Advertisement

  • Loading metrics

Open Access

Review

Human placental methylome in the interplay of adverse placental health, environmental exposure, and pregnancy outcome

Abstract

The placenta is the interface between maternal and fetal circulations, integrating maternal and fetal signals to selectively regulate nutrient, gas, and waste exchange, as well as secrete hormones. In turn, the placenta helps create the in utero environment and control fetal growth and development. The unique epigenetic profile of the human placenta likely reflects its early developmental separation from the fetus proper and its role in mediating maternal–fetal exchange that leaves it open to a range of exogenous exposures in the maternal circulation. In this review, we cover recent advances in DNA methylation in the context of placental function and development, as well as the interaction between the pregnancy and the environment.

Citation: Vlahos A, Mansell T, Saffery R, Novakovic B (2019) Human placental methylome in the interplay of adverse placental health, environmental exposure, and pregnancy outcome. PLoS Genet 15(8): e1008236. https://doi.org/10.1371/journal.pgen.1008236

Editor: Marisa S. Bartolomei, University of Pennsylvania, UNITED STATES

Published: August 1, 2019

Copyright: © 2019 Vlahos et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: TM is funded by an Australian Postgraduate Research Award. BN is funded by an NHMRC Australia CJ Martin Fellowship. MCRI is funded by a Victorian Government Infrastructure grant. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Establishing a successful pregnancy depends on a complex sequence of interactions between maternal and fetal cells, and their interlocutory signals [1]. This interaction takes place on many levels, from systematic endocrine factors to direct paracrine and juxtacrine contact between the placenta and maternal decidual cells. The placenta is a major transient endocrine organ that plays a key role in enabling many of these complex interactions. Placental trophoblasts extensively remodel the maternal decidua and uterine vasculature to ensure appropriate blood flow for the exchange of nutrients and waste between fetal and maternal circulations. In turn, this creates a unique environment at the fetal–maternal interface [2].

The extent to which the placenta invades the maternal decidua and interacts with the maternal circulation depends on the structure of the placenta. Evolutionarily, the placenta is one of the most diverse organs, evolving independently in multiple lineages and with different placental patterning [3]. The hemochorial human placenta is unique from other primates and model organisms like mice in that trophoblast invasion is more extensive and implantation is interstitial [4]. The placenta differentiates from the extraembryonic trophectoderm, giving rise to villous cytotrophoblasts (VCTs) that can proliferate or fuse to form the multinuclear syncytiotrophoblast (ST) layer, which acts as the barrier between maternal and fetal circulations, and the extra-villous trophoblasts (EVTs), which invade the decidua and remodel maternal spiral arteries [5]. Placental cell composition and function are dynamic over gestation, with a major change being the increased flow of maternal blood into the intravillous space around the transition from first to second trimester, which results in a change in oxygen concentration and extensive remodelling of villous structure [6, 7]. This interaction of maternal and fetal cells is facilitated by considerable adaptation of the maternal immune response, ensuring tolerance of the developing fetus while retaining capability for host cell defence and contributing to healthy development [8].

Compelling evidence from human epidemiological studies suggests that adverse exposures in utero can alter the intrauterine environment, with potential implications for fetal development and long-term offspring health [9]. Due to its role as interface between maternal and fetal circulations, the placenta is constantly exposed to exogenous factors that are present in the maternal circulation, with the potential for these exposures to leave a lasting molecular ‘footprint’ on the placenta. In keeping with this, it is not surprising that the extraembryonic origin, relatively short life span, and ongoing direct exposure to all exogenous signals are reflected in a unique epigenetic profile [10]. Here, we highlight and discuss emerging knowledge on the unique features of the human placental methylome (total DNA methylation profile) and summarise research linking in utero exposures, adverse placental health, and fetal outcomes to altered DNA methylation in the human placenta. We go on to discuss the potential impact of single-cell techniques on understanding the placental methylome and how this will pave the way for future use of DNA methylation in the placenta as a biomarker to diagnose placental syndromes and predict fetal outcomes.

DNA methylation remodelling in extraembryonic cells

Epigenetics refers to the molecular interactions that influence DNA packaging into chromatin, thereby regulating DNA accessibility and gene expression. Epigenetic modifications are added either directly to DNA (as in DNA methylation) or to the amino acid tails of histones that constitute the nucleosome around which DNA is coiled.

DNA methylation is the most widely studied and well-characterised epigenetic modification. It involves the addition of a methyl group to a cytoside within a cytoside-guanine dinucleotide (CpG site) [11]. CpG sites often cluster together in CpG islands close to the promoter and regulatory regions of genes, and within this context increased DNA methylation blocks access to the underlying DNA sequence, leading to reduced gene expression. During human preimplantation development, DNA methylation is highly dynamic (Fig 1A). Notably, at the blastocyst stage, the inner cell mass and trophectoderm undergo a wave of de novo DNA methylation, but to different extents, leading to differences in final global DNA methylation levels [12].

thumbnail
Download:
Fig 1. Unique epigenetic features of the human placenta.

(A) DNA methylation dynamics during early human development. After fertilisation, the sperm and egg undergo a wave of hypomethylation, except at imprinted genes. Around implantation, the inner cell mass of the blastocyst, which develops into the epiblast, and the trophoblast, which develops into the placenta, undergo de novo methylation to differing extents. (B) Unique features of the placenta methylome arising as a result of the remethylation differences in A. B (i) Global hypomethylation [15, 16], (ii) polymorphic imprinting [17], (iii) PMDs [18, 19], (iv) trophoblast differentiation dynamics [13], (v) epigenetic regulation of HLA-G [20], (vi) tumour suppressor promoter methylation [21, 22], (vii) silencing of chemokines by a repressive histone mark, H3K27me3 [23], (viii) control of parturition by H3K27me3 in decidua [23], (ix) interindividual variation increases with gestation [24], (x) TERRA promoter hypomethylation [25], (xi) trophoblast DNA methylation is sensitive to oxygen concentration [14], (xii) hypomethylated retrotransposons used as alternative promoters [26, 27]. C, cytosine nucleotide; CpG, cytoside-guanine dinucleotide; CT, cytotrophoblast; CXCL9, C-X-C motif chemokine ligand 9; CXCL10, C-X-C motif chemokine ligand 9; EVT, extra-villous trophoblast; HLA-G, major histocompatibility complex, class I, G; H3K27me3, histone 3 Lysine 27 trimethylation; meC, methylated cytosine; PMD, partially methylated domain; SINE, short interspersed nuclear element; ST, syncytiotrophoblast; TERRA, telomere RNA.

