Epigenetics and Reproductive Health (Volume 21) (Translational Epigenetics, Volume 21) [1 ed.] 0128197536, 9780128197530

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Translational Epigenetics Series Trygve O. Tollefsbol, Series Editor Transgenerational Epigenetics Edited by Trygve O. Tollefsbol, 2014

Epigenetic Mechanisms in Cancer Edited by Sabita Saldanha, 2017

Personalized Epigenetics Edited by Trygve O. Tollefsbol, 2015

Epigenetics of Aging and Longevity Edited by Alexey Moskalev and Alexander M. Vaiserman, 2017

Epigenetic Technological Applications Edited by Y. George Zheng, 2015 Epigenetic Cancer Therapy Edited by Steven G. Gray, 2015 DNA Methylation and Complex Human Disease By Michel Neidhart, 2015 Epigenomics in Health and Disease Edited by Mario F. Fraga and Agustin F. F Ferna´ndez, 2015 Epigenetic Gene Expression and Regulation Edited by Suming Huang, Michael Litt and C. Ann Blakey, 2015 Epigenetic Biomarkers and Diagnostics Edited by Jose Luis Garcı´a-Gim_enez, 2015 Drug Discovery in Cancer Epigenetics Edited by Gerda Egger and Paola Barbara Arimondo, 2015

The Epigenetics of Autoimmunity Edited by Rongxin Zhang, 2018 Epigenetics in Human Disease, Second Edition Edited by Trygve O. Tollefsbol, 2018 Epigenetics of Chronic Pain Edited by Guang Bai and Ke Ren, 2018 Epigenetics of Cancer Prevention Edited by Anupam Bishayee and Deepak Bhatia, 2018 Computational Epigenetics and Diseases Edited by Loo Keat Wei, 2019 Pharmacoepigenetics Edited by Ramo´n Cacabelos, 2019 Epigenetics and Regeneration Edited by Daniela Palacios, 2019

Medical Epigenetics Edited by Trygve O. Tollefsbol, 2016

Chromatin Signaling and Neurological Disorders Edited by Olivier Binda, 2019

Chromatin Signaling and Diseases Edited by Olivier Binda and Martin Fernandez-Zapico, 2016

Transgenerational Epigenetics, Second Edition Edited by Trygve Tollefsbol, 2019

Genome Stability Edited by Igor Kovalchuk and Olga Kovalchuk, 2016

Nutritional Epigenomics Edited by Bradley Ferguson, 2019 Prognostic Epigenetics Edited by Shilpy Sharma, 2019

Chromatin Regulation and Dynamics Edited by Anita GVondVor, 2016

Epigenetics of the Immune System Edited by Dieter Kabelitz, 2020

Neuropsychiatric Disorders and Epigenetics Edited by Dag H. Yasui, Jacob Peedicayil and Dennis R. Grayson, 2016

Stem Cell Epigenetics Edited by Eran Meshorer and Giuseppe Testa, 2020

Polycomb Group Proteins Edited by Vincenzo Pirrotta, 2016

Epigenetics Methods Edited by Trygve Tollefsbol, 2020

Epigenetics and Systems Biology Edited by Leonie Ringrose, 2017

RNA-Based Regulation in Human Health and Disease Edited by Rajesh Pandey, 2020

Cancer and Noncoding RNAs Edited by Jayprokas Chakrabarti and Sanga Mitra, 2017

Histone Modifications in Therapy Editor by Pedro Castelo-Branco and Carmen Jeronimo, 2020

Nuclear Architecture and Dynamics Edited by Christophe Lavelle and Jean-Marc Victor, 2017

Translational Epigenetics

Epigenetics and Reproductive Health Volume 21

Series Editor Trygve Tollefsbol

Series Volume Editors Nafisa Balasinor Priyanka Parte Dipty Singh

With immense gratitude, we dedicate this book to the memory of our mentor late Dr Harbans Singh Juneja who introduced us to research in the area of neuroendocrinology and this exciting world of the epigenome and its role in reproductive physiology.

Academic Press is an imprint of Elsevier 125 London Wall, London EC2Y 5AS, United Kingdom 525 B Street, Suite 1650, San Diego, CA 92101, United States 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom Copyright © 2021 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-0-12-819753-0 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Andre Gerhard Wolff Acquisitions Editor: Peter B. Linsley Editorial Project Manager: Megan Ashdown Production Project Manager: Maria Bernard Cover Designer: Miles Hitchen Typeset by TNQ Technologies

Contributors Ummet Abur Department of Medical Genetics, Faculty of Medicine, Ondokuz Mayıs University, Samsun, Turkey; Department of Multidisciplinary Molecular Medicine, Health Sciences Institute, Ondokuz Mayıs University, Samsun, Turkey Ahmed Hamed Arisha Department of Physiology, Faculty of Veterinary Medicine, Zagazig University, Zagazig, Egypt Brooke Armistead Michigan State University, Department of Obstetrics, Gynecology and Reproductive Biology, College of Human Medicine, Grand Rapids, MI, United States Kenneth I. Aston Andrology and IVF Laboratories, Department of Surgery, University of Utah School of Medicine, Salt Lake City, UT, United States N.H. Balasinor Neuroendocrinology, ICMR- National Institute for Research in Reproductive Health, Mumbai, Maharashtra, India Barbara Benassi Laboratory of Health and Environment, ENEA, Rome, Italy Douglas T. Carrell Andrology and IVF Laboratories, Department of Surgery, University of Utah School of Medicine, Salt Lake City, UT, United States Shrijeet Chakraborti Leighton Hospital, Mid Cheshire Hospitals NHS Foundation Trust, Crewe, Cheshire, United Kingdom Eugenia Cordelli Laboratory of Health and Environment, ENEA, Rome, Italy Kinjal Dave Mother and Child Health, Interactive Research School for Health Affairs (IRSHA), Bharati Vidyapeeth (Deemed to be) University, Pune, Maharashtra, India Sharvari Deshpande Neuroendocrinology, ICMR- National Institute for Research in Reproductive Health, Mumbai, Maharashtra, India Sascha Drewlo Michigan State University, Department of Obstetrics, Gynecology and Reproductive Biology, College of Human Medicine, Grand Rapids, MI, United States

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Contributors

Anthony R. Gostick Andrology and IVF Laboratories, Department of Surgery, University of Utah School of Medicine, Salt Lake City, UT, United States Sezgin Gunes Department of Multidisciplinary Molecular Medicine, Health Sciences Institute, Ondokuz Mayıs University, Samsun, Turkey; Department of Medical Biology, Faculty of Medicine, Ondokuz Mayıs University, Samsun, Turkey Nojan Hafizi Near East University, Institute of Health Sciences, Department of Medical Biology and Genetics, Nicosia, Cyprus Jinlian Hua College of Veterinary Medicine, Shaanxi Centre of Stem Cells Engineering & Technology, Northwest A&F University, Yangling, Shaanxi, China John Huntriss Discovery and Translational Science Department (DTSD), University of Leeds, Leeds, West Yorkshire, Great Britain Arif Hussain School of Life Sciences, Manipal Academy of Higher Education, Dubai, United Arab Emirates Hiroki Ikeda Nara Medical University, Department of Embryology, Kashihara, Nara, Japan Emma R. James Andrology and IVF Laboratories, Department of Surgery, University of Utah School of Medicine, Salt Lake City, UT, United States Timothy G. Jenkins Andrology and IVF Laboratories, Department of Surgery, University of Utah School of Medicine, Salt Lake City, UT, United States Eugenia Johnson Michigan State University, Department of Obstetrics, Gynecology and Reproductive Biology, College of Human Medicine, Grand Rapids, MI, United States Sadhana Joshi Mother and Child Health, Interactive Research School for Health Affairs (IRSHA), Bharati Vidyapeeth (Deemed to be) University, Pune, Maharashtra, India Shama Prasada Kabekkodu Department of Cell and Molecular Biology, Manipal School of Life Sciences, Manipal Academy of Higher Education, Manipal, Karnataka, India

Contributors

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Leena Kadam Department of Obstetrics and Gynecology, Wayne State University School of Medicine, Detroit, MI, United States Jyotdeep Kaur Department of Biochemistry, Postgraduate Institute of Medical Education and Research, Chandigarh, India Hisato Kobayashi Nara Medical University, Department of Embryology, Kashihara, Nara, Japan Hamid-Reza Kohan-Ghadr Michigan State University, Department of Obstetrics, Gynecology and Reproductive Biology, College of Human Medicine, Grand Rapids, MI, United States Takeo Kubota Faculty of Child Studies, Seitoku University, Chiba, Japan; Graduate School of Teacher Education, Seitoku University, Chiba, Japan Kazuki Kurimoto Nara Medical University, Department of Embryology, Kashihara, Nara, Japan Aatish Mahajan Department of Biochemistry, Postgraduate Institute of Medical Education and Research, Chandigarh, India Sweta Nair Neuroendocrinology, ICMR- National Institute for Research in Reproductive Health, Mumbai, Maharashtra, India Lakshmi Natarajan Independent Researcher, NJ, United States Francesca Pacchierotti Laboratory of Health and Environment, ENEA, Rome, Italy Priyanka Parte Department of Gamete Immunobiology, ICMR-National Institute for Research in Reproductive Health, Mumbai, Maharashtra, India Aniket G. Patankar Department of Gamete Immunobiology, ICMR-National Institute for Research in Reproductive Health, Mumbai, Maharashtra, India Sahar Qazi Department of Biochemistry, All India Institute of Medical Sciences, New Delhi, Delhi, India

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Contributors

Beenish Rahat Division of Intramural Research, Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, United States Albert Salas-Huetos Andrology and IVF Laboratories, Department of Surgery, University of Utah School of Medicine, Salt Lake City, UT, United States Sabita N. Saldanha Alabama State University, Department of Biological Sciences, Montgomery, AL, United States Divika Sapehia Department of Biochemistry, Postgraduate Institute of Medical Education and Research, Chandigarh, India Ashok Sharma Department of Biochemistry, All India Institute of Medical Sciences, New Delhi, Delhi, India Shefina Silas School of Life Sciences, Manipal Academy of Higher Education, Dubai, United Arab Emirates Isha Singh Department of Gamete Immunobiology, ICMR-National Institute for Research in Reproductive Health, Mumbai, Maharashtra, India Parampal Singh Department of Biochemistry, Postgraduate Institute of Medical Education and Research, Chandigarh, India Madhumitha Kedhari Sundaram School of Life Sciences, Manipal Academy of Higher Education, Dubai, United Arab Emirates Deepali Sundrani Mother and Child Health, Interactive Research School for Health Affairs (IRSHA), Bharati Vidyapeeth (Deemed to be) University, Pune, Maharashtra, India Padmanaban S. Suresh School of Biotechnology, National Institute of Technology, Calicut, Kerala, India Burak Tatar Department of Gynecologic Oncology, Health Sciences University Samsun Research and Training Hospital, Samsun, Turkey; Department of Multidisciplinary Molecular Medicine, Health Sciences Institute, Ondokuz Mayıs University, Samsun, Turkey Shilpa Thakur National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, United States

Contributors

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Sanu Thankachan School of Biotechnology, National Institute of Technology, Calicut, Kerala, India Pinar Tulay Near East University, Faculty of Medicine, Department of Medical Genetics, Nicosia, Cyprus; Near East University, DESAM Institute, Nicosia, Cyprus Eva Tvrda´ Department of Animal Physiology, Faculty of Biotechnology and Food Sciences, Slovak University of Agriculture in Nitra, Tr. A. Hlinku 2, Nitra, Slovakia Thejaswini Venkatesh Department of Biochemistry and Molecular Biology, Central University of Kerala, Kasaragod, Kerala, India Juqing Zhang College of Veterinary Medicine, Shaanxi Centre of Stem Cells Engineering & Technology, Northwest A&F University, Yangling, Shaanxi, China

Acknowledgment The Editorial team acknowledges the contributions by all the authors and co-authors to the chapters resulting in a comprehensive book on Epigenetics and Reproductive Health. We wish to thank the entire support team of Elsevier for the smooth publication of the book. We also thank Megan Ashdown for efficient coordination with the authors and editors.

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Introduction to epigenetics: basic concepts and advancements in the field Dipty Singh*, Kumari Nishi*, Kushaan Khambata*, N.H. Balasinor Neuroendocrinology, ICMR- National Institute for Research in Reproductive Health, Mumbai, Maharashtra, India

Introduction Reproduction is a fundamental process of replicating life which also contributes toward heritability of traits from one generation to the next [1]. The heritability is not only governed by genetic information harbored in gametes but also dictated by non-genetic factors such as epigenetics. Germ cell development and early embryo development are the crucial reproductive events when epigenetic landscape is architectured or maintained [2]. The developmental exposure to certain lifestyle and environmental factors may affect the phenotype of the next generation through remodeling of the epigenetic blueprint of gametes [3]. It is well understood that genetic factors contribute to risk of many diseases affecting human health. Evidence is slowly accumulating that genetic factors are not the only repository of all information in the health status of an organism; other external factors may also influence health via epigenetic modulations. Similarly, along with the genetic factors, epigenetic factors also influence reproductive health and fertility [3]. Therefore, understanding epigenetic modifications, basic cellular physiology and the underlying cause of a disease condition becomes important [4]. New advancements in mapping human epigenome has helped researchers to understand disease etiology in past three decades. This provides possibilities to interpret epigenetic code and develop better treatment strategies. This chapter presents an overview of key epigenetic mechanisms available techniques for epigenetic analysis and recent advances in epigenetic technology.

Epigenetics Definition Although all of more than 200 cell types in humans have the same DNA sequences, yet they exhibit different gene expression profiles and phenotypes. The phenotypic changes that occur in the cells during the course of development in a multicellular organism were originally described as an “epigenetic landscape” by developmental biologist Conrad Waddingtion. Later, Holliday defined “epigenetics” as nuclear inheritance which is not based on differences in DNA sequence. In its more modern version, epigenetics is molecularly and mechanistically described as the sum of modifications to the chromatin template that come together to establish and propagate different patterns of gene expression and silencing from the same genome. If “genetics” can be considered as words, “epigenetics” instructs how these words are read. Alternatively, if groups of genes can be thought of as computer hardware, epigenetic control can be compared with computer software. Thus, epigenetics is an additional regulatory layer that provides insights into how cellular events are coordinated [5]. *

Equal contribution.

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Epigenetic changes to the chromatin are brought about by three main mechanisms; namely, DNA methylation, post-translational histone modifications and non-coding RNAs. We attempt to provide a brief overview of these concepts in this chapter.

DNA methylation Since its discovery more than 35 years ago [6], DNA methylation at the cytosine residues is recognized as one of the prime epigenetic mechanisms regulating gene expression. Cytosine residues are converted by the addition of a methyl group to 5-methylcytosine (5mC) in the DNA template. It mainly occurs in dinucleotide sequence 50 CpG30 (i.e. CpG abbreviated for cytosine and guanine separated by phosphate group in the DNA). It can be transmitted by both the DNA strands, from mother to daughter cells during DNA replication and, can thus be inherited through cell division. Since DNA methylation patterns are heritable, they provide an epigenetic marking for the genome that is stable through multiple cell divisions, and thus contribute to form a cellular memory. CpGs are often found in clusters, and are called CpG islands. They are usually enriched at noncoding regions (e.g. centromeric heterochromatin) and interspersed repetitive elements (e.g. retrotransposons). CpG islands are also commonly found in the upstream region in the gene promoters where they regulate gene expression [7]. The methyl moiety of cytosines lies in the major groove of the DNA helix where many DNA-binding proteins interact with DNA. Thus, DNA methylation results in attraction or repulsion of various DNA-binding proteins. The binding of methyl-CpG binding domain (MBD) proteins to methylated CpGs recruits repressor complexes, (like HDACs, which abrogate activating histone acetylation marks) to the methylated promoter regions causing transcriptional silencing. Conversely, CpG methylation inhibits binding of certain transcriptional regulators like CTCF, thereby preventing transcription. Thus, in general, DNA methylation brings about gene repression or silencing and plays a crucial role in cellular differentiation, X-chromosome inactivation and genomic imprinting [8]. An important function of DNA methylation is to protect the genomic integrity by silencing transposable elements and ensure genomic stability. Although DNA methylation patterns are transmitted from cell to cell during cell division, they are not permanent. DNA methylation patterns can change throughout the lifetime of an individual. These changes can be a physiological response to environmental stimuli or may be associated with pathological processes like oncogenic transformation and aging [5].

DNA methylation: establishment and erasure In mammals, DNA methylation is brought about by DNA methyltransferases (Dnmts), which include three proteins belonging to two families that are structurally and functionally distinct. The first family includes the maintenance methyltransferase Dnmt1, which preferentially methylates hemimethylated CpG dinucleotides (i.e. DNA methylated at CpG in one of the two strands). Thus Dnmt1 is present at the replication fork and is responsible for semiconservative replicating DNA methylation patterns. The other family consists of de novo methyltransferases Dnmt3a and 3b, which lay down de novo DNA methylation patterns in early embryo development. The functions of Dnmt3a and 3b are guided by Dnmt3l, which itself lacks active methyltransferase activity, but is essential for sequence specific de novo methylation activities. DNA methylation marks can be removed by active demethylation process, which involves a family of DNA hydroxylases called ten-eleven translocation (Tet) proteins or by a passive demethylation process by inhibition of Dnmt1 during cell division [9].

Introduction to epigenetics

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The methyl group on CpG can be oxidized by Tet enzymes to 5-hydroxymethylcytosine (5hmC), which is another DNA modification found across the genome specifically at the transcription start sites and are enriched in the active chromatin regions, and are thus involved in regulation of gene expression. The Tet family of proteins can further oxidize 5hmC into 5-carboxylcytosine (5caC) and 5formylcytosine (5fC) utilizing ATP. These are less stable marks and activate base excision repair pathway, which ultimately leads to removal of the modified base and returns the 5mC to an unmethylated cytosine and thus promotes transcriptional activation [9].

Histone modifications In multicellular organisms, the DNA is packaged into a nucleoprotein complex called chromatin. The chromatin consists of DNA wrapped around a core of highly basic proteins called histones. The fundamental unit of the chromatin is called a nucleosome, which consists of 146 base pairs (bp) of DNA wrapped approximately twice around an octamer of core histones. Each nucleosome consists of two of each core histones, H2A, H2B, H3 and H4. In each nucleosome, the DNA and histone octamer core are associated with H1 linker protein. The histone core is bound to DNA by non-covalent ionic interactions between positively charged residues on the histone proteins and phosphate groups on the DNA [10,11].

Higher-order chromatin organization These nucleosomes form the building blocks of higher order chromatin structure. In its completely unfolded confirmation, the chromatin structures are visualised microscopically as 11-nm polymers with a “bead-on-string” appearance. According to the solenoid model of chromatin fiber, the nucleosomes are arranged into a helical array of about six to eight nucleosomes per turn with the histone H1 on the inside of the fiber. The linker DNA is bent to connect each nucleosome with the next one along the helical filament. This results in a more compacted 30 nm transcriptionally incompetent chromatin conformation. The chromatin can then be organized into larger looped domains (300e700 nm). The most condensed chromatin structure is formed at the time of chromosome formation in the metaphase of mitosis or meiosis, to permit faithful segregation of the genome [5]. Chromatin is a highly dynamic structure, and exists in many conformations. It is historically classified as euchromatin and heterochromatin. Euchromatin or “active” chromatin consists of coding and regulatory (e.g., promoters and enhancers) regions of the genome. It exists in an open, decompacted confirmation, favoring active transcription or “poised” for gene expression. Heterochromatin refers to the “inactive” regions of the genome existing in a closed, highly compacted state. It contributes to the great majority of the genome; and comprises of non-coding sequences and repetitive elements (e.g., retrotransposons, satellite repeats and LINEs). Heterochomatin can exists in two states: the permanently silenced “constitutive” heterochromatin, which generally found at pericentric and subtelomeric regions or as “facultative” heterochromatin, in which genes can be derepressed during a specific cell cycle or developmental stage [5]. The core histones are some of the most evolutionarily conserved eukaryotic proteins. They consist of a basic N-terminal domain and a histone-fold C-terminal domain. The histone-fold or globular domain heterodimerizes with a second histone (H3 with H4, H2A with H2B); and wraps DNA around to form nucleosome. The basic N-terminal “tail” domain lies outside the nucleosome and does not

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have any defined structure. Many residues in the histone tails, particularly in histones H3 and H4, are important sites of post-translational modifications (PTMs); although some residues in the more structured globular domains are also targeted. These modifications promote nucleosomal, and hence, chromatin variability. Acetylation and methylation of core histones, especially H3 and H4, were among the first covalent modifications studied. Besides these, several other histone modifications have been identified; namely, phosphorylation, ubiquitination, sumoylation, ADP-ribosylation, biotinylation, crotonylation, among others [5]. We shall take a look at some of them in a little detail.

Histone acetylation Histone can be acetlyated at the ε-amino groups of lysine (K) residues located at the N-termini. Although, all core histones can be acetylated in vivo, acetylation of H3 and H4 are most extensively studied [12]. H3 can be acetylated at lysine positions 9, 14, 18 and 23; while the lysine positions of 5, 8, 12 and 16 can be acetylated for H4. Addition of acetyl groups neutralizes the basic charge of histone tails and thus decreases affinity for DNA. It also alters histone-to-histone interactions between neighboring nucleosomes, and the interaction of histones with other regulatory proteins. Thus, histone acetylation creates an “open” chromatin environment favorable for gene transcription and is characteristically found in euchromatin regions [13]. Histone acetylation is highly dynamic and controlled by the opposing action of two types of enzymes: histone acetyltransferases and histone deacetylases (HDACs).

Histone methylation Histone can be methylated on the side chains of lysines (K) and arginines (R) residues. Unlike histone acetylation, methylation does not affect the charge of the histone protein. Additionally, lysines residues can be mono-, di- or tri-methylated, whereas arginines may be mono-, symmetrically or asymmetrically di-methylated [14]. This enhances the level of complexity offered by this modification. Several lysine residues on positions 4, 9, 27, and 36 of H3 and lysine 20 of H4, are preferred sites of methylation [15]. Sites of arginine methylation include H3R2, H3R8, H3R17, H3R26 and H4R3 (Young et al., 2010). As opposed to acetylation, which usually results in transcriptional activation, histone methylation can signal either activation or repression, depending on the sites of methylation. For example, methylation at H3K4 results in gene activation, while H3K27 methylation leads to gene silencing. Thus, these modifications are usually found in the regulatory regions of the genes like the promoter and enhancer elements. Methylation at H3K9 position leads to a closed chromatin conformation and is consequently associated with heterochomatin regions of the genome. Histone lysine methylation is also involved in diverse set of biological processes, such as heterochromatin-mediated transcriptional silencing, DNA damage response and X chromosome inactivation. Histone lysine methylation is tightly regulated by the enzymes methyltransferases (KMTs) and demethylases (KDMs) in order to maintain cell fate and genomic stability.

Other histone modifications Histone Phosphorylation: occurs on serine, threonine and tyrosine residues mainly in the N-terminal histone tails. Histone phosphorylation confers a negative charge to the histone, resulting in a more open chromatin conformation. It is therefore associated with gene expression and is involved in DNA damage repair and chromatin remodelling [16].

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ADP ribosylation: Histones can be reversibly mono- and poly-ADP ribosylated on glutamate and arginine residues, conferring a negative charge to the histone, and thus contribute to a relaxed chromatin state. These modifications increase upon DNA damage and are involved in the DNA damage response pathway [17]. Ubiquitylation and sumoylation: Histone ubiquitylation involves addition of large ubiquitin moiety (76-amino acid polypeptide) to lysine residues. Mono-ubiquitylation of histones can bring about either gene activation and repression, whereas polyubiquitinylation labels them for proteolytic degradation. Sumoylation, like ubiquitylation, results in the covalent attachment of small ubiquitin-like modifier molecules to histone lysines. Sumoylation can occur on all four core histones and inhibits acetylation or ubiquitylation of lysine residues, thereby causing gene repression [14].

The “Histone Code”- its writers, readers and erasers The covalent modifications occurring at multiple and specific sites on the histones give rise to remarkable nucleosomal diversity. Different combinations of histone modifications can modify the chromatin structure, which in turn regulate and determine the changes in gene expression. This concept was put forth as the “Histone Code Hypothesis”. According to this hypothesis, histones modifications provide a signaling platform to alter the chromatin states in order to bring about gene activation or silencing. The enzymes which establish these histone modifications are collectively referred to as “writers” of the histone code. They include enzymes like HATs, HMTs, histone kinases among others. In this scenario, the enzymes which remove the histone modifications like the HDACs, KDMs, are called “erasers” of histone codes. Besides altering the chromatin states, these modifications also serve as binding or recognition sites for the recruitment of several effector proteins termed as “readers” of the histone codes, which in turn recruit other co-regulator complexes to bring about further changes in the chromatin structure and hence the gene expression [18].

Non-coding RNAs The Human Genome Project revealed that only a small fraction of the human transcriptome (2%e5%) encoded for proteins while the functions of rest of the transcripts were unknown. Recent advances in sequencing technologies have revealed that in fact, three-quarters of the genome is transcribed [19] (with some studies estimating more than 90%). Although this view is hotly debated, it has challenged the long standing view point that most of the genome is not transcribed and considered as nonfunctional “junk” DNA. In recent years, the classification of RNA classes has undergone a major change, with it can be broadly classified into: protein-coding messenger RNAs and noncoding RNAs. The category of noncoding RNAs have now expanded beyond the well-known ribosomal and transfer RNAs (rRNA and tRNA), to include small interfering RNA (siRNA), microRNA (miRNA), circular RNA (circ RNA), PIWI-interacting RNAs (piRNAs), long non-coding RNAs (lncRNAs), promoterassociated RNAs (PARs), enhancer RNAs (eRNAs) and many others [20]. The functions of these ncRNAs have been under intense investigations and they are now known to play many important structural and regulatory roles. More importantly, noncoding RNAs have now emerged as a separate epigenetic mechanism regulating gene expression. The ncRNAs provide a scaffold for chromatinremodelling and -modifying enzyme complexes to bring about changes in the chromatin states through cis or trans mechanisms. They can also allow recruitment of factors involved in

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silencing (e.g. co-repressor) or activation of gene transcription. These ncRNAs are thought to provide the sequence specificity to guide the chromatin modifying complexes to their sites of action. As functions of more ncRNAs are being uncovered, they represent a key layer of epigenetic regulation [21].

RNA modifications RNA can be covalently modified by vast array of chemical additions to both its sugar and nucleotide groups [22]. The additions to the sugar backbone mainly protects the RNA molecule from degradation, the modifications at the nucleotide base confers novel regulatory functions. Specifically, the posttranslational addition of methyl group to N(6) position of adenosine (m(6)A) is the most common RNA modification in the coding and non-coding RNA. It occurs predominantly in the 30 UTRs and stop codons. RNA methylation controls various steps of mRNA metabolism, microRNA mediated decay, pre-microRNA processing, RNA poly adenylation. It also affects RNA secondary structure and regulates alternative splicing for a subset of mRNAs and lncRNAs, and it is also involved in translation and RNA degradation [22,23]. This modification is brought about by the METTL3 RNA methyltransferase complex and is erased by FTO (fat-mass and obesity associated) RNA demethylase. The dynamic activity of these enzymes regulates the level of this modification, which plays a crucial role in development, metabolism, and fertility [24].

Techniques for epigenetic analysis The primary methods in epigenetics engaged specific nuclease enzymes (e.g., restriction enzymes, DNase I, MNase) and later identification of DNA and histone modifying enzymes produced a big new data in the field. Past few years have witnessed development of new techniques which directly assess chromatin modification, interacting proteins, patterns of DNA methylation and nucleosomal occupancy. Active emergence of these new technologies to assess genome-wide DNA methylation patterns, chromatin structure in modern era has accelerated the speed of understanding of epigenetic mechanisms.Various methods developed to detect and measure epigenetic marks and assess their functions depending on genome wide or loci specific approaches have been discussed below.

DNA methylation A lot of advancements have been made in DNA methylation detection technologies starting from southern blot using restriction endonucleases to microarray-based epigenomics and methylation specific polymerase chain reaction to next generation sequencing based targeted or whole genome bisulfite sequencing [41,70]. The choice of method is important to get an unbiased answer to the research question. The selection of method depends on the study aim, amount and quality of DNA sample, availability of reagents and instruments, its cost effectiveness, how robust and simple the method is to meet the sensitivity and specificity criteria. Another criterion for choosing the method depends on whether information being sought is genome-wide or specific locibased. The method can also be chosen based on whether the candidate genes are known or not [4]. The methods used for DNA methylation analysis have been enlisted in Table 1.

Table 1 Different techniques to analyze DNA methylation S.No.

Class

Method

Principle

Features

Limitations

1.

Profiling Whole Genome Methylation (Target genes are not known)

HPLC-UV (high performance liquid chromatographyultraviolet) [4,44]

Hydrolysis of DNA into its constituent nucleoside bases, chromatographic separation of 5 mC and dC bases, measurement of fractions and 5 mC/ dC ratio calculation Similar to HPLC but quantitation is done with spiked internal standards and assessment using MS

“Gold standard” assay for quantifying the amount of deoxycytidine (dC) and methylated cytosines (5 mC) in a hydrolyzed DNA sample

Needs specialized laboratory equipment and large quantities (3 e10 mg) of the DNA

Smaller quantities of the hydrolyzed DNA required, no effect of poor quality DNA, costeffective highthroughput methylation analysis Quick and easy, inexpensive

Set-up cost is high; the internal standard procurement is costly

LC-MS/MS (Liquid chromatography coupled with tandem mass spectrometry) [68]

ELISA-Based Methods [65]

Can detect very low DNA methylation levels

Prone to high variability, suitable for the rough estimation of DNA methylation

Heavy amplification of target sequences is required to be detected by pyrosequencer, PCR amplification bias may happen

Introduction to epigenetics

Global DNA Methylation Assay-LINE-1 (long interspersed nuclear elements-1) [33]

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Uses a primary antibody raised against 5 Mc; and a labeled secondary antibody followed by colorimetric/ fluorometric detection Hybridization of fragmented DNA to biotinylated LINE-1 probes and immobilization to a streptavidin-coated plate. Quantification of methylated cytosines using an anti-5 mC antibody, HRPconjugated secondary antibody and chemiluminescent detection reagents

Continued

Table 1 Different techniques to analyze DNA methylationdcont’d Class

Principle

Features

Limitations

AFLP (Amplification fragment length polymorphism) and RFLP (Restriction fragment length polymorphism) [4] LUMA (luminometric methylation assay) [42]

PCR-based detection of differentially methylated fragments

Inexpensive, can be used for any species

Assesses only small percentage of global DNA methylation

Combined DNA cleavage by methylation-sensitive restriction enzymes, polymerase extension assay by Pyrosequencing Theoretically all C information is covered. Purified genomic DNA is sheared into fragments, which are then end-repaired; Atailing (adenine base addition at 30 end) and ligation of methylated adapters to the DNA fragments is done followed by sodium bisulfite treatment, PCR amplification, and sequencing. Based on enrichment of CpG-rich regions in near vicinity to the restriction enzyme’s recognition sequence Uses an antibody recognizing 5mC, followed by analysis of immunoprecipitated DNA by PCR Microarray on genome wide scale

Quantitative method, highly reproducible, easy to scale up

Requires high quality DNA

Gives genome-wide information and resolution at single nucleotide

Costly and requires intense calculation

Cost-effective

Sometimes there is a lack of coverage at intergenic and distal regulatory elements

Can be used for genome wide methylation pattern analysis during development time course and highly reproducible

Requires high quality and cross-reactive 5mC antibodies

Whole-genome bisulfite sequencing (WGBS) (BS-seq; MethylC-seq) [55]

Reduced-representation bisulfite sequencing (RRBS) [31] Methylated DNA immunoprecipitation (MeDIP) [35]

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Method

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S.No.

2.

To study changes in Differentially-Methylated Regions (Target genes are not known)

Bisulfite Sequencing [47]

Array or Bead Hybridization [4]

Endonuclease Digestion Followed by Sequencing [4]

3.

Bead Array [4]

PCR and Sequencing [4]

Pyrosequencing [51]

Incomplete conversion of unmethylated cytosine to uracil may produce bias

Can identify methylation of single locus

Can give false positive results due to crossreactivity

Used for methylated CpG islands isolation, can be combined with NGS. Bisulfite-free

High quality of DNA is required

Customization possible to profile up to 384 individual CpG sites, can be used for nonhuman species, can be used for non-human species For large methylation differences direct sequencing of PCR product suitable for low throughput projects, good technique for heterogeneous samples, detect even small differences in

Costly

Requires nested PCR to minimize unspecific amplification, Difficult genome assembly in presence of repetitive DNA; lack of proof reading

Continued

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Bisulfite-converted DNA amplified for the region of interest followed by sequencing Primer are designed, PCR products are obtained, and short-read pyrosequencing reaction (w100 bp) is performed. The level of

Single-nucleotide resolution

Introduction to epigenetics

DNA methylation analysis of Genes of Interest

“Gold standard” method in DNA methylation studies Bisulfite treatment of mediates deamination of cytosine into uracil, which are read as thymine, as determined by PCR-amplification and followed by Sanger sequencing No conversion of 5 mC residues to uracil which are read as cytosine. Methylated DNA fractions are used for hybridization with microarray Genomic DNA digestion by methylation-sensitive endonucleases at unmethylated sites followed by adapter ligation Extension-based assay

Table 1 Different techniques to analyze DNA methylationdcont’d Class

Method

PCR with High Resolution Melting [69]

COLD-PCR [30]

Features

methylation for each CpG site within the sequenced region is estimated based on the signal intensities for incorporated dGTP and dATP. Requires two pairs of primers, one pair for methylated and another for unmethylated DNA amplification. For each sample two qPCR reactions are performed followed by relative methylation calculations on the basis of the difference in Ct values Based on the differences in melting temperatures of methylated (high Tm) and unmethylated DNA (low Tm). Intercalating dye, e.g., SYBR or Eva green, fluoresces on binding to doublestranded DNA. With rise in temperature and dissociation of DNA strands there is loss of fluorescence. Lowering of denaturation temperature of the PCR so that only unmethylated fragments (less GC rich) are amplified efficiently and detected.

methylation (down to 5%)

Limitations

Quick method, multiple samples can be profiled

Assessment of methylation status of only one or two CpG sites at one point

Small methylation differences (5%e10%) are detected depending on the purity of PCR product.

Pure PCR product is necessary

Very low (0.1%) levels of methylated DNA can be detected

Denaturing gradient e.g. polyacrylamide is required

Introduction to epigenetics

Methylation-Specific PCR [39]

Principle

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S.No.

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Histone modifications and chromatin remodelling In past three decades a significant progress has been made in genome wide characterization of histone modifications and chromatin remodelling [43,50]. The major developments made in histone modification chromatin analysis such as improved high-throughput sequencing in combination with chromatin immunoprecipitation assay (ChIP) and DNA microarray (DNA chip) i.e. ChIP-on-chip, Chromatin Interaction Analysis by Paired-End Tag Sequencing (ChIA-PET), ChIP-Sequencing (ChIP-Seq) has helped in unraveling the human epigenome [47]. (1) ChIP: Chromatin immunoprecipitation, a powerful tool to analyze protein-DNA interactions allows to study the dynamics of histone methylation. The principle behind ChIP is the enrichment of the specific portion of DNA of interest (antigen) by immunoprecipitation followed by amplification of the enriched region by PCR so as to obtain sufficient quantity of the enriched fraction. Further, analysis is done by southern blotting, PCR or genome wide methods [53,54]. (2) ChIP-on-chip: There are two types of ChIP-on-chip method based on microarray contents i.e. (1) Promoter tiling arrays and (2) Genome tiling arrays. In promoter tiling arrays the probes are designed on specific genomic elements or promoters which may lead to loss of some relevant regions. The genome tiling array employs probes that cover entire genome thus, allowing global genome-wide analysis. Earlier, the ChIP-on-chip method was used to analyze histone modifications in yeast and Drosophila melanogaster. Recently, ChIP-on-chip was successful inanalysing histone modifications in human genome [29,64]. (3) ChIP-seq: Another method of analyzing histone methylation as well as chromatin remodelling is ChIP-seq which combines ChIP with next generation sequencing procedures. The ChIP-seq involves repair of DNA ends and ligation to a pair of adapters. The DNA is amplified and bound to the flow cell surface containing oligonucleotides. The adapter sequences ligated to DNA are recognized by these oligonucleotides. The genome analyzer reads each DNA sequence during solid-phase PCR processing and the resulting reads are mapped to a reference genome to find coordinates [26,63]. This technique enabled researchers to overcome low resolution and high noise problem arising from ChIP-chip technique [25]. (4) Mass Spectrometry: The mass spectrometry (MS) can be used for quantitative analysis of protein expression and differential expression of protein modifications [28,66]. Recently, to address the issue of inability of MS to map the modification patterns to specific promoter regions a combinatorial technique namely chromatin affinity purification along with MS (ChAP-MS) and Chromatin Proteomics (ChroP)/ChIP-MS) was developed. Therefore, now functionally distinct chromatin regions can be analyzed for the histone marks and binding proteins simultaneously [28,66]. Mass spectrometry strategies have been divided in to three categories on the basis of portion of histone analyzed viz. bottom-up, middle-down and top down. In traditional ‘bottom up’ approach the protease enzymes are used to cleave the target protein into smaller peptides (5e20 aa) before MS analysis. The analysis of intact protein is done by “top down” approach, while the “middle-down” approach which is a modified version of “top-down” method is used for the characterization of large peptides containing less than 50 N-terminal amino acid residues of histone tails [57]. (5) Chromosome Conformation Capture Technologies: Different chromosome conformation capture technologies have been developed to study three-dimensional structure of whole genome. These include chromosome conformation capture (3C), chromosome conformation capture on-

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chip (4C), Chromosome Conformation Capture Carbon Copy (5C), combined 3C-ChIP-cloning (6C), Genome Conformation Capture (GCC), and (ChIA-PET [37,62]. The 3C protocol involves crosslinking of cells by formaldehyde, cell lysis by hypotonic buffer and protease inhibitors, solubilization and digestion of chromatin by sodium dodecyl sulfate (SDS) followed by chromatin ligation under dilute conditions using ligase. Further, reverse cross linking and purification is done to generate 3C libraries. The 3C libraries are used to generate 5C libraries which are used to study and quantify 3D organization of chromatin at a particular locus at higher resolution and throughput [36,37]. (6) (ChIA-PET: The drawback of ChIP-chip technique is the designing of microarrays. To overcome this issue another strategy namely Chromatin Interaction Analysis by paired end tag ((ChIA-PET) a 3C based technique was developed. This strategy incorporates immunoprecipitated DNA “tags” cloned into a plasmid library andsequenced [45,46].

