker 156 may be involved. Another way in which the genome can influence the epigenome is via functional mutations within genes which contribute to the establishment or maintenance of epigenetic marks. For example, the 5, 10-methylenetetrahydrofolate reductase (MTHFR) gene encodes an enzyme critical for the supply of methyl donors for reactions such as DNA methylation. The C677T polymorphism within the human MTHFR gene results in reduced MTHFR activityReference Frosst, Blom and Milos 158 and is associated with genomic DNA hypomethylation in peripheral leucocytes.Reference Stern, Mason, Selhub and Choi 159 , Reference Friso, Choi and Girelli 160 Similarly, the R271Q polymorphism within the DNMT3L gene in humans is another example of a polymorphism resulting in DNA hypomethylation.Reference El-Maarri, Kareta and Mikeska 161 There is limited literature to suggest that histone modifications too may be dependent on the local genomic context.Reference Kadota, Yang and Hu 162 Consequently, in DOHaD studies utilizing genetically heterogeneous populations such as humans or outbred animals, it can be difficult to distinguish between epigenetic changes driven by the environment and those driven by genetic differences, unrelated to the environment.

In both humans and outbred animals, genetic differences can be accounted for, if not controlled for, by assaying genotype.Reference Kadota, Yang and Hu 162 – Reference Zhang, Cheng and Badner 165 In rodents, genetic differences can be minimized through the use of inbred strains. A recent study identified few genetic or epigenetic differences of gross magnitude between C57BL/6 littermates.Reference Oey, Isbel, Hickey, Ebaid and Whitelaw 166 However, even this approach is not without problems. Using inbred mice, Shea et al.Reference Shea, Serra and Carone 167 recently identified variation in DNA methylation at ribosomal DNA repeats in C57BL/6 sperm, which seemingly correlated with paternal diet. Upon further analysis, the difference in methylation was found to be an artifact of copy number variation at this repetitive element, and unrelated to paternal diet.Reference Shea, Serra and Carone 167 Taken together, the evidence supports the importance of considering the contribution of genetic variation to epigenetic variation, even in inbred animals.

Epigenetic modifications are cell- and tissue-specific

Although each individual tends to have minimal genetic variation across tissues, the epigenome is reflective of the different and often dynamic transcriptional identities of tissues, and even individual cells. As such, any given individual, despite having only one genome, can have numerous epigenomes.Reference Varley, Gertz and Bowling 12 , Reference Ohgane, Aikawa and Ogura 168 – Reference Leung, Jung and Rajagopal 173 When the methylomes of six regions from the brain as well as that of whole blood were compared in nine human donors by immunoprecipitation of methylated DNA followed by next generation sequencing, greater differences were observed between tissues within an individual than between the same tissue across individuals.Reference Davies, Volta and Pidsley 171 Even within tissues, cell-to-cell variation in DNA methylation has been reported.Reference Flanagan, Popendikyte and Pozdniakovaite 174 , Reference Farlik, Sheffield and Nuzzo 175

Similarly, when epigenetic changes are identified in one tissue following an early-life exposure, comparable changes are not assured in other tissues. Maternal smoking during pregnancy was associated with hypomethylation of the Aryl hydrocarbon receptor repressor (AHRR) gene in newborn cord blood mononuclear cells, but not in buccal epithelium or placental tissue.Reference Novakovic, Ryan and Pereira 176 This example of a tissue-specific response is not an isolated case, with methylation of long interspersed element 1 (LINE1) repeats, measured as a proxy for global DNA methylation, hypomethylated in the hypothalamus but not the striatum in offspring of a mouse model of prenatal maternal immune activation.Reference Basil, Li and Dempster 177 Furthermore, Kundakovic et al.Reference Kundakovic, Gudsnuk and Franks 178 observed an increase in DNA methylation within the estrogen receptor 1 (Esr1) gene in the prefrontal cortex, but not hypothalamus of male offspring from an inbred BALB/c mouse model of prenatal bisphenol A exposure. In the offspring from an ovine model of maternal undernutrition, despite the differential methylation of the glucocorticoid receptor gene in various regions of the brain, no evidence of differential methylation of this gene was identified in leucocytes.Reference Begum, Davies and Stevens 179 Therefore, caution is recommended particularly when inferring the epigenetic state of a disease-relevant, but inaccessible tissue based on the epigenetic state of another more accessible tissue.

The relatively accessible nature of whole blood makes it a commonly assayed tissue in many human studies of DNA methylation following an early-life environmental exposure.Reference Breton, Siegmund and Joubert 56 , Reference El Hajj, Pliushch and Schneider 180 , Reference Cardenas, Koestler and Houseman 181 There is however, a growing body of literature reporting considerable epigenetic heterogeneity within whole blood, reflective of its diverse cellular composition.Reference Jacoby, Gohrbandt, Clausse, Brons and Muller 182 , Reference Reinius, Acevedo and Joerink 183 When cellular composition was corrected for in silico in five published studies examining age-related DNA methylation changes, cellular composition explained a greater proportion of the reported epigenetic variation in these studies than did age.Reference Jaffe and Irizarry 184 Subsequently, Bauer et al.Reference Bauer, Linsel and Fink 185 found that a previously reported association between tobacco smoking and DNA methylation at the G protein-coupled receptor 15 (GPR15) locus was in fact an artifact of increased numbers of CD3+ T-cells in the smoking population. Therefore, any study of epigenetic changes in whole blood or white blood cells should consider the cellular composition of these samples.

There are a number of approaches available to reduce the confounding influence of cellular heterogeneity. One solution is to sort samples into individual cell types; however, this will significantly reduce the amount of tissue available for analysis. As an alternative to this, a number of computational methods have been developed to estimate and account for differences in cellular distributions in heterogeneous tissues such as blood and brain, utilizing previously defined DNA methylation signatures for each cell type.Reference Houseman, Accomando and Koestler 186 – Reference Yousefi, Huen and Quach 190 These methods have been applied in some studies examining DNA methylation following early-life exposures.Reference Kile, Houseman and Baccarelli 43 , Reference Silver, Kessler and Hennig 44 , Reference Cardenas, Koestler and Houseman 181 Future DOHaD studies will undoubtedly continue to assay blood and other heterogeneous tissues; however, the consideration of cellular heterogeneity should aid the meaningful interpretation of any epigenetic changes identified.

Epigenetic modifications are not necessarily stable over time

In addition to being influenced by genetics and cellular identity, the epigenome may also change with time. Global DNA methylation in the livers of male C57BL/6 mice was found to gradually decline from the ages of 6–24 months,Reference Singhal, Mays-Hoopes and Eichhorn 191 suggesting that aging can alter methylation profiles. A similar study of DNA methylation differences in lymphocytes between monozygotic twins ranging in age from 3 to 74 found that while younger twins had relatively few epigenetic differences, the magnitude of difference within twin pairs increased with age across multiple tissues.Reference Fraga, Ballestar and Paz 192 In this study, the contribution of genetic heterogeneity to epigenetic variation was reduced through the utilization of monozygotic twins.Reference Fraga, Ballestar and Paz 192 Similar trends in methylation differences were also observed in four separate tissue types, suggesting that the impact of cellular composition on the epigenetic profiles was minimal.Reference Fraga, Ballestar and Paz 192 It is possible, even likely, that in addition to aging, the observed methylation changes reflect differences in postnatal exposures between twins over life. Regardless of the underlying causes, these studies draw attention to the idea that DNA methylation patterns can change over time.