https://doi.org/10.1371/journal.pgen.1008236.g001

DNA methylation continues to be dynamic in trophoblast cells after implantation and throughout gestation, with the process of VCTs’ differentiation to ST and the acquisition of an invasive phenotype in EVT, both of which involving widespread DNA methylation changes [13]. Interestingly, culture in a low oxygen concentration induced opposing DNA methylation change at certain differentiation-associated differentially methylated regions (DMRs) and was associated with lower differentiation potential, an effect potentially modulated by the activator protein 1 (AP-1) transcription factor complex [14]. These findings highlight the intimate relationship between DNA methylation patterns, trophoblast differentiation, and the sensing of environmental signals.

Placental global hypomethylation is not specific to repetitive elements

The aforementioned differences in DNA methylation re-establishment result in the placenta exhibiting a unique, globally hypomethylated DNA methylation profile compared with normal somatic tissue (Fig 1B) [15]. High-performance liquid chromatography (HPLC) analysis of the total 5-methylcytosine content estimates that human placental tissue is on average 2.5%–3% methylated compared to approximately 4% in human cord blood (Fig 1B(i)). Repetitive elements, which cover approximately 35% of the human genome, are supressed with elevated levels of DNA methylation [28, 29]. Earlier studies of the placental methylome found that some repetitive elements, such as long interspersed element-1 (LINE-1) [30, 31]) and human endogenous reterovirus (HERV) [32], are key DMRs in placental tissue. Placental hypomethylation of these repetitive elements is thought to regulate placenta-specific functions. For example, some retrotransposons function as alternative promoters for placental-specific transcripts, such as potassium voltage-gated channel subfamily H member 5 (KCNH5) and interleukin 2 receptor, beta 1 subunit (IL2RB) (Fig 1B(xii)) [26, 27, 32]. Expression of an alternative form of KCNH5 may contribute to trophoblast invasiveness, a conclusion that is supported by the finding that the placenta-specific transcript and associated hypomethylated promoter occur frequently in melanoma cells, but never in healthy somatic cell types [33]. IL2RB is a subunit of the IL2 receptor and is expressed primarily by lymphocytes, and the function of the trophoblast specific-transcript is currently unknown [27]. In fact, despite the widespread potential for retrotransposon-driven novel transcription in the placenta, relatively little is known about its potential role in correct placental functioning.

Recent findings have dispelled the notion that retrotransposons are specifically hypomethylated in the placenta. In a landmark study, Schroeder and colleagues [18] showed that placental DNA hypomethylation mainly occurs within partially methylated domains (PMDs)—that is, large stretches (up to 100 kb) of gene-poor regions with reduced DNA methylation that are interspersed with regions of higher DNA methylation (Fig 1B(iii)). Using Methyl-C sequencing, it was estimated that PMDs covered approximately 37% of the placental genome and that genes within placental PMDs have placental-specific functions. An important observation was that LINE-1 elements located in a PMD showed the same level of methylation as the surrounding region (approximately 40%), while LINE-1 elements outside PMDs were not hypomethylated (approximately 80% methylation). Likewise, another study used reduced representation bisulfide sequencing (RRBS) to show that placental hypomethylation is enriched at CpG-poor intergenic regions and gene bodies, and that retrotransposons in these regions were, in fact, slightly more methylated than non-retrotransposon elements [34]. Taken together, this suggests that while the placenta may utilise unmethylated retrotransposon elements as alternative promoters, these regions do not solely explain the function global hypomethylation in the placenta. Further research into PMDs may reveal new insights into their function, although role of PMDs in placenta is yet to be determined.

Placental pseudomalignancy: An epigenetically regulated developmental mechanism co-opted by cancers?

The similarities between placentation and cancer were first pointed out in 1902 [35]. In the 1970s and 1980s, studies showed that the human placenta expressed high levels of oncogenes [36], while cancers expressed high levels of placental hormones, like human chorionic gonadotropin (hCG). Thus, this led to the hypothesis that tumorogenesis was a ‘pathological recapitulation’ of normal placental development [37]. Unlike cancers, placental trophoblasts restrict their proliferative and invasive phenotype, and have therefore been described as pseudomalignant. Trophoblasts activate similar molecular and epigenetic pathways observed in cancers [38], and it has been suggested that cancers co-opt placental-specific epigenetic programming [39].

In addition to the global hypomethylation and presence of PMDs, earlier studies showed that promoters of common tumour suppressor genes are monoallelically methylated in the placenta and trophoblasts (Fig 1B(vi)) [21, 22]. These promoters become completely methylated in choricarcinomas (trophoblast cancers), supporting the notion that these cancer-like programs are carefully regulated in the placenta. Similarly, cancer-like global DNA hypomethylation blocks in early gestation placenta are lost in later gestation, suggesting a necessary switch that limits placenta invasion and other tumour-like properties in later pregnancy [40].

Studying the establishment of these pseudomalignant methylomes during early implantation is difficult in humans. As such, we do not know how a completely hypomethylated human trophectoderm develops these features. An interesting mouse study that addressed this question comes from the Meissner lab, which showed that partially methylated promoter regions are established rapidly [19]. In embryonic stem cells, these regions are marked by a repressive histone modification, histone 3 lysine 27 trimethylation (H3K27me3), indicating that polycomb group of proteins are keeping these regions closed and thereby potentially allow DNA methyltransferases (DNMTs) to methylate DNA. An important question remains: are PMDs and tumour suppressor promoter methylation drivers of placental function or bystanders?