Methods to analyze methylation of RNA and ncRNA-species N6-methyladenosine (m6A) and other RNA modifications like N1-Methyladenosine (m1A), 20 -OMethylation (20 OMe/Nm) and 5-Methylcytosine (m5C) can be analyzed by purification of RNA by established protocols and detection of the type of RNA methylation [59]. The analysis and characterization of RNA depends on various factors such as modification types abundance, and the RNA sequence [56,59]. Various approaches to understand the RNA methylation are listed below. (1) Radioisotope incorporation assays: The radioisotope incorporation assay strategies utilize incorporating radioactive isotopes into RNA to estimate RNA methylation. The methyl donor, sadenosyl-methionine is labeled with tritium and addition of radioactive methyl group from donor to nucleoside takes place by methyltransferase activity which is measured by scintillation [27,52,56]. (2) Thin-layer chromatography: Two dimensional thin layer chromatography is another way of identifying most of RNA modifications. Two dimensional separation of RNA disperses nucleotides across cellulose substrate as per their charge and hydrophobicity. The drawback of 2D-TLC is that it provides a general transcriptome wide methylation status.The imaging of pattern with ultraviolet light can be then done. Site-specific cleavage and radioactive labeling followed by ligation-assisted extraction and thin-layer chromatography (SCARLET) can be used to study stoichiometry when the sequence is known [49]. (3) Mass spectrometry: The nucleotides are identified on the basis of mass-to-charge ratio by a standard comparative method using MS. Though the MS approach is similar to chromatographybased protocols, it does not require radioisotope or labeling. The MS strategies involve large amount of RNA and a priori-sequence information, which is a major drawback for MS based strategies [56,60]. (4) Bisulphite sequencing: Deamination of cytosine to uracil on sodium bisulphite treatment leads to a mutation in thymine when reverse transcription takes place and depicted in the final sequencing dataset. The methylated cytosine does not undergo deamination. This is the strategy behind bisulphite sequencing and provides information about RNA methylation to single base pair resolution. The drawback in bisulphite sequencing is the requirement of large RNA quantities and resistance of neighboring modifications as well as double stranded regions to bisulphite treatment [56,67].

Introduction to epigenetics xxxvii

(5) Antibody-based methods: Commercial antibodies against methylated RNA residues e.g. m6A, m1A and m5C are available. Moreover, antibodies specific to modifications along with NGS are being used to estimate m5C, m1A and m6A. The most common and established strategy to map these modification is “methylated RNA immunoprecipitation sequencing” (“MeRIP-seq”) [23].N6-Methyladenosine using m6A Crosslinking Immunoprecipitation Sequencing (m6A-CLIP Seq/mi-CLIP) is immunoprecipitation based method for high resolution mapping of as small as 1 mg of poly(A)-selected mRNA for m6A modification [32,48].

Analysis of epigenetic modulating enzymes and their functions Most common methods to study the expression and analysis of epigenetic modulators viz. DNMTs, HDACs and MeCPs are western blotting, ELISA, immunoprecipitation (ChIP) assays or coimmunoprecipitation. The co-immunoprecipitation method is generally used to study the interactions between epigenetic modulators [34]. Further, in vivo imaging methods are being used to study HDAC inhibitor pharmacokinetics [40], HDAC direct binding [38], HDAC activity [58]; Yeh et al., 2013; [61].

Single cell epigenomics The present knowledge of epigenetics has been derived from bulk measurements in population of cells which majorly associated it with active or repressed transcriptional states [72]. Such generalizations more often lead to an ambiguous answer of many complex biological questions. Epigenetic regulations can be more precisely studied at the single-cell level, where intercellular differences can be investigated and a deeper understanding of cellular functions and dysfunctions can be achieved [71,72]. Recent advances in single cell technologies convincingly demonstrated that seemingly homogenous cell population have difference in gene expression which can possibly be due to heterogeneity at epigenetic level. The emerging technology of single cell epigenomics is a powerful method to understand the gene regulation and associated molecular pathologies. This exciting technology may refine our existing understanding of epigenetic regulations [73]. However, the real potential of singlecell epigenetic studies can be appreciated through parallel evaluation of genomic, transcriptional, and epigenetic information. Collating different components of the epigenome into multi-omics measurements can add new layers of molecular connections among the genome and its functional output.

Methodologies for single cell epigenetic analysis Prerequisite of single cell epigenetic analysis is isolation of single cells from culture or dissociated tissue followed by cell lysis. This can be performed by means of manual manipulations, droplet encapsulation or fluorescence-activated cell sorting (FACS). Recently, developed microfluidics systems can also be used in which cells are trapped in chambers where lysis and RNA-seq library preparation can be done subsequently [72].

xxxviii Introduction to epigenetics

DNA methylation and other modification Today several DNA modificationsdlike methylation (5mC), hydroxymethylation (5hmC), and formylcytosine (5fC)dcan be probed in a single-cell by sequencing at single-nucleotide resolution [71]. Initially, genome-wide 5mC measurements in a single cell were performed using reduced representation bisulfite sequencing (scRRBS) method which was based on enrichment of CpG dense regions (such as CpG islands) and restriction digestion. Though it allows the measurement of lager fraction of promotor region CpG sites, it does not cover many important regulatory regions [72]. Further, technological developments in single-cell whole-genome methylation sequencing are based on a postbisulfite adapter-tagging (PBAT) method in which bisulfite modification is performed before library preparation. Another innovative approach for generating single-cell libraries using microfluidics has recently emerged. As compared to other methods, this technology has significantly increased the library preparation throughput, where cell specific barcoding is also performed before pooling the adaptertagged fragments. This technology enables methylation measurements in w50% of the CpG sites in single cell. This has allowed the detection of high variability between individual cell in distal enhancer methylation which is not usually captured by scRRBS. For single cell analysis of hydroxymethylated cytosine (5hmC), the currently established methods such as TET-assisted bisulfite sequencing (TAB-seq) and AbaSI (restriction enzyme) coupled with sequencing (Aba-seq) could potentially be adapted [72].

Histone modifications and transcription factor binding Histone marks are mapped by chromatin immunoprecipitation followed by sequencing (ChIP-seq), however performing ChIP-seq at the single-cell level is very challenging [71]. The single cell ChIP-seq has the limitation of background noise of nonspecific antibody pull-down, which increases with decreasing levels of target antigen. Recently, this has been substituted by micrococcal nuclease (MNase) digestion and barcoding to be effectively performed on thousands of cells. This approach uses droplet-based microfluidics technology and processes large numbers of cells parallelly [72]. ProteineDNA interactions in single cells can be studied by DamID, where a cell line expresses low levels of a fusion protein of Escheriichia coli deoxyadenosine methylase (Dam) and the protein of interest [74]. This Dam based technique methylates DNA on adenine residues next to sites of protein binding which is further cut by the methylation-sensitive restriction enzyme DpnI and ligation of sequencing adapters. Presently, this technique is limited by poor resolution (100 kb), however future optimizations may enable mapping of transcription factor binding sites in single cells. Additionally, single-cell DamID could also be used in genome-wide analysis of histone modifications by using Dam fusion with specific histone readers or modifiers [72,74].

Chromatin structure and chromosome organization The single cell evaluation for chromatin structure is based on the assay for transposase-accessible chromatin (ATAC-seq). This technique uses a Tn5 transposase enzyme for tagmentation which is a process of DNA fragmentation and adapter sequences attachment simultaneously. ATAC-seq gives single-cell resolution by applying a “combinatorial indexing” strategy, where tagmentation is carried

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out on 96 pools of a few thousand nuclei, and a unique barcode to every pool is introduced [72,74]. The second method for single-cell ATAC-seq has also been described by using a commercially available microfluidics device, which carry out the transposition reaction on individual cells. The resolution has been largely increased by using this combinatorial indexing method, which maps an average of 70,000 reads per cell. Ultimately, study of open chromatin genomic regions has been done in single cells by applying a DNase-seq approach to map regions that are DNaseI hypersensitive. Single cell DNase-seq provides an improved resolution of 300,000 mapped reads per cell, although with a very low mapping proficiency (2%) and even lower throughput [72]. Technological advancement has now made possible to assess chromosome conformation at the single-cell level with a HiC-based method in addition to chromatin organization. Hi-C, is considered as an extension of chromosome conformation capture (3C), which is capable of identifying genome-wide long range interactions (Berkum et al., 2010). Though single-cell HiC is presently limited in its resolution but still allows depiction of the individual chromosome organization such as compartmentalization and interchromosomal interactions [72,74].

Epigenome manipulation and editing The epigenome of eukaryotic cells is highly complex and strongly correlated with central cellular processes. Their dysregulation manifests in aberrant gene expression and disease. The epigenome editing holds a great promise of enhancing knowledge of epigenetics in gene regulation and enabling manipulation of cell phenotype for research or therapeutic purposes [75,79]. Recent advancements in genome engineering technologies use highly specific DNA-targeting tools to precisely manipulate epigenetic changes in a locus-specific manner, generating diverse epigenome editing platforms.

Epigenetic manipulation techniques Classic myriad of genetic techniques such as gene knockouts and individual domain deletions, point mutations, inducible expression constructs, ectopic expression of vectors, targeted knockdowns of a transcript, and various screens for gain or loss of function that manipulate genome structure or gene expression facilitate mimicking of the perturbation of discrete components of the epigenome [75]. These techniques have contributed greatly for creating the foundation of our current epigenetics knowledge. However, these methodologies also lead to global effects on the epigenome that may confound the experimental results [79].

Small-molecule inhibitors A subset of small-molecule inhibitors that can manipulate epigenetic marks are being applied in research and as anticancer drugs. Hallmarks of these drugs include the irreversible DNMT1 and DNMT3 inhibitors of azacitidine (5-azacitidine) and decitabine (5-aza-2 -deoxycytidine), alongside the histone deacetylase (HDAC) inhibitors suberoylanilide hydroxamic acid (SAHA) and romidepsin (depsipeptide or FK228). Additionally, there are numerous other small-molecule inhibitors that target specific epigenetic components, including histone-modifying enzymes are known. Although these compounds lack tissue/cell specificity, they have shown a remarkable efficacy in various models within a particular dose range [75].

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Targeted epigenome manipulations The three most important molecular tools that have been developed for targeted epigenome editing are zinc finger proteins, Transcription activator-like effectors (TALEs), and CRISPReCas systems.

Zinc finger proteins Zinc finger proteins are one of the most characterized systems used for the targeted sequence-specific nucleic acids manipulations. Zinc finger-related epigenome editing techniques includes joining programmable DNA binding zinc finger proteins designed to locate diverse sequences with catalytically active or scaffolding effector domains. The chimeric proteins thus alter gene expression profiles and serve as artificial transcription factors (ATFs) [77].

TALEs TALE has a central tandem amino acid repeat domain of approximately 33e35 residues in length which enables it for DNA recognition. The DNA specificity of TALEs is flexible which is encoded in its tandem repeat sequence. In contrast to zinc fingers, which require triplet sequence recognition sites, TALEs can target a single nucleotide at a time through its repeat variable di-residues (RVDs). This characteristic makes TALEs simpler to engineer and has enabled rational designing of artificial TALEs for epigenome editing applications [78].

CRISPR/Cas9 system The emerging epigenome editing tools are now more focused on the CRISPR platform due to the flexibility and ease of the CRISPR/Cas9 system. Multiple strategies have been applied for finding optimal targeting sites with the CRISPR/Cas9 system. Certain studies have manipulated several adjacent genes in the same region of the genome together, an approach that can work well for various novel epigenome editing tools. This kind of approach sometimes results in synergistic effects and intensifies the possibility of increased number of off-target sites, as well as convoluting steric effects, such as repression by catalytically inactive variants. Alternatively, CRISPR-based technique which screens with libraries of thousands of different gRNAs can be utilized to recognize the most potent gRNAs that alter transcription or protein expression. There are few drawbacks of this technique aside from cost and complexity. As it measures RNA or protein expression levels that may not correlate with changes to the epigenome. Overall, targeted epigenome editing techniques such as CRISPR/Cas9 system, offers unique platforms that allow easy and flexible targeting of many different domains possessing epigenetic activity [75,76]. The area of epigenome editing is still in infancy and only a small fraction of possible epigenetic targets has been investigated so far. With the advent of newer technology, understanding of epigenetic mechanisms responsible for modulating epigenetic marks will increase, and the power of epigenome editing to create a diversity of epigenetic states will grow exponentially.

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[48] Linder B, Grozhik AV, Olarerin-George AO, Meydan C, Mason CE, Jaffrey SR. Single-nucleotide-resolutionmapping of m6A and m6Am throughout the transcriptome. Nat Methods 2015;12:767e72. [49] Liu N, Parisien M, Dai Q, Zheng G, He C, Pan T. Probing N6-methyladenosine RNA modificationstatus at single nucleotide resolution in mRNA and long noncoding RNA. RNA 2013;19:1848e56. [50] Liu Y, Liao J, Lu Q. Laboratory methods in epigenetics. In: Epigenetics and Dermatology. Academic Press; 2015. p. 7e35. [51] Mahapatra S, Klee EW, Young CY, Sun Z, Jimenez RE, Klee GG, Tindall DJ, Donkena KV. Global methylation profiling for risk prediction of prostate cancer. Clin Cancer Res 2012;18:2882e95. [52] Martin SA, Moss B. Modification of RNA by mRNA guanylyltransferase and mRNA (guanine-7-) methyltransferase from vaccinia virions. J Biol Chem 1975;250:9330e5. [53] Massie CE, Mills IG. ChIPping away at gene regulation. EMBO Rep 2008;9:337e43. [54] Minard ME, Jain AK, Barton MC. Analysis of epigenetic alterations to chromatin during development. Genesis 2009;47:559e72. [55] Miura F, Ito T. Highly sensitive targeted methylome sequencing by post-bisulfite adaptor tagging. DNA Res 2014;22:13e8. [56] Mongan NP, Emes RD, Archer N. Detection and analysis of RNA methylation. F1000Research 2019:8. ¨ nder O ¨ , Sidoli S, Carroll M, Garcia BA. Progress in epigenetic histone modification analysis by mass [57] O spectrometry for clinical investigations. Expert Rev Proteomic 2015;12:499e517. [58] Reid AE, Hooker J, Shumay E, Logan J, Shea C, Kim SW, Collins S, Xu Y, Volkow N, Fowler JS. Evaluation of 6-([(18)F]fluoroacetamido)-1-hexanoicanilide for PET imaging of histone deacetylase in the baboon brain. Nucl Med Biol 2009;36:247e58. [59] Romano G, Veneziano D, Nigita G, Nana-Sinkam SP. RNA Methylation in ncRNA: classes, detection and molecular associations. Front Genet 2018;9:243. [60] Ross R, Cao X, Yu N, Limbach PA. Sequence mapping of transfer RNA chemical modifications by liquid chromatography tandem mass spectrometry. Methods 2016;107:73e8. [61] Sankaranarayanapillai M, Tong WP, Yuan Q, Bankson JA, Dafni H, Bornmann WG, Soghomonyan S, Pal A, Ramirez MS, Webb D, Kaluarachchi K, Gelovani JG, Ronen SM. Monitoring histone deacetylase inhibition in vivo: noninvasive magnetic resonance spectroscopy method. Mol Imaging 2008;7:92e100. [62] Sati S, Cavalli G. Chromosome conformation capture technologies and their impact in understanding genome function. Chromosoma 2017;126:33e44. [63] Schones DE, Zhao K. Genome-wide approaches to studying chromatin modifications. Nat Rev Genet 2008; 9:179e91. [64] Seifert M, Schneider R. Chromatin immunoprecipitation ChIP: wet lab meets InSilico. In: Mannhold R, Kubinyi H, Folkers G, editors. Epigenetic targets in drug discovery. Weinheim: Wiley-VCH; 2009. p. 139e60. [65] So MY, Tian Z, Phoon YS, Sha S, Antoniou MN, Zhang J, Wu RS, Tan-Un KC. Gene expression profile and toxic effects in human bronchialepithelial cells exposed to zearalenone. PLoS One 2014;9:3. [66] Soldi M, Bonaldi T. The ChroP approach combines ChIP and mass spectrometry to dissect locus-specific proteomic landscapes of chromatin. J Vis Exp 2014;86. [67] Squires JE, Patel HR, Nousch M, Sibbritt T, Humphreys DT, Parker BJ, Suter CM, Preiss T. Widespread occurrence of5-methylcytosine in human coding and non-coding RNA. Nucleic Acids Res 2012;40: 5023e33. [68] Thuc L, Kim KP, Fan G, Faull KF. A sensitive mass spectrometry method for simultaneous quantification of DNA methylation and hydroxymethylation levels in biological samples. Anal Biochem 2011;412:203e9. [69] Wojdacz TK, Dobrovic A, Hansen LL. Methylation-sensitive high-resolution melting. Nat Protoc 2008;3: 1903e8.

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[70] Yong WS, Hsu FM, Chen PY. Profiling genome-wide DNA methylation. Epigenetics Chromatin 2016;9(1): 26. [71] Kelsey G, Stegle O, Reik W. Single-cell epigenomics: Recording the past and predicting the future. Science 2017;358:69e75. [72] Clark SJ, Lee HJ, Smallwood SA, Kelsey G, Reik W. Single- cell epigenomics: powerful new methods for understanding generegulation and cell identity. Genome Biol 2016;17:72. https://doi.org/10.1186/s13059016-0944-x. [73] Verma M, Kumar V. Single-cell epigenomics: technology and applications single-cell omics, vol. 1; 2019. p. 215e29. [74] Lo PK, Zhou Q. Emerging techniques in single-cell epigenomics and their applications to cancer research. J Clin Genom 2018. https://doi.org/10.4172/JCG.1000103. [75] Holtzman L, Gersbach CA. Editing the Epigenome: reshaping the genomic landscape. Annu Rev Genom Human Genet 2018;19:43e71. [76] Enrı´quez P. CRISPR-mediated epigenome editing. Yale J Biol Med 2016;89:471e86. [77] Choo Y, Isalan M. Advances in zinc finger engineering. Curr Opin Struc Biol 2000;10:411e6. [78] Christian M, Cermak T, Doyle EL, Schmidt C, Zhang F, Hummel A, Bogdanove AJ, Voytas DF. TAL effector nucleases create targeted DNA double-strand breaks. Genetics 2010;186:757e61. [79] Klann TS, Black JB, Gersbach CA. CRISPR-based methods for high-throughput annotation of regulatory DNA. Curr Opin Struc Biol 2018;52:32e41.

CHAPTER

Epigenome reprogramming in the male and female germ line

1

Kazuki Kurimoto, Hiroki Ikeda, Hisato Kobayashi Nara Medical University, Department of Embryology, Kashihara, Nara, Japan

Introduction Germ cells form individuals through fertilization and play critical roles for heredity and generation of genetic diversity. Understanding of the mechanisms of germ cell formation is thus valuable for reproductive health. In some model organisms, including Drosophila melanogaster (fruit fly) and Danio rerio (zebrafish), germ cells are set aside at the onset of embryonic development (“preformation”), while in mammals the germ cells arise through differentiation during embryogenesis (“epigenesis”). Epigenesis is the more prevailing mechanism of germ cell formation among animals, and thus may be more ancient than preformation [1]. Because the developmental origin of the germ cell lineage produced by epigenesis segregates from the same progenitors as somatic cells, germ cells re-acquire the ability to form the next generation. Primordial germ cells (PGCs) play important roles in this process by resetting their epigenome (germline epigenome reprogramming). Key events of epigenome reprograming include genome-wide reduction of DNA methylation level, remodeling of histone modification, female X chromosome reactivation, and erasure of the genomic imprinting from parental origins followed by its re-establishment during the male and female gametogenesis. In the last decade, physiologically functional PGCs have been efficiently reconstituted from mouse pluripotent stem cells (mouse PGC-like cells: mPGCLCs) [2,3] (Box 1.1). The mPGCLCs are able to reconstitute the process of gametogenesis [4e7], and meiotic entry in an adhesive culture without gonadal somatic cells [8,9]. These in vitro germ cell reconstitution systems have provided a foundation for detailed experimental investigations into the mechanisms of mouse germ cell development [10e16]. Based on the success of mouse germ cell reconstitution, human pluripotent stem cells have been also induced into PGC-like cells (human PGCLCs: hPGCLCs) [17e19] (Box 1.2), and used for signaling and genetic studies of human PGC development [20e22]. hPGCLCs have been further differentiated into oogonia-like cells, through aggregation with ovarian somatic cells from mouse embryos [23]. These studies suggest that, in future, it might become possible to reconstitute human gametes in vitro, and their findings will make a contribution to reproductive biology and medicine. Studies of epigenome reprogramming of human and mouse germ lines have also relied on highthroughput sequencing methods for transcriptome and epigenome in small numbers of cells or

Epigenetics and Reproductive Health. https://doi.org/10.1016/B978-0-12-819753-0.00001-5 Copyright © 2021 Elsevier Inc. All rights reserved.

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Chapter 1 Epigenome reprogramming in the male and female

Box 1.1 Reconstitution of mouse germ cells in vitro In the last decade, a method reconstitute mouse germ cells in vitro has been established [2,3]. In this system, mouse pluripotent stem cells (mPSCs) cultured under a feeder-free, “2iþLIF” condition [96] are used as starting materials. These cells are induced into epiblast-like cells (EpiLCs) by activin and b-FGF. PGC-like cells (mPGCLCs) are induced from EpiLCs by a high concentration of BMP4. The physiological functions of mPGCLCs were demonstrated by transplantation assays resulting in fertile offspring [2,3]. Oocytes are also reconstituted in vitro [4] by combining mPGCLCs with a culture method for the entire folliculogenesis [97]. The dormant state of primordial follicles, which is observed in ovaries in vivo, has been also reproduced in the in vitro reconstituted ovaries [7]. Germline stem cells (GSCs) [60], containing spermatogonial stem cells, are derived from reconstituted testes in a gas-liquid interphase culture [98] of mPGCLCs and embryonic testicular somatic cells [5]. The mPGCLCs are also able to efficiently proliferate in an adhesive culture in the presence of forskolin and rolipram on feeder cells expressing stem cell factor (SCF) [8,9,99], and enter meiotic prophase by retinoic acid (RA) and BMP signaling [9,89,90]. Thus, germ cell reconstitution in vitro provided a useful experimental tool for the detailed investigation of the mechanism of germ cell development, in addition to opening the possibility of in vitro gametogenesis.

Box 1.2 Reconstitution of human germ cells in vitro Human PGCLCs (hPGCLCs) are induced directly from human PSCs [18,19] or through induction of incipient mesoderm/ primitive streak-like cells (iMELCs) [82]. The biological relevance of iMELCs is supported by an observation that, during the development of cynomolgus monkeys, genes for mesoderm (T and MSX2) are expressed albeit at low levels in PGCs and some other cells in the nascent amnion [82]. Xenogeneic aggregation of female hPGCLCs and murine ovarian somatic cells at E12.5 forms reconstituted ovaries [23]. During a long term-culture over 70 days, these hPGCLC-derived cells downregulate markers for early hPGCs and pluripotency and become positive for genes for the later PGC development (DAZL, DDX4) as well as some meiosis genes (SYCP3, REC8). At day 120, these cells are positive for STRA8, and show a gene expression profile similar to the RA-responsive cells observed in the human embryonic ovaries [95]. Thus, these cells are human oogonia-like cells that progress along the developmental pathway to meiotic oocytes.

single cells (Boxes 1.3 and 1.4) [24]. In this section, recent findings on epigenome reprograming during germ cell development in vivo and in vitro will be discussed.

Reprogramming of mouse germ cells The mouse has been most frequently used as a model organism for studies on mammalian germ cell development. The in vitro reconstitution systems have been also extensively used for experimental investigation of the mechanisms of germ cell development, particularly PGC development during the early developmental stages. The epigenome reprogramming of mouse germ cells will be discussed in this subsection.

Primordial germ cell development in mice In mice, around embryonic day (E) 6.0, at the time when gastrulation begins, mouse PGCs (mPGCs) are specified as about 10 cells in the most posterior part of epiblast, the cup-shaped simple pluripotent epithelium, from which all of the embryonic portion are derived (Fig. 1.1). This process occurs in

Reprogramming of mouse germ cells

5

Box 1.3 Methods for genome-wide DNA methylation profiling DNA methylation (i.e., methylation at the C-5 position of cytosine) is detectable at single-base resolution using in vitro sodium bisulfite treatment, which converts unmethylated cytosine to uracil (sequenced as thymine), but does not affect methylated cytosine. The combination of DNA sequencing and bisulfite reaction, called “bisulfite sequencing”, allows direct measurement of DNA methylation levels by calculating the ratio of methylated and unmethylated cytosine at each site [100]. The whole-genome bisulfite sequencing (WGBS) using high-throughput sequencing technology is currently considered the gold standard for a comprehensive and quantitative analysis of DNA methylation across the genome. Several library preparation strategies for single-cell DNA methylome profiling have been developed based on the post-bisulfite adapter tagging (PBAT) approach [101] and have been applied to study epigenetic heterogeneity in developing mammalian tissues and cells.

Box 1.4 Methods for genome-wide chromatin profiling and three-dimensional genomic structure Chromatin immunoprecipitation sequencing (ChIP-seq) has been used to survey genome-wide landscapes of protein-DNA binding profiles by combining immunoprecipitation of protein-bound DNA and high-throughput sequencing technologies [102]. Functions of genomic elements are inferred by chromatin states, such as histone modification and/or transcription factor-binding patterns, through integrated analyses of ChIP-seq data. HiC-seq is a comprehensive chromosome conformation capture (3C)-based technique [103], and measures the frequency of physical interactions between arbitrary genomic regions. To run a HiC-seq protocol, genomic DNA is crosslinked and digested by restriction enzymes, followed by formation of chimeric DNA fragments through the ligation of DNA fragments that are located in proximity in the three-dimensional genomic organization. The chimeric fragments are counted using the high-throughput sequencing. Together, ChIP-seq and HiC-seq are important techniques to elucidate the chromatin organization regulating gene expression.

response to bone morphogenetic protein 4 (BMP4) [25] produced by the extraembryonic ectoderm (ExE), an extraembryonic part of conceptus in intimate contact with the edge of the epiblast (Fig. 1.1). While essentially all epiblast cells are competent for the mPGC fate, BMP4 antagonists secreted from the anterior part of the visceral endoderm (anterior visceral endoderm: AVE) prevent the mPGC fate, thereby confining it to the posterior end of the epiblast [26]. BMP4 signaling also stimulates epiblast cells to secret WNT3, which acts as an autocrine factor essential for the mPGC fate [26]. Downstream of the BMP4 signaling, three transcription factors (TFs), BLIMP1 (also known as PRDM1), PRDM14 and TFAP2C, play critical roles in the mPGC specification [27e29]. At around E6.0, the precursors of mPGCs show a gene expression profile very similar to that of the epiblast, except for the start of BLIMP1 expression, as revealed by pioneering, quantitative single-cell transcriptome analyses [30,31]. Along with the progression of early gastrulation by E6.5, mPGC precursors show a strong activation of mesoderm markers, such as anterior Hox genes. Except for the expressions of BLIMP1 and a few other PGC markers, mPGC precursors at this stage is very similar to the surrounding early mesoderm cells. However, the mesoderm-like profile is only transiently activated, and such transient activation is immediately followed by a drastic downregulation by BLIMP1 by E7.5, accompanied by a global change from the mesoderm to the mPGC transcriptome [31,32].

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Chapter 1 Epigenome reprogramming in the male and female

Mouse

♂Birth extraembryonic ectoderm

mesoderm

PGC

ProSpermato- Spermatospermatogonia cytes gonia

inner cell mass

♀

epiblast visceral endoderm ectoderm

E3.5

E5.5 E5.5 5

Sperm

Birth

gonad

gut endoderm

Oogonia

E7.5 E7.5 5

Spermatids

E9.5 .5

Primordial follicle

E13.5

Antral follicle

P3

P10

MII Oocyte

P20

mPGC sspecifica specification ation (T, BLIMP1, BLIMP P1, PRDM14, TFAP2C) T TF Remodeling Remo odeling g off H3K27me3 H3K27m me3

DNA methylation in transcribed regions

♀

Enrichment Enric hment of promoter p promote bivalencyy Monotonous reduction of H3K9me2

Maternal imprint formation Broad domain of H3K4me3 Weak 3D genomic archtecture

methylat t tion Replication-coupled dilution of global CpG methylation (H3K27me3 and H3K9me3 can compensate DNA DN demethylation) d

♀

X chromosome reactivation

sivvely erased Imprint is comprehensively Transposons are retained d methylated Paternal imprint formation Massive de novo CpG methylation Transposons resists de novo methylation

♂

Sperm-specific genome archtecture

In vitro reconstituted systems ActA FGF2

BMP4, SCF

Plane culture

BMP4, RA

Meiocytes mPSCs

EpiLCs

mPGCLCs

mPGCLCs (proliferated) GSC-LCs

Reconstituted testes with embryonic testicular somatic cells GSC derivation culture Reconstituted ovaries with embryonic ovarian somatic cells In vitro growth & maturation

MII Oocyte

FIG. 1.1 Schematic representation for the epigenome reprogramming of mouse germ cells and their reconstitution in vitro. Currently known, major events of epigenome reprogramming are listed below the developmental stages in vivo. The corresponding in vitro germ cell reconstitution systems are also represented with colored circles; mPSCs (pale blue), EpiLCs (dirk blue), mPGCLCs (orange), mPGCLCs proliferated in the adhesive culture (pale orange), meiotic germ cells (meiocytes) (pink), GSC-LCs (cyan), and MII oocytes (pale pink).

Reprogramming of mouse germ cells

7

This transcriptome change involves downregulation of DNA methylation factors; DNMT3A, DNMT3B, and DNMT3L, which catalyze de novo methylation, and UHRF1 (also known as NP95), a factor required for the methylation maintenance. Interestingly, the expression level of DNMT1, which is recruited to the replication foci by UHRF1 and catalyzes DNA methylation of the new DNA strand, is not changed. Moreover, the histone methyltransferase GLP, which catalyzes methylation of histone H3 lysine 9 (K9) in complex with G9A, is also downregulated in mPGCs. Following these expression changes, mPGCs undergo genome-wide reprogramming of epigenetic marks; this includes drastic reductions of DNA methylation and di-methylation of the lysine 9 residue of histone H3 (H3K9me2). On the other hand, H3K27me3, which is catalyzed by Polycomb repressive complex 2 (PRC2), is upregulated in mPGCs. These changes occur in a genome-wide manner and are detectable with immunofluorescence [32], while changes in the genomic distributions of these modifications have been revealed by taking advantage of the in vitro reconstitution system of mPGCs and sequencing-based analytical methods for epigenome (Chromatin immunoprecipitation sequencing [ChIP-seq] and whole-genome bisulfite sequencing [WGBS]) as discussed below. The established mPGCs form a cluster of about 40 cells at the base of the allantois at around E7.5. Then, mPGCs migrate toward future gonads through the hindgut and dorsal mesentery. At around E10.5eE11.5, the mPGCs enter the genital ridges, (i.e., developing gonads) and start to proliferate there. At around E12.5, sex differentiation of the genital ridges becomes morphologically evident, and germ cells also acquire unambiguous sexual dimorphism in their gene expression profiles. The genome-wide erasure of DNA methylation continues until E13.5. In female embryos, X chromosomes, which are both activated in the inner cell mass (ICM) of blastocysts and randomly inactivated upon implantation, are again both reactivated during the mPGC development (reviewed in Ref. [33]). From E13.5 onward, the female germ cells cease mitosis and directly enter the prophase of the first meiosis (meiosis I), differentiating into primary oocytes. Then primordial follicles are formed through interaction with ovarian somatic cells, and are arrested at the meiotic prophase until they resume oogenesis upon puberty. Male germ cells cease mitosis at around E13.5, differentiating into prospermatogonia, and migrate to the basement membrane of the seminiferous tubules. After birth, they resume mitotic proliferation and differentiate into the type A spermatogonia, which provide abundant germ cells for spermatogenesis.

Induction of PGCs from mouse pluripotent stem cells Based on the findings of the signaling factors for the specification of mPGCs, PGC-like cells (mouse PGCLCs: mPGCLCs) have been induced from pluripotent stem cells (mouse PSCs [mPSCs]; i.e., mouse embryonic stem cells [mESCs] and/or mouse induced PSCs [miPSCs]) [2,3] (see also Box 1.1). In this induction procedure, the mPSCs are first induced into epiblast-like cells (EpiLCs). Then, in a floating aggregate of EpiLCs, mPGCLCs are induced by a high concentration of BMP4. The induction process of mPGCLCs recapitulates the specification process of mPGCs, including transient up-regulation of the gene expression profile for mesoderm. In fact, mPGCLCs show transcriptional and epigenetic features very similar to those of the migratory mPGCs at E9.5. The mPGCLCs are physiologically functional germ cells [2,3], and more recently, functional mouse oocytes have also been reconstituted from mPGCLCs [4,7]. These findings provide useful experimental tools for the studies of mammalian germ cell development, especially mPGCs at early developmental stages.

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Chapter 1 Epigenome reprogramming in the male and female

Epigenome reprogramming of mouse PGCs Epigenome reprogramming of germ cells begins as early as the specification stage, and its onset is coupled with the transcriptional program of the specification process. Reprogramming of histone modification and DNA methylation during this process have been analyzed using ChIP-seq and WGBS [10,12e15,34] (see also Boxes 1.3 and 1.4). The roles of key TFs has been also examined using their induced overexpression [11,35].

Transcriptional program for the mPGC fate Differentiation of the blastocyst ICM into the epiblast upon implantation undergoes a transition of pluripotency states from the naı¨ve to the primed state, and it is mimicked by differentiation of mPSCs into EpiLCs in vitro. During this transition, enhancer usage changes drastically, while the change of gene expression profile is relatively moderate [12,34]. This enhancer relocation also accompanies global change of the OCT4-binding sites mediated by OTX2, which is expressed in EpiLCs but not in mPSCs [34]. Downregulation of OTX2 is required for the mPGC specification [35], and this might relocate the binding sites of OCT4 in mPGCs into a state similar to that of mPSCs. The key TFs for the specification of mPGCs, BLIMP1, PRDM14, and TFAP2C, can directly induce mPGCLCs when they are inducibly expressed in the floating aggregates of EpiLCs [11]. This method bypasses the cytokine signaling, and the resulting TF-induced mPGCLCs contribute to fertile offspring. Interestingly, the mesoderm program, which is transiently up-regulated in mPGC specification, is not at all detected during the formation of the TF-induced mPGCLCs, and thus, is not essential for the mPGC formation itself. On the other hand, the enhancers of Blimp1 and Prdm14 as well as those of many mesoderm genes are directly activated by a key TF for the development of mesoderm, T (also known as BRACHYURY), through the recruitment of acetylation of H3K27 (H3K27ac) [10,12]. Thus, T drives the core transcriptional network for the specification of mPGCs in the physiological context [10]. This explains why mPGC specification proceeds under the mesoderm formation signals.

Remodeling of histone modification during mPGC development The N-terminal tails of histones are extensively modified by post-translational mechanisms, including methylation and acetylation of lysine residues of histone H3. These modifications mark the transcriptional states: e.g., H3K4me3 (active promoters), H3K9me2/me3 (heterochromatin), H3K27me3 (Polycombdependent repression), H3K27ac (active enhancer), and H3K36me3 (transcribed gene bodies). The genome-wide distribution of histone modifications during the mPGC specification has been analyzed by applying ChIP-seq (see also Box 1.4) to the in vitro reconstitution system (i.e., mPGCLC induction). In mESCs and EpiLCs, H3K27me3 is broadly distributed. When the mesoderm program is transiently activated during the mPGCLC induction from EpiLCs, the overall H3K27me3 level becomes down-regulated, but is locally retained or even up-regulated around genes associated with embryonic development [12]. In the established mPGCLCs, the global level of H3K27me3 is re-acquired, retaining the strong enrichment around the developmental genes. Bivalent promoters with H3K4me3 and H3K27me3 (i.e., transcriptionally poised promoters) are abundant in EpiLCs, decrease in number when the mesoderm program is transiently activated, and re-increase in the established mPGCLCs. BLIMP1 spreads H3K27me3 around a broad range of developmental genes, thereby regulating the drastic change in the gene expression pattern during mPGC specification [12]. On the other hand, another repressive histone mark, H3K9me2, is monotonously reduced without any marked change of its genomic distribution pattern.

Reprogramming of mouse germ cells

9

After the specification in vivo, mPGCs migrate into hindgut and then colonize genital ridges. The distribution of histone marks of the in vivo mPGCs at E11.5 onwards has been analyzed with ChIP-seq methods for small number of cells [36e39]. As in-vitro specification, abundant H3K4me3-H3K27me3 bivalent promoters are detected in germ cells at E11.5 and E13.5 [37], and thus, bivalent features seem to be maintained throughout the mPGC development. During this developmental process, Polycomb repressive complex 1 (PRC1) is essential for the maintenance of the pluripotency state of mPGCs and prevents premature induction of meiosis [40]. Hence, mPGCs acquire a strong repressive state with Polycomb: they show high-level H3K27me3 (PRC2) enriched with poised promoters, and require PRC1 to avoid premature differentiation.

Outline of DNA methylation reprogramming during mPGC development During the mouse life cycle, there are at least two developmental stages that undergo erasure of the genome-wide DNA methylation; preimplantation and germline development. After fertilization, DNA methylation is extensively erased from the zygote to blastocyst stages, and is re-acquired in the epiblast upon implantation. In the germ line, DNA methylation is erased during the developmental stages from the mPGC specification until around E13.5; at this stage, germ cells are comprehensively demethylated (w5% methylation). One of the most marked differences between these two demethylation waves is that genomic imprinting is erased in the germ line, while it is retained in the preimplantation embryos [41].

Details of DNA methylation reprogramming The DNA methylation erasure in the early mPGC development has been extensively investigated with WGBS (see also Box 1.3) using the in vitro reconstitution system [13e15]. During the differentiation of mESCs to EpiLCs, which recapitulates the epiblast formation upon implantation, the overall DNA methylation level becomes drastically upregulated and the methylation pattern shows a large-scale change. In contrast, mPGCLCs generally retain the DNA methylation pattern as it is established in EpiLCs, but progressively decrease the DNA methylation level during the induction process [13]. A similar trend is observed in the germ cell development until E13.5 in vivo; the methylation patterns are similar between the germ cells and the epiblast, but the methylation level of germ cells is progressively decreased, to a level much lower than that of mPGCLCs [13,42,43]. This demethylation kinetics supports the idea that the germ cells are subject to a passive demethylation mechanism that functions in a replication-coupled manner [44,45]. Imprinting regions and transposons are much more slowly demethylated than the other genomic regions during the early mPGC development, and subsequently, by E13.5, the imprinting regions are erased comprehensively while the transposons remain substantially methylated. The replication-coupled, passive DNA demethylation in mPGCs is partly due to the repression of UHRF1 at the mRNA level [31,45]. Regulation of UHRF1 is important because it binds to hemimethylated DNA and recruits DNMT1, the maintenance DNA methyltransferase, to the replication foci. In addition, cytoplasmic localization of UHRF1 is also observed in mPGCs [42]. STELLA/ DPPA3, which marks mPGCs and is essential for female fertility, recruits UHRF1 to the cytoplasm of oocytes, thereby maintaining the hypomethylated state of oocytes [46,47]. Proteolysis of UHRF1 is also involved in the DNA demethylation in the early embryos and mESCs [48]. Thus, the passive DNA demethylation in germ cells may also be explained by the cytoplasmic localization and/or proteolysis of UHRF1.

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Chapter 1 Epigenome reprogramming in the male and female

DNA hydroxymethylation 5-hydroxymethylcytosine (5hmC) is an oxidative derivative of 5-methylcytosine (5mC) catalyzed by the ten eleven translocation (TET) enzymes, and may be involved in DNA demethylation through enzyme-mediated active mechanisms. Because DNMT1 catalyzes DNA methylation of hemi-5hmC sites with much lower efficiency than that of hemi-5mC sites, 5hmC may also play a role in DNA demethylation in a replication-coupled manner [49]. During the in vitro mPGC specification, 5hmC exists in the genome at a level as low as 1%e3% of 5mC as shown by mass spectrometry and thus has only a little contribution, if any, to the genome-wide erasure of DNA methylation [13]. Interestingly, on the other hand, TET1 safeguards the lowly methylated state of the germ cells in genital ridges [50]. In line with this, TET1 deficient oocytes and sperm show stochastic and variable DNA hypermethylation in imprinting regions [51]. In addition, TET1 is involved in DNA demethylation of mESCs, and in the oscillation of the DNA methylation level when mESCs exit from the naı¨ve into the primed pluripotent state [52]. Thus, TET1 most likely regulates fine tuning of the DNA methylation reprogramming in the germ line.