The stability of epigenetic modifications over time also appears to be gene-dependent. When methylation of DNA isolated from saliva samples from adolescent monozygotic twin pairs were assayed before and after a period of several months, Levesque et al.Reference Levesque, Casey and Szyf 193 found that even in this relatively short period of time, the methylation of 46 genes was unstable, whereas 226 genes were identified to be temporally stable. Another study reported similar outcomes, with DNA methylation at five of eight candidate loci studied longitudinally over 11–20 years deemed stable in both whole blood and buccal cells.Reference Talens, Boomsma and Tobi 194 In this study, the authors were able to account for genetic heterogeneity by assaying for sequence variation, and for cellular heterogeneity using computational methods.Reference Talens, Boomsma and Tobi 194

Most DOHaD studies assay for epigenetic changes in samples collected at a single time-point,Reference Tobi, Lumey and Talens 38 , Reference Downing, Johnson and Larson 40 – Reference Kile, Houseman and Baccarelli 43 , Reference Tobi, Goeman and Monajemi 45 – Reference Thompson, Fazzari and Niu 47 , often far removed from the environmental exposure itself. The evidence presented above highlights the importance of assaying for epigenetic changes on multiple occasions, and demonstrating stability over time, especially if the epigenetic marks are proposed to confer a memory of the exposure or to serve as biomarkers.

The relationship between epigenetics and gene expression can be ambiguous

Although DNA methylation patterns are often linked with transcriptional activity, this is not always the case. Indeed, in eight human tissues, 5'UTR DNA methylation status was inversely correlated with transcription for only 37% of the 43 genes analyzed.Reference Eckhardt, Lewin and Cortese 195 Similarly, a paternal low-protein diet was associated with considerable changes in both mRNA and miRNA expression as well as DNA methylation in offspring livers; however, the genes at which promoter methylation was altered were not necessarily those which displayed differential expression.Reference Carone, Fauquier and Habib 196 In a separate study, the differential methylation of 181 gene promoters in pancreatic islets from patients with type 2 diabetes and healthy controls correlated with altered transcription for only 18% of the genes.Reference Volkmar, Dedeurwaerder and Cunha 197 It must be acknowledged that an epigenetic effect on adjacent locus expression at a different time-point cannot be excluded, nor an effect on the expression of other, more distantly located, genomic loci. In these instances however, such outcomes would need to be demonstrated experimentally.

A disconnect between transcriptional activity and DNA methylation has been observed at imprinted loci as well. The differential expression, but not DNA methylation, of a number of imprinted genes was identified in the livers of offspring exposed to gestational protein restriction.Reference Ivanova, Chen, Segonds-Pichon, Ozanne and Kelsey 145 Further, when altered expression, but not methylation of the imprinted paternally expressed 3 (Peg3) gene was observed in a mouse model of maternal undernutrition, the authors speculated that this was likely due to transcription factor-mediated mechanisms, rather than epigenetic mechanisms per se.Reference Radford, Isganaitis and Jimenez-Chillaron 146 It is however possible that other epigenetic marks, such as post-translational histone modifications, may still be contributing to these outcomes, even in the absence of changes to DNA methylation.

The finding that mRNA and protein levels are not always positively correlated,Reference Zhang, Ho, Vega, Burne and Chong 101 , Reference Chen, Gharib and Huang 198 , Reference Ghazalpour, Bennett and Petyuk 199 suggests that protein analyses may also be informative in epigenetic studies in DOHaD. The lack of correlation between mRNA and protein levels can potentially be explained by an epigenetic mechanism: miRNAs. In a model of gestational nutrient restriction, 23 miRNAs associated with the insulin-signaling pathway were identified to be differentially expressed in the liver of fetal lambs.Reference Lie, Morrison and Williams-Wyss 200 The expression of these miRNAs were then found to correlate with target protein but not mRNA levels,Reference Lie, Morrison and Williams-Wyss 200 suggesting that the miRNAs act on target protein translation rather than mRNA stability.

Our recent study which assayed DNA methylation, histone modifications and miRNA expression, as well as both mRNA and protein levels illustrates the informative potential of broader experimental designs. In this study, hippocampal tissue was assayed from adult male C57BL/6J mice following an early gestational ethanol exposure.Reference Zhang, Ho, Vega, Burne and Chong 101 Both a reduction in DNA methylation and increase in H3K4me3 (a marker of active chromatin) were observed at the promoter region of a vesicular glutamate transporter gene, Slc17a6, in ethanol-exposed mice. As would be predicted, these epigenetic changes correlated with an increase in Slc17a6 mRNA levels. However, when we assayed for the protein encoded by Slc17a6, there was a reduction in protein.Reference Zhang, Ho, Vega, Burne and Chong 101 We identified miR-467b-5p, a miRNA predicted in silico to target Slc17a6, to be differentially expressed in the same tissue, and experimentally validated this interaction using in vitro reporter assays. In this study, transcriptional output (mRNA) correlated with both promoter DNA methylation and histone modifications, following which at least one miRNA was proposed to regulate expression at a translational level, demonstrating that an early-life environmental exposure can exert complex, independent effects on gene expression.Reference Zhang, Ho, Vega, Burne and Chong 101

Finally, it is often difficult to discern whether epigenetic marks, such as DNA methylation and post-translational histone modifications which are associated with gene expression changes following an early-life environmental exposure, are a cause or a consequence of the change in transcription. As a result, the relationship between these epigenetic modifications and gene expression is often best described as correlative in nature.

Concluding remarks

The field of DOHaD has historically focused on characterizing the long-term health consequences of early-life environmental exposures. There is emerging evidence that these early-life exposures can affect the epigenome, which has the potential not only to impact gene expression and phenotype but may also be stably remembered for a lifetime. The assays available to investigate the epigenome are now well within the reach of all investigators. This review covers essential concepts in epigenetics which are relevant to the DOHaD field, and highlights potential pitfalls as well as key considerations for study design and interpretation. We look forward to an exciting new era of DOHaD studies which will bring us closer to understanding not only the impact of early-life environmental exposures on health and disease in later life, but also greater knowledge of the role of epigenetics in mediating such phenomena.

Acknowledgments

The authors would like to thank James Cuffe, Vicki Clifton and Neil Youngson for their thoughtful and constructive feedback during the preparation of this manuscript. The authors apologize to the authors of relevant studies that we were unable to discuss due to space constraints.

Financial Support

L.Y. was supported by an Australian Postgraduate Award and by Mater Research-UQ (Frank Clair Scholarship). S.C. was supported by an Australian Research Council Future Fellowship (FT100100333). This work was also supported by funding from the Mater Foundation. The Translational Research Institute is supported by a grant from the Australian Government.

Conflicts of Interest

None.