Novel imprinted regions in the placenta

During development, certain regions of DNA, called germline differentially methylated regions (gDMRs), maintain gamete DNA methylation during genome-wide reprogramming and throughout development [41]. These gDMRs can regulate expression at nearby genes, resulting in monoallelic parent-of-origin expression known as genomic imprinting [42]. There are multiple theories as to why genes evolved to be imprinted [43], but the prevailing hypothesis is that of the kinship theory [44]. In brief, imprinting allows for natural selection to maximise the fitness of the allele through maximising not just the health of the individual inheriting the allele but also the related individuals who will also inherit that allele from the same parent [42]. An example is a gene that regulates growth in utero, and consequently influences the fetal demand for maternal resources [45]. As multiple offspring from the same mother may have different fathers, maximising the kinship fitness bestowed by the maternally inherited allele is more likely to improve kinship fitness overall than the paternally inherited allele. As such, paternal alleles may evolve as silenced to avoid conflicts between maternal and paternal allelic expression [45]. Recent research indicates that the placenta has more gDMRs than previously thought [17, 46], which is consistent with the importance of the placenta in regulating fetal growth and maternal resource demand. However, it remains unclear whether the majority of placenta-specific gDMRs are functional in pregnancy, or whether they simply reflect the methylation status of a gamete. Hanna and colleagues [17] identified 144 gDMRs in the human placenta, in which most appeared to maintain maternal DNA methylation patterns (mDMRs). Similarly, Hamada and colleagues [46] performed combined whole-genome bisulfide sequencing and RNA-seq, which identified approximately 1,800 mDMRs in the human placenta, some of which were associated with potential paternal and maternal imprinted genes such as TP53 induced glycolysis regulatory phosphatase (TIGAR), sodium bicarbonate cotransporter 3 (SLC4A7), and zinc finger protein (ZFP90). The function of these imprinted genes in placenta or fetal development, however, remains to be elucidated. An interesting observation requiring follow-up study is that the DNA methyltransferase 1 (DNMT1) promoter itself is maternally imprinted in the human placenta [47]. As with several other aspects of human placental methylation, this promoter methylation is evolutionary conserved only in primates, not in other mammals, and does not seem to be required for global hypomethylation in the placenta [48, 49]. It is easy to speculate that maternal imprinting of DNMT1 is a mechanism to prevent the activation of growth-promoting genes, such as insulin-like growth factor 2 (IGF2), by DNMT1 overexpression [50]. In line with this, DNMT1 expression shows a modest positive correlation with placental growth [51]. However, there is no confirmed function for DNMT1 imprinting in the placenta. A limitation of using whole placental tissue to identify novel imprinted genes is that there may be variation between different cell types, which can lead to some imprinted regions being missed. Additionally, without identifying the specific cell types that show imprinting, it is hard to study the function of these novel imprinted regions.

In utero exposures alter the placenta methylation, which may impact placental and fetal health

DNA methylation is sensitive to environmental exposure but relatively stable once established. Consequently, studies have examined the influence of in utero exposures on DNA methylation in the placenta in the hope that these patterns could be used as potential biomarkers to diagnose placental disorders or even predict fetal outcomes. The overall level of placental tissue DNA methylation increases over gestation, as does the level of interindividual variation between placentas, potentially indicative of a response to a myriad of environmental exposures and stochastic events, or with a transition and variation in cell composition over time [24]. Many of these studies take the form of epigenome-wide association studies (EWAS), and generally use the Infinium HumanMethylation arrays [52, 53]. Mounting evidence suggests that exposures in utero influence the human placental methylome. In Tables 1 and 2, we summarise key EWAS studies using the human placenta since 2016. For a summary of studies predating 2016, please see [54]. Table 1 summarises emerging research linking in utero exposures and the human placental methylome. Given the key role of the placenta in fetal programming, altered DNA methylation profiles of the placenta may cause aberrant placental development and function, which may in turn influence fetal outcomes. Table 2 summarises emerging research linking the human placental methylome and pregnancy outcomes.

thumbnail
Download:
Table 1. Placenta epigenetics and environment.

https://doi.org/10.1371/journal.pgen.1008236.t001

thumbnail
Download:
Table 2. Placenta epigenetics, disease, and fetal outcome.

https://doi.org/10.1371/journal.pgen.1008236.t002

Over the past 5 years, there has been a shift from looking at maternal consumption (smoking, adiposity, and nutrition) towards exposures related to the natural and social environment, like plastics, chemicals, air pollution, other pollutants, and war trauma, reflecting the emerging broader scientific and societal interest in the impact of climate change and social issues on human health. We anticipate that as these climate change and societal issues continue to emerge, increased multidisciplinary research will be required to examine the impact of such exposures on many aspects of pregnancy, including placental development, epigenetic profile, and subsequent impact on placental and fetal health. We must also note that the majority of current studies are underpowered due to small sample size, and we urge that researchers consider EWAS power calculations and other considerations of study design prior to conducting such studies [55, 56]. It would be considerably beneficial to the field if the suppliers of the most commonly used platform made efforts to reduce the current excessive pricing of DNA methylation arrays relative to the current SNP-based genotyping equivalents.