Interplay between UHRF1 and H3K9me2 reduction UHRF1 interacts with H3K9me2/me3 through the tandem tudor domain (TTD) [53]. The G9A/GLP complex, an enzymatic complex for the H3K9 methylation, also plays a critical role for the DNA methylation in mESCs [54]. This enzyme catalyzes methylation of a lysine residue in a motif of DNA ligase 1; this motif mimics the N-terminal part of histone H3, and the methylated lysine residue in this motif binds to the TTD of UHRF1, thereby recruiting UHRF1 to the replicating DNA in mESCs [55]. These mechanisms may also explain the association between the monotonous down-regulation of H3K9me2 and DNA demethylation during the mPGC development.

Interplay between DNA demethylation and H3K9me3 During the period of genome-wide DNA reprogramming in mPGCs, retrotransposons are refractory to the demethylation wave, and retain partially methylated even in E13.5 germ cells. In germ cells, these transposons are enriched with H3K9me3 and H3K27me3 in an overlapped manner, while the other genomic regions generally bear these histone modifications in a mutually exclusive manner [39]. SETDB1, an enzyme catalyzing histone H3K9 methylation, is required for the silencing of many (but not all) transposons during germ cell development, and is essential for the DNA methylation in the H3K9me3-enriched regions, including the transposons silenced by SETDB1 [39]. An inducible knockout study of DNMT1, UHRF1, and SETDB1 in mESCs has demonstrated that the binding of UHRF1 to the newly replicated DNA blocks the SETDB1-mediated deposition of H3K9me3 [56]. If DNMT1 alone is inducibly knocked out, H3K9me3 is transiently depleted due to the prolonged binding of UHRF1 to the hemimethylated DNA in replication foci in the absence of DNMT1, leading to de-repression of retrotransposons [56]. The repressive state is recovered over time because UHRF1 no longer binds to un-methylated DNA after further DNA replication. Thus, it seems that DNA methylation and H3K9me3 serve as backups for each other. This mechanism may also exist in mPGCs, and may explain the down-regulation of UHRF1 for the replication-coupled DNA demethylation, and the reason why DNMT1 retains the expression level during this process. Conditional knockout of DNMT1 in early mPGCs significantly decreases the germ cell number in genital ridges [57]. The surviving germ cells show no de-repression of retrotransposons, consistent with the result that the repressive state in the DNMT1-knockout mESCs is recovered after the transient de-repression. Intriguingly, these surviving germ cells show premature expression of genes for oocytes and sperm. Analyses of acute phenotypes of knockout of DNMT1 and/or UHRF1 in mPGCs could

Reprogramming of mouse germ cells

11

help to elucidate the mechanisms of transposon repressions during the germline epigenome reprogramming.

Interplay between DNA demethylation and histone remodeling Interestingly, erasure of DNA methylation during mPGC specification is associated with histone marks H3K27ac and H3K27me3. In mESCs, the genomic domains of a few 10 kilobases covering w2.1% of the genome have very low methylation levels. These domains are enriched with H3K27ac and contain genes for pluripotency, such as Pou5f1. In contrast, during the mPGC specification, another type of genomic domains of a similar size covering w0.05% of the genome are rapidly demethylated. These domains are enriched with H3K27me3 and developmental genes (e.g., Hox genes), probably reflecting the differences between the regulations of epigenome reprogramming in early embryogenesis and in the germline development [13]. mPGCLCs and mPGCs are able to further proliferate in an adhesive culture system without gonadal somatic cells [8]. In this system, DNA methylation continues its progressive decline to a nadir of w6%, which is equivalent to the DNA methylation level of E13.5 germ cells, but, interestingly, the gene-expression profile and the genomic distribution of active enhancers are very similar to those of early mPGCs. Thus, the comprehensive DNA methylation erasure in mPGCs does not require gonadal signals, and occurs independent from the sex differentiation of germ cells, which starts at E12.5 in response to gonadal cues. Moreover, DNA demethylation of mPGCs itself does not result in global change of the transcriptional program [8,42]. Interestingly, during the adhesive-culture proliferation of mPGCLCs, H3K27me3 is upregulated, compensating for the DNA demethylation. Taken together, these studies demonstrate that Polycomb and SETDB1, which catalyze H3K27me3 and H3K9me3, respectively, serve as important mechanisms of gene repression during the mPGC development. These modifications interplay with DNA methylation. SETDB1 is required for the maintenance of DNA methylation at specific regions, including transposons. H3K27me3 is associated with rapid DNA demethylation, while it is upregulated compensating for DNA demethylation.

Epigenome in sexually differentiated germ cells in mice In embryonic testes and ovaries, mPGCs begin differentiation and at E12.5, the sexual dimorphism becomes evident in the transcriptome [58], while the epigenome reprogramming continues. At E13.5, germ cells are comprehensively demethylated, with their level of DNA methylation becoming lower than at any other point in their life cycle. Subsequently, during the sex specific differentiation, the DNA methylomes of male and female germ cells show obvious sexual dimorphisms.

Epigenome reprogramming and female germ cell pathway In the embryonic ovaries, female germ cells cease mitotic proliferation and enter the meiotic prophase. Throughout this developmental stage, the level of genomic DNA methylation is retained low. Only after birth, germ cells begin de novo DNA methylation during oogenesis. The feminization and meiotic initiation of mPGCLCs have been achieved by stimulating the mPGCLCs proliferated in the adhesive culture [8] with a combination of retinoic acid (RA) and BMP signaling [9] (see also Box 1.1). Cytological and transcriptomic features show that the in vitro female pathway faithfully recapitulate those of the in vivo counterparts from E12.5 to E14.5. Interestingly, if mPGCLCs do not experience proliferation on the adhesive culture, they do not enter the female pathway and meiosis in response to RA and BMP [9]. In these mPGCLCs, DNA methylation is not yet fully reprogrammed, and promoters for the later germ cell differentiation and/or

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Chapter 1 Epigenome reprogramming in the male and female

meiosis (e.g., Ddx4, Dazl, Scp1) are particularly highly methylated awaiting the comprehensive demethylation during the proliferation in genital ridges or in the adhesive culture [8,13]. Thus, the DNA demethylation may put the germline epigenome in a permissive state for the developmental pathway specific to female. This would explain why the BMP signaling functions in different ways between the mPGC specification and the female germ cell pathway.

Androgenic epigenome programming in male embryonic germ cells The male germ cells eventually establish the stem cell system for spermatogenesis and may undergo a more complex pathway than the female germ cells. In mouse embryonic testes, male germ cells arrest their mitotic proliferation and differentiate into prospermatogonia, which further differentiate into spermatogonia and resume mitotic proliferation after birth. The germ cells show a drastic elevation of overall DNA methylation levels throughout this differentiation process. In contrast to this global de novo DNA methylation, genes for spermatogenesis, which are not necessarily expressed in the spermatogonia themselves, retain a low DNA methylation level similar to the level at E13.5, preparing permissive states for the future spermatogenesis. Retrotransposons, of which DNA methylation is partially retained during the epigenome reprograming, also resist the initial de novo DNA methylation when mPGCs differentiate into the prospermatogonia. They retain a DNA methylation level similar to the lowest level at E13.5, resulting in expression of some of them [e.g., intracisternal A-particles (IAPs) and long interspersed elements (LINEs)]. This reflects the fact that the DNA methylation mechanism for the retrotransposons is different from that of the other portions of genomic DNA; de novo DNA methylation of retrotransposons is mediated by the piRNA pathway, in which the expressed retrotransposons are degraded by the PIWI protein and piRNA, and then used as guide RNAs for their DNA methylation [59]. Chromatin remodeling during spermatogenesis, which occurs after birth, will be discussed in Chapter 3 in this book.

Epigenome aberration during in vitro differentiation of male germ cells The differentiation into male germ cells has also been reconstituted in vitro, using reconstituted testes composed of mPGCLCs before the adhesive-culture proliferation and somatic cells from embryonic testes [5]. Cell lines very similar to germline stem cells (GSCs) [60] are derived from the reconstituted testes (GSC-like cells). The GSC-like cells show overall transcriptional and DNA methylation profiles similar to those of GSCs. However, only a limited fraction of GSC-like cells contain physiologically functional spermatogonial stem cells [5]. GSC-like cells show hypermethylation in promoters of as many as 3e5% of all genes, associated with aberrant expression of transposons and genes for meiosis. mPGCLCs used in this system did not experience proliferation on the adhesive culture and retained substantial genomic DNA methylation. Thus, insufficiency of epigenome reprogramming may explain the aberrant DNA methylome and the insufficient cellular activity of GSC-like cells as spermatogonial stem cells. In line with this, defects in MIWI2 (the piRNA pathway) and DNMT3L (de novo DNA methylation) during the male germ cell development, most likely during the androgenic programming, result in an aberrant transcriptome in spermatogonia, including de-repression of retrotransposons [61]. Another group has reported that a two-week co-culture of mPGCLCs and neonatal testicular somatic cells results in formation of functional spermatid-like cells [6]. In male mice, the first-wave spermatogenesis begins around postnatal day 10 (P10), producing round spermatids around P20. Thus, it would take about 30 days for mPGCLCs, which are highly similar to the E9.5 mPGCs, to differentiate into spermatids through the normal developmental pathway. Because the epigenome

Reprogramming of mouse germ cells

13

reprogramming in mPGCs and the androgenic programming in spermatogonia occur during this developmental period as discussed above, the in vitro spermatogenesis described above should be examined thoroughly for transcriptome and epigenome dynamics. Taken together, the in vitro reconstitution of sexually differentiated male and female germ cells highlights the importance of the appropriate epigenome reprogramming during the development of mPGCs.

Summary of the epigenome reprogramming of embryonic germ cell development The studies discussed above have investigated the epigenome regulations during the in vivo and in vitro development of mPGCs and their sex differentiation, and the results collectively suggest several key features of the epigenome reprogramming in the germ line. mPGCs show large-scale changes of histone modifications and erasure of DNA methylation to shape an epigenetic state for gene repression, depending on Polycomb and SETDB1, which catalyze H3K27me3 and H3K9me3, respectively. This repressive state is enriched with the poised transcriptional state with H3K4me3-H3K27me3 bivalency. Erasure of DNA methylation is compensated by H3K27me3 and H3K9me3, and only limitedly influences the transcriptome of mPGCs. Rather, the promoter demethylation prepares the late germline genes for expression in the appropriate timing. This epigenetic state persists until the beginning of sex-specific differentiation of germ cells. Then, germ cells respond to extrinsic signals, acquiring the sex-specific epigenome.

Oocytes form a unique epigenome during oogenesis De novo DNA methylation and genomic imprinting during oogenesis Mouse oogenesis begins at around P10. From the beginning of their growth to the germinal vesicle (GV) stage, oocytes gradually acquire de novo DNA methylation, including the establishment of imprint marks mediated by DNMT3A and DNMT3L. The de novo DNA methylation during oogenesis requires transcription and occurs in transcribed regions [62e64]. In line with this, transposons with long terminal repeats (LTR) expressed in oocytes are required for DNA methylation of a subset of imprinting regions, and the species differences of such LTR transposons explains a part of the species differences of genomic imprinting [65]. On the other hand, imprinting regions do not only acquire DNA methylation but also protect it from the extensive demethylation during the early embryogenesis after fertilization. The latter process is mediated by at least two DNA-binding proteins, ZFP57 and ZNF445 [66,67]. The evolutionary diversity of the target sequences of these proteins might also contribute to the generation of species differences in the imprinting regions.

Interplay between histone modification and de novo DNA methylation Histone modifications mediate the interaction between transcription and DNA methylation during oogenesis; H3K4me3 (active promoters) is downregulated prior to the de novo DNA methylation, while H3K36me3 (transcribed gene bodies) is upregulated during the methylation [68,69]. These relationships are underpinned by biochemical interactions between these modifications and DNA methyltransferases; H3K4me3 excludes DNMT3A and DNMT3L, while DNA methylation conversely inhibits deposition of H3K4me3; and H3K36me3 recruits DNMT3A and DNMT3B to genomic DNA [70e74].

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Chapter 1 Epigenome reprogramming in the male and female

Broad domains of H3K4me3 during oogenesis Recently, it has been revealed that oocytes acquire a unique histone modification profile during oogenesis, as demonstrated by low-input ChIP-seq analyses; mature oocytes have broad domains of H3K4me3 that are not associated with gene expression [74e77]. These domains cover about 22% of the genome, including putative enhancers and intergenic regions. This type of H3K4me3 distribution is not observed in the “usual” cell types, in which H3K4me3 marks transcriptionally active promoters and forms narrow peaks in genomic tracks of ChIP-seq. The H3K4me3 distribution gradually changes from the active promoters to the broad domains during oogenesis. This process is independent from transcription, and requires MLL2, an H3K4 methyltransferase [74]. These broad domains of H3K4me3 disappear at the start of the major zygotic gene activation (ZGA) after fertilization [75e77]. Then, H3K4me3 reverts to active promoters.

Histone replacement during oogenesis Oocytes show cell cycle arrest at the meiotic prophase I. HIRA, the chaperone of replicationindependent histone H3.3, is required for oocytes to establish proper genomic states [78]. Thus, the chromatin of oocytes is dynamic during oogenesis, and is maintained through the turnover of H3.3.

Three-dimensional organization during oogenesis Genomic DNA is not only packaged into chromatins, but also forms a three-dimensional structure. During maturation of oocytes, such a structure becomes weaker, as demonstrated by a single-cell HiC-seq analysis [79] (see also Box 1.4). In the metaphase of meiosis II, oocytes lose the three-dimensional structure of the genome [80], similarly to the case for mitotic metaphase. After fertilization, preimplantation embryos gradually recover the three-dimensional organization. Taken together, the findings discussed above show the formation of a unique epigenome of oocytes, including unique dynamics of histone modification and DNA methylation, as well as the unique three-dimensional structures of the genome. These oocyte-specific epigenetic features are extensively reprogrammed during early embryogenesis, signifying the formation of pluripotent cells from the totipotent fertilized egg. Details of genomic imprinting and epigenome reprogramming during embryogenesis will be discussed in detail in Chapters 2 and 6 of this book, respectively.

Epigenome reprogramming during the development of human germ cells Elucidation of developmental mechanisms of human germ cells, including epigenome reprogramming, is challenging because of the limitations of the experimental investigation of human embryos. This is especially the case for PGCs (human PGCs; hPGCs). Moreover, there are significant morphological differences between human and mouse embryos, making it difficult to infer the mechanisms of human development based on those of mice. Recent advances in the germ cell studies in human embryos using low-input genomics, achievements of in vitro reconstitution of hPGCs (hPGC-like cells; hPGCLCs), and embryological studies of surrogate models have significantly improved our knowledge of human germ cells. These studies show that development of human germ cells is in general similar to that of mice, but has greater developmental heterogeneity and significant species differences in signaling, transcriptional, and epigenetic mechanisms.

Epigenome reprogramming during the development of human germ cells

15

PGC specification and migration in human embryos After implantation of human embryos, epiblast and hypoblast form a bilaminar disc around week 2 (Wk2) (Fig. 1.2). A part of epiblast that attaches to the cytotrophoblast, which is derived from the polar trophectoderm, is directly differentiated into the amnion, forming an amniotic cavity. The hypoblast forms the roof and membranous wall of the primary yolk sac (also known as the Heuser’s membrane), which is soon replaced by the secondary yolk sac (also known as the definitive yolk sac) composed of additional hypoblast-derived cells. Then, gastrulation begins early in Wk3. The developmental stage in which hPGCs are first identified is around Wk3, at the time of which hPGCs are found in the endoderm of yolk sac. hPGCs are histologically identifiable with canonical

Human

Birth

inner nner cell mass syncytiocyto- trophoblast

PGCs Gonocyte

trophoblast

Pre-spermatogonia

Gonocyte proliferation

Spermatogonia

Birth epiblast

amnion

hypoblast

gut endoderm

gonad

2nd yolk sac

Wk1

Wk2

Oogonia

Wk3

Wk4

Wk7

Wk9

Oogania proliferation Heterogeneous differentiation

Wk20

Oogenesis

Puberty

hPGC specification (EOMES, SOX17, BLIMP1, TFAP2C)

Progressive global glo CpG demethylation similar to mPGCs

In vitro reconstituted systems ActA GSKi

hPSCs

BMP4

iMELCs

Xenogeneic reconstituted ovaries

hPGCLCs

Oogonia-like cells

BMP4

FIG. 1.2 Schematic representation for the epigenome reprogramming of human germ cell s and their reconstitution in vitro. Currently known, major events of epigenome reprogramming are listed below the developmental stages in vivo. Although the origin of human PGCs is unknown, it is most likely the nascent amnion. The corresponding in vitro germ cell reconstitution systems are also represented with colored circles. The color code for the circles for the PGCLC induction is the same as in Fig. 1.1: the circles for human PSCs, iMELCs (direct origin of PGCLCs), and hPGCLCs are colored pale blue, dirk purple, and orange, respectively.

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Chapter 1 Epigenome reprogramming in the male and female

morphological characteristics (reviewed in Ref. [81]). At Wk3, the number of hPGCs is as small as about 40, which is similar to the number of mPGCs immediately after specification. Then, from around Wk4, hPGCs migrate from the dorsal yolk sac into the hindgut endoderm. At Wk5eWk6, hPGCs enter genital ridges through the dorsal mesentery, colonize there, and undergo sex differentiation. The process of hPGC specification can be inferred from the process of PGC specification in cynomolgus monkeys, whose gastrulating embryos are very morphologically similar to human embryos. PGCs of cynomolgus monkeys are derived from the nascent amnion prior to gastrulation. In conceptuses of cynomolgus monkeys, BMP4 is secreted in the amnion itself and WNT3A is secreted in the cytotrophoblast [82]. Thus, the origin of hPGCs is most likely the nascent amnion. On the other hand, another model animal, pigs, which form a bilaminar embryonic disc prior to gastrulation, first show detectable PGCs in the midline of the posterior epiblast at the early primitive streak stage [83]. Thus, there is a substantial species difference in the PGC specification processes even among mammals with bilaminar disc-shaped embryos.

Reconstitution of early human PGCs and epigenome reprogramming in vitro Human pluripotent stem cells As described above, the human embryogenesis around gastrulation differs substantially from that of mice in terms of both morphology and time frame. Consistent with this, there are many differences between human and mouse PSCs. A single-cell transcriptome study with a 30 -sequencing method [84] has revealed that the post-implantation epiblast of cynomolgus monkeys shows characteristics very similar to PSCs of humans and cynomolgus monkeys [85]. This developmental coordinates of primate PSCs are different from the mPSCs, the in vivo counterpart of which is the ICM cells in blastocysts [86].

Induction of human PGC-like cells and their signaling and transcriptional regulation From the human PSCs, human PGCLCs (hPGCLCs) have been induced by BMP4 directly or via incipient mesoderm/primitive streak-like cells (iMELCs) [17e19] (see also Box 1.2). hPGCLCs are very similar to early PGCs of cynomolgus monkeys in terms of transcriptome [82], and thus, most likely recapitulate early hPGCs. In addition to BMP4, the hPGCLC induction also requires WNT signaling [20,22,83] as in the case of mPGC specification, and thus, the signaling principles of human and mouse PGC specification are similar to each other. On the other hand, SOX17 is an essential factor for hPGCLC induction directly upstream of BLIMP1 [17,18,20]. This is in stark contrast to the mPGC development, in which SOX17 is transiently expressed but is not essential [31,87]. For the expression of SOX17 in hPGCLCs, transient upregulation of EOMES, which is a TF required for the proper gastrulation in mice, is required [17,18], while T (BRACHYURY), which is a critical TF for the mesoderm development in mice and is the direct activator of Blimp1 and Prdm14 in the mPGC specification, does not seem to play any role in this process. PRDM14, which is critical for the mPGC specification [28], is expressed at low levels in hPGCLCs as well as in the PGCs of cynomolgus monkeys, and seems to play only a limited role [19,82]. Moreover, genes for the development of somatic lineages are progressively downregulated in hPGCLCs, in contrast to their rapid repression in mPGCs. These studies demonstrate the evolutionarily conserved principles and species diversities for the signaling and transcriptional mechanisms of mammalian PGC specification.

Epigenome reprogramming during the development of human germ cells

17

Reprogramming of human PGCs reconstituted in vitro Despite the species difference in the transcriptional mechanisms of mammalian PGC specification, hPGCLCs show gross properties of histone marks similar to those of mPGCs, as assayed with immunofluorescence (high H3K27me3 and low H3K9me2) [17,88]. This suggests that hPGC specification undergoes chromatin remodeling similar to that of mPGCs. On the other hand, the DNA methylation level of hPGCLCs is only slightly lower than that of human iPSCs (hiPSCs), suggesting that hPGCs undergo erasure of DNA methylation much more slowly than mPGCs, in line with that the germ cell development in humans occurs in a longer time span than that in mice.

Sex-specific differentiation of human germ cells Differentiation of female germ cells Human embryonic testes and ovaries begin sex differentiation at around Wk5eWk6 and Wk6eWk8, respectively [81]. In the female, hPGCs differentiate into oogonia, which are mitotically active and form clusters/cysts of dividing cells surrounded by simple squamous epithelia. At Wk10eWk11, some of the human oogonia differentiate into primary oocytes and enter meiosis, and the others continue their mitotic proliferation. During this period, folliculogenesis occurs, and even antral follicles are occasionally formed in the embryonic ovaries. These features of human female germ cell development are in stark contrast to those of mice, because mPGCs homogeneously cease mitotic proliferation and enter meiosis [89,90]. The human oogonia are mitotically active; the estimated number of female germ cells is about 300,000 at Wk9, and becomes as high as 7,000,000 at Wk20 (an w23-fold increase, corresponding to about 4.5 doublings) [81]. However, these cells are extensively eliminated through the perinatal germ cell loss, and only a part of the oogonia survive after birth.

Differentiation of male germ cells In the male, gonocytes, the male hPGCs in testes, form sex cords in combination with Sertoli cells from Wk7. Gonocytes are mitotically active and positive for pluripotency markers and hPGCs genes until Wk12. During Wk13eWk26, gonocytes show progressive differentiation into prespermatogonia (embryonic spermatogonia), and lose their mitotic activity. Then, spermatogonia migrate to the basement membrane of seminiferous tubules. This is also in contrast to mouse germ cell development, in which male germ cells homogenously enter mitotic arrest. The mitotic activity of human gonocytes is similar to that of oogonia; the estimated number of male germ cells is about 150,000 at Wk9, and becomes about 4,000,000 at Wk20 (an w27-fold increase, corresponding to about 4.8 doublings). In consideration of all the above, human germ cells develop in a generally similar manner to mouse germ cells, but they are much more heterogeneous and develop much more slowly [81].

Epigenome reprogramming during development of human germ cells in genital ridges Low-input DNA methylome analyses have revealed that the reprogramming of genomic DNA methylation in human gonadal germ cells from Wk5.5 to Wk20 occurs in a generally similar manner to those of mice in the corresponding stages [88,91e94]. The hPGCs enter genital ridges at around Wk5eWk7, and

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Chapter 1 Epigenome reprogramming in the male and female

by these developmental stages, hPGCs have already achieved substantial reprogramming of DNA methylation (20%e40% methylation). This reprogramming kinetics is similar to that in mice, in that mPGCs are substantially demethylated before they enter the genital ridges. The gonadal germ cells reach the lowest DNA methylation level (about 5%) at around Wk9eWk11 [88,91,92], at which time the oogonia and gonocytes are mitotically active and generally do not yet begin further differentiation into primary oocytes or prespermatogonia, respectively. This is also similar to mouse gonadal germ cell development, in that mouse germ cells reach the lowest DNA methylation levels by E13.5, before the female meiotic entry and the male mitotic arrest. The human germ cells in genital ridges show the DNA methylation pattern similar to those of hiPSCs and iMeLCs, which show developmental coordinates similar to postimplantation epiblast and nascent amnion, the putative origin of hPGCs (see also Box 1.2), but not to preimplantation embryos (blastocyst and naı¨ve human ESCs) [23]. The observation that human germ cells progressively decrease the DNA methylation level while presumably retaining the methylation patterns of epiblast and nascent amnion supports the idea that hPGCs may undergo DNA methylation reprogramming through a replication-coupled manner similarly to the case of mPGCs. In addition, similarly to the case in mice, human retrotransposons, especially those that are evolutionarily young (e.g., SINE-variable number of tandem repeats-Alu elements), escape the erasure of DNA methylation [23,88]. On the other hand, after epigenome reprogramming, germ cells may behave in considerably different manners between humans and mice. As discussed above, human oogonia and gonocytes are mitotically active; they have been estimated to increase 23e27-fold (4.5e4.8 doublings) over the 77 days of proliferation from Wk9 to Wk20 [81]; this means one mitosis per every 16e17 days. Thus, human germ cells seem to continue proliferation even after they are comprehensively reprogrammed, in contrast to mouse germ cells, which cease or arrest mitosis immediately after reprogramming. These species differences are probably due to the differences of developmental kinetics and heterogeneity between human and mouse germ cells.

Epigenome reprogramming in human oogonia reconstituted in vitro Because there are inaccessibility and/or difficulties in experimental investigation of human germ cells, a reconstitution system in vitro for the sex differentiation of hPGCs may help our understanding of the regulatory mechanisms of human germ cell development. In a recent study, female hPGCLCs were differentiated into oogonia-like cells when aggregated with somatic cells in mouse embryonic ovaries [23] (see also Box 1.2). The human oogonia-like cells show a gene expression profile similar to that of the RA-responsive cells in the human embryonic ovaries [95], and thus they progress along a developmental pathway to meiotic oocytes. The genome-wide DNA methylation level is as high as about 80% in hiPSCs and iMELCs, and slightly decreases in hPGCLCs [23]. The human oogonia-like cells progressively decrease the DNA methylation levels, and eventually at day 120, show DNA methylation level and pattern similar to those of gonadal germ cells at Wk7eWk10 [88,92]. Moreover, in the oogonia-like cells, parental imprinting and aberrant methylation derived from hiPSCs are erased. The oogonia-like cells also show progressive reactivation of X-chromosome, a hallmark of female germ cell development. These features indicate that the oogonia-like cells mimic bona fide epigenome reprogramming in the human germ line.

Glossary

19

Because experimental investigation for the development of human germ cells remains relatively poor, additional analyses of human germ cells in vivo and further improvements of human gametogenesis in vitro would contribute to our understanding of the mechanisms of human germ cell reprogramming.

Concluding remarks During the last decade, our knowledge of germ cell development has been greatly improved, at least partially due to the low-input genomics, including single-cell analyses, and the achievements of in vitro reconstitution of PGCs, together with many other physiological, genetic, signaling, and metabolic studies. The details of epigenome reprogramming in the germ line will contribute to our understanding of how germ cells properly shape their epigenome to regulate gene expression, and of how germ cells reset their epigenome for the development of the next generation and for the transmission of part of their epigenetic information. One of the major challenges in this field would be further progress in human in vitro gametogenesis, which will provide enormous opportunities for the experimental investigation of human gem cell development. In addition, research into the mechanisms that produce species differences in mammalian germ cell development would also improve our knowledge of the features specific to humans and the evolution of the germ line.

Glossary ChIP (chromatin immunoprecipitation) Methods to determine DNA fragments bound to a known protein through immunoprecipitation using specific antibodies. Epiblast A pluripotent simple columnar epithelium that arises in the embryo around implantation. Epiblast forms all of the embryo proper and part of the placental tissues. EpiLCs (epiblast-like cells) Culture cells induced from pluripotent stem cells by stimulation of activin and FGF2 signaling. EpiLCs arise only transiently, and around day 2 of induction, they show transcriptome and developmental potency very similar to those of the epiblast. GSCs (germline stem cells) Culture cells derived from spermatogonia. GSCs show a robust spermatogonial stem cell activity when transplanted into the seminiferous tubules of adult mice. HiC-seq A method to determine the three-dimensional genomic structure. Naı¨ve pluripotency A state of pluripotency that is similar to the state of the inner cell mass (ICM) of the preimplantation blastocyst. PGCs (primordial germ cells) The origin of all germ cell lineages. PGCs appear in the first trimester of the embryogenesis, and show no sexual dimorphisms except for the features of X chromosomes. PGCLCs (PGC-like cells) In vitro reconstitution of PGCs, induced from PSCs directly or indirectly. Pluripotency Cellular potential that can contribute to all cell types from the three germ layers in individuals. Primed pluripotency A state of pluripotency that is similar to the state of the postimplantation epiblast. PSCs (pluripotent stem cells) Culture cells that have self-renewal activity and pluripotency. WGBS (whole genome bisulfite sequencing) Methods to determine 5-methylatd cytosine at single base resolution in the whole genome, relying on the combination of bisulfite reaction and high-throughput sequencing.

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Chapter 1 Epigenome reprogramming in the male and female

Acknowledgments We thank Junko Komeda, a member of our laboratory, for her help with the laboratory management. This work was supported in part by Japan Society for the Promotion (JSPS) KAKENHI grants (JP18H0553, JP16H04720, and JP18K19295) and by funds from the Takeda Science Foundation.

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Chapter 1 Epigenome reprogramming in the male and female

[78] Nashun B, Hill PW, Smallwood SA, Dharmalingam G, Amouroux R, Clark SJ, et al. Continuous histone replacement by Hira is essential for normal transcriptional regulation and de novo DNA methylation during mouse oogenesis. Mol Cell 2015;60(4):611e25. [79] Flyamer IM, Gassler J, Imakaev M, Brandao HB, Ulianov SV, Abdennur N, et al. Single-nucleus Hi-C reveals unique chromatin reorganization at oocyte-to-zygote transition. Nature 2017;544(7648):110e4. [80] Ke Y, Xu Y, Chen X, Feng S, Liu Z, Sun Y, et al. 3D chromatin structures of mature gametes and structural reprogramming during mammalian embryogenesis. Cell 2017;170(2):367e381.e20. [81] De Felici M. In: Coticchio DFA G, De Santis L, editors. Origin, migration, and proliferation of human primordial germ cells. London: Springer-Verlag; 2013. [82] Sasaki K, Nakamura T, Okamoto I, Yabuta Y, Iwatani C, Tsuchiya H, et al. The germ cell fate of cynomolgus monkeys is specified in the nascent amnion. Dev Cell 2016;39(2):169e85. [83] Kobayashi T, Zhang H, Tang WWC, Irie N, Withey S, Klisch D, et al. Principles of early human development and germ cell program from conserved model systems. Nature 2017;546(7658):416e20. [84] Nakamura T, Yabuta Y, Okamoto I, Aramaki S, Yokobayashi S, Kurimoto K, et al. SC3-seq: a method for highly parallel and quantitative measurement of single-cell gene expression. Nucleic Acids Res 2015; 43(9):e60. [85] Nakamura T, Okamoto I, Sasaki K, Yabuta Y, Iwatani C, Tsuchiya H, et al. A developmental coordinate of pluripotency among mice, monkeys and humans. Nature 2016;537(7618):57e62. [86] Nichols J, Smith A. Naive and primed pluripotent states. Cell Stem Cell 2009;4(6):487e92. [87] Hara K, Kanai-Azuma M, Uemura M, Shitara H, Taya C, Yonekawa H, et al. Evidence for crucial role of hindgut expansion in directing proper migration of primordial germ cells in mouse early embryogenesis. Dev Biol 2009;330(2):427e39. [88] Tang WW, Dietmann S, Irie N, Leitch HG, Floros VI, Bradshaw CR, et al. A unique gene regulatory network resets the human germline epigenome for development. Cell 2015;161(6):1453e67. [89] Koubova J, Menke DB, Zhou Q, Capel B, Griswold MD, Page DC. Retinoic acid regulates sex-specific timing of meiotic initiation in mice. Proc Natl Acad Sci U S A 2006;103(8):2474e9. [90] Bowles J, Knight D, Smith C, Wilhelm D, Richman J, Mamiya S, et al. Retinoid signaling determines germ cell fate in mice. Science 2006;312(5773):596e600. [91] Gkountela S, Zhang KX, Shafiq TA, Liao WW, Hargan-Calvopina J, Chen PY, et al. DNA demethylation dynamics in the human prenatal germline. Cell 2015;161(6):1425e36. [92] Guo F, Yan L, Guo H, Li L, Hu B, Zhao Y, et al. The transcriptome and DNA methylome landscapes of human primordial germ cells. Cell 2015;161(6):1437e52. [93] Eguizabal C, Herrera L, de Onate L, Montserrat N, Hajkova P, Izpisua Belmonte JC. Characterisation of the epigenetic changes during human gonadal primordial germ cells reprogramming. Stem Cell 2016;34(9): 2418e28. [94] Guo H, Hu B, Yan L, Yong J, Wu Y, Gao Y, et al. DNA methylation and chromatin accessibility profiling of mouse and human fetal germ cells. Cell Res 2017;27(2):165e83. [95] Li L, Dong J, Yan L, Yong J, Liu X, Hu Y, et al. Single-cell RNA-seq analysis maps development of human germline cells and gonadal Niche interactions. Cell Stem Cell 2017;20(6):858e873 e4. [96] Ying QL, Wray J, Nichols J, Batlle-Morera L, Doble B, Woodgett J, et al. The ground state of embryonic stem cell self-renewal. Nature 2008;453(7194):519e23. [97] Morohaku K, Tanimoto R, Sasaki K, Kawahara-Miki R, Kono T, Hayashi K, et al. Complete in vitro generation of fertile oocytes from mouse primordial germ cells. Proc Natl Acad Sci U S A 2016;113(32): 9021e6. [98] Sato T, Katagiri K, Gohbara A, Inoue K, Ogonuki N, Ogura A, et al. In vitro production of functional sperm in cultured neonatal mouse testes. Nature 2011;471(7339):504e7.

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[99] Dolci S, Williams DE, Ernst MK, Resnick JL, Brannan CI, Lock LF, et al. Requirement for mast cell growth factor for primordial germ cell survival in culture. Nature 1991;352(6338):809e11. [100] Frommer M, McDonald LE, Millar DS, Collis CM, Watt F, Grigg GW, et al. A genomic sequencing protocol that yields a positive display of 5-methylcytosine residues in individual DNA strands. Proc Natl Acad Sci U S A 1992;89(5):1827e31. [101] Miura F, Enomoto Y, Dairiki R, Ito T. Amplification-free whole-genome bisulfite sequencing by postbisulfite adaptor tagging. Nucleic Acids Res 2012;40(17):e136. [102] Bernstein BE, Kamal M, Lindblad-Toh K, Bekiranov S, Bailey DK, Huebert DJ, et al. Genomic maps and comparative analysis of histone modifications in human and mouse. Cell 2005;120(2):169e81. [103] Dekker J, Rippe K, Dekker M, Kleckner N. Capturing chromosome conformation. Science 2002; 295(5558):1306e11.

CHAPTER

2

Genomic imprinting

Sharvari Deshpande
, Sweta Nair
, N.H. Balasinor Neuroendocrinology, ICMR- National Institute for Research in Reproductive Health, Mumbai, Maharashtra, India

Introduction In mammals, most of the autosomal genes are either biallelically expressed or repressed from the two parental alleles following the Mendelian law of inheritance. However, in a small fraction of the genes, one allele is selectively repressed in a parental origin-specific manner. This phenomenon is a known as genomic imprinting. Genomic imprinting is defined as monoallelic expression of a gene or chromosomal region depending on the parental origin of inheritance [108]. This phenomenon was first discovered by Surani and Solter and colleagues. Both independently demonstrated the functional nonequivalence of maternal and paternal genomes by pronuclear transplantation experiments in mice. They created gynogenetic and androgenetic embryos by transplanting either two female pronuclei or two male pronuclei respectively in enucleated eggs. Both these engineered embryos did not develop to term and were arrested at different developmental stages. The gynogenotes (which were equivalent to parthenogenotes) had recognizable embryo present with poor extraembryonic tissue, whereas the androgenotes had poor embryo development but the trophoblast proliferated excessively. These experiments demonstrated that diploidy is not sufficient, both maternal and paternal genes are essential for normal embryonic development and the two parental alleles are functionally non-equivalent [88,117]. This study was followed by identification of various imprinted genes that are involved in biparental contribution for normal embryo development [7,8,118]. These genes have been shown to play crucial roles in embryo development, placental function as well as in neurodevelopment and postnatal behavior [5,49,62,105] (Table 2.1). Till date, about 126 imprinted genes have been discovered in mouse [1], some of which have been conserved and imprinted in humans. A well-designed repository namely MetaImprint (http://bioinfo. hrbmu.edu.cn/MetaImprint), has been developed by Yan Zhang and his colleagues, which focuses on detail information on imprinted genes in eight mammalian species. In humans, uniparental gynogenotes and androgenotes are clinically recognized as ovarian teratoma and hydatidiform mole, respectively [93]. The complete hydatidiform mole arises from the fertilization of an enucleated egg either by a haploid sperm (followed by duplication of the paternal genome) or two haploid sperm (diandric diploidy). This trophoblastic disease is characterized by a


Equal contribution.

Epigenetics and Reproductive Health. https://doi.org/10.1016/B978-0-12-819753-0.00002-7 Copyright © 2021 Elsevier Inc. All rights reserved.

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Chapter 2 Genomic imprinting

Table 2.1 Imprinted genes and their knockout phenotypes. Imprinted genes

Knockout phenotype

References

Kcnq1ot1

10e20% reduction in body weight Motor dysregulation, learning deficit, seizures Memory or learning defects, small size Reduced placental growth and fetal growth restriction Placental and fetal overgrowth Lean, insulin resistant, enhanced fasting- and aging-related metabolism Embryo overgrowth and lethality after birth Placental and fetal overgrowth Placental and fetal growth abnormalities Fetal and post-natal developmental abnormalities Phenotypic variations in the offspring, placentomegaly and phenotypic defects in neonates Fetal growth restriction, bone related abnormalities, risk of post-natal lethality Growth retardation, skeletal abnormalities, reduced litter size and neonatal lethality Early embryo lethality due to abnormal placenta formation Early embryo lethality due to abnormal placenta formation

[85]

Ube3a Rasgrf1 Igf2

H19 Igf1r

Igf2r Grb10 Mest/Peg1 Gtl2/Meg3

Cdkn1c

Plagl1

Dlk1

Ascl2/Mash2

Peg10

[64] [12,63] [22,23]

[75] [44]

[126] [16] [74] [111]

[120,121,142]

[125]

[92]

[50]

[96]

Introduction

29

Table 2.1 Imprinted genes and their knockout phenotypes.dcont’d Imprinted genes

Knockout phenotype

References

Rtl1

Abnormal placentation, growth retardation, postnatal mortality Placentomegaly with expansion of spongiotrophoblasts Growth retardation of placenta and embryo and abnormal maternal behavior Embryo lethality Growth retardation after birth, dysfunction of sleep and food uptake Not known Not known Not known KO of imprinting control region leads to increase in the number of underweight neonates and also increases the risk of increases offspring mortality Not known Not known Placental overgrowth Not known Not known Not known

[112]

Ipl

Peg3

Gna/Nesp55 Magel2

Dio3 Ndn Ipw Snrpn

Znf127 Wt1 Phlda2 Nnat Znf331 Grb7

[45]

[76]

[137,138] [9]

[33]

[45]

completely androgenetic genome and results in reduced or absent fetal growth coupled with hyperplastic extraembryonic growth. In contrast, ovarian teratoma arises from the spontaneous activation of an oocyte resulting in the duplication of the maternal genome and is composed of a disorganized mass of differentiated embryonic tissues. These abnormalities indicate that even in humans, normal development proceeds only when a complete complement of both the parental genomes is present. These studies also suggested that genes expressed by the paternal genome are directed toward the development of the extraembryonic tissues, while those from the maternal genome are responsible for development of the embryo [93]. The parental specific expression of the imprinted genes is due to differential epigenetic marking mainly in the form of DNA methylation, on the two parental alleles during gametogenesis [6,106]. The parental specific marking on the DNA occurs at specific regions called the Differentially Methylated Regions (DMRs). DMRs are CpG-rich and often fulfill the criteria for CpG islands. The DMRs are a

30

Chapter 2 Genomic imprinting

part of the imprinting control region which controls the parental specific expression (reviewed by Ref. [5,28]). In addition, DMRs are also differentially marked by histone acetylation and methylation [29]. A recent study by Inoue and colleagues revealed that a few genomic loci are maternally imprinted because of the histone 3 lysine 27 trimethylation (H3K27me3)-dependent imprinting on the maternal allele and is independent of DNA methylation [61].