References

1. Hyatt, MA, Budge, H, Symonds, ME. Early developmental influences on hepatic organogenesis. Organogenesis. 2008; 4, 170–175.Google Scholar
2. Luyckx, VA, Brenner, BM. The clinical importance of nephron mass. J Am Soc Nephrol. 2010; 21, 898–910.Google Scholar
3. Waddington, CH. The pupal contraction as an epigenetic crisis in Drosophila. Proc Zool Soc Lond A. 1942; 111, 181–188.Google Scholar
4. Wu, C, Morris, JR. Genes, genetics, and epigenetics: a correspondence. Science. 2001; 293, 1103–1105.Google Scholar
5. National Institutes of Health. Overview of the Roadmap Epigenomics Project. National Institutes of Health, 2010. Retrieved 20 June 2016 from http://www.roadmapepigenomics.org/overview.Google Scholar
6. Egger, G, Liang, G, Aparicio, A, Jones, PA. Epigenetics in human disease and prospects for epigenetic therapy. Nature. 2004; 429, 457–463.Google Scholar
7. Godfrey, KM, Costello, PM, Lillycrop, KA. The developmental environment, epigenetic biomarkers and long-term health. J Dev Orig Health Dis. 2015; 6, 399–406.CrossRefGoogle ScholarPubMed
8. Bird, A. DNA methylation patterns and epigenetic memory. Genes Dev. 2002; 16, 6–21.CrossRefGoogle ScholarPubMed
9. Tomizawa, S, Kobayashi, H, Watanabe, T, et al. Dynamic stage-specific changes in imprinted differentially methylated regions during early mammalian development and prevalence of non-CpG methylation in oocytes. Development. 2011; 138, 811–820.Google Scholar
10. Ziller, MJ, Muller, F, Liao, J, et al. Genomic distribution and inter-sample variation of non-CpG methylation across human cell types. PLoS Genet. 2011; 7, e1002389.Google Scholar
11. Xie, W, Barr, CL, Kim, A, et al. Base-resolution analyses of sequence and parent-of-origin dependent DNA methylation in the mouse genome. Cell. 2012; 148, 816–831.Google Scholar
12. Varley, KE, Gertz, J, Bowling, KM, et al. Dynamic DNA methylation across diverse human cell lines and tissues. Genome Res. 2013; 23, 555–567.CrossRefGoogle ScholarPubMed
13. Guo, JU, Su, Y, Shin, JH, et al. Distribution, recognition and regulation of non-CpG methylation in the adult mammalian brain. Nat Neurosci. 2014; 17, 215–222.Google Scholar
14. Bestor, TH. Activation of mammalian DNA methyltransferase by cleavage of a Zn binding regulatory domain. EMBO J. 1992; 11, 2611–2617.Google Scholar
15. Okano, M, Bell, DW, Haber, DA, Li, E. DNA methyltransferases Dnmt3a and Dnmt3b are essential for de novo methylation and mammalian development. Cell. 1999; 99, 247–257.Google Scholar
16. Goll, MG, Bestor, TH. Eukaryotic cytosine methyltransferases. Annu Rev Biochem. 2005; 74, 481–514.Google Scholar
17. Hata, K, Okano, M, Lei, H, Li, E. Dnmt3L cooperates with the Dnmt3 family of de novo DNA methyltransferases to establish maternal imprints in mice. Development. 2002; 129, 1983–1993.CrossRefGoogle ScholarPubMed
18. Suetake, I, Shinozaki, F, Miyagawa, J, Takeshima, H, Tajima, S. DNMT3L stimulates the DNA methylation activity of Dnmt3a and Dnmt3b through a direct interaction. J Biol Chem. 2004; 279, 27816–27823.CrossRefGoogle ScholarPubMed
19. Gowher, H, Liebert, K, Hermann, A, Xu, G, Jeltsch, A. Mechanism of stimulation of catalytic activity of Dnmt3A and Dnmt3B DNA-(cytosine-C5)-methyltransferases by Dnmt3L. J Biol Chem. 2005; 280, 13341–13348.Google Scholar
20. Zhu, B, Zheng, Y, Angliker, H, et al. 5-Methylcytosine DNA glycosylase activity is also present in the human MBD4 (G/T mismatch glycosylase) and in a related avian sequence. Nucleic Acids Res. 2000; 28, 4157–4165.Google Scholar
21. Rai, K, Huggins, IJ, James, SR, et al. DNA demethylation in zebrafish involves the coupling of a deaminase, a glycosylase, and Gadd45. Cell. 2008; 135, 1201–1212.Google Scholar
22. Metivier, R, Gallais, R, Tiffoche, C, et al. Cyclical DNA methylation of a transcriptionally active promoter. Nature. 2008; 452, 45–50.CrossRefGoogle ScholarPubMed
23. Kangaspeska, S, Stride, B, Metivier, R, et al. Transient cyclical methylation of promoter DNA. Nature. 2008; 452, 112–115.Google Scholar
24. Ito, S, Shen, L, Dai, Q, et al. Tet proteins can convert 5-methylcytosine to 5-formylcytosine and 5-carboxylcytosine. Science. 2011; 333, 1300–1303.CrossRefGoogle ScholarPubMed
25. He, YF, Li, BZ, Li, Z, et al. Tet-mediated formation of 5-carboxylcytosine and its excision by TDG in mammalian DNA. Science. 2011; 333, 1303–1307.CrossRefGoogle ScholarPubMed
26. Pfaffeneder, T, Hackner, B, Truss, M, et al. The discovery of 5-formylcytosine in embryonic stem cell DNA. Angew Chem Int Ed Engl. 2011; 50, 7008–7012.Google Scholar
27. Bhutani, N, Burns, DM, Blau, HM. DNA demethylation dynamics. Cell. 2011; 146, 866–872.Google Scholar
28. Rougier, N, Bourc’his, D, Gomes, DM, et al. Chromosome methylation patterns during mammalian preimplantation development. Genes Dev. 1998; 12, 2108–2113.CrossRefGoogle ScholarPubMed
29. Carlson, LL, Page, AW, Bestor, TH. Properties and localization of DNA methyltransferase in preimplantation mouse embryos: implications for genomic imprinting. Genes Dev. 1992; 6, 2536–2541.Google Scholar
30. Howell, CY, Bestor, TH, Ding, F, et al. Genomic imprinting disrupted by a maternal effect mutation in the Dnmt1 gene. Cell. 2001; 104, 829–838.CrossRefGoogle ScholarPubMed
31. Esteller, M. Cancer epigenomics: DNA methylomes and histone-modification maps. Nat Rev Genet. 2007; 8, 286–298.CrossRefGoogle ScholarPubMed
32. Stein, R, Razin, A, Cedar, H. In vitro methylation of the hamster adenine phosphoribosyltransferase gene inhibits its expression in mouse L cells. Proc Natl Acad Sci U S A. 1982; 79, 3418–3422.Google Scholar
33. Bird, AP. CpG-rich islands and the function of DNA methylation. Nature. 1986; 321, 209–213.Google Scholar
34. Schubeler, D, Lorincz, MC, Cimbora, DM, et al. Genomic targeting of methylated DNA: influence of methylation on transcription, replication, chromatin structure, and histone acetylation. Mol Cell Biol. 2000; 20, 9103–9112.Google Scholar
35. Lorincz, MC, Dickerson, DR, Schmitt, M, Groudine, M. Intragenic DNA methylation alters chromatin structure and elongation efficiency in mammalian cells. Nat Struct Mol Biol. 2004; 11, 1068–1075.Google Scholar
36. Gelfman, S, Cohen, N, Yearim, A, Ast, G. DNA-methylation effect on cotranscriptional splicing is dependent on GC architecture of the exon-intron structure. Genome Res. 2013; 23, 789–799.Google Scholar