Exploiting the unique placental epigenome for noninvasive tracking of pregnancy health and outcome

Circulating fetal cell-free DNA in maternal blood is placental in origin, and has been used to noninvasively test for aneuploidy and disease-causing mutations [57, 58]. These tests depend on the detection of paternally inherited disease-causing SNPs that are not present in the mother. Early on, it was realised that placenta-specific DNA methylation patterns, such as the hypermethylation of the ras association domain-containing protein 1 (RASSF1A) promoter region [21], can be used as a positive control to rule out false negative results [59]. Since this pioneering work, Lo and colleagues used DNA methylation of fetal cell-free DNA for gestational dating, indicating that this technique has far-reaching potential [60]. However, replication of this finding is needed to confirm its usefulness in the diagnosis of gestational age. Recent studies have suggested that preeclampsia-specific DNA methylation patterns in the placenta, e.g., TIMP metallopeptidase inhibitor 3 (TIMP3) and RASSF1A promoters, can be detected in maternal circulation [61, 62]. However, no such test exists yet. It may also be possible to identify DNA methylation patterns associated with greater risk of preterm birth and small for gestational age (SGA) [63], or with their associated later morbidity [64, 65]. Such approaches have potential predictive utility through the analysis of circulating cell-free placenta-derived DNA, and are currently in development in several laboratories internationally [66, 67]. Key to this is identifying robust placenta methylation biomarkers associated with specific exposures and/or outcomes, which may not be feasible given the apparent low degree of methylation variation currently linked to most environmental exposures and outcomes (Table 1). It is now also possible to accurately reconstruct placental genome-wide DNA methylation by bisulfite sequencing of maternal plasma, which could allow the identification of epigenetic aberrations in the placenta without needing the placental sample itself [68].

Single-cell approaches may overcome issues associated with placenta heterogeneity

Placenta is a heterogeneous tissue comprised of many cell types [69], some with varying chromosome copy numbers [70]. These characteristics make it difficult to interpret DNA methylation analysis, as well as limit biomarker use and understanding of molecular pathways and networks. Furthermore, the composition of these cell populations and corresponding DNA methylation profile may change throughout pregnancy [24, 71]. For the time being, it is likely that cohort-based EWAS will continue to use whole placental tissue because of the technical difficulties associated with isolating specific cell types on a large scale. Here, cell-specific DNA methylation profiles will aid research by providing signatures that can be used to deconvolute data and remove cell-specific signatures as variables in statistical analysis, similar to the way whole blood data are analysed [72]. However, deconvolution tools are currently limited to the number of different cell subtypes they include in their modelling, and further work is necessary to account for rare cell subtypes in analysis.

Recent advances in genome sequencing and cell sorting technology have enabled the unbiased analysis of all cell types in the human placenta. This is highlighted by the publication of several papers reporting single-cell RNA-seq profiles of the human fetal–maternal interface at the first and third trimesters in the last 3 years [69, 73–77]. These studies detected new subsets of cytotrophoblasts and EVTs, as well as cell subtypes previously not observed in placental villi [69, 73]. Continued application of single-cell RNA-seq to human placenta-derived cell populations, particularly over gestation, will greatly improve our understanding of how cell composition changes over time and how individual cells respond to changing environments, such as a shift in oxygen concentration. However, techniques for profiling epigenetic marks from low cell numbers are now available and in wide use [78, 79], including single-cell techniques for DNA accessibility [80], protein expression/phosphorylation [81], and combinations of marks [82–84]. Given the relative ease of obtaining placental tissue, these techniques should be readily applied to purified placental cells. Finally, placental organoid models that recapitulate villi structure and can produce ST and EVT cells have now been successfully established [85, 86], and unlike trophoblast cell lines [87], the organoids resemble first trimester trophoblasts in terms of their genome-wide DNA methylation patterns [86]. These novel models in combination with state-of-the-art molecular techniques represent a very powerful approach to provide hitherto unparalleled insights into human trophoblast differentiation and function.

Conclusion

In this review, we highlighted the recent advances across the placental epigenetics field, which is now dominated by studies examining environmental influences. Given the close interaction between trophoblasts and immune cells, which make up to 40% of the feto-maternal interface, we anticipate the next few years will see epigenetic profiling of these resident immune cells. It will be interesting to see what PMDs look like in different placental cell types and how individual cells control the expression of hypomethylated retrotransposon promoters. Multi-omics approaches also have the potential to give us insight into the drivers and passengers involved in the interaction between the unique placental methylome and the environment.