Characteristics of imprinted genes Imprinted genes are often characterized by genomic clustering, regulation by antisense transcripts and presence of repeat elements near or within the DMR. Most of the imprinted genes are found in clusters, which allow coordinated regulation of genes in a given chromosomal region, although single imprinted genes also exist. Each cluster is under the control of a major cis-acting element, the imprinting control region (ICR) or Imprinting Center (IC), which is generally characterized by germline DMR or primary DMR. These imprinted clusters generally contain several protein-coding genes and one or more than one non-coding RNA. Genes in the cluster often have an antisense transcript which may serve to regulate the imprinted expression of their sense gene e.g. KCNQ1/KCNQ1OT1, UBE3A/UBE3A-AS. The short tandem repeat elements within the DMR of several imprinted genes are important for maintenance of their imprinted status [90]. Two imprinted gene clusters have been extensively described in humans, one on chromosome 11p15, which is linked to the pathogenesis of a fetal overgrowth syndrome, that is, BeckwithWiedemann Syndrome (BWS) and the other on chromosome 15q11-13, linked to Angelman Syndrome (AS) and Prader Willi Syndrome (PWS) (reviewed by Ref. [40,80]).

The life cycle of imprints In order to ensure that every generation receives the appropriate sex-specific imprints, the genome undergoes reprogramming. While inherited maternal and paternal ‘imprints’ in the somatic cells of the embryo are maintained and read, they are erased in the germline and new imprints established during gametogenesis according to the sex of the individual [108]. Despite the genome-wide reprogramming following fertilization, and DNA demethylation/remethylation occurring in the embryo, the imprints established in the gametes are maintained in the somatic cells of the embryo. The methylation at DMRs of imprinted genes is maintained by cis-and trans-acting factors that are responsible for recruitment of DNMT1 [78,94,107]. The imprints are ultimately read, resulting in parent-specific gene expression. In the primordial germ cells of the developing organism during the fetal stage, imprints are erased followed by establishment of new imprints depending on the sex of the embryo, thus completing the imprinting cycle (Fig. 2.1) [108]. Although the process of erasure is not completely understood, it appears that imprints are erased through a series of active and passive events, including those involving the action of the ten-eleven translocation (Tet) family which catalyze the oxidation of 5-methylcytosine to 5-hydroxymethylcytosine [51,135]. The establishment of imprints is initiated in the embryonic gonads and extends till meiosis in the adults. However, it occurs at different times in the male and female germlines. In males, it is initiated in the embryonic stage and completed before meiosis, whereas in the females, imprint acquisition occurs around the time of completion of the first meiotic division. The establishment of sex-specific imprint in

Introduction

31

FIG. 2.1 Imprinting cycle during mouse development.

the germ cells is through the action of the de novo DNA methyltransferase 3a (DNMT3A) and DNMT3L [11,53,67]. Recent studies have indicated that transcription through ICR sequences provides an access to the DNA methyltransferase proteins to the target site, leading to allele-specific DNA methylation in the germline [18,54]. It has been shown that not only DNMT3A, DNMT3B, and DNMT3L but also, histone demethylase i.e. KDM1B/AOF1 is involved in the establishment of new imprinting marks in the germline [19]. Certain PIWI-interacting RNAs (piRNA) are also involved in establishing DNA methylation imprinting at RasGrf1 ICR in male gametes [127]. Any errors during the process of maintenance, erasure and establishment of imprints can affect the transmission of appropriate imprinting to the subsequent generation. This could lead to developmental defects or deleterious post-natal consequences (reviewed by Ref. [26,122]). Some imprints are set not in the germline, but rather by immediate demethylation or de novo methylation, after fertilization. These are known as secondary DMRs. A primary DMR is established during gametogenesis and a secondary DMR develops during embryogenesis, most likely, due to a direct influence of a nearby primary DMR [79]. About 35 gametic/germline DMRs (gDMRs) have been identified in the human genome. The maternally methylated gDMRs are mainly intragenic and generally correspond to promoters, often of lncRNAs. gDMRs methylated on paternal chromosomes are intergenic and may function as insulators or enhancers (reviewed by Ref. [90]).

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Chapter 2 Genomic imprinting

Genomic imprinting in embryogenesis Nuclear transplantation experiments in mice demonstrated the distinct roles of the two parental genomes in embryogenesis. In addition, the first identified imprinted genes were shown to be essential for normal embryo growth. The most well-studied imprinted gene cluster that regulates embryo growth is the IGF2-H19 cluster, which has two imprinted genes, namely, IGF2 and H19. IGF2 is expressed from the paternal allele and is a positive regulator of embryo growth. Biallelic expression of Igf2 results in embryonic overgrowth, whereas its reduction leads to growth restriction [6,27,41]. H19, a non-coding RNA, is expressed from the maternal allele and is a negative regulator of embryo growth [46]. IGF2 binds to two receptors, IGF1r and IGF2r. Binding to IGF1r, IGF2 promotes embryo growth, however, IGF2 binding to IGF2r, a maternally expressed imprinted gene, targets it for lysosomal degradation [43]. Besides Igf2, Igf2r and H19, a number of imprinted genes are reported to have an effect on embryonic growth, namely Grb10, Peg1 (Mest), Gtl2 (Meg3), Cdkn1c, Plagl1 (Zac1) and Dlk1. Grb10 is maternally expressed in most murine tissues and acts as a growth restrictor. Maternal Grb10-knockout embryos exhibit overgrowth. Deletion of the Grb10 ICR results in its biallelic expression and significant undergrowth [16,113]. It has been shown that embryonic growth is regulated by Imprinted Gene Network (IGN) involving a number of imprinted genes, including Igf2, Peg1 (Mest), Gtl2 (Meg3), Cdkn1c, Plagl1 and Dlk1. Studies by Varrault et al. demonstrated that about 15 imprinted genes are coordinately regulated in multiple tissues forming an IGN. Zac1 (Plagl1) a paternally expressed imprinted gene regulates the IGN [125]. Deletion of Zac1 gene in mice results in intrauterine growth restriction and neonatal lethality. ZAC1 alters the expression of several imprinted genes, including Cdkn1c and Dlk1, and directly regulates the H19/Igf2 locus [125]. In addition to Zac1, H19 is a possible regulator of the IGN [46] and BMI1, a member of the Polycomb Repressive Complex 1 (PRC1) has also been shown to be implicated in the synchronized expression of multiple imprinted genes within this network [140].

Genomic imprinting in placentation Imprinting has an important role to play in placental development. In fact imprinting evolved with placentation [100]. Even though the placenta is a transient organ, it acts as the supply point of oxygen, growth factors and hormones for the fetus. A study by Arima et al. [3] showed that Dnmt3L is essential for the establishment of maternal imprinting in the embryo and the placenta. About 75 imprinted genes playing crucial roles in the growth and development of the placenta have been identified to be highly expressed in the placenta [95]. Maternally expressed imprinted gene, Mash2 was the first transcription factor shown to play important roles in the development of the trophoblast lineage [50]. In the study, Mash2 null embryos were found to die due to placental defects at Embryonic day (E)10. Deletion of another gene, Peg10, which is paternally expressed, had lethal effects on the embryos, owing to impaired placentation [96]. Rtl1, also a paternally expressed gene is essential for maintaining fetal capillaries. Its upregulation or deficiency, both results in neonatal lethality in mice [112]. The paternally imprinted genes, p57Kip2 and Ipl are involved in development of the labyrinthine and spongiotrophoblast layers of the placenta in rodents [45,120]. Knockout of DNMT3L itself resulted in placental abnormalities such as defective labyrinthine formation, reduced spongiotrophoblast and increased numbers of trophoblast giant cells. Growth of the placenta is known

Genomic imprinting in post-natal development

33

to be regulated by paternally expressed imprinted genes, Peg1 and Peg3 [25,74]. In addition, Chromosome 19 and Chromosome 14 miRNA clusters (C19MC and C14MC, respectively) are two imprinted miRNA gene clusters in humans known to play crucial roles in trophoblast functions such as, proliferation, invasion and migration. C19MC is a primate-specific microRNA cluster, which is one of the largest microRNA clusters in humans. It is paternally expressed and maternally imprinted. C14MC, which is, conserved across eutherian mammals, is a paternally imprinted cluster (reviewed by Ref. [84]). Imprinted genes are involved not only in maintaining the architecture of the placenta but also in regulating nutrient uptake. The uptake of monoamines is inhibited by deletion of the gene Slc22a3 while, nutrient uptake is inhibited by deletion of a placental isoform of Igf2 [22,144].

Genomic imprinting in post-natal development Many aspects of post-natal development such as, behavior at neonatal and adult stages and, brain functions are regulated by imprinted genes. In mice, suckling action of pups is affected by the expression of imprinted genes. An exonic deletion in Gnas contributes to impaired suckling in pups [99]. Deletion of paternally imprinted gene Magel2 also leads to impaired suckling accompanied by reduced appetite [109]. Lack of Peg3 expression results in impaired thermoregulation. This was evident by the inability to withstand cold conditions by the experimental animals [25]. Dlk, a paternally expressed imprinted gene regulates adipose tissue accumulation in the early post-natal period. Its deletion causes increased adiposity [21] and conversely, a rise in its levels results in utilization of triglycerides for energy generation under fasting conditions [14]. In the same locus, another paternally expressed gene, Dio3 affects brown adipose tissue development and heat generation in response to cold external conditions and hence, helps in postnatal survival [15]. The Plagl1 locus that is associated with transient neonatal diabetes in humans is also responsible for neonatal hyperglycaemia in mice when it is overexpressed [81,82]. Hypoinsulinemia occurs in adult mice upon deletion of a paternally expressed gene, RasGrf1 which also causes growth restriction in neonatal stages. Low insulin levels due to reduced beta cell numbers also cause glucose intolerance in these mice [20,42]. Grb10 has differential imprinting status in different tissues; it is paternally expressed in the brain and maternally expressed in peripheral tissues. Deletion of paternal Grb10 in the brain led to increase in social dominance [47], however, deletions of maternal Grb10 led to prenatal and postnatal (w130%) overgrowth, but with an abnormally small brain and large liver. Grb10 and another imprinted gene, Nesp55 are also important for reward receptivity functions of the brain. Loss of maternal allele of Nesp55 releases contentment for an immediately available but smaller-sized reward in mice [30]. On the other hand, loss of paternal Grb10 causes mice to forgo small rewards for a bigger reward [31]. Paternally expressed genes are implicated in maternal behavior. Peg1-deficient female mice showed apathy in the execution of maternal roles and reduced placentophagia [74]. Peg3 deficiency caused impaired ejection of milk from mammary glands of Peg3 mutant female mice, primarily due to increased apoptosis of oxytocin-producing neurons [76]. The gene disruption also reduces numbers of neurons in regions of brain responsible for olfactory and pheromone signaling and, sexual behavior [13]. The gene Grb10 has been found to play a role in mother-pup interactions. Deficiency of maternal allele of Grb10 in pups raised their nutritional demands while deletion of this allele in the mother did not lead to increased milk production [24].

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Chapter 2 Genomic imprinting

Genomic imprinting in disease conditions Infertility Infertility is an important medical and social issue affecting 15% of the couples worldwide. Several studies have shown association between aberrant methylation status of imprinted and non-imprinted genes and, male infertility. Marques et al. demonstrated that oligozoospermic individuals exhibit defective methylation of imprinted genes i.e. hypomethylation at H19 DMR and hypermethylation at MEST DMR [87]. In addition, aberrant imprinting at various other loci like SNRPN, PEG1, LIT1, PEG3, GTL2 and ZAC was reported in moderate to severe oligozoospermic individuals [71]. Several reports have also pointed out hypomethylation of the IGF2/H19 ICR in individuals with low sperm count and motility compared to normozoospermic individuals [52,101]. Further, Kobayashi et al. showed an association between imprinting errors in the sperm and in abortificant samples resulting from ARTs [70]. Studies in our laboratory have also shown aberrant imprinting at H19 ICR in normozoospermic individuals whose female partner was experiencing idiopathic recurrent spontaneous abortions [2]. Using high throughput technique, a handful of studies have shown defective methylation status of both imprinted and non-imprinted genes as well as repetitive elements in individuals with poor semen quality [56,116]. All these studies indicate that proper DNA methylation patterns in spermatozoa are crucial for normal sperm function, fertility and embryo growth (reviewed by Ref. [10]). However, the cause and occurrence of these methylation defects in spermatozoa are still unclear, which could be due to defects in erasure/establishment/maintenance of imprint marks. In addition, abnormal chromatin configuration during the spermiogenesis process, in which histones play a crucial role in establishment of methyl marks on the gametes could be one plausible reason of defective methylation at the imprinted loci (reviewed by Ref. [10]). Very few studies have shown a link between genomic imprinting defects and female infertility. Endometriosis, one of the leading causes of infertility, occurs in 10% of the women in their reproductive age [72]. Several studies have identified altered genome-wide DNA methylation profiles in endometriosis which includes genes associated with steroidogenesis and transcription factors [36,134]. However, few studies have identified aberrant imprinting pattern in this condition. Kobayashi et al. reviewed that out of 29 hypermethylated genes associated with endometriosis in the literature, 19 genes were residing near the known imprinted loci suggesting a plausible interaction of the downregulated genes linked with endometriosis susceptibility with the imprinted genes [69]. Ghazal et al. demonstrated that the expression of H19 gene was significantly reduced in the eutopic endometrium of women with endometriosis. This suggests that faulty H19 expression may contribute in the impairment of endometrial receptivity in women with endometriosis [48]. Another serious infertility condition that requires attention is PolyCystic Ovarian Syndrome (PCOS). Qin et al. observed significant increase in the expression of lncRNA H19 in peripheral blood leukocytes in PCOS group compared to controls [103]. This is the first evidence to show an association of aberrant imprinted gene expression with PCOS, however, the methylation pattern was not assessed in the PCOS condition [103]. Taken together, large scale imprinted gene profiling is required to understand the role of genomic imprinting in the pathophysiology of male and female infertility conditions.

Pregnancy-related disorders Aberrant expression of imprinted genes leads to “shallow” placentation which is one of the contributing factors for several human gestational diseases [91]. Disturbed imprinted gene expression leads to

Genetic and environmental influences on genomic imprinting

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placental insufficiency, which in turn causes growth retardation in the developing fetus (reviewed by Ref. [32]). Studies done of intra-uterine growth restriction (IUGR) and healthy placenta samples showed upregulation of PHLDA2 and CDKN1C; and downregulation of MEST, MEG3, GNAS, PLAGL1, and IGF2 in placenta from IUGR cases [59,89]. Diplas et al. also observed upregulation of PHLDA2, NNAT and PEG10 and, downregulation of PLAGL1 and ZNF331 in IUGR human placentas [34]. In mice, fetal growth restriction due to reduced placental junctional zone and depleted glycogen stores was observed in mice with overexpression of PHLDA2 [123]. In PHLDA2-null animals, an increase in placental size was noted [45]. Preeclampsia, a hypertensive disorder of pregnancy which involves proteinuria and/or elevated liver enzymes is a manifestation of inadequate trophoblast invasion and impaired spiral artery remodeling. Mice heterozygous for Cdkn1c deletion displayed insufficient trophoblast invasion with associated features similar to preeclampsia [66]. Hypermethylation of IG-DMR in umbilical vein epithelial cells was found in preeclampsia cases which was accompanied by upregulation of DLK1 and downregulation of MEG3 [139]. In another study, the transcript and protein levels of Phlda2 were found be significantly elevated in preeclamptic placentae as compared to the control. Overexpression of PHLDA2 in the JEG-3 cells arrested them in the G0/G1 phase and inhibited cell proliferation, invasion and migration while, knockdown reversed these effects [65]. Genome-wide linkage studies in Dutch and Icelandic preeclamptic women have identified imprinted loci, 10q21.3 and 2p13.1 respectively, that are maternally expressed [4,97]. MicroRNAs belonging to the paternally expressed primate-specific miRNA cluster on chromosome 19, C19MC have been implicated in a number of gestational diseases including preeclampsia, gestational hypertension, and IUGR cases, where they had a downregulated expression pattern [57]. Their role in preeclampsia by disrupting migration and invasive capacity of trophoblasts was elucidated by another study [133]. Two miRNAs belonging to the cluster, miR-520h and miR-518b were found to have predictive value for gestational hypertension [58]. miRNAs from another imprinted cluster, the maternally expressed C14MC were also found to be differentially expressed in preeclamptic placentae in addition, to those from C19MC and other non-imprinted clusters [143]. Zhang et al. carried out miRNA microarray analysis of cytotrophoblast and found that the majority of the downregulated miRNAs were from C19MC [141]. Among them, miR-515-5p was found to be notably downregulated during syncytiotrophoblast differentiation and upregulated in preeclamptic placentae [141]. A molar pregnancy or hydatidiform mole is a complication of pregnancy involving excessive proliferation of the trophoblasts. It may manifest into a highly invasive form known as choriocarcinoma. Rahat et al. found hypomethylation of SNRPN and PEG10 DMRs and hypermethylation of MEST DMR, in addition to increased transcript levels, in molar villi [104]. The same study also profiled the expression of these genes in preeclamptic villi, wherein all three genes showed downregulation [104].

Genetic and environmental influences on genomic imprinting So far, we have discussed how epigenetic marks control the imprinting patterns and how they are established during early embryo and placental development. Lately, studies have shown that these specific characteristics make genomic imprinting a potential target for genetic and environmental insults during early developmental stages and affect the health outcomes later in adult life. Herein, we review the literature regarding the impact of genetic and environmental influences on genomic imprinting.

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Chapter 2 Genomic imprinting

Genetic influences Loss or gain of imprinting due to deletion or uniparental disomy of a gene or chromosomal region has been linked to several human genetic disorders, cancers and certain neurological conditions (reviewed by Ref. [80]). Two different types of syndromes - AS and PWS are associated with deletion at the same chromosomal position i.e. 15q11-q13 (reviewed by Ref. [80]). A cluster of imprinted genes is located at this position namely, NDN, UBE3A, IPW, SNRPN and ZNF127 (reviewed by Ref. [80]). AS is triggered due to loss of function of either the maternal allele or due to duplication of the paternal allele in a region that spans UBE3A [124]. This imprinted gene has tissue specific imprinting pattern i.e. it is biallelically expressed in various tissues but is selectively silenced at the paternal allele in cerebellar and hippocampal neurons [124]. This syndrome is characterized by mental and motor retardation, epilepsy, hypotonia, absence of speech and ataxia. Loss of methylation within the ICR of the neighboring gene SNRPN is observed in this syndrome [124]. Only a small fraction of AS individuals (5%) has defective methylation at SNRPN ICR. PWS is caused either due to loss of function of the paternal allele or due to duplication of maternal allele within the SNRPN locus (reviewed by Ref. [80]). Individuals with PWS are of short stature and, suffer from mental retardation, hypogonadotropic hypogonadism, obesity and muscular hypotonia. This syndrome is also associated in some cases with gain of methylation at ICR within SNRPN locus. It results in loss of function of imprinted genes, mainly SNRPN, IPW, MAGEL2, ZNF127 and NDN, which are expressed in the same region (reviewed by Ref. [80]). Another imprinting syndrome, called Beckwith-Wiedemann syndrome (BWS) is associated with loss of function of the maternal allele at chromosome location 11p15 where a cluster of imprinted genes, mainly KCNQ1OT1, IGF2, KCNQ1, H19 and CDKN1C resides [83]. BWS is characterized by neonatal hypoglycaemia, visceromegaly, macroglossia, umbilical and abdominal wall abnormalities, exomphalos and indentations of the ear. Since, most of the defective imprinted genes in this condition are of developmental importance, children with this disorder are susceptible to cancers [83]. Two methylation defects have been observed in BWS individuals. (i) BWS imprinting center 1 (BWSIC1) defect: Gain of methylation is observed within the H19 DMR on the maternal allele due to which the H19 allele normally expressing from the maternal side is silenced and IGF2 gene is biallelically expressed. (ii) BWS imprinting center 2 (BWSIC2) defect: Loss of methylation within the KCNQ1OT1 locus, which is a KCNQ1 antisense transcript, on the maternal side. In this case, KCNQ1OT1 is biallelically expressed, while KCNQ1 and CDKN1C are silenced. Only 5%e10% individuals develop BWSIC1 defect while 40% individuals manifest BWSIC2 defect [83]. Monozygotic twinning is also more common among BWS individuals [128]. The twins are discordant for this syndrome, with the inflicted twin exhibiting loss of methylation followed by biallelic expression within KCNQ1OT1 locus [128]. Another condition, SRS is a growth disorder characterized by dwarfism, lateral asymmetry and low birth weight. Nearly one-tenth of the individuals exhibit maternal uniparental disomy on chromosome 7, while more than 38% show loss of methylation at ICR1 on chromosome 11p15 and 1% exhibit submicroscopic chromosomal aberrations. These evidences display lack of epigenotype/genotypephenotype correlation, and needs to be elucidated [37,102]. Evidences in literature point out the involvement of defective imprinting in the progression of certain types of cancers [124]. Tumors with imprinting defects such as, Wilms’ tumor exhibit loss of imprinting at the maternal allele resulting in biallelic expression of IGF2 and repression of H19 at

Environmental influences

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chromosome position 11p15, which is in close vicinity to the region associated with BWS (reviewed by Ref. [80]). Various reports suggest that other types of cancers, mainly choriocarcinoma, sporadic osteosarcoma, neuroblastoma, hepatoblastoma and rhabdomyosarcoma, also show loss of imprinting. Several imprinted genes act as tumor suppressors such as WT1 and IGF2R (reviewed by Ref. [80]).

Environmental influences Endocrine disruptors Endocrine disruptors (EDs) are natural or synthetic compounds that affect the endocrine axis of an organism. Several studies have shown that chronic exposure to endocrine disrupting chemicals affects the gene expression without involving the actual change in the DNA sequence. However, very few studies have shown the impact of EDs on imprinted genes in germ cells. Exposure to BPA, a synthetic estrogen, during late stages of meiosis in female mice resulted in alterations in the methylation and expression of imprinted genes in placental and embryonic tissues affecting embryo, placental and post-natal development [119]. This study revealed that chronic exposure to physiological doses of BPA during critical windows of oocyte, embryo and placental development can significantly perturb the methylation and expression pattern of developmentally important imprinted genes [119]. It remains to be seen whether the loss of imprinting and alteration in expression due to BPA exposure is due to direct effect on the embryo and placenta or, epigenetic changes which might have occurred in the oocyte prior to conception (reviewed by Ref. [38]). In vitro treatment with 2,3,7,8-tetra-chlorodibenzo-p-dioxin, an environmental contaminant, on pre-implantation embryos causes aberrant methylation of IGF2 and H19 genes [132]. Upon administration of Methoxychlor (MXC), a commonly used pesticide to adult male mice or pregnant dams significantly alters the methylation status of several imprinted genes in spermatozoa, liver, tail, and skeletal muscle of fathers as well as their offspring [115]. MXC treatment to pregnant mice significantly reduced the sperm concentration and altered methylation status of all the imprinted genes in the F1 offspring. Interestingly, MXC did not affect the imprinting pattern of somatic cells indicating that the effect may have occurred during the epigenetic reprogramming in gametic development [115]. Similar effects were observed upon exposure to antiandrogen vinclozolin, a fungicide, to pregnant dams [114]. Exposure of di (2-Ethylhexyl) Phthalate (DEHP), a dioxin, to pregnant dams led to cryptorchidism in F1 and F2 generation, suggesting transgenerational effect [17]. Dnmt expression and genome-wide methylation levels of various imprinted and non-imprinted genes were significantly altered in all the generations. These results suggest that DEHP disrupts the reproductive function of the offspring, which in turn affects the DNA methylation enzyme expression, leading to changes in imprinting pattern. These effects are then transgenerationally inherited in the form of cryptorchidism [17]. Taken together, these evidences further underline that endocrine disruptors significantly affect the imprinting pattern during critical windows of development, i.e. germ cell, embryo and placental development, thereby affecting the subsequent generations. A handful of studies have shown an association between ED exposure and aberrant imprinting in humans. La Rocca et al. showed a decrease in H19 methylation levels in placenta which was associated with high level of phthalates and its metabolites in urine of pregnant women in first trimester [73]. However, the methylation or the expression of imprinted genes did not correlate with the birth weight. Another study demonstrated significant association between anti-androgenic metabolites of phthalates

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Chapter 2 Genomic imprinting

and spermatozoal DMRs of imprinted genes in men undergoing IVF treatment [131]. The methylation status of 18 imprinted loci was altered indicating that paternal ED exposure may negatively influence epigenetic reprogramming during spermatogenesis, and hence, conception [131]. These evidences strongly match the animal studies. However, more studies are required as humans could be exposed to a mixture of EDs in a lifetime. Thus, delineating each of the EDs in humans will add more value to the existing data in the literature.

Assisted reproductive technologies induced defects About 1e2% live births in developed nations are attributed to ARTs [55]. These techniques are presumed to often lead to several imprinting errors in the developing embryo. These epigenetic errors arise during ovulation induction and in vitro culture of embryos [60]. A study by White et al. found significant methylation aberrations in imprinted genes, such as, H19, SNRPN and KCNQ1OT1 in good quality embryos [129]. Majority of Day-3 embryos (76%) and 50% blastocysts showed disturbed methylation patterns [129]. Studies in sheep and cattle have shown that in vitro culture of embryos resulted in large offspring syndrome (LOS). Sheep showing this syndrome had abnormal methylation and expression of Igf2r gene [136]. Mouse models have shown that embryo cultures result in aberrant methylation and expression of imprinted genes [35,68,86]. Methylation patterns of genes are also affected by the type of culture medium used. Doherty et al. used two different culture media to study their effects on methylation of the maternally expressed imprinted gene, H19 [35]. The study found that, while Kþsupplemented simplex optimized medium (KSOM) did not produce methylation aberrations, Whitten’s medium led to loss of methylation of paternal H19 allele and hence, biallelic expression of the H19 gene. Khosla et al. showed that fetal bovine serum when used as an additive to M16 medium affected growth of embryos and altered the methylation and expression of several imprinted genes such as, H19, Igf2, Grb7 and Grb10 [68]. Mouse embryos cultured in human tubal fluid which is a commonly used human embryo culture medium exhibited impaired imprinting at the H19 locus [77]. Thus, use of inappropriate culture conditions can have deleterious consequences for embryonic growth and development. Epigenetic disorders such as, BWS, AS and SRS are reported to occur with increased incidence due to ARTs [39,55]. The chances of premature birth and IUGR also get increased in pregnancies conceived by ARTs [98,110]. This may be partially attributed to epigenetic disturbances in the placenta whose effects may get reversed upon removal of placenta post-delivery, in the form of “postnatal catch-up growth” [130]. It is also possible that the imprinting error is inherent in the gametes and could be a cause of infertility, which is unmasked with ART. Thus, ART though presumed safe does pose substantial threat to the epigenetic machinery of the developing fetus.

Concluding remarks To summarize, genomic imprinting is a unique phenomenon in mammals important during fetal and placental development. The imprinting mark is established in the germ cells and maintained in the somatic and placental cells. Imprinting control region acts as core domain for most of the imprinted genes and regulates the expression of imprinted genes either from the paternal or maternal allele.

Concluding remarks

39

Imprinted genes play a crucial role in fetal growth and, placental development and post-natal development. Aberrant imprinting pattern due to errors in erasure, establishment and maintenance can lead to a variety of conditions like infertility, cancers, and neurological, metabolic, placental and ARTinduced defects (Fig. 2.2). Therefore, it is critical to study the mechanisms of establishment and maintenance of genomic imprinting in mammals. An association between ART and epigenetic defects is supported by experimental studies such as mouse embryo cultures, large offspring syndrome in ruminants and cases of BWS and AS children born by ARTs. In humans, conclusive demonstration of such an association and identification of its causes is yet to be done and can only be accomplished by large-scale and long-term follow-up studies. Such a study will help in determining the increased risk of ARTs on cancer and neurodevelopmental problems. Furthermore, studies are also needed to determine that “epigenetic defects are a cause of infertility, rather than a consequence of ART”. Such insights would help in diagnosis and treatment of underlying cause of infertility, thereby reducing the cost and trauma of undergoing ART procedures. It would also help in screening of patients who could be benefited by ARTs.

FIG. 2.2 Schematic representation of the impact of genetic or environmental insults (red arrows) on imprints during germ cells, embryo and placental development. Aberrant imprinting is transgenerationally inherited by the offspring as well as the subsequent generations thus causing serious complications in adult life.

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Chapter 2 Genomic imprinting

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CHAPTER

Chromatin remodeling of the male genome during spermiogenesis and embryo development

3

Isha Singh
, Aniket G. Patankar
, Priyanka Parte Department of Gamete Immunobiology, ICMR-National Institute for Research in Reproductive Health, Mumbai, Maharashtra, India

Introduction Spermatozoon (sperm) is a unique cell in context of its genetic and epigenetic organization. During the later stages of sperm formation, i.e. spermiogenesis, haploid male genome gets tightly packed before venturing into its journey to its final destination ethe oocyte. Chromatin compaction is a gradual process in which nucleosomal structure present at round spermatid stage undergoes remodeling such that somatic histones first get replaced by testis specific histone variants then transition proteins and finally with protamines. In round spermatids, DNA is wrapped around the nucleosome units which are histone octamers composed of dimer of each histone H2A, H2B, H3, and H4. Histone H1 binds to the DNA in between two nucleosomes and is thought to be involved in higher-order chromatin structure formation. Histone of nucleosome core particle (NCP) are divided into two parts: structured region made up of core and C-terminal region of histone, and N-terminal unstructured tail which protrudes out of the nucleosome and interacts with DNA. Approximately 146 bp DNA is wrapped around each nucleosome. The N terminal of histones is highly susceptible to undergo multiple Post Translational Modifications (PTMs) predominantly acetylation, methylation, phosphorylation, crotonylation and ubiquitination. These modifications influence chromatin architecture either directly by adding negative or positive charge leading to altered histone-DNA interaction, or indirectly by recruiting modification- specific chromatin remodeling factors [71]. The site of modification matters too. Histone H3 trimethylation at lysine 4 is found to be associated with transcriptionally active genome and conversely its trimethylation at lysine 27 leads to a repressive chromatin state. Nonetheless both modifications can be observed together in a bivalent promoter.




Equal contribution.

Epigenetics and Reproductive Health. https://doi.org/10.1016/B978-0-12-819753-0.00003-9 Copyright © 2021 Elsevier Inc. All rights reserved.

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Chapter 3 Epigenetic responsibilities of the father

Nuclear remodeling in sperm Eviction of somatic histones and accompanying events Hyperacetylation of histones During spermatogenesis, on completion of second meiotic division, three major events are initiated genome-wide: (A) histone hyperacetylation which essentially triggers the initiation of nuclear remodeling [24], (B) Transcription of several sperm specific genes, (C) Formation of sex body. In mouse, all four core histone species undergo acetylation in the spermatids between step 9 to 11 and gradually disappear as transition proteins (TPs) replace histones [57,95]. As evident from studies of different knockout genes affecting histone hyperacetylation, this process is indispensable for male fertility. In vitro studies have indicated that acetylation of histones is solely sufficient to replace them with protamines. However in vivo scenario indicates that multiple players are involved. The most explored chromatin remodeling factor highly expressed in meiotic male germ cell is BRDT. BRDT, a testis-specific protein containing two bromodomains, binds specifically to acetylated histone H4. Loss of the Bromodomain 1 of BRDT impairs spermiogenesis and absence of BRDT function blocks the entry of spermatocytes into the first meiotic division. In premeiotic and meiotic cells BRDT drives expression of specific spermatogenic genes and later controls the chromatin compaction of haploid male genome [23]. In a more recent study, BRDT has been shown to be involved in control of the global chromatin organization and histone modifications of the chromatin attached to the synaptonemal complex [52]. In male germ cells, acetylation appears to be more dynamic histone post translational modification which is finely tuned by stage specific action of histone acetyltransferases and deacetylases enzymes. This is evident from the TSA treated spermatogenic cells which show a drastic increase of histone acetylation in round spermatids suggesting that these deacetylases kept the histones in their deacetylated state before chromatin remodeling initiated. However, the same treatment failed to increase acetylation in condensing spermatids, suggesting that in these cells histones are still acetylated and they were not deacetylated before their eviction [28]. There are few knockout studies which support the importance of deacetylases in sperm function like SirT1, a NAD-dependent deacetylase sirtuin-1, knockout of which leads to lower H4 acetylation and abnormal histone to protamine transition. Dong et al. observed two critical components of the mammalian nucleosome acetyltransferase of H4 (NuA4) complexes- EPC1 and TIP60, to be co-expressed during meiotic phase of male germ cell formation. Elimination of either Epc1 or Tip60 led to aberrant spermatid development as a result of impaired histone hyperacetylation and abnormal histone retention [15]. Study by Hitoshi Shiota and Co-workers showed that Nuclear protein in testis (Nut) with the involvement of p300 and/or CBP enhances acetylation of H4 at both lysine 5 and lysine 8, thereby providing binding sites for the first bromodomain of Brdt. Knockout of Nut induces male sterility with spermatogenesis arrested at the histone-removal stage [91].

DNA strand breaks Another important event necessary for eviction of histones is action of an ATP-dependent enzyme that can cause DNA double strand break - Topoisomerase II (Top2), which changes DNA topography by removing knots and forming loops [55,103]. In elongating spermatid, H4 hyperacetylation dependent activity of top2b was found to be involved in both creating DNA strand breaks, and regulating transcription [42,45]. Meyer-Ficca et al. reported the presence of poly ADP-ribose (PAR) polymerases

Nuclear remodeling in sperm

49

PARP1 and PARP2 at steps 11 to 14 of rat spermiogenesis [58]. In their subsequent study they reported that DNA strand breaks trigger the activation of PARP1- a polymerase involved in DNA repair. Inhibition of PARP1 activation caused abnormal chromatin integrity with abnormal histone retention in mature sperm. These animals also showed impaired embryonic survival [58]. The hallmark for presence of DNA double stranded break is phosphorylated histone H2AX at Ser139 (gH2AX). It was observed that absence of testis specific kinase, TSSK6, blocks gH2AX formation. Sperm of tssk6-knockout mice showed increased histone retention (H3 and H4) and protamine 2 precursors. H2AX null mice were sterile and showed complete absence of the macrochromatin sex body, a global failure to inactivate gonosomal chromatin and defects in XeY synapsis. In another study by Jiang et al., histone acetyltransferase (HAT) MOF was found to be involved in H2AX phosphorylation and germ cell-specific deletion of Mof in spermatocytes caused global loss of H4K16ac [35].

Transcriptional surge It is generally accepted that sperm are transcriptionally and translationally inactive except for few isolated reports on translation of nuclear genes by mitochondrial ribosomes. It is thus mandatory for sperm to be prepared for its journey in female reproductive track before the transcription ceases. Though sperm acquires some proteins during epididymal maturation, the main gene expression event occurs at the round spermatid stage, post meiotic division. cAMP-responsive element modulator (CREM) has been reported to be master regulator of this transcriptional surge as Crem deficient mice showed declined transcription in post meiotic phases of sperm development and spermatid failed to differentiate into sperm [8,61]. Investigators have also observed essential role of proteins regulating CREM function. ACT (activator of CREM in testis) is a protein which interacts with CREM in post-meiotic stages of male germ cell development and help in enhancement of CREM-dependent transcription. The intracellular localization of ACT was found to be dependent on ACT-KIF17b interaction and is restricted to specific stages of spermatogenesis [51]. Interestingly, act knockout mice although showed reduced number of sperm in epididymis, they were fertile and all germ cell developmental stages were observed normal indicating that role of ACT is either bypassed or compensated by other protein. However, the investigators also observed involvement of ACT in expression of genes encoding structural components of the mature sperm. Another protein named CARM1 - a protein arginine methyltransferase (PRMT) has been firmly implicated in transcriptional regulation, negatively regulating the activity of CREM through methylation of p300 and restricting its binding to CREM activator, ACT. Coactivator p300 has both crotonyltransferase and acetyltransferase activities however p300 induced histone crotonylation directly enhances transcription at a greater level than histone acetylation. In vivo studies have indicated that p300 can also acetylate TP2 in its highly basic C-terminal domain, leading to significant reduction in its DNA condensation property as analyzed by circular dichroism and atomic force microscopy. Apart from CREM, few others proteins have also been investigated for their involvement in transcription regulation in spermatid. ChIP-Seq and RNA-Seq analysis using infertile Regulatory Factor X2 (RFX2) mouse model, showing complete arrest in spermatid development just prior to spermatid elongation, revealed that 139 genes were directly controlled by RFX2 [113]. Knockout of exclusive post meiotic gene on Y chromosome, Sly, indicated deregulation of large numbers of sex chromosome-encoded genes such as H2A variants and H3K79 methyltransferase DOT1L. In post-meiotic male germ cells SLY acts via its interaction with TBL1XR1 which is a member of SMRT/N-CoR repressor complex involved in gene regulation during sperm differentiation. Sly/

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mice show multiple sperm differentiation defects and are infertile [59]. Recently it has been shown that SOX30, a transcription factor of SOX family controls post meiotic gene expression. In round spermatid SOX30 regulates expression of post meiotic genes involved in sperm formation, maturation and capacitation [4]. TFIID is a general transcription factor required for transcription of most protein-coding genes by RNA polymerase II. Absence of TAF7L which is X-linked germ cell-specific paralogue of TAF7 and component of TFIID leads to downregulation of six gene transcripts (including Fscn1) and mice showed reduced sperm count and motility [114]. Earlier studies with human CDY and mouse CDYL proteins exhibited their in vitro histone acetyltransferase activity, especially on histone H4; Cdyl also was thought to be involved in histone to protamine transition via H4 hyperacetylation, as its expression and localization coincides with H4 hyperacetylation during spermatogenesis [43]. However, recent study demonstrated that Cdyl regulates the expression of sex chromosome-linked escaped genes in postmeiotic spermatogenic cells by acting as a crotonyl-CoA hydratase to convert crotonyl-CoA to b-hydroxybutyryl-CoA. Lack of cdyl leads to dysregulated histone replacement in the testis of Cdyl transgenic mice, however authors suggested this is not because of altered expression of transition proteins or protamines [48]. In a cell-based model of transcriptional activation, increasing or decreasing the cellular concentration of crotonyl-CoA leads to enhanced or diminished gene expression, respectively, and these crotonylated histones were present in regulatory region of active genes which again suggest involvement of histone crotonylation in active transcription [79]. In a study with BRWD1, the dual bromodomain-containing protein, investigators observed that BRWD1 promotes haploid spermatid-specific transcription and its absence leads to down-regulation of w300 mostly spermatid-specific transcripts in testis with protamines and transition protein transcripts being completely absent. At a global level however minor epigenetic changes in chromatin were observed suggestive of its selectivity toward some genomic loci. Brwd1 was also observed to be essential in female reproduction as its deletion caused severe chromosome condensation and structural defects associated with abnormal telomere structure. Gene expression changes at the germinal vesicle stage however were not that significant as in male counterpart [67]. Pygopus 2 (Pygo2) belongs to a family of evolutionarily conserved PHD finger proteins and thought to act as co-activators of Wnt signaling effector complexes composed of beta-catenin and LEF/TCF transcription factor. Pygo2 mutants show reduced levels of few postmeiotic genes including protamines, transition protein 2 and H1fnt along with altered pattern of histone H3 hyperacetylation [60]. Loss-of-function approach demonstrated that the mouse H3K9me2/ 1-specific demethylase JHDM2A (JmjC-domain-containing histone demethylase 2A, also known as JMJD1A) directly binds to and controls the expression of transition nuclear protein 1 (Tnp1) and protamine 1 (Prm1) gene [63]. Deficiency of SET domain-containing 2 (SETD2), the predominant histone methyltransferase which catalyzes H3K36me3, leads to complete loss of H3K36me3 and significantly decreases expression of thousands of genes, including acrosin-binding protein 1 (Acrbp1) and protamines in male germ cell [111]. Testis specific overexpression of the histone H3 lysine 4 (H3K4) demethylase KDM1A (also known as LSD1) severely impaired development and survivability of offspring. These defects persisted transgenerationally in the absence of KDM1A germline expression and were associated with altered RNA profiles in sperm and offspring. This altogether suggest that epigenetic inheritance of aberrant development can be initiated by histone demethylase activity in developing sperm, irrespective of changes in DNA methylation at CpG-rich regions [92]. Lu et al. found that ubiquitin ligase RNF8-dependent histone ubiquitination may induce H4K16 acetylation, which is an initial step in nucleosome removal. RNF8-dependent modifications also include trimethylation of H3K4, histone lysine crotonylation (Kcr), and incorporation of the histone variant H2AFZ. They observed RNF8-deficient mice could undergo meiotic sex chromosome

Nuclear remodeling in sperm

51

inactivation (MSCI) but were not able to remove nucleosome genome wide [50]. RNF8-dependent ubiquitination of histone H2A during meiosis establishes active epigenetic modifications, including dimethylation of H3K4 on the sex chromosomes.