37. Maunakea, AK, Chepelev, I, Cui, K, Zhao, K. Intragenic DNA methylation modulates alternative splicing by recruiting MeCP2 to promote exon recognition. Cell Res. 2013; 23, 1256–1269.Google Scholar
38. Tobi, EW, Lumey, LH, Talens, RP, et al. DNA methylation differences after exposure to prenatal famine are common and timing- and sex-specific. Hum Mol Genet. 2009; 18, 4046–4053.Google Scholar
39. Kaminen-Ahola, N, Ahola, A, Maga, M, et al. Maternal ethanol consumption alters the epigenotype and the phenotype of offspring in a mouse model. PLoS Genet. 2010; 6, e1000811.Google Scholar
40. Downing, C, Johnson, TE, Larson, C, et al. Subtle decreases in DNA methylation and gene expression at the mouse Igf2 locus following prenatal alcohol exposure: effects of a methyl-supplemented diet. Alcohol. 2011; 45, 65–71.Google Scholar
41. Bekdash, RA, Zhang, C, Sarkar, DK. Gestational choline supplementation normalized fetal alcohol-induced alterations in histone modifications, DNA methylation, and proopiomelanocortin (POMC) gene expression in beta-endorphin-producing POMC neurons of the hypothalamus. Alcohol Clin Exp Res. 2013; 37, 1133–1142.Google Scholar
42. Laufer, BI, Mantha, K, Kleiber, ML, et al. Long-lasting alterations to DNA methylation and ncRNAs could underlie the effects of fetal alcohol exposure in mice. Dis Model Mech. 2013; 6, 977–992.Google Scholar
43. Kile, ML, Houseman, EA, Baccarelli, AA, et al. Effect of prenatal arsenic exposure on DNA methylation and leukocyte subpopulations in cord blood. Epigenetics. 2014; 9, 774–782.Google Scholar
44. Silver, MJ, Kessler, NJ, Hennig, BJ, et al. Independent genomewide screens identify the tumor suppressor VTRNA2-1 as a human epiallele responsive to periconceptional environment. Genome Biol. 2015; 16, 118.Google Scholar
45. Tobi, EW, Goeman, JJ, Monajemi, R, et al. DNA methylation signatures link prenatal famine exposure to growth and metabolism. Nat Commun. 2014; 5, 5592.Google Scholar
46. Amarasekera, M, Martino, D, Ashley, S, et al. Genome-wide DNA methylation profiling identifies a folate-sensitive region of differential methylation upstream of ZFP57-imprinting regulator in humans. FASEB J. 2014; 28, 4068–4076.Google Scholar
47. Thompson, RF, Fazzari, MJ, Niu, H, et al. Experimental intrauterine growth restriction induces alterations in DNA methylation and gene expression in pancreatic islets of rats. J Biol Chem. 2010; 285, 15111–15118.CrossRefGoogle ScholarPubMed
48. Kovacheva, VP, Mellott, TJ, Davison, JM, et al. Gestational choline deficiency causes global and Igf2 gene DNA hypermethylation by up-regulation of Dnmt1 expression. J Biol Chem. 2007; 282, 31777–31788.Google Scholar
49. Gardebjer, EM, Anderson, ST, Pantaleon, M, Wlodek, ME, Moritz, KM. Maternal alcohol intake around the time of conception causes glucose intolerance and insulin insensitivity in rat offspring, which is exacerbated by a postnatal high-fat diet. FASEB J. 2015; 29, 2690–2701.Google Scholar
50. Stover, PJ. One-carbon metabolism-genome interactions in folate-associated pathologies. J Nutr. 2009; 139, 2402–2405.Google Scholar
51. Ngai, YF, Sulistyoningrum, DC, O’Neill, R, et al. Prenatal alcohol exposure alters methyl metabolism and programs serotonin transporter and glucocorticoid receptor expression in brain. Am J Physiol Regul Integr Comp Physiol. 2015; 309, R613–R622.Google Scholar
52. Drake, AJ, O’Shaughnessy, PJ, Bhattacharya, S, et al. In utero exposure to cigarette chemicals induces sex-specific disruption of one-carbon metabolism and DNA methylation in the human fetal liver. BMC Med. 2015; 13, 18.Google Scholar
53. Garro, AJ, McBeth, DL, Lima, V, Lieber, CS. Ethanol consumption inhibits fetal DNA methylation in mice: implications for the fetal alcohol syndrome. Alcohol Clin Exp Res. 1991; 15, 395–398.Google Scholar
54. Haycock, PC. Fetal alcohol spectrum disorders: the epigenetic perspective. Biol Reprod. 2009; 81, 607–617.Google Scholar
55. McKay, JA, Waltham, KJ, Williams, EA, Mathers, JC. Folate depletion during pregnancy and lactation reduces genomic DNA methylation in murine adult offspring. Genes Nutr. 2011; 6, 189–196.Google Scholar
56. Breton, CV, Siegmund, KD, Joubert, BR, et al. Prenatal tobacco smoke exposure is associated with childhood DNA CpG methylation. PLoS One. 2014; 9, e99716.Google Scholar
57. Cheung, P, Allis, CD, Sassone-Corsi, P. Signaling to chromatin through histone modifications. Cell. 2000; 103, 263–271.CrossRefGoogle ScholarPubMed
58. Luger, K, Mader, AW, Richmond, RK, Sargent, DF, Richmond, TJ. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature. 1997; 389, 251–260.Google Scholar
59. Lanzuolo, C, Orlando, V. The function of the epigenome in cell reprogramming. Cell Mol Life Sci. 2007; 64, 1043–1062.Google Scholar
60. Paulson, JR, Taylor, SS. Phosphorylation of histones 1 and 3 and nonhistone high mobility group 14 by an endogenous kinase in HeLa metaphase chromosomes. J Biol Chem. 1982; 257, 6064–6072.Google Scholar
61. Davie, JR, Murphy, LC. Level of ubiquitinated histone H2B in chromatin is coupled to ongoing transcription. Biochemistry. 1990; 29, 4752–4757.CrossRefGoogle ScholarPubMed
62. Wondrak, GT, Cervantes-Laurean, D, Jacobson, EL, Jacobson, MK. Histone carbonylation in vivo and in vitro. Biochem J. 2000; 351(Pt 3), 769–777.Google Scholar
63. Poirier, GG, de Murcia, G, Jongstra-Bilen, J, Niedergang, C, Mandel, P. Poly(ADP-ribosyl)ation of polynucleosomes causes relaxation of chromatin structure. Proc Natl Acad Sci U S A. 1982; 79, 3423–3427.CrossRefGoogle ScholarPubMed
64. Shiio, Y, Eisenman, RN. Histone sumoylation is associated with transcriptional repression. Proc Natl Acad Sci U S A. 2003; 100, 13225–13230.Google Scholar
65. Bannister, AJ, Kouzarides, T. Regulation of chromatin by histone modifications. Cell Res. 2011; 21, 381–395.Google Scholar
66. Shi, X, Hong, T, Walter, KL, et al. ING2 PHD domain links histone H3 lysine 4 methylation to active gene repression. Nature. 2006; 442, 96–99.Google Scholar
67. Hassan, AH, Prochasson, P, Neely, KE, et al. Function and selectivity of bromodomains in anchoring chromatin-modifying complexes to promoter nucleosomes. Cell. 2002; 111, 369–379.Google Scholar