References

  1. 1. Fowden A.L., et al., The placenta and intrauterine programming. J Neuroendocrinol, 2008. 20(4): p. 439–50. pmid:18266944
  2. 2. Huppertz B., The feto-maternal interface: setting the stage for potential immune interactions. Semin Immunopathol, 2007. 29(2): p. 83–94. pmid:17621696
  3. 3. Griffith O.W. and Wagner G.P., The placenta as a model for understanding the origin and evolution of vertebrate organs. Nat Ecol Evol, 2017. 1(4): p. 72. pmid:28812655
  4. 4. Carter A.M., Animal models of human placentation—a review. Placenta, 2007. 28 Suppl A: p. S41–7.
  5. 5. Red-Horse K., et al., Trophoblast differentiation during embryo implantation and formation of the maternal-fetal interface. J Clin Invest, 2004. 114(6): p. 744–54. pmid:15372095
  6. 6. Burton G.J. and Jauniaux E., The cytotrophoblastic shell and complications of pregnancy. Placenta, 2017. 60: p. 134–139. pmid:28651899
  7. 7. Jauniaux E., et al., Onset of maternal arterial blood flow and placental oxidative stress. A possible factor in human early pregnancy failure. Am J Pathol, 2000. 157(6): p. 2111–22. pmid:11106583
  8. 8. Mor G. and Cardenas I., The immune system in pregnancy: a unique complexity. Am J Reprod Immunol, 2010. 63(6): p. 425–33. pmid:20367629
  9. 9. McMillen I.C. and Robinson J.S., Developmental origins of the metabolic syndrome: prediction, plasticity, and programming. Physiol Rev, 2005. 85(2): p. 571–633. pmid:15788706
  10. 10. Novakovic B. and Saffery R., The ever growing complexity of placental epigenetics—role in adverse pregnancy outcomes and fetal programming. Placenta, 2012. 33(12): p. 959–70. pmid:23102655
  11. 11. Jones P.A., Functions of DNA methylation: islands, start sites, gene bodies and beyond. Nat Rev Genet, 2012. 13(7): p. 484–92. pmid:22641018
  12. 12. Monk M., Boubelik M., and Lehnert S., Temporal and regional changes in DNA methylation in the embryonic, extraembryonic and germ cell lineages during mouse embryo development. Development, 1987. 99(3): p. 371–82. pmid:3653008
  13. 13. Novakovic B., et al., Variable DAXX gene methylation is a common feature of placental trophoblast differentiation, preeclampsia, and response to hypoxia. FASEB J, 2017. 31(6): p. 2380–2392. pmid:28223336
  14. 14. Yuen R.K., et al., Hypoxia alters the epigenetic profile in cultured human placental trophoblasts. Epigenetics, 2013. 8(2): p. 192–202. pmid:23314690
  15. 15. Ehrlich M., et al., Amount and distribution of 5-methylcytosine in human DNA from different types of tissues of cells. Nucleic Acids Res, 1982. 10(8): p. 2709–21. pmid:7079182
  16. 16. Gama-Sosa M.A., et al., Tissue-specific differences in DNA methylation in various mammals. Biochim Biophys Acta, 1983. 740(2): p. 212–9. pmid:6860672
  17. 17. Hanna C.W., et al., Pervasive polymorphic imprinted methylation in the human placenta. Genome Res, 2016. 26(6): p. 756–67. pmid:26769960
  18. 18. Schroeder D.I., et al., The human placenta methylome. Proc Natl Acad Sci U S A, 2013. 110(15): p. 6037–42. pmid:23530188
  19. 19. Smith Z.D., et al., Epigenetic restriction of extraembryonic lineages mirrors the somatic transition to cancer. Nature, 2017. 549(7673): p. 543–547. pmid:28959968
  20. 20. Ferreira L.M., et al., A distant trophoblast-specific enhancer controls HLA-G expression at the maternal-fetal interface. Proc Natl Acad Sci U S A, 2016. 113(19): p. 5364–9. pmid:27078102
  21. 21. Chiu R.W., et al., Hypermethylation of RASSF1A in human and rhesus placentas. Am J Pathol, 2007. 170(3): p. 941–50. pmid:17322379
  22. 22. Novakovic B., et al., Specific tumour-associated methylation in normal human term placenta and first-trimester cytotrophoblasts. Mol Hum Reprod, 2008. 14(9): p. 547–54. pmid:18708652
  23. 23. Nancy P., et al., H3K27me3 dynamics dictate evolving uterine states in pregnancy and parturition. J Clin Invest, 2018. 128(1): p. 233–247. pmid:29202469
  24. 24. Novakovic B., et al., Evidence for widespread changes in promoter methylation profile in human placenta in response to increasing gestational age and environmental/stochastic factors. BMC Genomics, 2011. 12: p. 529. pmid:22032438
  25. 25. Novakovic B., et al., DNA methylation mediated up-regulation of TERRA non-coding RNA is coincident with elongated telomeres in the human placenta. Mol Hum Reprod, 2016. 22(11): p. 791–799. pmid:27604461
  26. 26. Macaulay E.C., et al., Hypomethylation of functional retrotransposon-derived genes in the human placenta. Mamm Genome, 2011. 22(11–12): p. 722–35. pmid:21874386
  27. 27. Cohen C.J., et al., Placenta-specific expression of the interleukin-2 (IL-2) receptor beta subunit from an endogenous retroviral promoter. J Biol Chem, 2011. 286(41): p. 35543–52. pmid:21865161
  28. 28. Walsh C.P., Chaillet J.R., and Bestor T.H., Transcription of IAP endogenous retroviruses is constrained by cytosine methylation. Nat Genet, 1998. 20(2): p. 116–7. pmid:9771701
  29. 29. Yoder J.A., Walsh C.P., and Bestor T.H., Cytosine methylation and the ecology of intragenomic parasites. Trends Genet, 1997. 13(8): p. 335–40. pmid:9260521
  30. 30. Cotton A.M., et al., Inactive X chromosome-specific reduction in placental DNA methylation. Hum Mol Genet, 2009. 18(19): p. 3544–52. pmid:19586922
  31. 31. Perrin D., et al., Specific hypermethylation of LINE-1 elements during abnormal overgrowth and differentiation of human placenta. Oncogene, 2007. 26(17): p. 2518–24. pmid:17043645
  32. 32. Reiss D., Zhang Y., and Mager D.L., Widely variable endogenous retroviral methylation levels in human placenta. Nucleic Acids Res, 2007. 35(14): p. 4743–54. pmid:17617638