Incorporation of testis specific histone variants Hyperacetylation of histones leads to eviction of somatic histones and incorporation of testis-specific histone variants. In silico and in vitro analysis of nucleosome containing testis specific histone variant showed fewer histone-DNA contacts suggestive of an open chromatin structure, characteristic of transcriptionally active genome [65]. Excepting histone H4, testis variants have been reported till date for all core histones H2A, H2B, H3, and linker histone H1. Testis specific histone variants and their knockout phenotypes have been reviewed elsewhere [66]. Van der Heijden et al. [31] observed exclusive incorporation of the H3.3 variant post somatic nucleosomal eviction to be generally associated with transcriptional activity. They also observed that nucleosomal exchange leads to selective acquisition of specific histone marks. Proteasomes containing the activator PA200 catalyze the polyubiquitin-independent degradation of histones. Deletion of PA200 in mice abolishes acetylationdependent degradation of somatic core histones during DNA double-strand breaks and delays core histone disappearance in elongated spermatids [73]. CTCF regulates chromatin compaction necessary for packaging of the paternal genome into mature sperm [33]. Chromodomain helicase DNA-binding protein 5 (Chd5) has a key role in DNA compaction. It is involved in H4 hyperacetylation, histone variant expression, and removal and replacement of the histones with nucleoprotamines. Chd5 deficiency in mice leads to defective sperm chromatin compaction and infertility [46]. Low expression of Chd5 has also been observed in the testis of infertile men by the same group. In elongating spermatids of Chd5 KO mice, histone H4 hyperacetylation was greatly decreased and histones are retained in sperm [110].

Histone replacement by protamines During spermiogenesis, testis-specific histones get replaced by transition proteins (TP). Mammals, including mouse, rat, human, ram, and boar, predominantly have two types of transition proteins, viz., transition protein 1 (TP1) and transition protein 2 (TP2) [96]. Both TP1 and TP2 are encoded by single- copy genes, Tnp1 and Tnp2, respectively [76]. TP1 is a 6200 Da protein and contains about 20% arginine and 20% lysine, distributed uniformly, and no cysteine [39]. TP2 is a 13,000 Da protein with about 10% arginine, 10% lysine, and 5% cysteine [9]. It has a highly basic C-terminal domain and an N-terminal domain that forms zinc fingers [56]. Immediately after its synthesis, Transition protein 2 (TP2), has been shown to undergo phosphorylation catalyzed by sperm-specific isoform of protein kinase A (Cs-PKA). Phosphorylation was found necessary for its NLS dependent nuclear transport. TP1 is abundantly expressed and shows high sequence similarity across various mammalian species as compared to TP2 [29,41]. TP1/ and TP2/ knockout mice have been shown to be less fertile than normal mice and show abnormal chromatin condensation [109]. TP1 and TP2 double knockout mice are sterile, and spermatogenesis is severely impaired suggesting their important role in spermiogenesis [108]. In case of mammals, protamines are of two types protamine 1 (P1) and protamine 2 (P2). The presence of P1 in association with sperm DNA can be observed in nearly all vertebrates, whereas P2 is present only in primates, many rodents, and a subset of other placental mammals [6]. The number of

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protamine genes and copies present per haploid genome varies from species to species. PRM1 and PRM2 are first detected during mouse spermiogenesis at step 12 and step 14, respectively, and thereafter remain in mature sperm [5]. Protamines are arginine-rich, small, basic, major nucleoproteins in sperm. They are synthesized in late-stage spermatid. Around 80%e85% sperm DNA is compact due to protamination. Mammals have single-copy genes of P1 and P2, located on chromosome 16 [77]. P1and P2 are products of gene Prm1 and Prm2, respectively. The precursor protein of Prm2 undergoes proteolytic processing at its N-terminus to give rise to p2, p3, and p4 which differ in 3e4 residues at Nterminus. The arginine-rich DNA-anchoring domains by which protamines bind with the negatively charged DNA and the multiple serine and threonine residues that can be used as phosphorylation sites form the structural elements of protamines [6]. The cysteine residues allow disulfide bond formation and thus link two adjacent protamines, which leads to further compaction of DNA. Two subunits of CAF1, a canonical histone H3/H4 chaperone that mediates replication-coupled nucleosome assembly, CAF1-p180 and CAF1-p75 are reported to have different functions during spermatogenesis in which CAF1-p75 act as a protamine-loading factor [115]. In Drosophila however, ProtA and ProtB were found to be replaceable and Mst77F which is spermatid-specific histone H1-like protein was essential for male fertility. In these flies their role was thought to be involved only in paternal DNA damage response [75]. Sperm lacking Prm1 can generate offspring despite being abnormally shaped and having destabilized DNA, decondensed chromatin and a reduction in mitochondrial membrane potential [99]. Though the Prm2 heterozygous male mice are fertile with sperm displaying normal head morphology and motility, in Prm2-deficient sperm, however, DNA-hypercondensation is reported and acrosome formation is severely impaired [86]. Thus, by the time sperm are ready for their voyage into the female reproductive tract they have heterochromatin with majority of sperm DNA being protamine bound and coiled into toroids, and a fairly smaller amount of paternal DNA histone bound, approximately 4%e10% in humans and 1% in mouse [10,26]. These retained histones are positioned on specific sequences in the sperm and are associated with genes involved in early embryogenesis (HOX cluster) and regulatory functions [3,22,26]. Several researchers have investigated nucleosomal positioning of the genes in sperm using various approaches and have arrived at differing conclusions (Table 3.1).

Table 3.1 Nucleosomal positioning in the sperm and accessibility to gene regulatory elements. Source of spermatozoa Healthy human donors

Fertile men

Methods used Southern hybridization, estimation of percentage of telomeric DNA released Comparative genome hybridization

Nuclesomal positioning and inference drawn telomeric DNA is · the micrococcal nuclease sensitive

·

and hence is associated with nucleosomes gene regulatory regions, like promoter sequences and sequences recognized by CCCTC-binding factor (CTCF).

References [107]

[3]

53

Nuclear remodeling in sperm

Table 3.1 Nucleosomal positioning in the sperm and accessibility to gene regulatory elements.dcont’d Source of spermatozoa Proven fertile men

Methods used MNase digestion followed by ChIP (for modified nucleosomes)/ mononucleosomal purified DNA - high-throughput sequencing, high-density promoter-tiling array/MeDIP and bisulphite sequencing for DNA methylation

Nuclesomal positioning and inference drawn

· 4% nucleosomal retention in sperm of fertile men at loci important for · Retention embryo development (HOX)

· · · · · ·

Spermatozoa from normospermic fertile men; Spermatozoa from caudal epididymis of CD1 and C57BL/6J mice.

Cross-linking done; ChIP for H3K4me2 and H3K27me3; ChIP-chip

· · ·

· ·

including embryonic transcription factors and signaling pathway components; at the promoters of miRNAs and at imprinted genes TH2B associated with 0.3% of gene promoters H2A.Z nucleosomes poor in GO categories H3K4me2 and H3K27me3 enriched significantly at promoters for developmental transcription factors; H3K9me3 high at pericentric regions developmental promoters hypomethylated in sperm H3K4me3 mark at paternally expressed genes and non-coding RNAs Detected w10% of histone retention in human sperm and w1% in mouse sperm Genome-wide nucleosomal distribution with a slight enrichment at regulatory region H3K4me2- marked genes involved inspermatogenic and housekeeping functions (e.g., PRM1, PGK2, BRDT RPS3, SFRS6, DICER and PRMT5 and TSH2B) H3K27me3- marked genes involved in developmental functions (SOX2, CDX2, GATA6 and BMP4, BRACHYURY (T) and HOX) H3K4me2 marked promoters were significantly hypomethylated; Evolutionary conservation of H3K4me2 as well as the H3K27me3 marks between mouse and human spermatozoa.

References [26]

[10]

Continued

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Chapter 3 Epigenetic responsibilities of the father

Table 3.1 Nucleosomal positioning in the sperm and accessibility to gene regulatory elements.dcont’d Source of spermatozoa

Methods used

Proven fertile men

Analysis of data from [26]

Nuclesomal positioning and inference drawn

· base composition determines ·

Mouse sperm (C57BL/ 6J mice)

bisulfite conversion and highthroughput sequencing of sperm DNA associated with nucleosomes

· · ·

· E14 ESC lines; 10 week old C57/6J mice

MNase digestion followed by sperm ChIP-Seq; Fluorescence in situ hybridization (FISH)

· · ·

nucleosome retention in human sperm in both gene-rich and gene-poor regions The retention of nucleosomes at GC-rich sequences especially at transcription start sites. Genome-wide nucleosomal overrepresentation at promoter regions and exons, underrepresentation at introns and repeat regions CpG dense regions associated with nucleosome in sperm; methylation inversely correlated with nucleosomes retention in sperm Paternal ICRs of maternally expressed genes such as H19, Dlk1, Meg3 are methylated and without nucleosome whereas ICRs regulating maternally imprinted genes like Kcnq1, Snrpn and Peg10 are unmethylated and have nucleosomes. histone variant H3.3 enriched over H3$1/H3.2 proteins protein in CpG-island (CGIs) only extensive MNase digestion produces a minor portion of nucleosome sized fragments that aligned with CpG- rich promoters histone retention specially over gene deserts histone localization at chromocentre that is occupied by repeat elements rather than the CpG- rich fraction of developmental promoters (fluorescence microscopy).

References [102]

[17]

[11]

Nuclear remodeling in sperm

55

Table 3.1 Nucleosomal positioning in the sperm and accessibility to gene regulatory elements.dcont’d Source of spermatozoa Fertile bulls of holstein-Friesian breed; Fertile men

Methods used MNase digestion followed by sperm ChIP-Seq

Nuclesomal positioning and inference drawn

· 2.9% of human and 13.45% of

bovine sperm genome is associated with nucleosomes significant nucleosome enrichment in distal intergenic and intron regions, repetitive DNA elements-like centromere repeats, LINE1, and SINE Absence of putative nucleosome binding site in the HOX cluster The enrichment reported at repetitive elements results from redundant use of reads with multiple mapping positions in the genome enrichment of H3K4me3 at TSSs of developmental genes with CpG rich promoters H4 localization majorly to intergenic regions, whereas modified histones enriched at specific genomic elements (H3K4me3 at CpG-rich promoters; H3K9me3 in satellite repeats) histone association with gene promoter regions, smaller number of H3-binding genes in HRCS than in the swim-up sperm histone H3 in gene-rich regions inconsistent with the results reported by Refs. [11,82].The reason for this discrepancy is unknown. The H3 associated genes of HRCS, showed enrichment of genes involved in neural cell differentiation Nucleosomes form Nucleosomeal Domains (NDs) in human sperm that are specifically localized proximal to the Post Acrosomal Region (PAR) Protamine 1 in the core chromatin Infertile samples showed no NDs

References [82]

· ·

Fertile bulls; fertile men

Reanalysis of [82] data

Mouse sperm from 10to 15-week-old ICR mice;

nucleoplasmin (NPM) treatment: sperm chromatin crosslinking; ChIP-seq

11e12 weeks C57BL/6 male mice

Crosslinking ChIP (X-ChIP) of histone-to-protamine replacement-completed sperm (HRCS); No MNase digestion

·

[78]

[104]

·

· ·

[105]

· ·

Proven fertile men; men with known sterility

3D localization of nucleosomes in sperm by structured illumination microscopy (SIM)

· · ·

[112]

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Chapter 3 Epigenetic responsibilities of the father

From being sperm nucleus to paternal pronucleus Genome-wide reprogramming at fertilization Zygotic Genome Activation At fertilization both the gametes are transcriptionally quiescent with the paternal chromatin highly condensed due to the presence of protamines. Thus, although sperm delivers RNA to the oocyte which are translated by the oocytes’s translation machinery [18], the de-condensation of male nuclear chromatin and organization of the gametes is critical for initiation of zygotic transcription and replication. Initiation of zygotic transcription, as signaled by RNA polymerase II mediated incorporation of BrUTP first occurs in the paternal pronucleus at the late one-cell embryo in mice [2]. This is followed by major burst of transcription occurring in the mid-to-late two-cell stage, which is termed as major Zygotic Genome Activation (ZGA) [19]. Oocyte takes up the intricate task of remodeling the highly condensed sperm chromatin to accessible and transcriptionally active chromatin of male pronucleus [54]. On its entry in the egg, the paternal chromatin relaxes with the help of oocyte’s endogenous molecules. The decondensation of sperm nucleus is assured by the tripeptide glutathione (y-glutamyl-cysteinyl-glycine; GSH), a reducing agent present in mammalian oocytes. Glutathione reduces the intra-/inter-disulphide bonds between protamine residues. Post disulfide bond disruption, the sperm nucleus protamines are substituted by the maternal histones aided by nucleoplasmin [68,69]. Nucleoplasmin, an acidic nuclear protein, found in oocyte of Xenopus laevis and mice is acknowledged to decondense the sperm chromatin [70]. It is a pentamer with a patch of negatively charged amino acid residues containing a polyglutamic acid tract in the C-terminal of each of its five subunits. It is proposed that nucleoplasmin bind histones with these negatively charged regions acting together [14,16]. Therefore, nucleoplasmin removes protamines and transfers maternal histones (H1FOO, H3.3, TH2A/TH2B) to DNA and helps in nucleosomal assemblage in the male pronucleus [44]. Loss of HIRA, a maternal histone chaperone that helps in assembling H3.3 on paternal genome post fertilization leads to failure of nucleosome assembly on male pronucleus [49]. This ultimately leads to developmental arrest. Shinagawa et al., have demonstrated the incorporation of maternal TH2A/B in the male pronucleus, failure to do so leads to decrease in the litter size [90]. The transition from nucleoprotamine to nucleohistone architecture generates a somatic chromatin state which is conducive to ZGA. The association of histone with the 3- and g-globin genes, which are expressed early in embryonic yolk sac, points that the presence of histones leads to formation of a permissive chromatin structure for 3- and g-globin transcription during early development and may mark these genes for early expression in the embryo [21]. Also, it has been shown that sperm derived histones contribute to paternal chromatin in zygote [32]. These findings raise the likelihood that specific nucleosomal positioning at specific loci might influence zygotic gene expression, probably by conserving histone modifications at these genes or regions [26,30,37,78,82,100]. Differential experimental practices might lead to some variations in loci thought to be associated with these retained histones [80]. None the less these studies and the observations of altered histone retention in infertile men compared to fertile controls [27] jointly suggest that paternal chromatin may effect embryonic gene activation.

Is sperm merely a vector for the paternal genome?

57

Methylation - demethylation Genome-wide reprogramming at fertilization also includes global DNA demethylation of both the mammalian gametes. Upon fertilization, the terminally differentiated paternal genomes transit from a hypermethylated to a hypomethylated state. Firstly, the paternal pronucleus undergoes active de-methylation independent of replication, whereas, the maternal pronucleus experiences a step-wise replication-dependent passive demethylation [53,83]. This global demethylation erases all the methylation mark till the blastocyst stage with the exception of parental imprints whose methylation marks are preserved. After ZGA, followed by implantation and initiation of lineage separation, the embryonic inner cell mass (ICM) derived cells genome becomes hypermethylated, whereas the genome of trophectodermal (TE), extra-embryonic cells, remains hypomethylated.

Is sperm merely a vector for the paternal genome? Even after the discovery and acclaimed establishment of the presence of spermatozoal RNA its functional significance to fertilization and early embryonic development remained vague. Using zona free hamster oocytes and human sperm penetration assay it was established that sperm contributes sperm specific RNA to the oocyte at fertilization. qRT-PCR showed presence of protamine-2 and clusterin transcripts in spermatozoa, and zygotes but not in hamster oocytes thus establishing delivery of RNA from the paternal gamete to the maternal gamete [64]. This work highlights the importance of sperm RNA in zygotic and embryonic development. Furthermore, one can conclude that one of the hidden reasons behind idiopathic male infertility and the limited success of parthenogenesis and SCNT techniques might be sperm RNA. Rassoulzadegan et al., studies on the Kit gene in mice depicted epigenetic (non-Mendelian) inheritance via transmission of RNA among gametes and zygotes for a paramutation (heritable epigenetic modification)-like phenomenon. In wild type mice, kit is transcribed in spermatogonia and not in spermatids and sperm. They did a transgene (lacZ) insertion at the Kit locus that led to a whitetail phenotype due to no KIT protein. However mating heterozygotes for mutant Kit allele led to the lesser than expected Mendelian ratio of phenotypically wild-type mice. Subsequent analysis showed that generation of wild-type progenies was indeed in the expected Mendelian ratio, although most had sustained the mutant white tail tip phenotype. Notably, this mutant phenotype (KitMut) can be inherited by succeeding generation from the wild-type mice [74]. The proposed reason for this was higher RNA levels in the sperm of mice heterozygotes and in paramutated males. Additional study by the same group presented that microinjection of miR-124 RNA in the male pronucleus of one-cell embryo leads to overgrowth in body size. This giant’ phenotype is maintained postnatally and is subsequently inherited over numerous generations. The enhanced growth rate was in fact established even at morula to blastocyst stage. Further analysis revealed Sox9 among upregulated transcripts in the variant embryos. This high mobility group (HMG)-box transcription factor is involved in many terminal differentiation processes [72,101]. ChIP analysis revealed changes in the epigenetic state as increased H3K9me2 and H3K9me3 at a regulatory element in the upstream region of the Sox9 promoter was observed. Role of Sox9 in embryonic growth was established by the

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observation of overgrown embryos expressing a Sox9 DNA transgene. Increased Sox9 expression in the paramutants was not linked to an alteration in miR-124 expression, but to the creation of a distinct heritable epigenetic modified structure in the promoter region of Sox9 [25]. These studies clearly point out that the paternal gamete is not merely a vector to transfer the haploid genome. Such studies prompted the need to evaluate sperm’s epigenetic potential in regulating embryo development, especially the role of sperm-borne non-coding RNAs.

The “not-so-small” importance of sperm small RNA population Gaining from the high throughput sequencing data from independent laboratories, the small RNA population in mature murine sperm comprises of tRNA-derived small RNAs (tsRNAs), microRNAs (miRNAs), Piwi-interacting RNAs (piRNA), and rRNA-derived small RNAs (rsRNAs) [12,20,74,88]. Sperm small RNAs composition changes dynamically during the epididymal maturation with tsRNA fractions increasing over piRNA fraction. Additionally, tissue-specific metabolic labeling of RNAs showed RNA transfer from the epididymis to maturing sperm, during the caput to caudal transit, via epididymosomes. This data demonstrates soma to germ-line transfer of RNA [89].

Sperm miRNAs MicroRNAs (miRNAs) represents endogenous 21e22-nucleotides (nt) noncoding small RNAs. They are generated by the processing of precursor miRNAs (pre-miRs) by 2 endoribonucleases Drosha and Dicer into mature miRNAs after a series of processing steps. They are “multivalent” i.e. they regulate a number of genes post transcriptionally. Numerous studies have revealed the significance of spermborne RNA. Intracytoplasmic sperm injection (ICSI) of sperm having altered miRNA and endo-siRNA from a Germline specific Dicer and Drosha conditional knockout (cKO) mice in wild-type (WT) eggs, displayed significant reduction in pre-implantation developmental potential of the embryos, which could be overcome by supplementing WT sperm-derived total or small RNAs into ICSI embryos [106]. Thus, highlighting the importance of sperm borne miRNA and endo-siRNA in regulating the preimplantation embryo development. The first cleavage division in mouse zygote requires microRNA-34c which targets Bcl-2. Zygotic miR-34c is sperm-borne as shown by: (i) similar levels of miR-34c in the zona-bound sperm and zygotes but not in the oocytes; (ii) presence of both the precursor and mature forms of miR-34c in sperm; and (iii) a-amanitin treated oocytes did not show reduced zygotic miR-34c. The deduction was more reinforced by the absence of miR-34c in ethanol-activated parthenogenetic oocytes. Further, Western blotting showed that miR-34c modulates B-cell leukemia/lymphoma 2 (Bcl-2) expression in the zygote [47]. Upon fertilization, sperm-borne miR-34c targets Bcl-2 and p27 and thus promotes entry in S-phase and first cleavage. Injecting recombinant Bcl-2 protein or miR-34c inhibitor before S-phase prevents first cleavage. A very recent study in bovine model by Alves et al., highlights paternal contribution of spermborne miR-216b to pre-implantation embryo development. They spotted differences in relative expression levels of 9 out of 380 sperm miRNAs in frozenethawed semen from high (HF) and low

Father’s extra-genomic contribution to the zygote

59

fertility (LF) bulls. miR-216b was relatively lower in HF bull sperm cells. However, the higher comparative level of its target gene, K-RAS in HF two-cell embryos, leads to an increased first cleavage rate and blastocyst cell number. K-RAS is related to cell proliferation, whereas, studies of cancer tumors delineate the role of miR-216b as inhibiting cell growth and proliferation. Thus, HF embryos showed increased proliferation. Additionally, the group reported a higher level of miR-216b in polyspermy zygotes thus indicating that sperm distributes miR-216b to zygotes [1]. Studies by Salas-Huetos evaluated 736 miRNAs and identified 57 miRNAs differentially expressed in sperm of normozoospermic infertile individuals with normal semen parameters compared to normozoospermic proven fertile individuals with normal seminal characteristics. Using in silico analysis, these miRNAs were predicted to modulate the expression of 8606 target genes. These predicted 8606 targets were significantly enriched in biological processes linked to embryonic morphogenesis and chromatin modification suggesting the epigenetic influence of sperm miRNAs on sperm function and embryo development [81].

Sperm non-coding RNAs The possible role of RNAs as epigenetic elements was maintained by experiments showing inheritance of epigenetic traits in mice mediated by RNA. Deep sequencing technology to evaluate murine sperm RNA generated RNA sequences including mRNAs, rRNAs, piRNAs, and miRNAs. Two novel small noncoding mature sperm RNAs identified as sperm RNAs (spR) 12 and 13 were studied. These were encoded from a 40 kb piRNA cluster on chromosome 17, but they differ in length (20e21 nt) as well as sequences as compared to known typical micro- and pi-RNAs. Their resistance to periodateoxidation-mediated reaction, points that they must be undergoing terminal post-transcriptional modification. They were detected by quantitative RT-PCR in the gametes as well as in very early developmental stages [38].

Father’s extra-genomic contribution to the zygote Mammals have bisexual mode of reproduction. At fertilization, the paternal gamete contributes not only its imprinted haploid genome but a number of sperm specific molecular signatures which are essential for embryo development. The molecular gifts of sperm at fertilization are still unfolding. The ejaculated sperm undergoes capacitation, binds the egg’s zona pellucida, and undergoes acrosome reaction. Spermeegg fusion is mediated by interaction of sperm’s Izumo1 with the corresponding egg receptor Juno, a GPI-anchored Folate receptor 4 (Folr4) present on the egg surface that is vital for female fertility [7]. Fusion of the plasma membrane occurs at the equatorial segment of the sperm head, and the zona pellucida of the egg. The point of sperm entrance states the embryonice abembryonic axis. Fusion of the sperm leads to the “oocyte activation” by the release of sperm-specific phospholipase C, PLCz in the oocyte [84]. PLCz triggers intracellular Ca2þ oscillations initiated by targeting membrane-bound Phosphatidylinositol 4,5-bisphosphate (PIP2) leading to inositol 1,4,5trisphosphate (IP3)- facilitated release of intracellular Ca2þ in the oocyte [98]. Oocyte activation on sperm entry in the oocyte leading to resumption of oocyte’s second meiotic division, release of second polar body and initiation of syngamy. It also leads to cortical granule reaction which prevents polyspermy.

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Apart from the sperm-borne oocyte activating factor, centrosomes are also contributed by the sperm except in rodents [85,93]. During gametogenesis the centrioles are destroyed and centrosomes disintegrated in the maturing oocyte. This is to ensure proper number of centrioles in the zygote. Sperm centrosome recruits the Microtubule organizing center (MTOC) components and thus leads to the formation of functional mitotic spindle poles for the embryonic cell divisions. However, centriole and centrosome are maternally inherited in rodents. A recent investigation has illustrated that during the epididymal maturation the murine sperm centrioles, first distal and then proximal, gets destroyed in the mid-to- lower corpus [94]. In contrast to paternal inheritance of centrioles, embryo’s mitochondrial inheritance is exclusively maternal. Sutovsky et al., have shown in their studies that sperm mitochondria is ubiquitinated throughout spermatogenesis, especially in the round spermatid stages. During spermiogenesis and epididymal maturation, these ubiquitinated mitochondria get concealed by the inter- and intra-disulfide-bridges between protamines. However, post fertilization egg’s glutathione induced disulphide bond reduction exposes these ubiquitinated mitochondria. It is been shown that sperm mitochondrial ubiquitination increases in the egg cytoplasm [97]. This ubiquitin tagged mitochondria are selectively proteolyzed by proteasome or lysosome. This ubiquitination is a mechanism of ear marking the paternal mitochondria to remove heteroplasmy. From the discovery of RNA in sperm [87] and absence of sperm RNA elements (SRE) correlating to idiopathic infertility in males [36], it can be concluded that sperm RNA is important for pre-implantation embryo development and transfer of epigenetic information to the offspring [12]. In silico analysis of RNA sequencing data from sperm RNA, MII oocytes and pre-cleavage stage zygotes revealed 5 potential paternal RNAs (Hdac11, Fbxo2, Map1lc3, Pcbp4 and Zfp821) whose translated products interact with maternal co-factors to bring about a successful maternal to zygotic transition (MZT) [62]. Transcription factors like STAT4, PT32 are paternally derived, and STAT4 is presumed to be involved in the onset of zygotic transcription [34]. Paternal RNAs like clusterin, A-kinase anchoring protein 4 (AKAP4), forkhead box G1B (FOXG1B), wingless-type MMTV integration site family member 5A (WNT5A) are important for fertilization, differentiation and embryo’s morphogenetic patterning [40]. The first cleavage division in mouse requires sperm borne-microRNA-34c [47]. Furthermore, sncRNAs including miRNAs and tsRNAs are being shown to transfer paternal epigenetic traits trans generationally [13,20]. These RNAs hence define the unique paternal contribution to the zygote.

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FIG. 3.1 Spermatozoal contributions to fertilization and zygotic development.

In conclusion, molecular contributions of sperm are regularly associated with early embryonic development. These encompass transfer of sperm DNA, mRNAs, proteins and non-coding RNAs such as microRNAs (miRNAs), PIWI-interacting RNAs (piRNA), tRNAs and associated modifications to the embryo (Fig. 3.1). These signatures seem to be linked to sperm quality and male fertility potential [40,47,84]. In spite of all these insights in the complex world of sperm RNA, one question still haunts us: what is the basis of differential gene expression in germ cells of infertile men and fertile ones? Apart from the environmental, epigenetic intervention leading to the alteration of RNA profiles in infertile men, probably methylation and histone modifications may lead to these discrepancies in expression profiles.

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[105] Yoshida K, et al. Mapping of histone-binding sites in histone replacement-completed spermatozoa. Nat Commun 2018. https://doi.org/10.1038/s41467-018-06243-9. [106] Yuan S, et al. Sperm-borne miRNAs and endo-siRNAs are important for fertilization and preimplantation embryonic development. Development (Cambridge) 2016. https://doi.org/10.1242/dev.131755. [107] Zalenskaya IA, Morton Bradbury E, Zalensky AO. Chromatin structure of telomere domain in human sperm. Biochem Biophys Res Commun 2000. https://doi.org/10.1006/bbrc.2000.3917. [108] Zhao M, et al. Transition nuclear proteins are required for normal chromatin condensation and functional sperm development. Genesis 2004;38(4):200e13. https://doi.org/10.1002/gene.20019. [109] Zhao M, et al. Targeted disruption of the transition protein 2 gene affects sperm chromatin structure and reduces fertility in mice. Mol Cell Biol 2001;21(21):7243e55. https://doi.org/10.1128/mcb.21.21.72437255.2001. [110] Zhuang T, et al. CHD5 is required for spermiogenesis and chromatin condensation. Mech Dev 2014;131(1): 35e46. https://doi.org/10.1016/j.mod.2013.10.005. Elsevier Ireland Ltd. [111] Zuo X, et al. The histone methyltransferase SETD2 is required for expression of acrosin-binding protein 1 and protamines and essential for spermiogenesis in mice. J Biol Chem 2018;293(24):9188e97. https:// doi.org/10.1074/jbc.RA118.002851. American Society for Biochemistry and Molecular Biology. [112] Zhang MZ, et al. In the human sperm nucleus, nucleosomes form spatially restricted domains consistent with programmed nucleosome positioning. Biology Open 2019. https://doi.org/10.1242/bio.041368. [113] Kistler WS, et al. RFX2 is a major transcriptional regulator of spermiogenesis. PLoS Genet. 2015;11(7). https://doi.org/10.1371/journal.pgen.1005368. Public Library of Science. [114] Cheng Y, et al. Abnormal sperm in mice lacking the Taf7l gene. Mol Cell Biol. 2007;27(7):2582e9. https:// doi.org/10.1128/mcb.01722-06. American Society for Microbiology. [115] Doyen CM, et al. Subunits of the histone chaperone CAF1 also mediate assembly of protamine-based chromatin. Cell Rep 2013;4(1):59e65. http://doi.org/10.1016/j.celrep.2013.06.002. Cell Press.

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Epigenetic regulation in stem cells

4

Juqing Zhanga, Ahmed Hamed Arishab, Jinlian Huaa College of Veterinary Medicine, Shaanxi Centre of Stem Cells Engineering & Technology, Northwest A&F University, Yangling, Shaanxi, China; bDepartment of Physiology, Faculty of Veterinary Medicine, Zagazig University, Zagazig, Egypt

a

During a life course, in order to maintain self-balance during development and adaptation, the body retains some undifferentiated primitive cells, which are generally named stem cells. Once needed, these cells can divide into new cells in the normal way of development. As an special organism ages, this cellular property changes and cells often reach a state of cellular exhaustion where they lose the ability to differentiate. Eventually, this will disturb the well-balanced cell replacement, tissue regeneration and maintaining organ function. The definition of stem cells is constantly being revised and supplemented. Most biologists and physicians approve that stem cells are cells that have the capacity to self-renew as well as the ability to generate differentiated cells [1,2]. Because of the limitations of people’s initial understanding of stem cells, the terminology in many literatures remains confusing. All of these complicated the concept of stem cells. Stem cells usually have the following characteristics. First of all, cells can undergo self-renewing to produce at least one offspring which is identical to the parent cell and maintain the stem cell pool. Second, through lineage commitment and differentiation, more differentiated progenitor cells, precursor cells and terminal differentiated cells are produced. Third, after being transplantation into the body, stem cells has a homing ability to a given tissue, differentiate into the cell type of this tissue in a specific environment and have corresponding functions [3]. Simply speaking, stem cells, is a kind of undifferentiated cells with multidimensional differentiation potential and self-replication ability that has the ability to form various tissues and organs of mammals. For the potential of regenerating various tissues and organs, they are believed to be the best materials for regenerative medicine [4].

History of stem cell research In the process of life evolution, organisms are responsible for maintaining the continuity of races which requires the production of offspring that have characteristics similar to those of their parents at a minimal cost. In the mid-nineteenth century, it was found that all animals and plants developed from cells and were composed of cells and cell products. Furthermore, growth and development of any organism depend on proliferative cells in the body. In 1896, the foundation underlying of concept of

Epigenetics and Reproductive Health. https://doi.org/10.1016/B978-0-12-819753-0.00004-0 Copyright © 2021 Elsevier Inc. All rights reserved.

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stem cells was first proposed by Wilson [5]. At that time, stem cells were thought to be only the primitive cells capable of producing progeny cells. In 1958, Stevens transplanted mouse embryos into the testis or kidney of mice and obtained teratoma. Subsequently, embryonic carcinoma cells (ECCs) was isolated. Later on, many scholars have shown that ECCs are very similar to cells that develop into mouse embryos [6]. In 1981, the mice embryonic stem cells (mESCs) cell line was first established by Evans’ team [7]. This kind of cell has the potential to proliferate indefinitely, self-renew and differentiate into three embryonic layers in vitro. In the following 20 years, ES-like cell lines of pigs, cows, rabbits, goats, mink, hamsters and primates were established. Since the 1990s, the technology of isolating and culturing stem cells from various sources has been continuously improved. An increasing number of scientists are devoting themselves to stem cell research. In 1998, Thomson’s and Gearhart’s group established human embryonic stem cell lines from early embryos and primordial germ cells (PGC) respectively which play great impetus to regenerative medicine [8]. At the end of the 20th century, it was found that adult stem cells can differentiate into the cells and/or tissues in certain microenvironment [9]. It seems that stem cells have strong plasticity. In August 2006, Yamanaka’s team introduced 24 transcription factors into mouse fibroblasts, and finally determined that, at least, four transcription factors were needed to reprogram fibroblasts into induced pluripotent stem cells (iPSCs) which are similar to ES cells [10]. Then, regarding iPSCs, scientists did a lot of research and made breakthroughs. IPSCs obtained from patient cells can be used to treat their own diseases with minimal immune rejection. It also provides great convenience for the study of the origin of early life. IPSCs has become one of the greatest biological achievements in the 21st century [11].

Types of stem cells Stem cells are generally round or oval in shape, small in size, relatively large in nucleus, mostly euchromatin, and have high telomerase activity. According to the developmental potential, there are four types: totipotent stem cell, pluripotent stem cell, multipotent stem cell and unipotent stem cell [12]. The development and differentiation of stem cells are accompanied by the origin and development of life. Generally speaking, the differentiation potential of stem cells progressively decreases. During ontogeny, most of cells change from totipotency to pluripotency or multipotency and then to unipotency, at last, turning to terminally differentiated cells. The zygote forms the early embryo through cleavage. During this period, cells have the ability to develop into both a trophoblast and a complete fetus which was called expanded stem cells (EPSCs) in vitro. In 2017, Deng’s team first reported such cells in vitro. The early embryo further develops into blastocyst containing ICM which have the ability to develop into a complete fetus [13]. Individuals continue to develop. Some of the epiblast cells migrate to form PGC and the rest continue to differentiate becoming adult stem cells or terminally differentiated cells. Furthermore, according to their sources, the term stem cells encompass many distinct types including embryonic stem cells (ESCs), tissue specific progenitor stem cells (TSPSCs), mesenchymal stem cells (MSCs), umbilical cord stem cells (UCSCs), bone marrow stem cells (BMSCs), and induced pluripotent stem cells (iPSCs) (Fig. 4.1).

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FIG. 4.1 The division of cellular pluripotency corresponds to developmental status in vivo. EPSC, Expanded stem cell: A kind of cell line have the ability to develop into both a trophoblast and a complete fetus; ICM, Inner cells mass:Cells from ICM of an embryo are pluripotent and can differentiate into the cells forming the three germ layers; ES, Embryo stem cell: Embryonic stem cell lines with the same developmental capacity as ICM.

Induced pluripotent stem cells (iPSCs) Plasticity of cell fate is an important issue in cell biology research. Since the 1960s, as the development of somatic cell nuclear transplantation, cell fusion and overexpression of transcription factors, researchers has increasingly recognize that somatic cells in animals retain complete genetic information and the fate of cells can be changed through reprogramming [14]. In 2006, Yamanaka’s team introduced reprogramming factors Oct4, Sox2, Klf4 and c-Myc into mouse fibroblasts by reverse transcription virus, and obtained induced pluripotent stem cells (iPSCs) which is similar to mouse ES cells. Then, human iPSCs line was rapidly established. With the further development of research, the safety of gene introduction, tissue sampling and induction culture have been improved [15].

Mechanism on the self-renewal of PSCs The development of multicellular organisms is a result of selective gene expression. Each cell contains all the genetic information of the organism. However, not all genes are active at the same time, some are active and some are not. Differential gene expression determines cell fates. The maintenance of pluripotent state depends on the stable expression of pluripotent genes. We will highlight three main factors, transcription factors, signal transduction pathways and epigenetics.

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Core transcription factors OCT4 POU transcription factor is a DNA-binding protein that activates gene transcription and has ciselements in the promoter or enhancer regions [16]. Mouse Oct4 protein has 352 amino acids that belongs to POUⅤ proteins. This gene is expressed only in oocyte, ICM and PGC and is considered as a marker of pluripotency [17]. If the expression of Oct4 is absent, blastocysts development was compromised [18,19]. If Oct4 is knocked out in ES cells, the pluripotency will not be maintained. Moreover, most of the Oct4 action sites are located at the enhancers at the distal end of the gene. For example, in mice Rex1 is regulated by Oct4. ICM differentiates into the epiblast when the expression of Rex1 is significantly reduced [20]. Until now, Oct4 has been identified as the core transcription factor of pluripotent stem cells in many species. Similar to Oct4, transcription factor Sox2 is also associated with the pluripotency of ESCs and iPSCs [21].