68. Strahl, BD, Allis, CD. The language of covalent histone modifications. Nature. 2000; 403, 41–45.Google Scholar
69. Vermeulen, M, Mulder, KW, Denissov, S, et al. Selective anchoring of TFIID to nucleosomes by trimethylation of histone H3 lysine 4. Cell. 2007; 131, 58–69.Google Scholar
70. Cao, R, Wang, L, Wang, H, et al. Role of histone H3 lysine 27 methylation in Polycomb-group silencing. Science. 2002; 298, 1039–1043.Google Scholar
71. Bernstein, BE, Mikkelsen, TS, Xie, X, et al. A bivalent chromatin structure marks key developmental genes in embryonic stem cells. Cell. 2006; 125, 315–326.Google Scholar
72. Stevens, A, Begum, G, Cook, A, et al. Epigenetic changes in the hypothalamic proopiomelanocortin and glucocorticoid receptor genes in the ovine fetus after periconceptional undernutrition. Endocrinology. 2010; 151, 3652–3664.Google Scholar
73. Wang, X, Gomutputra, P, Wolgemuth, DJ, Baxi, LV. Acute alcohol exposure induces apoptosis and increases histone H3K9/18 acetylation in the mid-gestation mouse lung. Reprod Sci. 2010; 17, 384–390.Google Scholar
74. Zinkhan, EK, Fu, Q, Wang, Y, et al. Maternal hyperglycemia disrupts histone 3 lysine 36 trimethylation of the IGF-1 gene. J Nutr Metab. 2012; 2012, 930364.Google Scholar
75. Davison, JM, Mellott, TJ, Kovacheva, VP, Blusztajn, JK. Gestational choline supply regulates methylation of histone H3, expression of histone methyltransferases G9a (Kmt1c) and Suv39h1 (Kmt1a), and DNA methylation of their genes in rat fetal liver and brain. J Biol Chem. 2009; 284, 1982–1989.Google Scholar
76. Mehedint, MG, Niculescu, MD, Craciunescu, CN, Zeisel, SH. Choline deficiency alters global histone methylation and epigenetic marking at the Re1 site of the calbindin 1 gene. FASEB J. 2010; 24, 184–195.Google Scholar
77. Suter, MA, Abramovici, AR, Griffin, E, et al. In utero nicotine exposure epigenetically alters fetal chromatin structure and differentially regulates transcription of the glucocorticoid receptor in a rat model. Birth Defects Res A Clin Mol Teratol. 2015; 103, 583–588.Google Scholar
78. Tyler, CR, Weber, JA, Labrecque, M, et al. ChIP-Seq analysis of the adult male mouse brain after developmental exposure to arsenic. Data Brief. 2015; 5, 248–254.Google Scholar
79. Yang, KF, Cai, W, Xu, JL, Shi, W. Maternal high-fat diet programs Wnt genes through histone modification in the liver of neonatal rats. J Mol Endocrinol. 2012; 49, 107–114.Google Scholar
80. Wang, KC, Chang, HY. Molecular mechanisms of long noncoding RNAs. Mol Cell. 2011; 43, 904–914.Google Scholar
81. Nagano, T, Mitchell, JA, Sanz, LA, et al. The air noncoding RNA epigenetically silences transcription by targeting G9a to chromatin. Science. 2008; 322, 1717–1720.Google Scholar
82. Bonasio, R, Tu, S, Reinberg, D. Molecular signals of epigenetic states. Science. 2010; 330, 612–616.Google Scholar
83. Kim, VN, Han, J, Siomi, MC. Biogenesis of small RNAs in animals. Nat Rev Mol Cell Biol. 2009; 10, 126–139.Google Scholar
84. Iwasaki, YW, Siomi, MC, Siomi, H. PIWI-interacting RNA: its biogenesis and functions. Annu Rev Biochem. 2015; 84, 405–433.Google Scholar
85. Landgraf, P, Rusu, M, Sheridan, R, et al. A mammalian microRNA expression atlas based on small RNA library sequencing. Cell. 2007; 129, 1401–1414.Google Scholar
86. Huang, Y, Zhang, JL, Yu, XL, et al. Molecular functions of small regulatory noncoding RNA. Biochemistry (Mosc). 2013; 78, 221–230.Google Scholar
87. Bartel, DP. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell. 2004; 116, 281–297.Google Scholar
88. Yekta, S, Shih, IH, Bartel, DP. MicroRNA-directed cleavage of HOXB8 mRNA. Science. 2004; 304, 594–596.Google Scholar
89. Bagga, S, Bracht, J, Hunter, S, et al. Regulation by let-7 and lin-4 miRNAs results in target mRNA degradation. Cell. 2005; 122, 553–563.Google Scholar
90. Olsen, PH, Ambros, V. The lin-4 regulatory RNA controls developmental timing in Caenorhabditis elegans by blocking LIN-14 protein synthesis after the initiation of translation. Dev Biol. 1999; 216, 671–680.Google Scholar
91. Pillai, RS, Bhattacharyya, SN, Artus, CG, et al. Inhibition of translational initiation by let-7 microRNA in human cells. Science. 2005; 309, 1573–1576.Google Scholar
92. Djuranovic, S, Nahvi, A, Green, R. miRNA-mediated gene silencing by translational repression followed by mRNA deadenylation and decay. Science. 2012; 336, 237–240.Google Scholar
93. Saraiya, AA, Li, W, Wang, CC. Transition of a microRNA from repressing to activating translation depending on the extent of base pairing with the target. PLoS One. 2013; 8, e55672.Google Scholar
94. Vasudevan, S, Tong, Y, Steitz, JA. Switching from repression to activation: microRNAs can up-regulate translation. Science. 2007; 318, 1931–1934.CrossRefGoogle ScholarPubMed
95. Lewis, BP, Burge, CB, Bartel, DP. Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets. Cell. 2005; 120, 15–20.Google Scholar
96. Xie, X, Lu, J, Kulbokas, EJ, et al. Systematic discovery of regulatory motifs in human promoters and 3’ UTRs by comparison of several mammals. Nature. 2005; 434, 338–345.Google Scholar
97. Brennecke, J, Stark, A, Russell, RB, Cohen, SM. Principles of microRNA-target recognition. PLoS Biol. 2005; 3, e85.Google Scholar
98. Krek, A, Grun, D, Poy, MN, et al. Combinatorial microRNA target predictions. Nat Genet. 2005; 37, 495–500.Google Scholar
99. Wang, LL, Zhang, Z, Li, Q, et al. Ethanol exposure induces differential microRNA and target gene expression and teratogenic effects which can be suppressed by folic acid supplementation. Hum Reprod. 2009; 24, 562–579.Google Scholar
100. Soares, AR, Pereira, PM, Ferreira, V, et al. Ethanol exposure induces upregulation of specific microRNAs in zebrafish embryos. Toxicol Sci. 2012; 127, 18–28.Google Scholar
101. Zhang, CR, Ho, MF, Vega, MC, Burne, TH, Chong, S. Prenatal ethanol exposure alters adult hippocampal VGLUT2 expression with concomitant changes in promoter DNA methylation, H3K4 trimethylation and miR-467b-5p levels. Epigenetics Chromatin. 2015; 8, 40.Google Scholar
102. Maccani, MA, Marsit, CJ. Exposure and fetal growth-associated miRNA alterations in the human placenta. Clin Epigenetics. 2011; 2, 401–404.Google Scholar
103. Balaraman, S, Lunde, ER, Sawant, O, et al. Maternal and neonatal plasma microRNA biomarkers for fetal alcohol exposure in an ovine model. Alcohol Clin Exp Res. 2014; 38, 1390–1400.Google Scholar