  33. 33. Macaulay E.C., et al., Retrotransposon hypomethylation in melanoma and expression of a placenta-specific gene. PLoS ONE, 2014. 9(4): p. e95840. pmid:24759919
  34. 34. Chatterjee A., et al., Placental Hypomethylation Is More Pronounced in Genomic Loci Devoid of Retroelements. G3 (Bethesda), 2016. 6(7): p. 1911–21.
  35. 35. Beard J., Embryological Aspects and Etiology of Carcinoma. The Lancet, 1902. 159(4112): p. 1758–1761.
  36. 36. Adamson E.D., Oncogenes in development. Development, 1987. 99(4): p. 449–71. pmid:2822372
  37. 37. Conway F.J., A proposed model of cancer as the inappropriate expression of non-body introns. J Theor Biol, 1983. 100(1): p. 1–24. pmid:6220182
  38. 38. Ferretti C., et al., Molecular circuits shared by placental and cancer cells, and their implications in the proliferative, invasive and migratory capacities of trophoblasts. Hum Reprod Update, 2007. 13(2): p. 121–41. pmid:17068222
  39. 39. Lorincz M.C. and Schubeler D., Evidence for Converging DNA Methylation Pathways in Placenta and Cancer. Dev Cell, 2017. 43(3): p. 257–258. pmid:29112847
  40. 40. Nordor A.V., et al., The early pregnancy placenta foreshadows DNA methylation alterations of solid tumors. Epigenetics, 2017. 12(9): p. 793–803. pmid:28678605
  41. 41. Guo H., et al., The DNA methylation landscape of human early embryos. Nature, 2014. 511(7511): p. 606–10. pmid:25079557
  42. 42. Reik W. and Walter J., Genomic imprinting: parental influence on the genome. Nat Rev Genet, 2001. 2(1): p. 21–32. pmid:11253064
  43. 43. Wilkins J.F. and Haig D., What good is genomic imprinting: the function of parent-specific gene expression. Nat Rev Genet, 2003. 4(5): p. 359–68. pmid:12728278
  44. 44. Tucci V., et al., Genomic Imprinting and Physiological Processes in Mammals. Cell, 2019. 176(5): p. 952–965. pmid:30794780
  45. 45. Ubeda F. and Wilkins J.F., Imprinted genes and human disease: an evolutionary perspective. Adv Exp Med Biol, 2008. 626: p. 101–15. pmid:18372794
  46. 46. Hamada H., et al., Allele-Specific Methylome and Transcriptome Analysis Reveals Widespread Imprinting in the Human Placenta. Am J Hum Genet, 2016. 99(5): p. 1045–1058. pmid:27843122
  47. 47. Das R., et al., DNMT1 and AIM1 Imprinting in human placenta revealed through a genome-wide screen for allele-specific DNA methylation. BMC Genomics, 2013. 14: p. 685. pmid:24094292
  48. 48. Ng H.K., et al., Distinct patterns of gene-specific methylation in mammalian placentas: implications for placental evolution and function. Placenta, 2010. 31(4): p. 259–68. pmid:20167366
  49. 49. Novakovic B., et al., DNA methylation-mediated down-regulation of DNA methyltransferase-1 (DNMT1) is coincident with, but not essential for, global hypomethylation in human placenta. J Biol Chem, 2010. 285(13): p. 9583–93. pmid:20071334
  50. 50. Biniszkiewicz D., et al., Dnmt1 overexpression causes genomic hypermethylation, loss of imprinting, and embryonic lethality. Mol Cell Biol, 2002. 22(7): p. 2124–35. pmid:11884600
  51. 51. Mukhopadhyay A., et al., Placental expression of DNA methyltransferase 1 (DNMT1): Gender-specific relation with human placental growth. Placenta, 2016. 48: p. 119–125. pmid:27871462
  52. 52. Dedeurwaerder S., et al., Evaluation of the Infinium Methylation 450K technology. Epigenomics, 2011. 3(6): p. 771–84. pmid:22126295
  53. 53. Pidsley R., et al., Critical evaluation of the Illumina MethylationEPIC BeadChip microarray for whole-genome DNA methylation profiling. Genome Biol, 2016. 17(1): p. 208. pmid:27717381
  54. 54. Januar V., et al., Epigenetic regulation of human placental function and pregnancy outcome: considerations for causal inference. Am J Obstet Gynecol, 2015. 213(4 Suppl): p. S182–96.
  55. 55. Saffari A., et al., Estimation of a significance threshold for epigenome-wide association studies. Genet Epidemiol, 2018. 42(1): p. 20–33. pmid:29034560
  56. 56. Tsai P.C. and Bell J.T., Power and sample size estimation for epigenome-wide association scans to detect differential DNA methylation. Int J Epidemiol, 2015. 44(4): p. 1429–1441. pmid:25972603
  57. 57. Chiu R.W., et al., Noninvasive prenatal diagnosis of fetal chromosomal aneuploidy by massively parallel genomic sequencing of DNA in maternal plasma. Proc Natl Acad Sci U S A, 2008. 105(51): p. 20458–63. pmid:19073917
  58. 58. Lo Y.M., et al., Prenatal diagnosis of fetal RhD status by molecular analysis of maternal plasma. N Engl J Med, 1998. 339(24): p. 1734–8. pmid:9845707
  59. 59. Chan K.C., et al., Hypermethylated RASSF1A in maternal plasma: A universal fetal DNA marker that improves the reliability of noninvasive prenatal diagnosis. Clin Chem, 2006. 52(12): p. 2211–8. pmid:17068167
  60. 60. Jiang P., et al., Gestational Age Assessment by Methylation and Size Profiling of Maternal Plasma DNA: A Feasibility Study. Clin Chem, 2017. 63(2): p. 606–608. pmid:27979959
  61. 61. Tsui D.W., et al., Quantitative aberrations of hypermethylated RASSF1A gene sequences in maternal plasma in pre-eclampsia. Prenat Diagn, 2007. 27(13): p. 1212–8. pmid:17994635
  62. 62. Yuen R.K., et al., DNA methylation profiling of human placentas reveals promoter hypomethylation of multiple genes in early-onset preeclampsia. Eur J Hum Genet, 2010. 18(9): p. 1006–12. pmid:20442742
  63. 63. Makaroun S.P. and Himes K.P., Differential Methylation of Syncytin-1 and 2 Distinguishes Fetal Growth Restriction from Physiologic Small for Gestational Age. AJP Rep, 2018. 8(1): p. e18–e24. pmid:29472990
  64. 64. Tilley S.K., et al., Placental CpG methylation of infants born extremely preterm predicts cognitive impairment later in life. PLoS ONE, 2018. 13(3): p. e0193271. pmid:29513726