Nanog In 2003, scientists discovered a protein called Nanog that can maintain the pluripotency of ICM and ESCs to a large extent. Nanog is expressed in ES cells, embryonic germ cells and embryonic cancer cells, but not in hematopoietic stem cells, parietal endoderm, fibroblasts, adult tissues or differentiated ES cells [22]. Further research showed that Nanog can be continuously expressed in ES cells and endoderm cells [23], but its expression is down-regulated during implantation stage. The pluripotency of ES cells decreases without Nanog which differentiates into ectoderm. The core pluripotent factors of embryonic stem cells interact to form a network and maintain pluripotency [24]. In mouse embryonic stem cells, Oct4, Sox2 and Nanog interact to form a huge transcriptional activation complex [25]. This transcriptional activation complex binds to the upstream of the regulation sequence of pluripotent gene’s, such as rex1 klf4, and recruits other transcriptional activators to activate the pluripotent gene [26].

Signal transduction pathways The establishment and maintenance of cellular pluripotent state requires the establishment and maintenance of extensive gene expression regulatory network. Transcription factors associated with specific pluripotent states of cells recruit large numbers of transcription activators, which form a large complex that recognizes specific chromatin locations and activates downstream genes which are often associated with specific biological processes that are regulated by complex signaling pathways. For example, pluripotent genes activate the genes at the node of the pluripotent signaling thus maintaining the cell’s pluripotent state. Up till now, at least three signal transduction pathways have been found to play an important role in maintaining ES cell pluripotency.

LIF-STAT pathway The mES cells cultured in vitro will differentiate when LIF is absent in the culture medium. In recent years, extracellular signaling pathways have been shown to regulate ES cell proliferation or

Wnt pathway

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FIG. 4.2 LIF-STAT3 pathway.

differentiation. Signal transducer and transcription3 (STAT3) and extracellular signal-regulated kinase (ERK) plays an important role in self-renewal and differentiation of stem cells [27]. These two signaling pathways determine cell differentiation or proliferation. Extracellular signals bind to receptors that located on cell surface and activate the receptor-coupled tyrosine kinase (JAK), which phosphorylates tyrosine on STAT3 and ERK. The main molecule acting on STAT3 signaling pathway is gp130, and the molecule acting on ERK pathway is SHP-2 (Fig. 4.2).

Wnt pathway Wnt is a family of protein signaling molecules, which is involved in the regulation of gene expression, cell proliferation and differentiation. Wnt/beta-catenin pathway which is also be called the classical Wnt signaling pathway [28]. When Wnt binds to its receptor proteins, the spatial structure changes, leading to the suppression of glycogen synthase kinace-3b(GSK-3b). GSK-3b can make the phosphorylation of b-catenin, which can be recognized by the E3 ubiquitin ligase and leads to ubiquitin degradation. Activation of Wnt signal can maintain the pluripotency of ES cells in human and mouse. Inhibition of Wnt signaling pathway may also interfere with the maintenance of ES omnipotence by LIF. As a powerful signaling molecule, Wnt plays a regulatory role in a variety of stem cells, including hematopoietic stem cells, neural stem cells and skin stem cells. Wnt signal transduction can play different functions under different conditions. It can promote differentiation or maintain cell pluripotency. However, Wnt signaling alone is not sufficient to support pluripotency.

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TGF-b pathway The TGF-b signaling pathway is involved in the regulation of apoptosis, proliferation, senescence, inflammation, cell fate, and tissue repair during both the developmental and adult life. The TGF-b family consists of more than 30 growth factors including TGF-bs and BMPs. The TGF-b pathway can also be regulated by Wnt pathway. Repression of the TGF-b signaling during the reprogramming of mouse embryonic fibroblasts into iPSCs with SB4315342, a small molecule inhibitor specific to TGFb signaling pathway, via modulating the FGF/MEK/ERK pathway by reducing ERK phosphorylation in ES cells [29]. In the maintenance of pluripotent stem cells, SB431542 functions by inhibiting fibroblast growth factor/MEK/ERK signaling pathway. Many signal proteins of Smad family are effectors of TGF-b signaling pathway [30]. In addition, many new stem cell signaling pathways have been reported recently involving BMP signaling pathway that promotes cell pluripotency and selfrenewal potential by activating Smad family signaling molecules [31].

Epigenetics Epigenetics is a branch of genetics that studies the hereditary changes of gene expression without altering the nucleotide sequence of genes [32]. Mammalian cells contain all the genetic information of the individual. In eukaryotes, gene expression regulation is much more complex than that in prokaryotes, because the transcription machinery recognizes chromatin templates rather than exposed DNA. The binding of transcription machinery to chromatin is essentially a molecular interaction [33]. Each binding site of transcription factor has a given pattern; these patterns can be known as Motif. These motifs often have many copies in the genome. However, some chromatins are highly concentrated, which limits the binding of transcriptional complexes to these motifs. Therefore, whether the chromatin is accessible become a key problem. When chromatin is open, transcription factors are more likely to be enriched. Chromatin is shut down and transcription factors may even lose their binding capacity. The study of the open state of chromatin belongs to epigenetics. In conclusion, cell reprogramming is also epigenetic reprogramming [34]. There are many phenomena of epigenetics, such as DNA methylation, genomic imprinting, maternal effects, gene silencing, nucleolar dominance, dormant transposon activation and RNA editing. Among them, DNA methylation, histone modification and chromatin remodeling are the hotspots of current research.

DNA methylation There is a wide range of methylation in mammalian genomes, especially in the cytosine sites of CpG dinucleotides, which are symmetrically methylated on complementary double strands of DNA [35]. This methylation pattern changes periodically during embryonic development. This suggests that DNA methylation plays an important role in cell fate determination. Shortly after fertilization, male prokaryotic cells rapidly and actively demethylate. During the period from 2 cells to blastocyst, DNA methyltransferases are largely removed from the nucleus, and the genomes of both male and female parents are actively demethylated. Of course, DNA methylation is also retained as a genetic imprint. During differentiation, DNA methylation is critical for maintaining both stem cell identity and lineage

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commitment. In ESCs, global DNA hypomethylation plays essential role in their blocking and hence in silencing some pluripotency factors while favoring the expression of differentiation-associated markers [36]. Furthermore, ESCs maintains significantly higher levels of non-CpG methylation mediated by DNMT3A and DNMT3B [37,38]. Yet, the relation between non-CpG methylation and pluripotency in either ESCs and iPSCs remains not fully clear [39]. ESCs posses a well-organized and stable epigenomes inspite of their high DNA methylation turnover rates [40]. Regarding HSCs differentiation, while DNMT1 protects from the premature activation of differentiation programs [41,42]. DNMT3A and DNMT3B represses the self-renewal gene network during differentiation [43]. DNA methylation is a major factor limiting the efficiency of somatic cell nuclear transfer. On the other hand, DNA demethylation is an important event in the reprogramming of iPSCs. IPSCs has been shown to differentiate into an original cell lineage despite possising some donor cells residual DNA methylation signatures [44e46]. Previous studies have shown that DNA demethylation reagent 5-azacyitidine can improve induction efficiency in reprogramming. Under certain conditions, DNA demethylase TET1 can replace Oct4 to achieve somatic cell reprogramming in mice [47]. The importance of DNA methylation to cell physiology can also be well illustrated by the disease caused by errors in the process of establishing and reading these markers.

Histone modification Although DNA methylation is very important for the correct expression of many genes, in some diseases, it cannot fully explain some problems of transcriptional regulation. Besides DNA methylation, several other evolutionary mechanisms are important for transcriptional regulation in eukaryotic cells such as histone modification. Without histone binding, the eukaryotic genome cannot be assembled and folded effectively. Structurally, histone modification occurs on the N-terminal tail of histone protruding from the nucleosome. These modifications include methylation, acetylation, phosphorylation, ubiquitination and polyribosylation. In general, specific histone methylation such as methylation of the ninth lysine occurring in H3, makes nucleosomes more dense, thus rejecting the binding of transcription factors. At the same time, there will be some molecular which can recognize histone methylation that inhibits gene transcription, accelerating heterochromatin formation. On the contrary, histone acetylation can make chromatin more porous, which is conducive to the recruitment of more transcription factors making gene activation [48]. In early embryos, H3K27 acetylation levels are high, and in adult tissues, H3K9 methylation levels are high. Therefore, covalent modification of histones must be temporary so that the transcriptional state of genes can be manipulated. Histone methylation is also dynamically regulated by histone methylation complexes and demethylation complexes that is the same as histone acetylation. These enzymes balance with each other and co-regulate cell homeostasis. H3K4me3 and H3K27me3 catalyzed by trithorax group (TrxG) and the Polycomb group (PcG) proteins respectively are vital for the regulation of ESCs via association with promoter regions of genes regulating differentiation [49].

Chromatin remodeling Except for covalently modifying histones, there is another evolutionary conservative mechanism regulating chromatin conformation. This mechanism is mainly performed by chromatin remodeling

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complexes dependent on ATP hydrolysis such as SWI/SNF. SWI/SNF itself has no ability to bind DNA, but can be recruited into promoters by specific transcription factors, using the energy produced by ATP hydrolysis to change the position of nucleosomes. SWI/SNF complex has been proved to be synergistic with histone acetylation [50]. Transcription activators can recruit HAT to specific sites of DNA and acetylate histones. Then, acetylated lysine residues can be used as binding sites for bromo domain which was a common domain as core factors of complexes have. Chromatin remodeling complexes promote nucleosomes that bind to them, making histones slipping and even causing exposed DNA regions, Increasing the probability of binding transcription factors (Fig. 4.3).

Three-dimensional structure of genome With the development of chromatin conformation capture (3C), enhancer-bound proteins were found to be able to cross-link not only with DNA, but also with other proteins. If the protein binds to other DNA, a special spatial structure of DNA-protein-DNA will be formed at this site. The known longrange interactions can reach several Mbs. 4C, 5C and High C are the derivatives of 3C technology, and the last is the one that can be enriched in almost all interactions with high throughput [51]. This interaction explains some biological laws and disease occurrence at three-dimensional level that is closely related to the identification of stem cells. It has been recognized that genomes and proteins are not disorderly arranged in the nucleus. They can interact to form more advanced structures to regulate gene expression more accurately. In embryonic stem cell (ESC), both the Polycomb repressive complexes PRC1 and PRC2 are essential for the pluripotent state induced via silencing of lineage-specifying developmental regulator genes1. Recently, these PRCs were suggested to function via controlling the spatial genome organization [52e55]. This spatial network contains the four Hox gene clusters and early developmental transcription factor genes. For instance, evaluating the 3D conformation of the murine HoxB locus in mouse ESCs revealed that, while, the transcription factor CTCF, essential for the formation of

FIG. 4.3 Effect of epigenetic modification on gene expression in ESCs.

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chromatin loops and enhancerepromoter interactions, occupancy alone was not sufficient to predict the in vivo folding, homotypic interactions between active and Polycomb-repressed promoters cooccurring in the same DNA fiber sufficiently explained the HoxB folding patterns [56].

Conclusion Transcription factors, signal transduction pathways and specific epigenetic landscapes work together to determine cell fate and maintain cell pluripotency. In-depth understanding of the nature of pluripotency is crucial for developing methodologies to induce or convert somatic cells into PSCs. This can be considered a cornerstone for future applications in the field of regenerative medicine and disease prevention.

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CHAPTER

5

Aberrant epigenetics and reproductive disorders

Ummet Abura, b, Sezgin Gunesb, c a

Department of Medical Genetics, Faculty of Medicine, Ondokuz Mayıs University, Samsun, Turkey; bDepartment of Multidisciplinary Molecular Medicine, Health Sciences Institute, Ondokuz Mayıs University, Samsun, Turkey; c Department of Medical Biology, Faculty of Medicine, Ondokuz Mayıs University, Samsun, Turkey

Epigenetic alterations are heritable and reversible DNA modifications of both gene expression and activity without any alteration in DNA sequences. DNA methylation, histone tail modifications and chromatin remodeling and non-coding RNAs nc(RNAs) regulation are major epigenetic mechanisms. Reproductive system is highly vulnerable to environmental factors including season and temperature alterations and exposure to chemical agents and life style alterations such as diet, smoking, and use of alcohol. Recent studies have demonstrated the association between epigenetic alterations and reproductive function [1]. Therefore, in this chapter we have reviewed the current scientific data regarding epigenetic modifications and reproductive system.

Epigenetic alterations and reproductive system Reproductive system is highly vulnerable to environmental conditions including season, and exposure to chemical agents and life style alterations such as diet, smoking, and use of alcohol. In this regards, recent studies have demonstrated the association between epigenetic alterations including methylation of DNA, modifications of histone, and non-coding RNAs and reproductive function [1].

Role of epigenetic alterations in female infertility Polycystic ovary syndrome Polycystic ovary syndrome (PCOS) is a highly heterogenous and complex endocrine and metabolic disorder in women affecting reproductive function at reproductive age [2]. Recent studies have shown a relation between epigenetic aberrations and PCOS [3]. Aberrant methylation of several genes such as aromatase (CYP19A1) [4], EPHX1 [5] and luteinizing hormone chorionic gonadotropin receptor (LHCGR). [5], yes-associated protein (YAP1) [6], and follistatin (FST) [7] in distinct tissues have been found to be associated with PCOS. Recently, two studies have highlighted the significance of DNA alterations in global DNA methylation alterations in granulosa cells (GCs) of patients with PCOS [8,9]. Sagvekar and colleagues have studied differentially methylated regions in 6486 CpG sites of 3840 genes in GCs of women with PCOS and controls. Hypomethylation and hypermethylation have been found in 2977 CpGs Epigenetics and Reproductive Health. https://doi.org/10.1016/B978-0-12-819753-0.00005-2 Copyright © 2021 Elsevier Inc. All rights reserved.

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representing 2063 genes and 2509 CpGs within 1777 genes, respectively. Methylation alterations have been also observed in ncRNAs are associated ovarian functions and dysregulation in PCOS [8]. Pan and colleagues identified using RNA-seq, 92 differentially expressed genes (DEGs) GCs from 110 patients with PCOS undergoing in vitro fertilization compared to those of 119 women with normal ovulatory cycles. Additionally, 25% decrease in global DNA methylation and hypomethylation of several gene promoters of GCs related to lipid and steroid synthesis in PCOS women have been reported [9]. A recent study investigated the potential roles of circular RNAs (circRNAs) obtained from exosomes of follicular fluids in patients with PCOS. Bioinformatic analysis has detected 245 downregulated and 167 upregulated circRNAs in follicular fluids of PCOS women compared to that in non-PCOS women. Functional analysis findings have suggested chronic inflammation, bacterial infection and oxidative stress pathways could be the target of these circRNAs [10].

Endometriosis The endometrium is a highly dynamic tissue and normally undergoes cyclic rounds starting with proliferation, followed by differentiation, degradation, and round up by regeneration. These rounds are regulated along remarkable alterations in gene expression that arise in response to alterations in circulating concentrations of the progesterone and estradiol hormones [11,12]. Across the menstrual cycle between cycle stages thousands of genes are differentially methylated that lead to alterations of methylation profile in human endometrium [13,14]. Endometriosis is a common and complex chronic inflammatory disease with the functional endometrial tissue presence and growth outside the uterine cavity. This growth is mainly in the ovaries, pelvic peritoneum and rectovaginal septum and leads to reduced fertility in women of reproductive age [15,16]. A number of studies revealed the role of genetic factors to the susceptibility to endometriosis [15,17]. The incidence of the disease is higher in women with patients of endometriosis among the relatives than in sporadic cases [15]. Recently, a study has investigated tissue-specific genotype-methylation profile of endometrium and blood samples obtained from women at different menstrual cycle stages. Large methylation alterations that were not seen in blood have been reported in endometrium using 27,262 DNAm probes across the menstrual cycle [12].

DNA methylation The development of endometriosis is induced by two driving forces which are hypoxia and inflammation. These two forces regulate DNA methylation patterns via the enzymes called DNA methyltransferases (DNMTs) and promote epigenetic remodeling [18]. Hypoxia leads to downregulation of DNMT1 through miR-148a and finally results in global hypomethylation. However, inflammation stimulates elevation in DNMT3a loci-specific hypermethylation of DNA and expression is regulated by hypoxia and inflammation regulates through microRNAs (miRNAs). Additionally, DNMT3a expression is suppressed by blocking of the prostaglandin E2 (PGE2) path suggesting that DNMT3a level might be stimulated or retained by an inflammation pathway [19]. The third key component is estrogen pathway that is important in the development and maintenance

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of endometriosis. Recent studies have demonstrated aberrant methylation of promoter and/or introns of several genes of steroid hormone pathway including steroidogenic acute regulatory protein (StAR), aromatase (CYP19), estrogen receptor (ER) b12, cyclo-oxygenase (COX)-2 and steroidogenic factor (SF)-1 [20e24].

Modification of histone code Histones are basic, small and positively charged proteins rich in arginine and lysine residues localized in the nucleus of both somatic cells and gametes. These proteins are modified post-translationally on their carboxyl- and amino-terminal tails via acetylation, methylation, phosphorylation and ubiquitination [25]. These histone tail modifications alter the binding capacity of RNA polymerase and transcription factors to DNA, therefore cause changes in gene expression and activity [26,27]. Acetylation of lysine (K) residues of Histone 3 (H3) and Histone 4 (H4) causes to activation of transcription through leading to loose of chromatin structure and facilitating the binding of transcriptional factors to DNA [28,29] in somatic cells and spermatozoa. Conversely, deacetylation of histone residues leads to transcriptional silencing, and correlates with methylation of histones in general [30]. Histone acetyl transferases (HATs) and histone deacetylases (HDACs) are key enzymes involved in histones acetylation via a process that leads opening of the DNA and activation of the gene expression and removal of the acetyl groups leading to silencing of gene expression, respectively [26,31]. HDAC1 and/or HDAC2 levels have been found to be upregulated in endometriotic stromal cells [32e34]. HDAC1 and HDAC2 expression is stimulated by the steroid hormones, Estradiol (E2) and Progesterone (P4), leading to down regulation of ER a and development of endometriosis [19].

Short non-coding RNAs and endometriosis MicroRNAs are short about 20e25 nucleotide in length, endogenous, single stranded and non-coding RNAs. miRNAs play important regulatory roles in post-transcriptional gene expression level via various epigenetic mechanisms. miRNAs downregulate gene expression through pairing with mRNA causing either degradation or translational repression of mRNA [26]. Recent studies using genomewide analysis of the miRNA expression profile demonstrated that dysregulated miRNAs have critical roles in the pathogenesis of endometriosis [35,36]. miRNAs follow three different pathways during the development of endometriosis. miRNAs mediated development and progression of endometriosis pathways are related with hyperactivated inflammatory responses, steroidogenic regulation and hypoxic responses. The principle role of steroid hormones in the pathogenesis of endometriosis is well-known. Aberrantly suppressed miR-23a and miR-23b expression levels have been observed in the endometrium of women with endometriosis compared to those of healthy women. These data suggest that higher levels of miR-23a lead to inhibition of SF-1 expression and therefore low steroidogenic activity could be maintained in normal endometrium. miR-148a is responsible for global passive demethylation, its level aberrantly increase in the ectopic stromal cells and later this global DNA hypomethylation is induced by microenvironmental hypoxia. In addition, hypermethylation-mediated suppression of miR-196b and miR-503 depresses the genes involved in the induction of apoptosis, inhibition of proliferation, and expression of angiogenesis, thus promoting the initiation of endometriosis. Additionally, inflammatory cytokine-induced miR-302a modulates DNA methylation by inducing an inflammatory response [19].

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HDAC inhibitor in the treatment of endometriosis Histone acetylation becomes an attractive therapeutic target due to its critical role in the transactivation of central genes in the developing of endometriosis. Studies investigating the use of HDAC inhibitors in the treatment of endometriosis demonstrate that these inhibitors lead to apoptosis, cell cycle arrest, and therefore decrease the lesion size in vivo. In addition, reactivation of several candidate tumor suppressor genes including death receptor 6 and C/EBPa are reported to be evaluated as therapeutic target. HDAC inhibitors lead to hyperacetylation of histone result in inhibition of the DNA damage responses and mitosis. Although HDAC inhibitors rise as promising candidates for the treatment of the disease, investigations are necessary to maximize their therapeutic effect and minimize their cytotoxic and adverse effects [19].

Premature ovarian failure Premature ovarian failure (POF) commonly called primary ovarian insufficiency (POI) (POF1: MIM 311360, POF2: MIM 300511; POF3: MIM 608996) is a heterogeneous and complex disorder and characterized by the loss of normal function of the ovaries, premature reduction of ovarian follicles before 40 years of age, with low estrogen and high gonadotrophin level [37,38]. The frequency of POF at the age of 20, 30, and 40 is 1/10,000, 1/1000, and 1/100, respectively [39]. Several causes are reported to be associated with POF such as X-chromosome abnormalities including monosomy or trisomy X, X-chromosome partial deletions, X; autosome translocations, monogenic defects such as mutations of FMR1 on Xq27.3, DIAPH2 on Xq21.33, FOXL2 on 3q22.1, BMP15 on Xp11.22, NOBOX on 7q35, NR5A1 on 9q33.3, infections and autoimmune diseases [38]. Several studies indicated that aberrant epigenetic modifications including microRNAs and long noncoding RNAs induce development of POF [40e42]. A recent study revealed that miR-15b stimulates POF in mice by inhibiting Klotho expression. The expression level of miR-15b is induced by cyclophosphamide in ovarian granulosa cells of mice. Overexpressed miR-15b demonstrated higher levels of ROS and aberrant energy metabolism resulting in autophagy dysfunction compared to controls [40]. Xiong et al. [41] showed that cyclophosphamide promotes inhibition of proliferation of ovarian granulosa cells and plays role in the POF pathogenesis by inducing the endogenous lncRNA-Meg3 expression and activating the p53-p66Shc-p16 pathway [41]. Various histone modifications have been found to play a role in the modification of the FMR1 gene. CGG repeats of FMR1 gene have been found to be hypermethylated and hypomethylated at H3K9 and H3K4, respectively with low levels of histone acetylation in cells with the FMR1 full mutations. Therefore, increase in the CGG repeat number leads to alterations in the chromatin structure resulting in variations in FMR1 gene in POF [42].

Male infertility and DNA methylation The methylation of cytosines in CpG-islands (CGIs) is called DNA methylation. CGIs are short interspersed DNA sequences rich in C þ G and are located in the regulatory regions or most of the promoters of constitutive and developmental genes, and various tissue-specific genes [43]. Generally, hypermethylation of these DNA sequences repress the gene expression by preventing the access of polymerases and transcription factors to the promoters. Rather, gene expression is enhanced by hypomethylation [44].

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In recent studies, decreased expression of DNMT3B has been indicted in germ cells of men with bilateral spermatogenic arrest [45]. Additionally, aberrant methylation patterns have been shown of DNMT3L gene in oligozoospermic patients [46]. The deletion of Dnmt3a or Dnmt3l genes in male germ cells in various animal studies have indicated that this deletion causes meiotic arrest in spermatocytes and loss of paternal imprints in spermatogonia [47].

Methylation of imprinted genes and global/genome-wide and gene-specific methylation Imprinted genes are biallelic genes expressed monoallelically depending on their parental origin. All methylation marks of primordial germ cells (PGCs) are demethylated by passive or active demethylation and then are differentially labeled by methylation of DNA in primordial germ cells (PGCs) depending on parental origin during germline development [48]. DNA demethylation may be performed passively or actively. The passive demethylation is performed via DNMT1 on newly synthesized DNA strands during DNA replication. However, active DNA demethylation does not require replication of DNA and methylation is mediated by one or more enzymes. This process of demethylation continues with a particular remethylation process of type I spermatocytes and spermatogonia. Therefore, signature of the father is transmitted to his child/children by spermatozoa [49]. Igf2/H19, Rasgrfl, Dlk-Gtl2, and Zdbf2 are generally highly methylated paternally imprinted genes in the spermatozoa. For example, the H19 is a 2.7 kilobase gene with five exons and four introns mapped to 11p15.5. H19 encodes a putative 29K protein and an untranslated cytoplasmic RNA involved in protein transport/synthesis and in RNA processing. H19 DMR is unmethylated in the maternal allele, thus permitting expression of H19 and preventing the admission from the enhancer of IGF2 gene. However, H19 DMR methylation causes expression of IGF2 gene from the paternal allele. MEST, ZAC1, PEG3, SNRPN are maternally imprinted genes and these genes are generally unmethylated in spermatozoa and methylated in the oocyte therefore these genes are expressed through the paternal allele [50e53]. MEST encodes a family member of the alpha/beta hydrolase localized on 7q32 and is imprinted with monoallelic paternal pattern during fetal development. MEST involved in folding of alpha/beta-hydroxylase and is important in the development of fetal mesoderm [54]. Recently, strong association have been reported between aberrant DNA methylation and male infertility in different group of infertile patients (Table 5.1). A recent meta-analysis investigated the aberrations of sperm DNA methylation of imprinted genes between the idiopathic infertile men and fertile controls in 24 studies. In this meta-analysis, the methylation levels of MEST and SNRPN DMR were found to be significantly hypermethylated in infertile men then fertile men. In addition, H19 DMR methylation level has been found to be lower in infertile males compared to fertile men and the sperm concentration and motility reported to be two of the most affected semen parameters [59]. Marques and colleagues investigated the H19 and MEST DMRs methylation pattern in semen samples of infertile and fertile men. The findings have been demonstrated hypomethylation in H19 DMR in patients with OAT [77]. Similar to previous studies, hypomethylation in the H19 DMRs and hypermethylation in the SNRPN DMRs have been observed in oligozoospermic, asthenozoospermic, teratozoospermic men compere to normozoospermic men [62]. On the other hand, methylation changes of such genes in sperm DNA is suggested may adversely affect the assisted reproductive technology (ART) success as well as male fertility. Indeed, a study of

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Table 5.1 The relationship between male infertility and DNA methylation of spermatozoa. Gene

Groups (sample size)

Results

Reference

Genome-wide differential methylation analyses

Men with OA (38) and NZ (26)

[55]

Global

Infertile patients (42) and fertile donors (19) 94 men with normal and abnormal sperm parameters Men with OZ (30) and men with NZ (62)

/ Significantly differential methylation using deep sequencing, e.g. BCAN, EBF3, HOXB1, GATA3 and TCERG1L genes / Hypermethylation at APCS gene in oligozoospermic patients / No global methylation change

Global Global H19, MEST

H19, MEST, SNRPN, LINE-1

Meta-analysis of 24 studies

MEST, P16 H19, LINE-1, GNAS, FAM50B H19

AS (46) NZ (49)

/ Global sperm DNA hypomethylation methylation in OZ patients / No-methylation alterations in H19 and MEST of DMR in DNA / Hypomethylation at H19 in infertile men / Hypermethylation at MEST and SNRPN in infertile men / No difference in LINE-1 methylation levels between infertile men and controls / Hypomethylation at the P16 and MEST DMRs in AS patients

H19, LINE-1, SNRPN

Men with OZ (48), AS (52),TZ (55) and NZ (50)

LINE1, H19

Infertile cases (23) controls (11)

H19, SNRPN

Infertile patients OA (39),AT (36) and NZ (50)

H19, GNAS, DIRAS3,DNMT1, DNMT3A, DNMT3B, DNMT3L KCNJ5, MLPH, SMC1b

Men with NZ (39), moderate OZ (45), severe OZ (51) and fertile controls (59)

/ Hypomethylation at H19 DMR CpG 1, 3 and 6 in the infertile group / Hypomethylation at H19 in OZ and AZ patients / Hypermethylation at SNRPN in patients with AZ and TZ No significance at LINE1, and H19 methylation between cases and controls / Hypomethylation at H19 in the OA and AT group / Hypermethylation at SNRPN in the OA and AT groups / No significant association between aberrant methylation patterns

Subfertile cases (57) fertile controls (21)

/ Hypomethylation at KCNJ5 and MLPH

Infertile patients (15) and control group (15)

[56] [57] [58]

[59]

[60]

[61]

[62]

[63]

[64]

[65]

[66]

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Table 5.1 The relationship between male infertility and DNA methylation of spermatozoa.dcont’d Gene

PRICKLE2, ALS2CR12, ALDH3B2, PTGIR TYRO3, CGb, FAM189A1 PRRC2A, ANXA2, MAPK8Ip3, GAA

Groups (sample size)

Results

Males with reduction in fecundity (55) Proven fertile males (56)

/ Hypermethylation at SMC1b in cases / Hypomethylation at all CpGs for the PRICKLE2 and ALS2CR12 / Hypomethylation at ALDH3B2 and PTGIR / Hypomethylation at all CpGs of TYRO3, CGb and FAM189A1

Males with reduced fecundity (65) and proven fertile males (43) Subfertile males (50) and proven fertile male (28)

ALS2CR12, GAA, UBE2G2

Subfertile males (72) an proven fertile male controls (64)

MTHFR

44 oligozoospermic men 9 fertile men

BRCA1 BRCA2

Men with OAT (73) and NZ (20)

MLH1, MSH2

Men with OZ (10) and NZ (29)

VDR

Infertile men (69) and controls (37) Men with AS (25) and NZ (27)

VDAC2

MAEL

Men with NOA (26) and obstructive azoospermia and NZ (12)

/ Hypermethylation at the PRRC2A in the patients / The significantly different DNA methylation levels in the MAPK8Ip3 ANXA2 GAA gene were in the case group compared to the controls / A significant difference in the methylation level at ALS2CR12, GAA and UBE2G2 genes between cases and controls / No significant DNA methylation alterations in spermatozoa associated with the MTHFR C677T No significant differences between the promoter methylation status of either BRCA1 or BRCA2 genes / Hypermethylation at MLH1 gene in oligozoospermic men / No association between MSH2 methylation and oligozoospermia / Hypermethylation at VDR infertile group / Hypermethylation of VDAC2 promoter CpGs and positive correlation with low sperm motility / Hypermethylation of 26 consecutive CpGs in the MAEL promoter in patients with NOA

AS, Asthenozoospermic; AT, asthenoteratozoospermia; NZ, Normozoospermia; OA, oligoasthenozoospermia; OAT, oligoasthenoteratozoospermia; OZ, oligozoospermia; TZ, Teratozoospermia.

Reference

[67]

[68]

[69]

[70]

[71]

[72]

[73]

[74] [75]

[76]

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Chapter 5 Aberrant epigenetics and reproductive disorders

human placentas obtained from intracytoplasmic sperm injection (ICSI), in vitro fertilization (IVF), and natural conception indicated that H19 expression (mRNA) levels were significantly elevated in placentas of both ICSI and IVF compared to natural conception. This study supports that H19 gene hypomethylation may possibly associated with the entering of ART programs [78]. On the other hand, some studies have demonstrated an elevation in Angelman syndrome incidence in children born after ART [79]. Similarly, Manning and colleagues has shown abnormal DNA methylation causes to Angelman syndrome cases born after ART, not microdeletions in the SNRPN gene [80]. In conclusion, the etiology of these imprinting gene abnormalities in idiopathic infertility remains unclear.

Global/genome-wide and gene-specific DNA methylation In addition to imprinting genes, methylation of non-imprinting genes is found to be associated with global sperm DNA methylation of sperm DNA and methylation of these genes may be a risk factor for male infertility. Montjean and colleagues have investigated the global sperm DNA methylation status in patients with oligozoospermia (OZ) and normozoospermia (NZ). Global sperm DNA hypomethylation has been detected in patients with OZ [58]. Urdinguio and colleagues investigated the global sperm DNA methylation patterns between infertile and fertile males. They showed alterations in the methylation pattern of sperm genomic DNA in 2752 CGI and hypomethylation in the spermatozoa genome [81]. In another global sperm DNA methylation study, Schutte and colleagues have observed hypermethylation in spermatogenesis-related genes of infertile men [82]. On the contrary, Jenkins and colleagues have found no significant difference in the global sperm DNA analysis. Moreover, significant regional methylation changes at 772 CGIs have been detected between poor quality and high quality sperm samples [83]. Methylenetetrahydrofolate reductase (MTHFR) gene plays a critical role in both DNA synthesis and methylation processes. A significant correlation has been reported among the methylation of the MTHFR gene and infertile patients with NOA or poor sperm quality and recurrent spontaneous miscarriages [26]. Discoidin domain receptor 1 (DDR1) is a tyrosine kinase receptor that have critical roles in the cell proliferation, cell morphogenesis, differentiation and apoptosis of postmeiotic germ cells. Ramasamy and colleagues found a significant relationship between DDR1 promoter methylation and DDR1 expression levels in patients with non-obstructive azoospermia NOA compared to fertile controls [84] (Table 5.1).

Histone modifications and male infertility Recently, the relationship between histone tail modification and spermatogenesis has been investigated in details. In one study, acetylation status of H3K4Ac and H4K5Ac and methylation profile of H3K4Me, H3K4Me3, H3K9Me2, H3K79Me2 and H3K36Me3 in normal and abnormal human sperm samples were investigated. The findings have demonstrated heterogeneity among the histone modifications and marks on H3K4Me1, H3K9Me2, H3K4Me3, H3K79Me2, and H3K36Me3 in poor human sperm samples [85]. Yuen and colleagues developed a knockout mouse model for the histone variant H3.3-encoding gene, H3f3b. They found that the loss of H3f3b gene induces abnormalities in testes morphology and spermatozoa therefore leading to infertility [86]. In addition, Vieweg and colleagues demonstrated the loss of histone H4 lysine 12 acetylation (H4K12ac) in selected developmentally important promoters of subfertile patients [87]. The histone post-translational

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modifications in normal and abnormal sperm samples were investigated and significant alterations in post-translational modification of histones were observed in abnormal sperm samples compared to those normal sperm samples [88]. These findings suggest the importance of histone alterations in normal sperm function and fertility. To evaluate the role of histone tail modifications in human fertility further studies are necessary.

Role of protamination in male infertility Protamines (Ps) are smaller proteins compared to histones, arginine-rich, and replace 85%e90% of the histones packing the sperm DNA in late haploid phase of spermatogenesis. Transition of protamines from histones allows the sperm DNA to take up less space in the nucleus thus lead to tight condensation of sperm nucleus and enhance the motility of spermatozoa. In addition, protamination defends the sperm genome from oxidation, degradation and various harmful molecules in the female reproductive system [27,89]. P1 and P2 are two classes of protamines that are expressed equally in fertile human spermatozoa thus P1/P2 ratio is equal to one in a fertile male. However, P1/P2 protamine ratio is not equal to one in most the infertile men. Incorrect processing of protamine transcripts results in elevation of the production of immature P1 and P2 precursors reported to be associated with male subfertility [90,91]. Although the importance of protamines is still widely discussed, alterations in the protamine ratio is suggested to be associated with a variety of phenotypic features in infertile men including reduced sperm function and count, poor embryonic quality and miscarriages [92,93]. On the other hand, 10%e15% of the sperm genome remains packaged by histones and this content of histones is important for the early embryonic development. In conclusion, anomalies in protamine content or deviations from P1/P2 ratio are suggested to affect epigenetic data inherited by the paternal DNA.

Non-coding RNAs and male infertility ncRNAs can be classified into two major groups according to their length as short ncRNAs and long ncRNAs (lncRNAs). Short ncRNAs are RNA molecules that regulate gene expression after transcription through epigenetic mechanisms. MicroRNAs (miRNAs), small-interfering RNAs (siRNAs) and piwi-interacting RNAs (piRNAs) are three major classes of short ncRNAs. piRNAs are only present in round spermatids and in spermatocytes at the pachytene stage whereas endo-siRNAs and miRNAs are abundantly expressed in male germ cells throughout spermatogenesis [94]. Aberrant miRNA expression has been reported in subfertile patients with NOA and asthenozospermia [95e97]. Abu Halima and colleagues investigated miRNA profile of seminal plasma samples of 12 oligoasthenozoospermic subfertile men and 12 normozoospermic men using microarray analysis. Higher miR-765 and miR-1275 and lower miR-15a expression levels have been found in oligoasthenozoospermic men compared with the normozoospermic men [98]. Earlier the same group had investigated the miRNA profile of semen samples of 9 each of asthenozoospermic, oligoasthenozoospermic and normozoospermic men using miRNA microarray and validated with PCR. In sperm of asthenozoospermic men, 50 miRNAs were found to be upregulated and 27 miRNAs were downregulated. In oligoasthenozoospermic men 42 miRNAs were reported to be upregulated and 44 miRNAs downregulated [99]. Salas-Huetos and colleagues examined the expression levels of 736 miRNAs in spermatozoa from 10 normozoospermic fertile men. Three (miR-374b-5p, miR-532-5p and miR-564) of 48 miRNAs with

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Chapter 5 Aberrant epigenetics and reproductive disorders

stable expression have been suggested as fertility biomarkers [100]. miR-202-5p haS been shown in Sertoli cells of normal fertile men; however, miR-202-5p haS not been observed in testicular tissue samples of azoospermic men with Sertoli cell only syndrome (SCO syndrome) [101]. Yang and colleagues investigated short RNA transcriptome in testicular samples of 3 normal males using next generation sequencing. They detected 5 novel and 770 known miRNAs and 20121 piRNAs in testicular tissue indicating that the human testis has a complex population of small ncRNAs. In addition, the highest expression was reported to be the of expression miR-34c-5p, miR-103a-3p, miR-202-5p, miR-508-3p, miR-509-3-5p miRNAs and let-7 family members [102]. piRNAs profile of seminal plasma samples was investigated from 211 infertile men and 91 fertile controls. The findings revealed that piR-31068, piR-31925, piR43771, piR-30198 and piR-43773 can be potential molecular biomarkers to differentiate asthenozoospermic and azoospermic individuals from controls [103]. In summary, it has been clearly demonstrated that ncRNAs have great potential as molecular biomarkers in male infertility. Further research is needed to fully understand their complex interactions.

Conclusion Extensive studies have been performed to investigate the role of epigenetic mechanisms in reproductive disorders. Studies reporting aberrant methylation, histone modifications and abnormalities in ncRNAs in reproductive disorders in both females who undergo hormone therapy and males with infertility are also available. Still currently there are no epigenetic signatures for early diagnosis and screening of reproductive disorders. Appropriate screening and treatment algorithms for reproductive disorders including PCOS, endometriosis and male infertility are important issues both for gynecologist and andrologist. Epigenetics is a promising area which could be helpful to understand the pathology of these reproductive disorders.