104. Vrijens, K, Bollati, V, Nawrot, TS. MicroRNAs as potential signatures of environmental exposure or effect: a systematic review. Environ Health Perspect. 2015; 123, 399–411.CrossRefGoogle ScholarPubMed
105. Zhang, Y, Fatima, N, Dufau, ML. Coordinated changes in DNA methylation and histone modifications regulate silencing/derepression of luteinizing hormone receptor gene transcription. Mol Cell Biol. 2005; 25, 7929–7939.Google Scholar
106. Nan, X, Ng, HH, Johnson, CA, et al. Transcriptional repression by the methyl-CpG-binding protein MeCP2 involves a histone deacetylase complex. Nature. 1998; 393, 386–389.Google Scholar
107. Jones, PL, Veenstra, GJ, Wade, PA, et al. Methylated DNA and MeCP2 recruit histone deacetylase to repress transcription. Nat Genet. 1998; 19, 187–191.Google Scholar
108. Eden, S, Hashimshony, T, Keshet, I, Cedar, H, Thorne, AW. DNA methylation models histone acetylation. Nature. 1998; 394, 842.Google Scholar
109. Zhao, Q, Rank, G, Tan, YT, et al. PRMT5-mediated methylation of histone H4R3 recruits DNMT3A, coupling histone and DNA methylation in gene silencing. Nat Struct Mol Biol. 2009; 16, 304–311.Google Scholar
110. Stewart, KR, Veselovska, L, Kim, J, et al. Dynamic changes in histone modifications precede de novo DNA methylation in oocytes. Genes Dev. 2015; 29, 2449–2462.Google Scholar
111. Vire, E, Brenner, C, Deplus, R, et al. The Polycomb group protein EZH2 directly controls DNA methylation. Nature. 2006; 439, 871–874.Google Scholar
112. Ooi, SK, Qiu, C, Bernstein, E, et al. DNMT3L connects unmethylated lysine 4 of histone H3 to de novo methylation of DNA. Nature. 2007; 448, 714–717.Google Scholar
113. Cedar, H, Bergman, Y. Linking DNA methylation and histone modification: patterns and paradigms. Nat Rev Genet. 2009; 10, 295–304.Google Scholar
114. Rose, NR, Klose, RJ. Understanding the relationship between DNA methylation and histone lysine methylation. Biochim Biophys Acta. 2014; 1839, 1362–1372.Google Scholar
115. Du, J, Johnson, LM, Jacobsen, SE, Patel, DJ. DNA methylation pathways and their crosstalk with histone methylation. Nat Rev Mol Cell Biol. 2015; 16, 519–532.Google Scholar
116. Han, L, Witmer, PD, Casey, E, Valle, D, Sukumar, S. DNA methylation regulates microRNA expression. Cancer Biol Ther. 2007; 6, 1284–1288.Google Scholar
117. Sun, BK, Deaton, AM, Lee, JT. A transient heterochromatic state in Xist preempts X inactivation choice without RNA stabilization. Mol Cell. 2006; 21, 617–628.Google Scholar
118. Schmitz, KM, Mayer, C, Postepska, A, Grummt, I. Interaction of noncoding RNA with the rDNA promoter mediates recruitment of DNMT3b and silencing of rRNA genes. Genes Dev. 2010; 24, 2264–2269.Google Scholar
119. Tomikawa, J, Shimokawa, H, Uesaka, M, et al. Single-stranded noncoding RNAs mediate local epigenetic alterations at gene promoters in rat cell lines. J Biol Chem. 2011; 286, 34788–34799.Google Scholar
120. Holz-Schietinger, C, Reich, NO. RNA modulation of the human DNA methyltransferase 3A. Nucleic Acids Res. 2012; 40, 8550–8557.Google Scholar
121. Di Ruscio, A, Ebralidze, AK, Benoukraf, T, et al. DNMT1-interacting RNAs block gene-specific DNA methylation. Nature. 2013; 503, 371–376.CrossRefGoogle ScholarPubMed
122. Zhou, J, Yang, L, Zhong, T, et al. H19 lncRNA alters DNA methylation genome wide by regulating S-adenosylhomocysteine hydrolase. Nat Commun. 2015; 6, 10221.Google Scholar
123. Watanabe, T, Tomizawa, S, Mitsuya, K, et al. Role for piRNAs and noncoding RNA in de novo DNA methylation of the imprinted mouse Rasgrf1 locus. Science. 2011; 332, 848–852.Google Scholar
124. Kuramochi-Miyagawa, S, Watanabe, T, Gotoh, K, et al. DNA methylation of retrotransposon genes is regulated by Piwi family members MILI and MIWI2 in murine fetal testes. Genes Dev. 2008; 22, 908–917.Google Scholar
125. Braconi, C, Huang, N, Patel, T. MicroRNA-dependent regulation of DNA methyltransferase-1 and tumor suppressor gene expression by interleukin-6 in human malignant cholangiocytes. Hepatology. 2010; 51, 881–890.Google Scholar
126. Huang, J, Wang, Y, Guo, Y, Sun, S. Down-regulated microRNA-152 induces aberrant DNA methylation in hepatitis B virus-related hepatocellular carcinoma by targeting DNA methyltransferase 1. Hepatology. 2010; 52, 60–70.Google Scholar
127. Barski, A, Jothi, R, Cuddapah, S, et al. Chromatin poises miRNA- and protein-coding genes for expression. Genome Res. 2009; 19, 1742–1751.Google Scholar
128. Wu, SC, Kallin, EM, Zhang, Y. Role of H3K27 methylation in the regulation of lncRNA expression. Cell Res. 2010; 20, 1109–1116.Google Scholar
129. Guil, S, Esteller, M. DNA methylomes, histone codes and miRNAs: tying it all together. Int J Biochem Cell Biol. 2009; 41, 87–95.Google Scholar
130. Peschansky, VJ, Wahlestedt, C. Non-coding RNAs as direct and indirect modulators of epigenetic regulation. Epigenetics. 2014; 9, 3–12.Google Scholar
131. Rinn, JL, Kertesz, M, Wang, JK, et al. Functional demarcation of active and silent chromatin domains in human HOX loci by noncoding RNAs. Cell. 2007; 129, 1311–1323.Google Scholar
132. Tsai, MC, Manor, O, Wan, Y, et al. Long noncoding RNA as modular scaffold of histone modification complexes. Science. 2010; 329, 689–693.Google Scholar
133. Noonan, EJ, Place, RF, Pookot, D, et al. miR-449a targets HDAC-1 and induces growth arrest in prostate cancer. Oncogene. 2009; 28, 1714–1724.Google Scholar
134. Wong, CF, Tellam, RL. MicroRNA-26a targets the histone methyltransferase Enhancer of Zeste homolog 2 during myogenesis. J Biol Chem. 2008; 283, 9836–9843.Google Scholar
135. Messerschmidt, DM, Knowles, BB, Solter, D. DNA methylation dynamics during epigenetic reprogramming in the germline and preimplantation embryos. Genes Dev. 2014; 28, 812–828.Google Scholar
136. Li, E. Chromatin modification and epigenetic reprogramming in mammalian development. Nat Rev Genet. 2002; 3, 662–673.Google Scholar
137. Santos, F, Dean, W. Epigenetic reprogramming during early development in mammals. Reproduction. 2004; 127, 643–651.Google Scholar
138. Morgan, HD, Santos, F, Green, K, Dean, W, Reik, W. Epigenetic reprogramming in mammals. Hum Mol Genet. 2005; 14(Spec No 1), R47–R58.Google Scholar