  65. 65. Mhatre M., et al., 909: Methylation profile of circulating cfDNA at term demonstrates relatively low placental contribution to hypomethylated cfDNA. American Journal of Obstetrics and Gynecology, 2019. 220(1, Supplement): p. S587–588.
  66. 66. Della Ragione F., et al., Differential DNA methylation as a tool for noninvasive prenatal diagnosis (NIPD) of X chromosome aneuploidies. J Mol Diagn, 2010. 12(6): p. 797–807. pmid:20847278
  67. 67. Jensen T.J., et al., Whole genome bisulfite sequencing of cell-free DNA and its cellular contributors uncovers placenta hypomethylated domains. Genome Biol, 2015. 16: p. 78. pmid:25886572
  68. 68. Sun K., et al., Noninvasive reconstruction of placental methylome from maternal plasma DNA: Potential for prenatal testing and monitoring. Prenat Diagn, 2018. 38(3): p. 196–203. pmid:29334402
  69. 69. Suryawanshi H., et al., A single-cell survey of the human first-trimester placenta and decidua. Sci Adv, 2018. 4(10): p. eaau4788. pmid:30402542
  70. 70. Kasak L., et al., Extensive load of somatic CNVs in the human placenta. Sci Rep, 2015. 5: p. 8342. pmid:25666259
  71. 71. Benirschke, K., Burton, G.J., Baergen, R.N., Pathology of the Human Placenta, ed. Sixth. 2012.
    • 72. Houseman E.A., et al., DNA methylation arrays as surrogate measures of cell mixture distribution. BMC Bioinformatics, 2012. 13: p. 86. pmid:22568884
    • 73. Liu Y., et al., Single-cell RNA-seq reveals the diversity of trophoblast subtypes and patterns of differentiation in the human placenta. Cell Res, 2018. 28(8): p. 819–832. pmid:30042384
    • 74. Nelson A.C., et al., Single-cell RNA-seq reveals cell type-specific transcriptional signatures at the maternal-foetal interface during pregnancy. Nat Commun, 2016. 7: p. 11414. pmid:27108815
    • 75. Pavlicev M., et al., Single-cell transcriptomics of the human placenta: inferring the cell communication network of the maternal-fetal interface. Genome Res, 2017. 27(3): p. 349–361. pmid:28174237
    • 76. Tsang J.C.H., et al., Integrative single-cell and cell-free plasma RNA transcriptomics elucidates placental cellular dynamics. Proc Natl Acad Sci U S A, 2017. 114(37): p. E7786–E7795. pmid:28830992
    • 77. Vento-Tormo R., et al., Single-cell reconstruction of the early maternal-fetal interface in humans. Nature, 2018. 563(7731): p. 347–353. pmid:30429548
    • 78. Lara-Astiaso D., et al., Immunogenetics. Chromatin state dynamics during blood formation. Science, 2014. 345(6199): p. 943–9. pmid:25103404
    • 79. Schmidl C., et al., ChIPmentation: fast, robust, low-input ChIP-seq for histones and transcription factors. Nat Methods, 2015. 12(10): p. 963–5. pmid:26280331
    • 80. Buenrostro J.D., et al., Single-cell chromatin accessibility reveals principles of regulatory variation. Nature, 2015. 523(7561): p. 486–90. pmid:26083756
    • 81. van Eijl R., et al., Single-Cell ID-seq Reveals Dynamic BMP Pathway Activation Upstream of the MAF/MAFB-Program in Epidermal Differentiation. iScience, 2018. 9: p. 412–422. pmid:30466065
    • 82. Angermueller C., et al., Parallel single-cell sequencing links transcriptional and epigenetic heterogeneity. Nat Methods, 2016. 13(3): p. 229–232. pmid:26752769
    • 83. Clark S.J., et al., scNMT-seq enables joint profiling of chromatin accessibility DNA methylation and transcription in single cells. Nat Commun, 2018. 9(1): p. 781. pmid:29472610
    • 84. Gerlach J.P., et al., Combined quantification of intracellular (phospho-)proteins and transcriptomics from fixed single cells. Sci Rep, 2019. 9(1): p. 1469. pmid:30728416
    • 85. Haider S., et al., Self-Renewing Trophoblast Organoids Recapitulate the Developmental Program of the Early Human Placenta. Stem Cell Reports, 2018. 11(2): p. 537–551. pmid:30078556
    • 86. Turco M.Y., et al., Trophoblast organoids as a model for maternal-fetal interactions during human placentation. Nature, 2018.
    • 87. Novakovic B., et al., Wide-ranging DNA methylation differences of primary trophoblast cell populations and derived cell lines: implications and opportunities for understanding trophoblast function. Mol Hum Reprod, 2011. 17(6): p. 344–53. pmid:21289002
    • 88. Grindler N.M., et al., Exposure to Phthalate, an Endocrine Disrupting Chemical, Alters the First Trimester Placental Methylome and Transcriptome in Women. Sci Rep, 2018. 8(1): p. 6086. pmid:29666409
    • 89. Alexander J., et al., Offspring sex impacts DNA methylation and gene expression in placentae from women with diabetes during pregnancy. PLoS ONE, 2018. 13(2): p. e0190698. pmid:29470513
    • 90. Brunst K.J., et al., Cumulative lifetime maternal stress and epigenome-wide placental DNA methylation in the PRISM cohort. Epigenetics, 2018. 13(6): p. 665–681. pmid:30001177
    • 91. Morales E., et al., Genome-wide DNA methylation study in human placenta identifies novel loci associated with maternal smoking during pregnancy. Int J Epidemiol, 2016. 45(5): p. 1644–1655. pmid:27591263
    • 92. Cardenas A., et al., Placental DNA Methylation Adaptation to Maternal Glycemic Response in Pregnancy. Diabetes, 2018. 67(8): p. 1673–1683. pmid:29752424
    • 93. Fa S., et al., Assessment of global DNA methylation in the first trimester fetal tissues exposed to maternal cigarette smoking. Clin Epigenetics, 2016. 8: p. 128. pmid:27924165
    • 94. Loke Y.J., et al., Time- and sex-dependent associations between prenatal alcohol exposure and placental global DNA methylation. Epigenomics, 2018. 10(7): p. 981–991. pmid:29956547
    • 95. Kertes D.A., et al., BNDF methylation in mothers and newborns is associated with maternal exposure to war trauma. Clin Epigenetics, 2017. 9: p. 68. pmid:28680507