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[34] Colon-Diaz M, et al. HDAC1 and HDAC2 are differentially expressed in endometriosis. Reprod Sci 2012; 19(5):483e92. [35] Braza-Boils A, et al. MicroRNA expression profile in endometriosis: its relation to angiogenesis and fibrinolytic factors. Hum Reprod 2014;29(5):978e88. [36] Hirakawa T, et al. miR-503, a microRNA epigenetically repressed in endometriosis, induces apoptosis and cell-cycle arrest and inhibits cell proliferation, angiogenesis, and contractility of human ovarian endometriotic stromal cells. Hum Reprod 2016;31(11):2587e97. [37] Cordts EB, et al. Genetic aspects of premature ovarian failure: a literature review. Arch Gynecol Obstet 2011;283(3):635e43. [38] Okten G, et al. Disruption of HDX gene in premature ovarian failure. Syst Biol Reprod Med 2013;59(4): 218e22. [39] Ghahremani-Nasab M, et al. Premature ovarian failure and tissue engineering. J Cell Physiol 2019;235(5): 4217e26. [40] Liu T, et al. miR-15b induces premature ovarian failure in mice via inhibition of alpha-Klotho expression in ovarian granulosa cells. Free Radic Biol Med 2019;141:383e92. [41] Xiong Y, et al. Cyclophosphamide promotes the proliferation inhibition of mouse ovarian granulosa cells and premature ovarian failure by activating the lncRNA-Meg3-p53-p66Shc pathway. Gene 2017;596:1e8. [42] Eslami H, et al. Epigenetic aberration of FMR1 gene in infertile women with diminished ovarian reserve. Cell J 2018;20(1):78e83. [43] Zhu J, et al. On the nature of human housekeeping genes. Trends Genet 2008;24(10):481e4. [44] Giacone F, et al. Epigenetics of male fertility: effects on assisted reproductive techniques. World J Mens Health 2019;37(2):148e56. [45] Adiga SK, et al. Reduced expression of DNMT3B in the germ cells of patients with bilateral spermatogenic arrest does not lead to changes in the global methylation status. Mol Hum Reprod 2011;17(9):545e9. [46] Kobayashi H, et al. DNA methylation errors at imprinted loci after assisted conception originate in the parental sperm. Eur J Hum Genet 2009;17(12):1582e91. [47] Bourc’his D, Bestor TH. Meiotic catastrophe and retrotransposon reactivation in male germ cells lacking Dnmt3L. Nature 2004;431(7004):96e9. [48] Reik W, Walter J. Genomic imprinting: parental influence on the genome. Nat Rev Genet 2001;2(1):21e32. [49] Boissonnas CC, Jouannet P, Jammes H. Epigenetic disorders and male subfertility. Fertil Steril 2013;99(3): 624e31. [50] Kaneda M. Genomic imprinting in mammals-epigenetic parental memories. Differentiation 2011;82(2): 51e6. [51] Arnaud P. Genomic imprinting in germ cells: imprints are under control. Reproduction 2010;140(3): 411e23. [52] Brannan CI, et al. The product of the H19 gene may function as an RNA. Mol Cell Biol 1990;10(1):28e36. [53] Zhang Y, Tycko B. Monoallelic expression of the human H19 gene. Nat Genet 1992;1(1):40e4. [54] Zheng HY, et al. Assisted reproductive technologies do not increase risk of abnormal methylation of PEG1/ MEST in human early pregnancy loss. Fertil Steril 2011;96(1):84e89.e2. [55] Sujit KM, et al. Genome-wide differential methylation analyses identifies methylation signatures of male infertility. Hum Reprod 2018;33(12):2256e67. [56] Camprubi C, et al. Spermatozoa from infertile patients exhibit differences of DNA methylation associated with spermatogenesis-related processes: an array-based analysis. Reprod Biomed Online 2016;33(6): 709e19. [57] Jenkins TG, et al. Teratozoospermia and asthenozoospermia are associated with specific epigenetic signatures. Andrology 2016;4(5):843e9. [58] Montjean D, et al. Sperm global DNA methylation level: association with semen parameters and genome integrity. Andrology 2015;3(2):235e40.

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[83] Jenkins TG, et al. Intra-sample heterogeneity of sperm DNA methylation. Mol Hum Reprod 2015;21(4): 313e9. [84] Ramasamy R, et al. Integrative DNA methylation and gene expression analysis identifies discoidin domain receptor 1 association with idiopathic nonobstructive azoospermia. Fertil Steril 2014;102(4):968e973 e3. [85] La Spina FA, et al. Heterogeneous distribution of histone methylation in mature human sperm. J Assist Reprod Genet 2014;31(1):45e9. [86] Yuen BT, et al. Histone H3.3 regulates dynamic chromatin states during spermatogenesis. Development 2014;141(18):3483e94. [87] Vieweg M, et al. Methylation analysis of histone H4K12ac-associated promoters in sperm of healthy donors and subfertile patients. Clin Epigenet 2015;7:31. [88] Schon SB, et al. Histone modification signatures in human sperm distinguish clinical abnormalities. J Assist Reprod Genet 2019;36(2):267e75. [89] Torregrosa N, et al. Protamine 2 precursors, protamine 1/protamine 2 ratio, DNA integrity and other sperm parameters in infertile patients. Hum Reprod 2006;21(8):2084e9. [90] Nanassy L, Carrell DT. Abnormal methylation of the promoter of CREM is broadly associated with male factor infertility and poor sperm quality but is improved in sperm selected by density gradient centrifugation. Fertil Steril 2011;95(7):2310e4. [91] Aoki VW, Liu L, Carrell DT. Identification and evaluation of a novel sperm protamine abnormality in a population of infertile males. Hum Reprod 2005;20(5):1298e306. [92] Nasr-Esfahani MH, et al. Effect of protamine-2 deficiency on ICSI outcome. Reprod Biomed Online 2004; 9(6):652e8. [93] de Mateo S, et al. Protamine 2 precursors (Pre-P2), protamine 1 to protamine 2 ratio (P1/P2), and assisted reproduction outcome. Fertil Steril 2009;91(3):715e22. [94] Song R, et al. Male germ cells express abundant endogenous siRNAs. Proc Natl Acad Sci U S A 2011; 108(32):13159e64. [95] Wang C, et al. Altered profile of seminal plasma microRNAs in the molecular diagnosis of male infertility. Clin Chem 2011;57(12):1722e31. [96] Tang D, et al. Altered miRNA profile in testis of post-cryptorchidopexy patients with non-obstructive azoospermia. Reprod Biol Endocrinol 2018;16(1):78. [97] Lian J, et al. Altered microRNA expression in patients with non-obstructive azoospermia. Reprod Biol Endocrinol 2009;7:13. [98] Abu-Halima M, et al. Altered micro-ribonucleic acid expression profiles of extracellular microvesicles in the seminal plasma of patients with oligoasthenozoospermia. Fertil Steril 2016;106(5):1061e1069.e3. [99] Abu-Halima M, et al. Altered microRNA expression profiles of human spermatozoa in patients with different spermatogenic impairments. Fertil Steril 2013;99(5):1249e1255.e16. [100] Salas-Huetos A, et al. New insights into the expression profile and function of micro-ribonucleic acid in human spermatozoa. Fertil Steril 2014;102(1):213e222.e4. [101] Dabaja AA, et al. Possible germ cell-Sertoli cell interactions are critical for establishing appropriate expression levels for the Sertoli cell-specific MicroRNA, miR-202-5p, in human testis. Basic Clin Androl 2015;25:2. [102] Yang Q, et al. MicroRNA and piRNA profiles in normal human testis detected by next generation sequencing. PLoS One 2013;8(6):e66809. [103] Hong Y, et al. Systematic characterization of seminal plasma piRNAs as molecular biomarkers for male infertility. Sci Rep 2016;6:24229.

CHAPTER

Epigenetic reprogramming in the embryo

6 John Huntriss

Discovery and Translational Science Department (DTSD), University of Leeds, Leeds, West Yorkshire, Great Britain

Introduction Preimplantation embryo development is marked by numerous fascinating milestones that are required for the development of the conceptus. These include the meeting and fusion of the gametes, fertilisation and the completion of meiosis in the oocyte, the transition in control from the maternal (oocyte) transcriptome to an embryonic one via embryonic gene activation, the switch from totipotency to pluripotency, the differentiation of the trophectoderm and several changes in embryonic morphology during a phase of cell division and growth. Among these developmental processes are the underlying epigenetic changes that are essential for the developing conceptus. The epigenetic reprogramming during preimplantation development erases the majority of the gametic epigenetic patterns in order to allow the embryo to re-establish an epigenetic profile that is fit for early development and for allowing differentiation thereafter to numerous cell types during embryonic growth. In this chapter we explore some of the main features, mechanisms and mediators of arguably one of the most dynamic phases in the epigenetic lifecycle.

Epigenetics, a brief overview A brief overview of epigenetic regulation and epigenetic marks and their relevance to the early embryo is provided here. Epigenetics can be viewed as another ‘layer’ of information that complements the information presented in the DNA sequence. Epigenetic programming is essentially a marking system that regulates gene expression and hence ultimately controls the phenotype and function of the cell. Epigenetic programming is essential for normal development, and therefore, if epigenetic processes are disrupted, this may lead to disease [51]. Epigenetic information is an important determinant of the phenotypic variability of cells within an organism. Therefore, cells from two different tissues will have differing epigenetic modifications. During gametogenesis and early development, many changes in epigenetic marks occur, associated with the requirement to remodel the cell types and their functions. It is also important to note that many external factors can exert an influence upon epigenetic information in a cell including environment (exposures), age, and diet as examples.

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Epigenetic marks that are relevant to preimplantation development An epigenetic mark may be a chemical modification of the DNA bases themselves (e.g. the addition of a methyl group to DNA), or a modification that leads to a change in chromatin structure, for example, a chemical modification of a histone tail. During normal mammalian development, the growth of the fetus is regulated by genetic information that is inherited from both the sperm and the oocyte. Apart from the clear differences that are associated with the X and Y chromosomes, the parental genetic contributions to the embryo also differ via a system of ‘epigenetic’ marks. Epigenetic marks have an effect on the way that DNA is ‘read’ by the cell, as they influence interactions with transcription complexes and other regulatory factors and can cause chromatin to be either open or closed, ultimately controlling gene expression. It is important to note that epigenetic marks are typically used in combination. In many situations, both DNA methylation and the histone modification ‘code’ are likely to contribute to the overall process of epigenetic regulation and hence the phenotype of a cell. Indeed, there are other epigenetic regulators that must be considered, particularly the regulatory RNA species [79], which may be used in conjunction with methylation and the histone modifications.

DNA methylation One of the most well studied epigenetic marks is DNA methylation, (5-Methylcytosine or 5 mC) which is formed when methyl group is added to the fifth carbon of the DNA base cytosine. DNA is frequently methylated at CpG sites (50 -cytosine-phosphodiester bond-guanine-3), where a cytosine is positioned next to a guanine nucleotide in the DNA sequence. Methylation of cytosine of the CpG dinucleotides within DNA is essential for vertebrate development [60]. The CpG sequence is palindromic and the methylation will also be present on the cytosine on the complimentary strand. As DNA is replicated, the DNA methyltransferase 1 enzyme DNMT1, a maintenance methyltransferase, acts upon the hemi-methylated sequence and methylates the cytosine on the newly synthesised strand. DNA methylation plays a critical role in a number of biological processes that are unique or highly regulated in gametes and preimplantation embryos, including genomic imprinting [59], X-chromosome inactivation, the expression of genes according to lineage, suppression of repetitive element sequences and genome stability [74]. Accordingly, the disruption of the DNA methylation processes can cause disease [42]. DNA methylation (5-methylcytosine) is an epigenetic mark that regulates chromatin and gene expression [134]. The location of where the DNA methylation occurs within a gene can play an important role in the activity of the gene. For example, an ummethylated gene promoter region is often a feature of an active gene, whereas DNA methylation of cytosines at CpG sites within a promoter region of a gene often leads to transcriptional silencing (the methylation here makes the gene inactive). DNA methylation at CpG sites within a promoter can silence a gene since it can recruit transcriptional repressors [121]. In contrast, methylation of cytosines at CpG sites within the gene body can activate expression of a gene and this has been observed in oocytes and in the placenta for example [41,54,103]. DNA methylation can also occur in other sequence contexts [106]. For example, non-CpG methylation, where DNA is methylated in different sequence contexts (CpA, CpT, or CpC), may be a particularly important mark in oocytes [110].

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5-Hydroxymethylcytosine (5hmC) Another epigenetic mark, which is particularly important in gametogenesis and preimplantation development is 5-hydroxymethylcytosine (5hmC) [61]. Although 5hmC has been traditionally viewed as an intermediate in the demethylation process (when methyl groups are removed from DNA-see later), more recent evidence suggest that 5hmC can in fact be a stable epigenetic mark [15] and may in fact serve as an epigenetic mark itself that is particularly important in gametes and embryos.

Histone modifications The epigenetic modification of histone proteins plays an important role during early embryonic development. Histones are important in DNA packaging, and they can be covalently modified by a number of post-translational modifications that can significantly affect whether chromatin conformation is open or closed. Chromatin conformation is affected by the chemical modification of histone tails especially those of H3 and H4 histones by methylation, acetylation, phosphorylation and other chemical modifications. The combination of modifications is called the ‘histone code’ [50]. Open chromatin is accessible to DNA replication and transcription (gene expression), and “closed” chromatin is not accessible [12].

Epigenetic reprogramming in the zygote and preimplantation embryo: an overview The mature gametes (oocyte and sperm) are highly specialized cells with highly defined functions. Picture for example the unique morphology, motility and function of a sperm cell in comparison to other cell types and it clearly demonstrates how specialised cell phenotype can be and that epigenetic information is a major driver in this process. The high degree of cellular specialization is accompanied and administered by epigenetic information unique to that cell type. Therefore, around the time of fertilisation and during the stages of preimplantation development thereafter, the epigenetic information that defined the gametes must be erased from the paternal and maternal genomes in a process called erasure. This event is necessary to remove the epigenetic features that are unique to the gametes such that a pluripotent state can be established in the embryo, a process that is required for embryo-specific patterns of gene expression that drive development. After the preimplantation stages, during gastrulation, a new pattern of epigenetic information is created, a process is called establishment. It is these processes of erasure and establishment that drive critical developmental transitions during preimplantation development and the development of the conceptus thereafter in a process called epigenetic reprogramming. In addition to the significant epigenetic reprogramming that occurs during preimplantation development, extensive epigenetic reprogramming is also required in the formation of primordial germ cells (erasure) and during gametogenesis (establishment) (for a review see Ref. [78]. See Fig. 6.1 for an overview of these changes in DNA methylation during gametogenesis and preimplantation development. At present, the epigenetic mark that is most amenable to researchers is DNA methylation and accordingly our understanding of epigenetic programming during embryonic development has focused on this modification in particular.

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FIG. 6.1 General pattern of DNA methylation reprogramming during gametogenesis and preimplantation development showing the major phases. Blue line represents DNA methylation in the male germline (or after fertilization, the male genome). Red signifies the female germline/ genome. The two short periods of de novo methylation in the zygote and mid-preimplantation stages are informed by recent studies with RRBS or WGBS as detailed in section 7. The image shows overall (global) methylation only, therefore certain sequences (for example imprinted genes) may show a different pattern over the stages shown. The placing of images representing the various stages of development are not precise as information is from taken both mouse and human studies and from a number of different methodologies.

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The gametes and their epigenetic characteristics It is important to note that at the point of fertilisation, the epigenetic states of the parental genomes in the male and female gametes are strikingly different. Therefore, in the zygote extensive epigenetic programming occurs that is an essential process to allow onwards development and the transition of the maternal and paternal genomes from a gametic epigenetic state to an embryonic state. Firstly, the majority of the paternal genome (single copy (1C), as delivered by the mature sperm, is associated with protamines rather than histones. Protamines are a unique feature of the sperm chromatin that are understood to be important for the tight condensation and packaging of sperm DNA [5]. At the time of fertilization therefore, the paternal genome has only a small number of regions of the genome that are associated with histones. The residual regions of the sperm genome that remain associated with histones include regions that are poised for readiness in early development [56,77,127]. The sperm haploid genome is highly methylated with approximately 80%e90% of the CpG sites being methylated [72,90] while the DNA methylation levels of the maternal genome in the fertilised oocyte are approximately half of that observed in sperm [45,96,111]. Another epigenetic difference between the pronuclei is that the acetylated histones H3K27ac, H4K5ac and H4K16ac can only be detected in the paternal pronucleus in mouse zygotes [2,40,114]. In contrast, the maternal genome is associated with H3K9me3 in centromeric major satellites, H3K9me2 in minor satellites and numerous other histones and their modifications (H4ac, H3K4me1, H3K9me2/3, H3K27me1, H4K20me3) [2,53,104,108]. Interestingly, histone H3 modifications appear to play an important role in regulating DNA methylation in oocytes whereby the CpG islands (CGIs) that become methylated have reduced amounts of H3K4me2 and H3K4me3, while in contrast, H3K36me3 is increased at the CGIs that become methylated in growing oocytes [115]. Knockout experiments performed on the H3K4 demethylases KDM1A and KDM1B show that demethylation of H3K is an essential requirement for the establishment of DNA methylation at CGIs in oocytes.

The zygote: a hotbed of epigenetic activity Upon fertilisation, the sperm DNA decondenses and the protamines in sperm chromatin are rapidly replaced with histones. In contrast to paternal genome, the maternal genome from the oocyte is arrested at metaphase II and its genome is packaged with histones. After fertilization the maternal genome completes meiosis. After fertilisation, the formation of the maternal and the paternal pronuclei is an essential process that is divided into 6 stages, namely PN0 through to PN5. At PN0 the sperm has entered and the oocyte has completed meiosis. By PN1 the process of epigenetic remodeling is initiated, whereby the paternal pronulei starts to undergo demethylation, a process that continues through to PN2 [28]. Maternal histones replace protamines before DNA replication takes place.

Regulation of DNA methylation in the zygote By the end of pronuclear maturation (PN4/PN5), DNA methylation (5MeC) is removed from the male pronucleus. In the mouse, the male pronucleus is demethylated within 4 h of fertilization and loss of methylation in the embryo continues thereafter up to the morula stage [135]. It should be noted that there is currently significant debate about the demethylation process that occurs on the pronuclei,

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especially as to whether the process is active or passive (or both). The rapid demethylation of the paternal pronucleus is understood to be largely driven by an active mechanism although there may be also be evidence that it may also happens to a degree by a passive mechanism, that is, DNA replication without maintenance methylation [1,33,34]. In addition, demethylation of the maternal genome, once thought to occur exclusively by a passive mechanism may also be driven at least in part by active mechanisms. It is clear however that in the mouse zygote, the paternal genome appears to undergo rapid demethylation while in contrast, the maternal genome at these stages remains relatively unchanged, undergoing more gradual demethylation during the cleavage stages. The difference in DNA demethylation events on the pronuclei is likely to be determined by differential initial epigenetic states of the maternal and paternal pronuclei. In the zygote, developmental pluripotency-associated protein Dppa3/PGC7/Stella is important in protecting the maternal genome from excessive active demethylation caused via Tet3 mediated oxidation. This includes protection of maternal (oocyte-derived) methylation at imprinted gene regions, marks which are established in the female germline and that need to be retained so that the parental alleles can be appropriately recognised in the developing embryo [83]. Dppa3/PGC7/Stella binds to H3K9me2 in maternal chromatin and protects Tet3mediated conversion of 5mC to 5hmC. This mechanism whereby Dppa3/PGC7 binds to H3K9me2 may also protect paternal DMRs such as Rasgrf1 and H19 [84]. The histones that are retained in the sperm (see above) and their associated epigenetic marks are thought to be important in protecting paternal imprints from demethylation and for activation of paternal genes (H3K4 me3) [37,120].

Regulation of histones and histone modifications in the zygote During pronuclear maturation, significant chromatin reorganisation occurs in the paternal pronucleus. In addition to the genome-wide loss of DNA methylation, histone acquisition occurs toward the end of PN maturation. A critical process of de novo nucleosome assembly occurs during paternal PN formation and this is dependent on Hira (Histone Cell Cycle Regulator) and Histone H3.3 and leads to the establishment of de novo nucleosomes on the paternal pronucleus (the nucleosome is a fundamental component of chromatin and is comprised of two copies each of H2A, H2B, H3 and H4). Of the maternal histones, H3.3, as opposed to H3.1 or H3.2, becomes incorporated into the paternal pronucleus by virtue of the actions of its chaperone Hira and this appears to be a critical event in the establishment of chromatin on the paternal genome during the oocyte to embryo transition [49,65]. Histone H2A and H2B variants may also be important in genomic remodeling and sperm decondensation. The histone H2A variant H2A.Z may play an important role in conjunction with histone H3.3, acting to enable gene expression [39]. The acquisition of histones by the male pronucleus leads an accumulation of the epigenetic marks carried with them including H3K9me1 and H3K9me2, and H3K27me2/3. In contrast, between pronuclear stages PN0 to PN5 the maternal pronucleus which is rich in histone epigenetic marks, remains relatively unchanged [3,97]. There is also differential marking between the maternal and paternal pronuclei with respect to the histone 3 lysine 9 (H3K9) methylation state [55,66]. The phosphorylated Histone H2A variant gH2A.X is more prominent in the paternal pronucleus than the maternal pronucleus but becomes depleted after the first cell division [82,125]. By the end of PN5 therefore, the epigenetic configuration of the male and female pronuclei are strikingly different and this difference is referred to as ‘epigenetic asymmetry’. Other histones that may be important in reprograming are the TH2A and TH2B histones that were originally identified as testis-specific H2A/H2B variants. These histones are expressed in sperm, oocytes and one cell

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embryos are also a focus of research into understanding reprogramming [109]. In the blastocyst, the primary differentiation event leads to the creation of the trophectoderm and the inner cell mass (ICM) which are differentially marked by histone modifications. As early as the cleavage stage, there are differences in histone arginine (H3R26) methylation that may be important in regulating the developmental trajectory of cells to either ICM or TE lineage. Thus, the blastomeres with higher H3R26me are more likely to become ICM cells [119]. In the blastocyst, H3K27me1, me2, and me3 are present predominantly in the ICM [78]. The ICM and trophectoderm cell lineages of the mouse blastocyst are differentially marked by Histone H3 lysine 27 methylation at key developmental genes [21]. Histone modifications may be important in regulating specific genes associated with lineage specifying genes. Thus, H3K4me3 and H3K27me3 are found at ICM-specific genes while TE-specific genes are marked and repressed with H3K27Me3 in the ICM lineage [92].

Detailed mapping of methylation changes in preimplantation development DNA methylation is highly dynamic during gametogenesis and preimplantation development. Our understanding of how DNA methylation is regulated during gametogenesis and preimplantation development has been driven and informed by a range of different methods of DNA methylation analysis. Until recently, and as described above, the most commonly used method of assessing DNA methylation in preimplantation embryos was by immunohistochemistry-based analysis of DNA methylation using antibodies directed against 5-methylcytosine. These studies were able to show general DNA methylation patterns and revealed that genome-wide erasure of DNA methylation occurs during preimplantation development [57]. However, these methods were unable to provide information on which sequences and genomic regions undergo methylation changes. More recently, nextgeneration sequencing-based technologies have allowed the exact patterns of DNA methylation reprogramming to be revealed in extremely fine detail. These technologies include reduced representation bisulphite sequencing (RRBS), whole genome bisulphite sequencing (WGBS) and variants of these methods that allow genome-wide methylation analysis in an individual embryo and in some cases, individual cells. This work culminated in 2014 with the publishing of a series of important papers detailing the methylation dynamics of the human preimplantation embryo [33,34,113, 88]. This work has added a considerable amount of extra detail to our understanding of epigenetic programming in preimplantation development. The assessment of DNA methylation in mouse preimplantation development by RRBS revealed (i) major reduction in methylation between the sperm and zygote, particularly at Long Interspersed Elements (LINEs), (ii) gradual reduction during mid-preimplantation and (iii) a major increase from the blastocyst ICM to the D6.5 post-implantation embryo [112]. Smallwood et al [111] used RRBS to identify that over 100os CGIs were methylated in mouse oocytes and the extent of preimplantation reprogramming of the CGIs was demonstrated by the fact that only around 15% of these CGIs retained 40% methylation in blastocysts. Gou et al. [33,34] performed genome-wide DNA methylation of human zygotes and preimplantation embryos using RRBS, to reveal that overall, the patterns of DNA methylation changes were in fact similar to the observations deduced from immunohistochemistry-based analysis on preimplantation embryos. Therefore, RRBS revealed that methylation decreases over preimplantation development and then de novo methylation occurs later in embryonic development as differentiation

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happens. However, using these novel sequencing techniques, it has been possible for the first time to identify exactly which regions of the genome undergo reprogramming and to extract significantly more detail on the reprograming process. In contrast to the mouse where genome-wide demethylation occurs by the 1 cell stage, in humans, it was observed to be mostly complete by the 2-cell stage. In the human zygote, both pronuclei were demethylated but the paternal pronucleus was demethylated to a greater extent. Furthermore, methylation profiles of individual transposable elements such as short interspersed nuclear elements (SINEs) and long interspersed nuclear elements (LINEs) differed across preimplantation development. Another RRBS study by Ref. [113] using RRBS analysis of human preimplantation embryos reached similar conclusions to Refs. [33,34] again noting differential methylation of the various retrotransposon classes throughout human preimplantation development. Okae et al [88] used WGBS to profile human oocytes, blastocysts and sperm and their data indicated that in humans, the maternal genome is demethylated to a lesser extent than in the mouse, with the blastocyst apparently retaining many methylation marks from the oocyte, while the paternal genome was globally demethylated. In agreement with the finding of [34] the study also showed differential methylation patterns between retrotransposon classes. Thus, the SINE-VNTR-Alu (SVA) class of retrotransposon (SVAs), particularly the SVA_A subfamily, and some LTRs containing CpG-rich VNTRs were more protected from demethylation compared to SINEs and LINEs and were thus more highly methylated in blastocysts. Zhu et al [132] used single cell DNA methylation profiling of human preimplantation embryos by post-bisulfite adaptor tagging (PBAT) DNA methylome analysis to investigate DNA methylation programming of the human preimplantation embryo, an approach allowing greater detail than previous studies and accordingly a more complex DNA methylation pattern was revealed for human preimplantation development. Three waves of demethylation were observed: (1) Wave 1: 10e12h after fertilization when methylation of the paternal genome decreased from 82.0% in the sperm to 52.9% in the early male pronucleus (PN). At the same time, methylation decreased slightly from 54.5% in the mature oocyte to 50.7% in the early female PN; (2) Wave 2: Global demethylation from the late zygote to the two-cell stage, (methylation decreased 49.9%e40.4%); (3) Wave 3: eight-cell to the morula stage, where methylation decreased 47.0%e35.1%. Uniquely, and in contrast to any prior reports using immunohistochemistry-based methods, this study also detected 2 robust waves of de novo methylation, particularly at repeat elements, (SINEs, long interspersed nuclear elements (LINEs), and long terminal repeats (LTRs). These waves occurred at the early male pronuclear to the mid-pronuclear stage and then again at the four-cell to the eight-cell stage. These periods of de novo methylation suggest that DNA methylation programming in early embryos is a balancing act between genomewide demethylation (particularly on the paternal genome) with targeted remethylation at a restricted number of regions, particularly the repeat regions. The authors suggest that the de novo DNA methylation occurring at these repeat elements in the zygote and the mid-preimplantation stages is likely to function in suppressing the transcription and mobilisation of these elements, thus acting to minimise their effects in causing genomic instability.

Mediators of DNA methylation in gametes and preimplantation embryos There are several important factors that regulate the process of DNA methylation in gametes and in preimplantation embryos. A number of enzymes are involved in regulating DNA methylation. The DNA methyltransferases (Dnmts), are crucial to the regulation of DNA methylation in the oocyte and

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during the preimplantation stages. Dnmts add a methyl group on to the 5-position of the cytosine ring. DNA methyltransferases include Dnmt1, Dnmt3a, and Dnmt3b, all of which are essential for murine development [17]. Dnmt1, Dnmt3a, and Dnmt3b DNA methyltransferases have different functions. Dnmt1 is mostly involved in maintenance methylation, and is recruited to hemi-methylated DNA by accessory protein UHRF1 [13]. In contrast, Dnmt3a and Dnmt3b are mostly active as de novo methyltransferases (they can newly methylate cytosines, without a requirement for hemi-methylated DNA). Functional variety of Dnmts is enhanced by the expression of alternative transcript variants of these Dnmts in the oocyte/early embryo [32]. For example, Dnmt3b has over 30 isoforms. Different Dnmts and their various isoforms work cooperatively in regulating DNA methylation. In addition to Dnmt1, Dnmt3a, and Dnmt3b, Dnmt3L (Dnmt3-like) that is expressed in germ cells, preimplantation embryos and embryonic stem cells, has no methyltransferase activity, but appears to cooperate with the de novo methyltransferases Dnmt3a and Dnmt3b in the methylation process. In the mouse, a further DNA methyltransferase, termed Dnmtc is important for suppression of retrotransposons [7]. One further DNA methyltransferase, Dnmt2 appears to be involved in transfer RNA (tRNA) methylation and its role in DNA methylation is less clear [30]. The expression of DNMTs and their role in epigenetic regulation is critical and if disrupted can lead to disease such as cancer [131]. The oocyte itself is of fundamental importance in the regulation of DNA methylation patterns that are required for onwards development in the preimplantation embryo, the developing conceptus and the placenta. Thus, most imprinted genes inherit methylation imprints in the female germline. In the mouse, primary imprinting in the female germline is progressively established from the primordial through to the antral stages of follicle development [6,67,86,87]. Further indications of the importance of the process of DNA methylation the oocyte in are revealed by the observations in mouse oocytes that methylation by Dnmt3 methyltransferases is essential for trophoblast development [16]. Similar conclusions have been reported in humans, showing that oocyte-derived methylation marks persist into the placenta [102]. Dnmt3L (Dnmt3-like), has no methyltransferase activity, but is required for the establishment of maternal methylation imprints during murine oogenesis [14] but its role in human imprinting is less clear. Dnmt3L is also required for retaining hypomethylation of developmental genes in embryonic cells [85] and retrotransposon suppression in germ cells [64]. Cooperation of Dnmt3 family members, is important in the establishment of maternal genomic imprints and other methylation patterns in the oocyte [26,93,111,116]. Non-CpG methylation is accumulated in oocytes, particularly in gene bodies via Dnmt3a-Dnmt3L interaction [110] and appears to be an important of hallmark of oocyte maturation, including in human oocytes [129]. After implantation and beyond, DNA methylation is established in the epiblast by the de novo methyltransferases, Dnmt3a and Dnmt3b [89] and in the mouse, methylation is established within two days of implantation [83]. DNMT3B plays a major role in methylating CpG islands and both DNMT3A and DNMT3B methylate other regions of the genome including repetitive elements [4]. In mice, Dnmt3a2 is an important Dnmt3a isoform for de novo methylation [18].

The subcortical maternal complex is a key effector of epigenetic programming in the oocyte/early embryo Maternal Effect Genes that are expressed in the oocyte, are essential for the correct regulation of epigenetic programming early embryonic development. A number of these proteins are members of the subcortical maternal complex (SCMC), a multiprotein complex that is present in mammalian

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oocytes that is understood to have multiple functions including spindle formation and positioning, the regulation of translation, and epigenetic reprogramming of early embryos [11]. Therefore, disruption of the function of the SCMC, for example by virtue of a deleterious genetic mutation in one of its constituent members can lead to serious developmental consequences. In the mouse, the SCMC contains at least 4 proteins including Mater/Nlrp5, Filia/KHDC3, Floped (Ooep) and Tle6 and appears to be necessary for development beyond the 2-cell stage [58]. Crucially, the murine SCMC is required for formation of the cytoplasmic F-actin meshwork that controls the central position of the spindle and hence symmetric division of the zygote [130]. The complex is found in the subcortex of oocytes and early embryos. In mice the SCMC is composed of at least four proteins including Mater (Maternal Antigen That Embryos Require), Floped (Factor Located in Oocytes Permitting Embryonic Development-official name is Ooep), Tle6 (Transducin-Like Enhancer of Split 6) and Filia (official name Khdc3) as reported by Ref. [130]. The SCMC is required for the symmetric division of mouse zygotes, by regulating F-actin dynamics. In humans and sheep the known SCMC comprises of NLRP5 (official gene symbol for Mater), OOEP, TLE6 and KHDC3L proteins [10,133]. Proteomic studies have shown that SCMC proteins are among the most abundant in the oocyte proteome [122]. The SCMC proteins have been demonstrated to interact with each other [133]. The SCMC appears to be of critical importance in regulating epigenetic information. In humans, mutations in NLRP7 and KHDC3L genes, both of which are expressed in oocytes and members of the SCMC, leads to an inability to correctly establish and/or maintain DNA methylation at imprinted loci [80,95]. This condition, is called familial biparental hydatidiform mole (FBHM). In these recurrent molar pregnancies of biparental origin, the process of genomic imprinting [68] is disrupted. Furthermore, mutations in another oocyte gene NLRP5, that is also a member of the SCMC, are associated with Multilocus Imprinting Disturbance (MLID) in which there is a failure in maintenance of multiple imprinting marks [25]. To date there has been limited progress in understanding how the disruption of SCMC proteins can lead to epigenetic defects. That is, how the SCMC itself is connected mechanistically the processes of DNA methylation, histone modification and/or other epigenetic processes and what role does complex plays in shaping the epigenome. Only very recent evidence has reported that indicates that the murine NLRP2 protein, itself recently identified as part of the SCMC, may direct the subcellular localization of DNMT1 [70]. Therefore, deficiency in the SCMC components may affect interaction of the complex with key epigenetic regulators that are essential in the imprinting process. It is also possible that disruption of the SCMC may affect transcriptional processes in the oocyte, particularly transcription through IG DMRs, a mechanism that is required for imprint establishment [20]. It is also conceivable that asymmetric division of the zygote, caused by deficiencies in the SCMC could distort embryonic pre-patterning and/or lead to significant deficiencies in essential maternal gene transcripts or proteins in the affected blastomere(s) such as key epigenetic regulators and/or cell fate regulators in cleavage-stage embryo.

Demethylation events in the zygote and preimplantation embryo Perhaps the most striking feature of all the epigenetic reprogramming events in the zygote and during preimplantation development is the removal of DNA methylation (demethylation) and this has been a particular focus of study in recent years. Essentially, this erasure process removes the epigenetic features that were associated with the gametes in order that the totipotent state of the embryo can be created. The maternal and paternal genome appear to be demethylated to different extents, using

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different mechanisms and over a different time scale. Thus the maternal genome becomes demethylated mostly by passive demethylation. This is where DNA replication occurs without maintenance methylation occurring on the new strand, and is understood to be caused the exclusion of DNMT1 from the nucleus by UHRF1 (a hemi-methylated-CG-binding protein) and the Subcortical Maternal Complex [69]. In the mouse, rapid demethylation occurs from the paternal genome shortly after fertilisation by virtue of an active process. This active process is mediated in part by the TET3 (Tet Methylcytosine Dioxygenase 3), an enzyme that is expressed in oocytes and embryos and that can demethylate DNA by converting 5-methylcytosine into 5-hydroxymethylcytosine [31,47,124]. The ten-eleven translocation (TET) family of dioxygenases can act to cause further oxidation of 5hydroxymethylcytosine to form 5-formylcytosine (5-fC) and 5-carboxycytosine (5-caC) [100]. The role of the other ten-eleven translocation (TET) family members (TET1, TET2) in the preimplantation demethylation event is less clear although the reduced litter sizes and reduced body sizes observed in knockout experiments have indicated that TET1 may have some roles in embryonic development and in the gametes [23]. Other mediators of the active DNA methylation process may include members of the Base Excision Repair (BER) pathway. The demethylation could be mediated by deamination of methylated cytosine by either the AID enzyme (activation-induced deaminase) or the APOBEC1 enzyme (apolipoprotein B mRNA-edition enzyme catalytic polypeptide 1), which could induce subsequent conversion of the cytosine to thymidine. The BER could be activated by recognition of the T:G mismatch and act to replace the thymidine with an unmethylated cytosine. Another candidate for active demethylation is the GSE (gonad-specific expression gene) [38] As described above, there is some more recent evidence for active demethylation on the maternal genome too [33,105]. Interestingly, a small number of genomic regions, do not undergo the demethylation process and are thus able to retain methylation. There are regions within imprinted genes (genes that are expressed from a single parental allele [8] for example that do not undergo demethylation. Imprinted genes have different methylation patterns (in Differentially Methylated Regions, DMRs) on the maternal and paternal alleles that play an important regulatory role. DMRs typically consist of stretches of differentially methylated CpG sites that are close to an imprinted gene, and this epigenetic information regulates allele-specific gene expression. A germline DMR will therefore have a different methylation pattern in the sperm than in the oocyte, and this differential marking needs to be protected from demethylation so that it can be recognised in the preimplantation embryo. One critical player in maintaining DNA methylation at the imprinted genes is an oocyte-specific form of Dnmt1, termed Dnmt1o [75] that is synthesized and stored in the cytoplasm of the oocyte and is used to maintain maternal methylation imprints in 8-cell stage mouse embryos a time when genome-wide erasure of DNA methylation is occurring at other regions of the genome [44]. Other factors in addition to the aforementioned UHRF1 are critical in this process: Trim28/Kap1 recruits Dnmt1 [76] and ZFP57 [62,98] is required for targeting these factors to imprinted gene regions. Moreover, in addition to regulating imprinting, Dnmt1o is also required for other roles in placental methylation since Dnmt1o deficiency causes placental DNA hypomethylation [73].

Species-specific differences in reprogramming The investigation of a variety of animal models has revealed that there are species-specific differences in preimplantation reprogramming. Originally, immunohistochemistry-based studies revealed that DNA methylation programming in ovine and rabbit preimplantation embryos differed significantly

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from other species (bovine, human, mouse). Therefore, in the ovine and rabbit zygote, there was no significant demethylation in either of the parental pronuclei [9]. Furthermore, only limited demethylation of the ovine embryonic genome occurs subsequently between the two- and eight-cell stages with no de novo methylation observed at/after the blastocyst stage. Similar results have been observed in goat zygotes [46]. Selective demethylation of the male pronucleus has been reported in bovine, mouse, human and pig zygotes [9]. In the ovine zygote, neither pronuclei undergo significant demethylation [9,48] although those results have been contested [71]. In contrast, partial asymmetric demethylation occurs in bovine zygotes [9,91], whereas full asymmetric demethylation occurs in mouse and humans [9,24,48]. Preimplantation methylation programming also differs: sheep embryos do not undergo major demethylation in cleavage stages [9]. Bovine embryos undergo de novo methylation between the 8e16 cells stages [9,24] or possibly earlier but no obvious de novo methylation is seen in sheep embryos. These observations are based on immunohistochemistry-based studies. The example of the sheep embryo alone raises fascinating questions with respect how pluripotency is established with such limited methylation erasure [94] and emboldens the need to investigate the mechanisms of developmental DNA methylation reprogramming in other mammals.