139. Reik, W, Dean, W, Walter, J. Epigenetic reprogramming in mammalian development. Science. 2001; 293, 1089–1093.Google Scholar
140. Oswald, J, Engemann, S, Lane, N, et al. Active demethylation of the paternal genome in the mouse zygote. Curr Biol. 2000; 10, 475–478.Google Scholar
141. Monk, M, Boubelik, M, Lehnert, S. Temporal and regional changes in DNA methylation in the embryonic, extraembryonic and germ cell lineages during mouse embryo development. Development. 1987; 99, 371–382.Google Scholar
142. Bhutani, N, Brady, JJ, Damian, M, et al. Reprogramming towards pluripotency requires AID-dependent DNA demethylation. Nature. 2010; 463, 1042–1047.Google Scholar
143. Liu, Y, Balaraman, Y, Wang, G, Nephew, KP, Zhou, FC. Alcohol exposure alters DNA methylation profiles in mouse embryos at early neurulation. Epigenetics. 2009; 4, 500–511.Google Scholar
144. Chen, PY, Ganguly, A, Rubbi, L, et al. Intrauterine calorie restriction affects placental DNA methylation and gene expression. Physiol Genomics. 2013; 45, 565–576.Google Scholar
145. Ivanova, E, Chen, JH, Segonds-Pichon, A, Ozanne, SE, Kelsey, G. DNA methylation at differentially methylated regions of imprinted genes is resistant to developmental programming by maternal nutrition. Epigenetics. 2012; 7, 1200–1210.Google Scholar
146. Radford, EJ, Isganaitis, E, Jimenez-Chillaron, J, et al. An unbiased assessment of the role of imprinted genes in an intergenerational model of developmental programming. PLoS Genet. 2012; 8, e1002605.Google Scholar
147. Lecomte, V, Youngson, NA, Maloney, CA, Morris, MJ. Parental programming: how can we improve study design to discern the molecular mechanisms? Bioessays. 2013; 35, 787–793.Google Scholar
148. Kawasaki, Y, Lee, J, Matsuzawa, A, et al. Active DNA demethylation is required for complete imprint erasure in primordial germ cells. Sci Rep. 2014; 4, 3658.CrossRefGoogle ScholarPubMed
149. Sasaki, H, Matsui, Y. Epigenetic events in mammalian germ-cell development: reprogramming and beyond. Nat Rev Genet. 2008; 9, 129–140.Google Scholar
150. Lucifero, D, Mann, MR, Bartolomei, MS, Trasler, JM. Gene-specific timing and epigenetic memory in oocyte imprinting. Hum Mol Genet. 2004; 13, 839–849.Google Scholar
151. Geuns, E, De Rycke, M, Van Steirteghem, A, Liebaers, I. Methylation imprints of the imprint control region of the SNRPN-gene in human gametes and preimplantation embryos. Hum Mol Genet. 2003; 12, 2873–2879.Google Scholar
152. Okae, H, Chiba, H, Hiura, H, et al. Genome-wide analysis of DNA methylation dynamics during early human development. PLoS Genet. 2014; 10, e1004868.CrossRefGoogle ScholarPubMed
153. Heijmans, BT, Kremer, D, Tobi, EW, Boomsma, DI, Slagboom, PE. Heritable rather than age-related environmental and stochastic factors dominate variation in DNA methylation of the human IGF2/H19 locus. Hum Mol Genet. 2007; 16, 547–554.Google Scholar
154. Schilling, E, El Chartouni, C, Rehli, M. Allele-specific DNA methylation in mouse strains is mainly determined by cis-acting sequences. Genome Res. 2009; 19, 2028–2035.Google Scholar
155. Zhi, D, Aslibekyan, S, Irvin, MR, et al. SNPs located at CpG sites modulate genome-epigenome interaction. Epigenetics. 2013; 8, 802–806.Google Scholar
156. Banovich, NE, Lan, X, McVicker, G, et al. Methylation QTLs are associated with coordinated changes in transcription factor binding, histone modifications, and gene expression levels. PLoS Genet. 2014; 10, e1004663.Google Scholar
157. McClay, JL, Shabalin, AA, Dozmorov, MG, et al. High density methylation QTL analysis in human blood via next-generation sequencing of the methylated genomic DNA fraction. Genome Biol. 2015; 16, 291.Google Scholar
158. Frosst, P, Blom, HJ, Milos, R, et al. A candidate genetic risk factor for vascular disease: a common mutation in methylenetetrahydrofolate reductase. Nat Genet. 1995; 10, 111–113.Google Scholar
159. Stern, LL, Mason, JB, Selhub, J, Choi, SW. Genomic DNA hypomethylation, a characteristic of most cancers, is present in peripheral leukocytes of individuals who are homozygous for the C677T polymorphism in the methylenetetrahydrofolate reductase gene. Cancer Epidemiol Biomarkers Prev. 2000; 9, 849–853.Google Scholar
160. Friso, S, Choi, SW, Girelli, D, et al. A common mutation in the 5,10-methylenetetrahydrofolate reductase gene affects genomic DNA methylation through an interaction with folate status. Proc Natl Acad Sci U S A. 2002; 99, 5606–5611.Google Scholar
161. El-Maarri, O, Kareta, MS, Mikeska, T, et al. A systematic search for DNA methyltransferase polymorphisms reveals a rare DNMT3L variant associated with subtelomeric hypomethylation. Hum Mol Genet. 2009; 18, 1755–1768.Google Scholar
162. Kadota, M, Yang, HH, Hu, N, et al. Allele-specific chromatin immunoprecipitation studies show genetic influence on chromatin state in human genome. PLoS Genet. 2007; 3, e81.Google Scholar
163. Kerkel, K, Spadola, A, Yuan, E, et al. Genomic surveys by methylation-sensitive SNP analysis identify sequence-dependent allele-specific DNA methylation. Nat Genet. 2008; 40, 904–908.Google Scholar
164. Haycock, PC, Ramsay, M. Exposure of mouse embryos to ethanol during preimplantation development: effect on DNA methylation in the H19 imprinting control region. Biol Reprod. 2009; 81, 618–627.Google Scholar
165. Zhang, D, Cheng, L, Badner, JA, et al. Genetic control of individual differences in gene-specific methylation in human brain. Am J Hum Genet. 2010; 86, 411–419.Google Scholar
166. Oey, H, Isbel, L, Hickey, P, Ebaid, B, Whitelaw, E. Genetic and epigenetic variation among inbred mouse littermates: identification of inter-individual differentially methylated regions. Epigenetics Chromatin. 2015; 8, 54.Google Scholar
167. Shea, JM, Serra, RW, Carone, BR, et al. Genetic and epigenetic variation, but not diet, shape the sperm methylome. Dev Cell. 2015; 35, 750–758.Google Scholar
168. Ohgane, J, Aikawa, J, Ogura, A, et al. Analysis of CpG islands of trophoblast giant cells by restriction landmark genomic scanning. Dev Genet. 1998; 22, 132–140.Google Scholar
169. Shiota, K, Kogo, Y, Ohgane, J, et al. Epigenetic marks by DNA methylation specific to stem, germ and somatic cells in mice. Genes Cells. 2002; 7, 961–969.Google Scholar