    • 96. Basak S., Srinivas V., and Duttaroy A.K., Bisphenol-A impairs cellular function and alters DNA methylation of stress pathway genes in first trimester trophoblast cells. Reprod Toxicol, 2018. 82: p. 72–79. pmid:30352284
    • 97. Abraham E., et al., Pregnancy exposure to atmospheric pollution and meteorological conditions and placental DNA methylation. Environ Int, 2018. 118: p. 334–347. pmid:29935799
    • 98. Maghbooli Z., et al., Air pollution during pregnancy and placental adaptation in the levels of global DNA methylation. PLoS ONE, 2018. 13(7): p. e0199772. pmid:29979694
    • 99. Everson T.M., et al., Cadmium-associated differential methylation throughout the placental genome: epigenome-wide association study of two US birth cohorts. Environmental health perspectives, 2018. 126(1): p. 017010. pmid:29373860
    • 100. Reichetzeder C., et al., Increased global placental DNA methylation levels are associated with gestational diabetes. Clinical epigenetics, 2016. 8(1): p. 82.
    • 101. Zhao Y., et al., Third trimester phthalate exposure is associated with DNA methylation of growth-related genes in human placenta. Sci Rep, 2016. 6: p. 33449. pmid:27653773
    • 102. Green B.B., et al., Epigenome-Wide Assessment of DNA Methylation in the Placenta and Arsenic Exposure in the New Hampshire Birth Cohort Study (USA). Environ Health Perspect, 2016. 124(8): p. 1253–60. pmid:26771251
    • 103. Kim S., et al., Prenatal exposure to persistent organic pollutants and methylation of LINE-1 and imprinted genes in placenta: a CHECK cohort study. Environment international, 2018. 119: p. 398–406. pmid:30005188
    • 104. Kingsley S.L., et al., Maternal residential proximity to major roadways, birth weight, and placental DNA methylation. Environment international, 2016. 92: p. 43–49. pmid:27058926
    • 105. Mitsuya K., et al., Alterations in the placental methylome with maternal obesity and evidence for metabolic regulation. PLoS ONE, 2017. 12(10): p. e0186115. pmid:29045485
    • 106. Neven K.Y., et al., Placental promoter methylation of DNA repair genes and prenatal exposure to particulate air pollution: an ENVIRONAGE cohort study. The Lancet Planetary Health, 2018. 2(4): p. e174–e183. pmid:29615218
    • 107. Schmidt R.J., et al., Self-reported pregnancy exposures and placental DNA methylation in the MARBLES prospective autism sibling study. Environmental epigenetics, 2016. 2(4): p. dvw024. pmid:28781890
    • 108. Shorey-Kendrick L.E., et al., Vitamin C Prevents Offspring DNA Methylation Changes Associated with Maternal Smoking in Pregnancy. Am J Respir Crit Care Med, 2017. 196(6): p. 745–755. pmid:28422514
    • 109. Ma M., et al., Preeclampsia is associated with hypermethylation of IGF-1 promoter mediated by DNMT1. Am J Transl Res, 2018. 10(1): p. 16–39. pmid:29422991
    • 110. Martin E., et al., Epigenetics and Preeclampsia: Defining Functional Epimutations in the Preeclamptic Placenta Related to the TGF-beta Pathway. PLoS ONE, 2015. 10(10): p. e0141294. pmid:26510177
    • 111. Herzog E.M., et al., Early- and late-onset preeclampsia and the tissue-specific epigenome of the placenta and newborn. Placenta, 2017. 58: p. 122–132. pmid:28962690
    • 112. van den Berg C.B., et al., Early- and late-onset preeclampsia and the DNA methylation of circadian clock and clock-controlled genes in placental and newborn tissues. Chronobiol Int, 2017. 34(7): p. 921–932. pmid:28613964
    • 113. Paauw N.D., et al., H3K27 acetylation and gene expression analysis reveals differences in placental chromatin activity in fetal growth restriction. Clin Epigenetics, 2018. 10: p. 85. pmid:29983832
    • 114. Chen P.-Y., et al., Prenatal growth patterns and birthweight are associated with differential DNA methylation and gene expression of cardiometabolic risk genes in human placentas: a discovery-based Approach. Reproductive Sciences, 2018. 25(4): p. 523–539. pmid:28693373
    • 115. Banister C.E., et al., Infant growth restriction is associated with distinct patterns of DNA methylation in human placentas. Epigenetics, 2011. 6(7): p. 920–7. pmid:21758004
    • 116. Khot V.V., et al., Variable methylation potential in preterm placenta: implication for epigenetic programming of the offspring. Reproductive Sciences, 2017. 24(6): p. 891–901. pmid:27678102
    • 117. Leavey K., et al., Epigenetic regulation of placental gene expression in transcriptional subtypes of preeclampsia. Clin Epigenetics, 2018. 10: p. 28. pmid:29507646
    • 118. Paquette A.G., et al., Regions of variable DNA methylation in human placenta associated with newborn neurobehavior. Epigenetics, 2016. 11(8): p. 603–13. pmid:27366929
    • 119. Price E.M., et al., Profiling placental and fetal DNA methylation in human neural tube defects. Epigenetics & chromatin, 2016. 9(1): p. 6.
    • 120. Roifman M., et al., Genome-wide placental DNA methylation analysis of severely growth-discordant monochorionic twins reveals novel epigenetic targets for intrauterine growth restriction. Clinical epigenetics, 2016. 8(1): p. 70.
    • 121. Schroeder D.I., et al., Placental methylome analysis from a prospective autism study. Mol Autism, 2016. 7: p. 51. pmid:28018572
    • 122. Wilson S.L., et al., Mining DNA methylation alterations towards a classification of placental pathologies. Human molecular genetics, 2017. 27(1): p. 135–146.
    • 123. Konwar C., et al., DNA methylation profiling of acute chorioamnionitis-associated placentas and fetal membranes: insights into epigenetic variation in spontaneous preterm births. Epigenetics Chromatin, 2018. 11(1): p. 63. pmid:30373633