170. Yagi, S, Hirabayashi, K, Sato, S, et al. DNA methylation profile of tissue-dependent and differentially methylated regions (T-DMRs) in mouse promoter regions demonstrating tissue-specific gene expression. Genome Res. 2008; 18, 1969–1978.Google Scholar
171. Davies, MN, Volta, M, Pidsley, R, et al. Functional annotation of the human brain methylome identifies tissue-specific epigenetic variation across brain and blood. Genome Biol. 2012; 13, R43.Google Scholar
172. Roadmap Epigenomics, C, Kundaje, A, Meuleman, W, et al. Integrative analysis of 111 reference human epigenomes. Nature. 2015; 518, 317–330.Google Scholar
173. Leung, D, Jung, I, Rajagopal, N, et al. Integrative analysis of haplotype-resolved epigenomes across human tissues. Nature. 2015; 518, 350–354.Google Scholar
174. Flanagan, JM, Popendikyte, V, Pozdniakovaite, N, et al. Intra- and interindividual epigenetic variation in human germ cells. Am J Hum Genet. 2006; 79, 67–84.Google Scholar
175. Farlik, M, Sheffield, NC, Nuzzo, A, et al. Single-cell DNA methylome sequencing and bioinformatic inference of epigenomic cell-state dynamics. Cell Rep. 2015; 10, 1386–1397.Google Scholar
176. Novakovic, B, Ryan, J, Pereira, N, et al. Postnatal stability, tissue, and time specific effects of AHRR methylation change in response to maternal smoking in pregnancy. Epigenetics. 2014; 9, 377–386.Google Scholar
177. Basil, P, Li, Q, Dempster, EL, et al. Prenatal maternal immune activation causes epigenetic differences in adolescent mouse brain. Transl Psychiatry. 2014; 4, e434.Google Scholar
178. Kundakovic, M, Gudsnuk, K, Franks, B, et al. Sex-specific epigenetic disruption and behavioral changes following low-dose in utero bisphenol A exposure. Proc Natl Acad Sci U S A. 2013; 110, 9956–9961.Google Scholar
179. Begum, G, Davies, A, Stevens, A, et al. Maternal undernutrition programs tissue-specific epigenetic changes in the glucocorticoid receptor in adult offspring. Endocrinology. 2013; 154, 4560–4569.Google Scholar
180. El Hajj, N, Pliushch, G, Schneider, E, et al. Metabolic programming of MEST DNA methylation by intrauterine exposure to gestational diabetes mellitus. Diabetes. 2013; 62, 1320–1328.Google Scholar
181. Cardenas, A, Koestler, DC, Houseman, EA, et al. Differential DNA methylation in umbilical cord blood of infants exposed to mercury and arsenic in utero. Epigenetics. 2015; 10, 508–515.Google Scholar
182. Jacoby, M, Gohrbandt, S, Clausse, V, Brons, NH, Muller, CP. Interindividual variability and co-regulation of DNA methylation differ among blood cell populations. Epigenetics. 2012; 7, 1421–1434.Google Scholar
183. Reinius, LE, Acevedo, N, Joerink, M, et al. Differential DNA methylation in purified human blood cells: implications for cell lineage and studies on disease susceptibility. PLoS One. 2012; 7, e41361.Google Scholar
184. Jaffe, AE, Irizarry, RA. Accounting for cellular heterogeneity is critical in epigenome-wide association studies. Genome Biol. 2014; 15, R31.Google Scholar
185. Bauer, M, Linsel, G, Fink, B, et al. A varying T cell subtype explains apparent tobacco smoking induced single CpG hypomethylation in whole blood. Clin Epigenetics. 2015; 7, 81.CrossRefGoogle ScholarPubMed
186. Houseman, EA, Accomando, WP, Koestler, DC, et al. DNA methylation arrays as surrogate measures of cell mixture distribution. BMC Bioinformatics. 2012; 13, 86.Google Scholar
187. Guintivano, J, Aryee, MJ, Kaminsky, ZA. A cell epigenotype specific model for the correction of brain cellular heterogeneity bias and its application to age, brain region and major depression. Epigenetics. 2013; 8, 290–302.Google Scholar
188. Koestler, DC, Christensen, B, Karagas, MR, et al. Blood-based profiles of DNA methylation predict the underlying distribution of cell types: a validation analysis. Epigenetics. 2013; 8, 816–826.Google Scholar
189. Langevin, SM, Houseman, EA, Accomando, WP, et al. Leukocyte-adjusted epigenome-wide association studies of blood from solid tumor patients. Epigenetics. 2014; 9, 884–895.Google Scholar
190. Yousefi, P, Huen, K, Quach, H, et al. Estimation of blood cellular heterogeneity in newborns and children for epigenome-wide association studies. Environ Mol Mutagen. 2015; 56, 751–758.Google Scholar
191. Singhal, RP, Mays-Hoopes, LL, Eichhorn, GL. DNA methylation in aging of mice. Mech Ageing Dev. 1987; 41, 199–210.Google Scholar
192. Fraga, MF, Ballestar, E, Paz, MF, et al. Epigenetic differences arise during the lifetime of monozygotic twins. Proc Natl Acad Sci U S A. 2005; 102, 10604–10609.Google Scholar
193. Levesque, ML, Casey, KF, Szyf, M, et al. Genome-wide DNA methylation variability in adolescent monozygotic twins followed since birth. Epigenetics. 2014; 9, 1410–1421.Google Scholar
194. Talens, RP, Boomsma, DI, Tobi, EW, et al. Variation, patterns, and temporal stability of DNA methylation: considerations for epigenetic epidemiology. FASEB J. 2010; 24, 3135–3144.Google Scholar
195. Eckhardt, F, Lewin, J, Cortese, R, et al. DNA methylation profiling of human chromosomes 6, 20 and 22. Nat Genet. 2006; 38, 1378–1385.Google Scholar
196. Carone, BR, Fauquier, L, Habib, N, et al. Paternally induced transgenerational environmental reprogramming of metabolic gene expression in mammals. Cell. 2010; 143, 1084–1096.Google Scholar
197. Volkmar, M, Dedeurwaerder, S, Cunha, DA, et al. DNA methylation profiling identifies epigenetic dysregulation in pancreatic islets from type 2 diabetic patients. EMBO J. 2012; 31, 1405–1426.Google Scholar
198. Chen, G, Gharib, TG, Huang, CC, et al. Discordant protein and mRNA expression in lung adenocarcinomas. Mol Cell Proteomics. 2002; 1, 304–313.Google Scholar
199. Ghazalpour, A, Bennett, B, Petyuk, VA, et al. Comparative analysis of proteome and transcriptome variation in mouse. PLoS Genet. 2011; 7, e1001393.Google Scholar
200. Lie, S, Morrison, JL, Williams-Wyss, O, et al. Impact of embryo number and maternal undernutrition around the time of conception on insulin signaling and gluconeogenic factors and microRNAs in the liver of fetal sheep. Am J Physiol Endocrinol Metab. 2014; 306, E1013–E1024.Google Scholar
Figure 0

Table 1 Strategies to address common challenges in epigenetic studies

Figure 1

Table 2 A selection of recent DOHaD publications that have addressed two or more of the challenges highlighted in this review. Black boxes denote the challenge addressed by the study