RNA as a mediator of epigenetic events in gametes and preimplantation embryos RNA also plays a significant role in epigenetic reprogramming during gametogenesis and preimplantation development and this complex area can only briefly be described here. In addition to delivering DNA to the oocyte, the sperm nucleus itself harbors many RNA species including messenger (mRNAs), short non-coding RNAs (sncRNAs) including Piwi-interacting RNA (piRNAs), microRNAs (miRNAs) and tsRNAs (tRNA-derived small RNAs) [19,107]. These RNAs play important roles in gametogenesis and fertilisation [35]. For example, piRNAs play an essential role in fertility by suppressing retrotransposons in the germ line and do so via an epigenetic mechanism [29]. microRNAs (miRNAs), which are endogenous noncoding single-stranded RNA molecules that are approximately 22 nucleotides in length, play critical roles in many processes during gametogenesis and preimplantation development [43]. miRNAs regulate gene expression and indeed, a typical miRNA may regulate the expression of hundreds of target genes. Accordingly, the disruption of microRNAs (miRNAs) that function in reproductive processes may lead to infertility [63,117,126]. Further complexity of mammalian preimplantation development has been revealed using a method called single-cell universal poly(A)-independent RNA sequencing [27]. This method allows the sequencing of both polyadenylated and non-polyadenylated RNA transcripts from single cells. The study revealed the expression of 2891 circular RNAs (circRNAs) in mouse preimplantation embryos. Circular RNAs (circRNAs) are a newly described class of non-polyadenylated non-coding RNAs, most of which are actually formed by the exons of coding genes, and some even originate from introns. CircRNAs are believed to have important cellular functions such as the binding and repressing of microRNA (miRNAs), which are themselves important in modulating gene expression. In human oocytes and preimplantation embryos, over 10,000 circRNAs were identified, arising from nearly 3000 hosting genes [22]. The use of highly sensitive RNA sequencing methodologies has allowed the detailed profiling of the expression of non-coding RNAs in gametes and preimplantation embryos. These include long

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noncoding RNAs (lncRNAs), which are untranslated RNAs of 200 nucleotides or more in length which have diverse functions in mammalian cells [81] including the regulation of gene expression and epigenetic information. Significantly [128], used sensitive RNA-sequencing to detect over 18,000 lncRNAs in human preimplantation embryos of which 2733 were novel lncRNAs. The detection of such a large proportion of the known and novel human lncRNAs in early embryos suggests they may have critical functions in early development. In the same study, 7214 known lncRNAs and 1487 novel lncRNAs were identified in human MII oocytes. The importance of long non-coding RNAs in the early embryo is illustrated by the case of the lncRNA Xist/XIST (X-inactive specific transcript) which is expressed in both mouse and human preimplantation embryos and is required for the regulation of X chromosome inactivation (XCI). This is the process that achieves dosage compensation for the X chromosome between males and females [101]. Another form of lncRNA are the Long intergenic noncoding RNAs (lincRNAs). Essentially these are lncRNAs that do not extend into protein-coding genes [99] and these lincRNAs appear to be critical in early embryonic development. To exemplify their importance, the long intergenic noncoding RNA LincGET is essential for early preimplantation development, acting as a transcription factor and as a regulator of RNA splicing [123]. Other types of lncRNA are important in early development. In a study by Ref. [36]; directional RNA-sequencing revealed that more than 1000 lncRNA/mRNA gene pairs were expressed at zygotic gene activation (ZGA) and may play essential functions in the activation of zygotic genes. These bidirectional promoter-associated noncoding RNAs (pancRNAs) are poly(A)þ RNAs that are associated with the upregulation of their cognate gene transcripts and are capable of regulating epigenetic information by causing changes in local DNA methylation [118]. To conclude, RNAs are critically important contributors to the epigenetic regulation of reproductive processes and hence fertility.

Conclusions The preimplantation stages are a very dynamic phase of development with respect to epigenetic programming and our knowledge is improving, but is far from complete. Interpretations can be limited by the technology/method used during analysis and the nature of the reprogramming varies according the species of the embryos being studied. The use of better methodology can have profound impacts on our interpretations of how epigenetic information is regulated during mammalian preimplantation development. Thus, the enhanced sensitivity and detail afforded by sequencing-based methylation analysis is already providing a much more comprehensive picture of epigenetic reprogramming during preimplantation development. The finding of waves of de novo methylation may have ramifications for example in the formulation of culture media that is used for human assisted reproduction, suggesting the requirement for inclusion of an adequate supply of methyl donors. Future research using NGSbased methods will focus on how chromatin is regulated during preimplantation development, and how the transcriptome and phases of embryonic gene activation work alongside the changes in the epigenome. In addition, use of gene knockdown and knockout experiments for example will assist our understanding how epigenetic mediators contribute to the mechanisms of reprogramming in preimplantation development. Another focus will be to study whether infertility and/or assisted reproduction in humans can cause detrimental effects on epigenetic reprograming, what mechanisms are likely to be disrupted and what measures can be used to reduce risks.

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CHAPTER

Epigenetic regulation during placentation

7

Divika Sapehiaa,
, Shilpa Thakurb,
, Beenish Rahatc, Aatish Mahajana, Parampal Singha, Jyotdeep Kaura a

Department of Biochemistry, Postgraduate Institute of Medical Education and Research, Chandigarh, India; bNational Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, United States; c Division of Intramural Research, Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, United States

The placenta The placenta is a transient feto-maternal organ and its formation takes place during pregnancy. It acts as an interface between mother and fetus. It assists in carrying out vital functions of the fetus such as nutrition, protection, respiration and excretion including feto-maternal tolerance, which allows the immune system of the mother to tolerate the genetically different fetus.

Description of the placenta and placental cells Structure of human placenta The term placenta comes from Latin word plakuos which means “flat cake” because of its gross anatomical appearance. By the end of pregnancy, the diameter of placenta is about 15e20 cm, thickness is 2e3 cm and it weighs nearly 500 g, which is equal to 1/6 of the fetal weight and is delivered with fetus at the time of birth. During period of gestation it provides nutrition from about 100 to 150 maternal uterine spiral arteries situated in the basal plate. Besides providing nutrition, placenta also plays a role in gas exchange, removal of wastes and comprises of hematopoietic stem cells [1]. Both fetal and maternal components are involved in the formation of placenta. The former comprises of placental disc, fetal membranes (i.e., amniotic and chorionic membranes) and the umbilical cord, whereas the later is derived from the maternal endometrium and is termed as the decidua.

Placental cells Placenta is made up of three different layers, with following different types of cells in each layer: 1. Cytotrophoblast and Syncytiotrophoblast cells form an envelope around the surface of the villous tree in placenta and are washed by the blood of the mother within the intervillous space.


The authors have contributed equally.

Epigenetics and Reproductive Health. https://doi.org/10.1016/B978-0-12-819753-0.00007-6 Copyright © 2021 Elsevier Inc. All rights reserved.

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2. Mesenchymal, Hoffbauer (macrophages derived from mesenchyma) and Fibroblasts cells that are present in villous core stroma. 3. Fetal vascular cells like Pericytes and Endothelial cells which are vascular smooth muscle cells.

Cytotrophoblasts and syncytiotrophoblasts Outer layer of a blastocyst is formed by the trophoblast cells which supplies nutrients to the developing embryo. This layer further differentiates into two types of cells i.e., cytotrophoblasts (undifferentiated) and syncytiotrophoblasts (fully differentiated). An undifferentiated cytotrophoblastic stem cell will differentiate into an extra villous cytotrophoblast intermediate and into an interstitial cytotrophoblast. Interstitial cytotrophoblasts continuously differentiate into syncytiotrophoblasts and form syncytium during the course of development.

Villous core stroma cells Early villous core fetal cells are derived from mesenchymal stem cells. These cells differentiate into hemangioblastic cell cords, which are considered to be the precursors of capillary endothelial cells and hematopoietic stem cells in early stage of placental development [2]. Hofbauer cells are antigenpresenting cells in the placenta villous stroma and were discovered by Hofbauer in 1903. These cells help in differentiation of trophoblasts and angiogenesis by producing different growth factors and cytokines such as fibroblast growth factor (FGF), vascular endothelial growth factor (VEGF) and vasculotropin [3e6].

Pericytes and endothelial cells Perivascular cells (also known as pericytes) possess dendritic processes that cover fetal capillary endothelium and venules. Microvascular integrity of placenta and vessel stability is maintained by the pericytes [7] by restricting endothelial cell migration and proliferation which is induced by the growth factors [8,9]. Microvascular endothelial cells of the placenta regulate the angiogenic activity during the course of pregnancy. Angiogenic activity is increased during the first and second trimesters as it is necessary for rapid expansion of placenta, but as the pregnancy approaches to term, microvascular cells stop angiogenesis that causes placental growth arrest. Thus, placental microvascular endothelial should function properly for successful pregnancy.

How does the placenta form? The formation of placenta occurs when the outer trophoblastic covering of a developing blastocyst comes in contact with the uterine mucosa. This newly formed connection between the mother and the fetus grows continuously at a high rate during pregnancy and also mediates exchange between the two.

Human placental development from fertilization to full term Early placenta formation Fertilisation to 7 day: Around 6e7 days after conception the development of human placenta starts by the attachment of the blastocyst to the uterine surface epithelium known as implantation. At this stage, the blastocyst is formed which consists of two main parts one is a single layered epithelial outer cover trophoblast, and other is inner cell mass, known as embryo blast [10]. The precursor of the epithelial parts of feto-maternal barrier of placenta is trophoblast, whereas embryo blast contributes in the formation of placental mesenchyme and the fetal vascular Fig. 7.1 (a). Due to continuous proliferation

The placenta

119

FIG. 7.1 Stages of early placental development (A) Implantation- Blastocyst consisting of blastocoel (fluid- filled cavity) and embryoblast, implantation takes place here when trophoblast cells which are in direct contact to the uterine epithelium differentiate and their syncytial fusion develop synctiotrophoblast. Synctiotrophoblast cells penetrate uterine epithelium for implanting into maternal tissue. (b) Onset of lacunar period- Confluent vacuoles starts appearing in the syncytiotrophoblastic mass at the implantation pole, further in the lacunar stage(L) fluid-filled spaces lacunae develop in the mass of Synctiotrophoblast (S). (C) Villous stage- Lacunae grow together to generate intervillous space (IVS), cytotrophoblast (C) begins to penetrate into the syncytiotrophoblast (S) and the trophoblast shell into the maternal decidua to form extra villous trophoblast (EVT). After few days, the extraembryonic mesenchyme (M) starts to penetrate the syncytiotrophoblast as well and displaces the cytotrophoblast further leads to the development of the secondary villi. (created with biorender.com).

inner cellular layer of trophoblast and syncytial fusion of few daughter cells there is a rapid and enormous increase in volume of the syncytiotrophoblastic mass (prelacunar period) Fig. 7.1 (b). Syncytiotrophoblastic mass can invade maternal capillaries and rupture the uterine wall for establishing a linking interface between embryonic extracellular fluid and maternal blood. The

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Chapter 7 Epigenetic regulation during placentation

formation of syncytiotrophoblastic first occurs at the invading pole of the blastocyst and subsequently allows the entry of the blastocyst in the maternal endometrial stroma [11]. Day 8 to 12: On day 8 after conception, a system of confluent vacuoles starts appearing in the syncytiotrophoblastic mass at the implantation pole. The appearance of these vacuoles is a sign of the onset of the lacunar period lasting from day 8 to 13th after conception. Now the lacunae are surrounded by the pillars of syncytiotrophoblast and the lamellae known as trabeculae. This system of lacunae and trabeculae is covered by two layers: one facing the endometrium (basal layer) known as trophoblastic shell and other facing the blast ocystic cavity (super facial layer) called primary chorionic plate. Day 12: Starting at the 12th day after conception at the primary chorionic plate, dividing cytotrophoblasts grow into the syncytial trabeculae and finally enters the trophoblastic shell. The branches of cytotrophoblasts which protrude into the lacunae and end blindly are called primary villi while their parent trabeculae from which they are derived, are known as anchoring villi because they connect the villous system with the trophoblastic shell. The expanding lacunar system along with the appearance of the first primary villi, is called the intervillous space [12] Fig. 7.1 (c). On the 12th day in parallel to these events, the cells of the trophoblastic shell invade the maternal endometrium vessels as well as uterine glands. At this stage and trophoblast plugs can be observed in spiral arteries due to the progression of trophoblastic invasion in the developing placental bed. Trophoblastic plugs have the ability to block the entry of blood cells in the uteroplacental circulation particularly at early pregnancy stages. Meanwhile, the blastocyst is now surrounded by endometrial stroma from all sides completely embedded in the endometrium [13].

Transition to the primitive villous tree After Day 12: At day 14 after conception, mesenchymal cells begin spreading out from the embryonic disk forming a loose network of branching cells, the extraembryonic mesenchyme. The movement of mesenchymal cells adds one more layer in the primary chorionic plate. So, the primary chorionic plate now consists of following three layers: • • •

the newly formed and added mesenchymal layer, a middle layer made up of cytotrophoblast, and a layer of syncytiotrophoblast which is facing the intervillous space.

Day 15 to 20: From the 15th to 20thday cells of extraembryonic mesenchyme divide into the center of the primary villi and establish a connective tissue core and transform them into secondary villi. The parts of the anchoring villi which is joined to trophoblastic shell remain merely trophoblastic because mesenchymal differentiation never reaches the trophoblastic shell. These trophoblastic cell columns are consisting of a voluminous cytotrophoblastic core [14] In addition to this, in a process known as “Trophoblast Invasion” this cytotrophoblastic core deeply erode into the endometrium and form an admixture with maternal tissue componentsdthe so-called Junctional zone [15]. On the appearance of first mesenchyme within first few hours inside the primary villi, some of the mesenchymal cells transforms into macrophages that secrete angiogenic growth factors. After day 20: Around 20th day inside the villi, the first fetal capillaries appears [16]. Thus, from the third week onwards, placental villi are changed into tertiary villi by the action of these capillaries. The tertiary villi already consist of the all the basic constituents of the placental barrier like mesenchyme, trophoblastic epithelial layers and vascular networks.

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Meanwhile, the fetal vascularised allantois comes in the vicinity of the chorionic plate [17]. Allantoic vessels divides over the villi which are originating directly from the chorionic plate and anastomose with the local intravillous capillaries. In this way around the end of the fifth week a complete fetoplacental circulation is established [18].

Establishment of the maternofetal barrier Because of the appearance of tertiary villi and the intervillous space the maternofetal barrier is defined now. The maternofetal barrier separating fetal blood from intervillous space is made up of the following layers: (1) (2) (3) (4) (5)

a layer of continuous syncytiotrophoblast, covering the intervillous space a complete layer of cytotrophoblast initially (in the 1st trimester) a trophoblastic basal layer layer derived from the extraembryonic mesoderm; and fetal endothelium layer, only in the last trimester.

This layering system at the feto-maternal barrier is constant factor throughout pregnancy and barrier undergoes many changes in the following months of pregnancy [18], the thickness of the two trophoblast layers decreases from more than 15 mm to a mean of 4.1 mm [19].

Placental blood circulation and spiral artery remodeling Placenta is unique well vascularised organ which is derived from the close relationship between mother and fetus, so the blood circulation of placenta is composed of both fetal and maternal circulatory systems. Based on this placental blood circulation is divided into two separate systems: fetalplacental and maternal placental circulation systems.

Maternal-placental circulation systems Maternal circulation or Uteroplacental circulation is mostly developed after the end of the first trimester. The maternal blood enters the placenta through the basal plate endometrial arteries (spiral arteries), traversing into the intervillous spaces, and flows around the villi. The maternal blood perfusing around the villi exchange nutrients and oxygen with fetal blood and drains back through the placenta intervillous space into the venous openings in the basal plate, to returns into the maternal systemic circulation through uterine veins. The flow of maternal blood into the placenta is propelled by maternal arterial pressure because of the low-resistance nature of uteroplacental vessels, which adapt and accommodate the massive increase in uterine perfusion during rest of the course of pregnancy [20]. The volume of maternal blood during pregnancy starts increasing from 6e8th week to up to a maximum approximately at 32e34 weeks and then relatively remains constant until term. In general, as compared to a non-pregnant state maternal blood (plasma) volume in pregnancy is increased up to 40%e50% near term. In a study by Gowland et al. in human placenta from 20 weeks of gestational age maternal blood perfusion was analysed up to term using echo planar imaging (EPI) and it was found that the average perfusion rate in normal pregnancy was about 176  24 mL/100 g/min [21].

Spiral artery remodeling A key event in process of placental development is when blood vessels are remodeled in the lining of the uterus for increasing the blood supply to placenta, through a process known as spiral artery

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remodeling. In spiral artery remodeling, cytotrophoblast cells invade the space mostly around the spiral artery and replace cells which normally lining the vessel. This remodeling results into the larger vessels which optimizes the blood supply to the placenta. The placental blood flow increased throughout pregnancy and reaches to about 600e700 mL/min (80% of the uterine perfusion) at term. There is also a huge change in the blood pressure from uterine arteries to intervillous spaces, it is about only 10 mmHg in intervillous space, 70 mmHg in spiral arteries and 80e100 mmHg within uterine arteries. This gradient of blood pressure between uterine arteries and placental intervillous space allow the maternal blood to exchange nutrients and gases with fetal blood in 2e3 min.

Fetal placental circulation system Fetal blood flows via umbilical arteries, in order to reach the villous capillaries through finer vessels of chorionic plate. The fetal surface is connected to blood circulation by two umbilical arteries (UA), which bring deoxygenated blood to the terminal villi. These terminal villi of placenta that are in direct contact with oxygen-rich maternal blood in the intervillous space, where the gas exchange takes place. The umbilical arteries are the strong muscular vessels, having a mean diameter of about 1.1e4.2 mm between 15 and 40 weeks’ of pregnancy (ultra-sonographically measured) [22] and their size increases up to approximately 2.5 mm at term [23]. Umbilical arteries are the longest blood vessels in the human usually do not divide along the length of the umbilical cord, but they only branch over the fetal surface of placenta to form chorionic plate arteries, to provide blood supply to the terminal villi. The oxygenated nutrient-rich blood returns to the fetus via a single umbilical vein.

Placental physiology The placenta performs a number of vital physiologic functions and the most crucial one is delivery of oxygen and nutrients and the removal of waste from the developing fetus. The growth of the fetus is dependent on this complex interaction between mother and fetus. Some of the well-known physiological functions of placenta are:

Feto-maternal exchange through placenta The exchange processes in placenta occur via classic membranous transport mechanisms through passive and simplified transport as explained below: In Passive transport, where the movement occurs without energy consumption, movement of fat-dissolvable and non-polar molecules follow simple diffusion. Diffusion of oxygen, carbon dioxide, fats occurs from the side with the higher concentration to the lower, until a balance is achieved. Except water which enters the placenta through specialized pores by osmosis [24] known as aquaporines or water channels [25]. In Simplified transport, transition is there from lower to higher solute concentration, with the help of transport molecules (e.g., glucose) [26] or using energy. Transport against a concentration gradient through the cellular membrane using energy (Naþ/Kþ or Caþþ) occurs in active transport whereas in vesicular transport macro-molecules are captured by microvilli are absorbed or repelled in the cells by Endocytosis or Exocytosis. The placental exchange surface is enlarged from 5 m2 at 28 weeks to roughly 12 m2 shortly before delivery [27].

Breathing function The placenta can play the role of “fetal lungs”, but is actually 15 times less efficient than the real lungs (with equivalent weight of tissue). The respiratory function of the placenta makes possible removal of

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carbon dioxide and supply of oxygen to the fetus [28]. The exchange of carbon dioxide and oxygen takes place between the blood of the umbilicales which is oxygen-poor and maternal oxygen-rich blood. The diffusion of oxygen occurs from the maternal into the fetal circulation system (PO2 maternal > PO2 fetal) and the partial pressure of carbon dioxide which is elevated in fetal blood, follows a reversed gradient. The oxygen saturated blood returns via the umbilical vein to the fetus, while oxygen-poor maternal blood flows back into the uterine veins.

Nutritive and excretory functions The maternal transfer of nutrients guarantees the energy required for fetal growth. Water a universal solvent diffuses by osmolar gradient into the placenta [25]. Its exchange increases during the gestation up to the 35th week (3.5 L/day) and helps in the transfer of other electrolytes too. Glucose is the main source of energy for fetus and passes via simplified transport into placenta. The sugar concentration in fetal blood is 2/3 of the mother’s and totally dependent on it. Placenta can store and synthesize glycogen at the trophoblastic level in order to meet the requirement of local glucose through glycogenolysis. However, important alterations in metabolism of carbohydrates are there in mother during the gestation to cope with the fetal needs. Due to the impaired insulin sensitivity in tissues, pregnancy is “diabetogenous” for mother [29]. Water-soluble vitamins are also transported through the placental membrane to fetus whereas there is a quite low amount of the fat-soluble vitamins (A, D, E and K) in the fetal circulation [30].

Endocrinal function The placenta act as a large endocrine gland especially due to the presence of syncytiotrophoblast cells. At the beginning of the pregnancy the corpus luteum ensure synthesis of estrogen and progesterone is further maintained by the human chorion-gonadotropin (HCG). But with further progression of time the activity of the corpus luteum decreases at the onset of eighth week and replaced entirely by placenta at the end of the first trimester. Throughout the gestation the hormone concentration in the maternal blood is maintained by the cooperative function of hypophysial and fetal suprarenal as well as placental hormones [31,32].

Epigenetic mechanisms in placental development Epigenetic can be defined as heritable alterations in the gene expression which are independent of any changes in the DNA sequence [33]. These epigenetic modifications can occur through different mechanisms which include DNA methylation, the action of non-coding RNAs and histone modifications [34]. All of these mechanisms have been reported to regulate the processes of normal placental development and alterations in either of these mechanisms can lead to abnormal placentation or pregnancy-related disorders [33,34].

DNA methylation DNA methylation is the most-studied epigenetic modification. Usually, DNA methylation is associated with transcriptional repression, however, in certain cases, it can also lead to transcriptional activation [35]. Placenta is known as a hypomethylated tissue, characterized by low DNA methylation levels in comparison to other healthy tissues [36]. During early stages of mammalian development, nonimprinted genes undergo global methylation changes which are associated with active and passive

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phases of DNA demethylation followed by de novo methylation. This global reprogramming occurs after fertilization and the de novo methylation takes place only in the ICM and not in trophectoderm, which stays hypomethylated. These methylation differences are maintained throughout the pregnancy which makes embryo hypermethylated in comparison to placenta [34]. Despite the hypomethylation of placenta, DNA methylation has been reported to play crucial roles at different stages of placental development and there are numerous studies where alterations in DNA methylation status in the placenta have been associated with pregnancy complications [37e42]. The analysis of DNA methylation of placenta is difficult owing to the presence of various types of cells. The most common part of placenta that has been used for methylation studies is chorionic villi, which consists all types of trophoblast cells and inner cell mass (ICM)- derived cells (extraembryonic mesoderm and endoderm progenitor cells) [35,36]. The different cell types of placenta have been characterized by different methylation profiles as reported in a study where DNA methylation of cytotrophoblasts (CTB), fibroblasts, and whole placental villi were compared among each other. The study showed that methylation profile of CTB and placental villi was relatively similar in comparison to fibroblasts [43]. Besides this, differences in DNA methylation have been observed among various types of trophoblasts and were correlated with trophoblast differentiation. Analysis of global DNA methylation in three trophoblast populations- (1) side population trophoblasts (trophoblasts stem cell population); (2) CTB (highly proliferative progenitor population); and (3) extra-villous trophoblasts, EVT (terminally differentiated population)-isolated from first trimester placenta, showed distinctive methylation profile of each of these populations. In comparison to CTB and/or EVT, side populations have differential methylation of the genes and miRs, which play roles in differentiation and cell cycle regulation. Moreover, the study identified 41 genes which were hypomethylated and were characterized by higher expression in EVT in comparison to CTB. These genes have function in epithelial to mesenchymal transition and/or in metastatic cancer pathways, which demonstrate their role in trophoblast differentiation and invasion [44]. Another study has shown differences in the 5-methyl cytosine (5-mC) levels between CTB and STB [45]. Overall, differences in DNA methylation have been observed among different cell types of placenta which point toward the significance of this epigenetic modification in the placentation process. The importance of DNA methylation in normal placental development has been proven through mice studies where knock out of DNMT1 and DNMT3L functions lead to severe developmental defects. The targeted mutation of the DNMT enzyme caused abnormal development of embryos which did not survive past mid-gestation [46]. Knock out of DNMT3L in mice lead to alterations in the maternal methylation imprints in both embryos and placenta. Moreover, loss of DNMT3L in these mice lead to defective labyrinth formation, overproduction of trophoblast giant cells, poor attachment of chorion layer with the ectoplacental cone, poor vessel outgrowth, and reduced spongiotrophoblast layer formation. The study also reported arrest of trophoblast proliferation without inducing apoptosis [47]. All these observations highlight the important of DNMTs and methylation in theplacental development. DNA methylation in the placenta is affected by gestational age. Analysis of global DNA methylation in the placentas of different gestational stages reported an increase in methylated cytosine levels with gestational age, with highest methylation in third-trimester and lowest in the first-trimester [48,49]. This gestational age-dependent increase in methylation levels might be responsible for the changes in cell composition and/or differentiation of the placental cells over time to perform certain functions [35]. For instance, a group of genes which were observed to be differentially methylated among the three trimesters are known to play a crucial role in the regulation of immune system [48].

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The variations in their methylation status with gestational age suggest their involvement in the modulation of the immune functions in the placenta during pregnancy. Effects of DNA methylation on normal placental functions have been studied through various in-vitro and in vivo studies. Differentiation of rat trophoblast cells have been associated with changes in DNA methylation patterns [50]. Besides, DNA methylation has also been shown to regulate important trophoblastic functions like invasion, migration and epithelial to mesenchymal transition in trophoblast cells [51,52]. Inhibition of DNA methylation has been shown to reduce the proliferation of trophoblast cells and loss of placental mass in rats [53,54]. Moreover, DNA methylation has been demonstrated as an important epigenetic mechanism in the regulation of various placental genes like Maspin [55], CREB5 [56], lncRNA MEG8 [57], tissue inhibitor of metalloproteinase 3 (TIMP3) [58], matrix metalloproteinase 9 (MMP9) [59], Oct4 [60], LINE1 elements [61], adenomatous polyposis coli (APC) [62], c-myc [63], telomerase (TERT) [63], and Nanog [64]. These genes play crucial roles in placental development/functions and alterations in their expression have the ability to cause abruption in normal placentation process which can lead to placental disorders.

Non-coding RNAs ncRNAs are the RNA molecules that are not translated into proteins. The function of these RNAs is to regulate gene expression at transcriptional and/or post-transcriptional level. The ncRNAs which are involved in epigenetic regulation of genes include microRNAs (miRs), long non-coding RNAs (lncRNAs), small-interfering RNAs (siRNAs) and Piwi-interacting RNAs (piRNAs) [65]. To date, the majority of the studies in regard to the role of ncRNAs in placental development have been focused on miRs and lncRNAs.

miRs miRs are small single-stranded RNA molecules that play an important role in regulation of gene expression in a variety of biological processes. To date, there have been several studies suggesting the role of miRs during different processes of placental development which includes trophoblast proliferation, differentiation, invasion, migration, vasculogenesis, and angiogenesis [66].

miRs in trophoblast proliferation The proliferation of trophoblast cells in a regulated fashion is very crucial for normal placental development. Compelling proof of the involvement of miRs in the trophoblast proliferation comes from a study where Dicer-the miR processing enzyme-was targeted in the first-trimester placental tissue. The study also reported Dicer expression in CTB, but not in syncytiotrophoblast (STB), which are terminally differentiated trophoblast cells. Knockdown of Dicer in first trimester placental tissue resulted in a global reduction of miR levels. The silencing of this enzyme promoted proliferation of CTB cells, suggesting the role of miRs in regulation of trophoblast proliferation. In addition to this, knockdown of Dicer resulted in increased expression of ERK and SHP-2 levels, both of which can enhance mitogenic signaling [67]. The exact function of most of these miRs in trophoblast proliferation is not known. There are many studies where the role of different miRs in placental development has been examined (Table 7.1). miR-378a-5p has been shown to promote trophoblast proliferation. The primary target of this miR is Nodal, which is a member of transforming growth factor-b (TGF- b) family. Nodal is known to inhibit proliferation, migration, and invasion of trophoblast cells while promoting their

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Table 7.1 The table summarizes the role of various miRs, their targets and role in placental development. miRs miR-675

miR-376c miR-378a-5p

miR-141 miR-155 miR-144 miR-377 let-7a miR-195

miR-20a

miR-101 miR-128a miR-29b

miR-30a-3p miR-519d-3p Chromosome 19 miR cluster (C19MC) miR-17-92 cluster

Targets Nodal Modulator-1 (NOMO1), Insulin-like growth factor-1 (IGF-1) Activin receptor-like kinase 5 (ALK5), ALK7 Nodal, cyclin G2 (CCNG2)

Leukemia Inhibitory Factor (LIF) Cyclin D1 Titin MYC/ERK MYC/ERK Flavin adenine dinucleotidedependent oxidoreductase domain-containing protein 1 (FOXRED1), pyruvate dehydrogenase phosphatase regulatory subunit (PDPR), Type II receptor for ActivinA and Nodal (ActRIIA) Forkhead Box Protein A1, Vascular endothelial growth factor (VEGF) Endoplasmic Reticulum Protein 44 (ERp44) BCL2 Associated X (Bax) Myeloid cell leukemia sequence 1 (MCL1), Matrix metalloproteinase 2 (MMP2), Vascular endothelial growth factor A (VEGFA) and Integrin b1 (ITGB1) Insulin-like growth factor 1 (IGF-1) MMP2 Multiple targets Multiple targets

Regulated processes

References

Trophoblast proliferation

[72,73]

Trophoblast proliferation and invasion Trophoblast proliferation, invasion, migration and differentiation Trophoblast proliferation

[71]

Trophoblast proliferation and invasion Trophoblast proliferation Trophoblast proliferation Trophoblast proliferation Trophoblast proliferation and invasion

Trophoblast proliferation, invasion, and placental angiogenesis Trophoblast proliferation

[70,78]

[98] [99] [100] [75] [75] [79,101]

[91,94]

[102]

Trophoblast proliferation Trophoblast proliferation, invasion and placental angiogenesis

[103] [84]

Trophoblast proliferation and invasion Trophoblast invasion and migration Trophoblasts differentiation and migration Trophoblasts differentiation

[104]

[77,105] [76]

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Table 7.1 The table summarizes the role of various miRs, their targets and role in placental development.dcont’d miRs miR-106a-363 cluster miR-210 miR-204 miR-34a

miR-135b miR-21

miR-290 cluster

Targets

Regulated processes

Multiple targets Ephrin-A3 and HomeoboxA9 MMP9 Notch1, Jagged 1 pathway, plasminogen activator inhibitor-1 (PAI-1), MYC C-X-C motif chemokine 12 (CXCL12) Phosphatase and tensin homolog (PTEN), Programmed cell death 4 (PDCD4) Multiple targets

Trophoblasts differentiation Trophoblast migration and invasion Trophoblast invasion Trophoblast invasion

[76] [85]

Trophoblast invasion

[106]

Trophoblast proliferation, migration, and invasion

[80]

Trophoblast proliferation and placental vasculature Trophoblast invasion

[74]

miR-582-3p

Estrogen receptor a (ESRa), SMAD family member 2 (Smad2) Endocrine gland-derived vascular endothelial growth factor (EG-VEGF) EG-VEGF

miR-520g

MMP-2

miR-520c-3p miR-200c miR-20b miR-136 miR-126

MMP-2 VEGF VEGF VEGF Phosphoinositide-3-Kinase Regulatory Subunit 2 (PIK3R2) Cyclin E1, VEGF-A

miR-18a

miR-346

miR-16

References

[87] [81e83]

[107,108]

Trophoblast invasion and migration

[86]

Trophoblast invasion and migration Trophoblast invasion and migration Trophoblast invasion Placental angiogenesis Placental angiogenesis Placental angiogenesis Placental angiogenesis

[86]

[90] [94] [94] [95] [97]

Placental angiogenesis

[96]

[89]

apoptosis [68,69]. miR-378a-5p has been shown to promote the growth of trophoblast cells in first-trimester placental tissues [70]. In addition to miR-378a-5p, there are several other miRs that target TGF-b/nodal signaling. miR-376c promotes proliferation and invasion of trophoblast cells by targeting activin receptor-like kinase 5, ALK5 (type I receptor for TGF-b), and ALK7 (type I receptor for Nodal) [71]. miR-675 also modulates nodal signaling by targeting Nodal Modulator-1 (NOMO-1) [72]. Besides modulating Nodal signaling, miR-675 also targets insulin-like growth factor-1 (IGF-1) receptor as silencing of miR-675 in mice promoted expression of IGF-1 which was associated with an increase in the size of placenta [73].

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Paikari et al. [74] have identified the significant role of miR-290 cluster in placental development. Deletion of miR-290 cluster in mice resulted in an up-regulation of multiple targets regulated by this cluster which was accompanied by the loss of trophoblast progenitor cells and reduction in the size of the placenta [74]. miR-377 and let-7a also play a critical role during placental development. The expression of these miRs is higher in term placenta in comparison to the first-trimester placenta. Overexpression of these miRs resulted in decreased CTB proliferation in first-trimester placental tissue explants [75]. Besides these, there are other miRs, have been shown to regulate trophoblast proliferation and are listed in Table 7.1 along with their targets.

miRs in trophoblast invasion, migration, and differentiation Trophoblast differentiation, invasion, and migration are highly regulated processes during pregnancy and multiple studies have reported the involvement of miRs in these processes. One of the important processes in placentation is differentiation ofCTB to STB. Studies have shown involvement of C19MC, miR-17-92, and miR-106a-363 clusters in this differentiation process [66]. Expression levels of multiple members of these miR clusters were observed to be downregulated during this differentiation process [76,77]. Besides these, miR-378a-5p has also been shown to play a role in STB differentiation [78]. Studies have shown the involvement of multiple miRs in the regulation of migration and invasion of trophoblast cells. Some of these miRs promote invasion/migration, whereas, others inhibit these processes. The miRs which have been shown to improve migration and/or invasion of trophoblast cells include miR-378a-5p [70], miR-195 [79], miR-376c [71], and miR-21 [80]. Other miRs such as miR-34a [81e83], miR-29b [84], miR-210 [85], miR-346, miR-582-3p [86], miR-204 [87], miR-519d-3p [88], miR-520g [89], miR-520c-3p [90] and miR-20a [91] have been reported to inhibit migration and/or invasion of trophoblast cells. The potential role of these miRs in trophoblast invasion/ migration has been identified through in-vitro studies and therefore, these observations need to be verified through in-vivo studies. The identified targets of these miRs are listed in Table 7.1.

miRs in placental vascularization and angiogenesis The human placenta is a highly vascularized organ and the process of vasculogenesis, and angiogenesis is tightly regulated. De-regulation of these processes can lead to placental pathologies. The formation of proper placental vascular network and angiogenesis is essential for materno-fetal exchange, placental development and fetal growth [92]. The first strong evidence regarding the role of miRs in placental angiogenesis comes from a study where the miR processing enzyme, Dicer was targeted. The deficiency of this enzyme caused not only embryonic lethality but also lead to defective angiogenesis [93]. In a recent study, deletion of miR-290 cluster resulted in reduction in the area of vasculature along with disorganization of vascular labyrinth in the mice placenta. This was also associated with the thickening of theplacental maternofetal barrier and reduction in the diffusional exchange capacity [74]. miRs have also been shown to regulate the expression of vascular endothelial growth factor (VEGF) which plays a crucial role in vasculogenesis as well as angiogenesis in the placenta. miR-200c, miR-20a, and miR-20b have been shown to target VEGF in placental trophoblast cells [94,95]. Other miRs which might also play a negative role in placental angiogenesis by targeting VEGF include miR-16 [96] and miR-29b [84]. On the contrary, miR-126 has been reported to promote angiogenesis in mice placenta [97]. Taken together, these studies dictate the importance of miRs in

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placental angiogenesis, more so, than vasculogenesis. More studies are, therefore, required to understand the role of miRs in vasculogenesis. Also, the above-mentioned studies are preliminary in nature and further evaluation of these miRs is necessary to understand their complete role in the regulation of placental vasculogenesis and angiogenesis.

Long non-coding RNAs (lncRNAs) lncRNAs are transcribed RNAs which do not encode for any protein and are more than 200 nucleotides in length. They play a crucial role in a broad range of processes which includes regulation of gene expression, epigenetic modifications, post-transcriptional modifications, RNA splicing, RNA editing, imprinting and X-chromosome inactivation [65]. At present, the understanding of the roles of lncRNAs in placental development is limited. Based on the available reports, lncRNAs have been shown to play critical role in trophoblast proliferation, migration, and invasion. Recently, RNA-seq analysis was performed on human term placenta which led to the discovery of 15,819 lncRNAs and functional analysis of these lncRNAs led to the identification of 2021 target genes [109]. This study is preliminary in nature and therefore requires lot of research to identify the role of these lncRNAs in the placental development. The most studied lncRNA studied in the placenta is H19. H19 is a paternally imprinted gene which is present downstream of IGF2. The deregulation of methylation status and expression of H19 has been associated with placental pathologies. H19 can regulate placental development through at least two different pathways, both of which involve miRs [110]. H19 serves as a source of miR-675 which express exclusively in the placenta and plays crucial role in regulating trophoblast proliferation and invasion [72]. On the contrary, H19 has binding sites for let-7 family of miRs which play important role in developmental processes and cancer by targeting genes which are involved in proliferation, differentiation, cell-cycle progression and apoptosis [111]. The exact mechanism and role of let-7 miRs in placental development are not yet known. Metastasis associated lung adenocarcinoma transcript 1 (MALAT-1) is another lncRNA whose altered expression levels are associated with two placental pathologies (PE and Placenta previa increta/ percreta, I/P) [112]. MALAT-1 expression was observed to be downregulated in PE patients which had poor trophoblast invasion [113]. On the contrary, MALAT-1 levels were upregulated in I/P patients which had very invasive placentation [114]. These observations indicate the role of MALAT-1 in trophoblast invasion. In an in-vitro study performed on human trophoblastic choriocarcinoma cell lines (BeWo, JAR and JEG-3), silencing of MALAT-1 resulted in the reduction of invasive potential of these cells [114]. In another study, knockdown of MALAT-1 levels in JEG-3 cells resulted in cell cycle arrest, reduction in cellular proliferation, invasion and migration which was associated with increased apoptosis of the cells [113]. These studies indicate the importance of MALAT-1 in normal placental development as its altered expression levels are associated with pregnancy-related disorders. SPRY4-IT1 is a highly expressing lncRNA in normal placental tissues and an increase in its levels has been reported in PE patients [115]. In a study utilizing HTR-8/SVneo cell line (First-trimester human trophoblast cells), SPRY4-IT1 has been shown to play a role in cellular proliferation, apoptosis, and migration [112]. Over-expression of SPRY4-IT1 in these cells promoted cell apoptosis along with reduction in cellular proliferation and migration. On the other hand, knockdown of this lncRNA promoted cellular proliferation and migration while decreasing cell apoptosis. Also, the study showed that over-expression of SPRY4-IT1 might affect spiral artery network formation which is important for normal placental development [115]. The findings of this study suggest an important role of SPRY4-IT1 in regulation of trophoblast functions.

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Similar to SPRY4-IT1, altered levels of RPAIN, MEG3, TUG-1, and PVT1 are associated with PE [116e119]. In comparison to normal controls, MEG3, PVT-1, TUG-1 levels were observed to be downregulated whereas RPAIN levels were upregulated in PE patients. Overexpression of RPAIN in HTR-8/SVneo cells resulted in suppression of cellular proliferation and invasion whereas apoptosis was promoted [116]. In contrary to this, overexpression of MEG-3 in HTR-8/SVneo and JEG3 cells resulted in a reduction of apoptosis whereas migration potential was increased [117]. Knockdown of TUG-1 and PVT-1 levels inhibited cellular proliferation and promoted apoptosis in HTR-8/SVneo and JEG-3 cells [118,119]. Other lncRNAs that have been shown to be de-regulated in PE patients includes FLT1P1, LOC284100, and CEACAMP8 [120]. All of these lncRNAs were observed to be upregulated in PE patients. The exact role of these lncRNAs in placenta is not known but they might be associated with regulation of lipid metabolism [120]. MIR503HG and LINC00629 are the long-intergenic ncRNAs (lincRNAs) which are also expressed in human placenta [112]. MIR503HG is present exclusively on placenta whereas LINC00629 expresses in other reproductive tissues besides placenta [121]. Both lincRNAs are positively correlated in their expression and over-expression of these lincRNAs has been shown to be associated with a reduction in cellular and invasion of JEG-3 cells [121]. This suggests their potential role in regulation of cell migration and invasion during placental development however more studies are required to establish their functional importance in normal placentation. Altogether, the studies accessing the role of lncRNAs in normal placentation are rather limited and most of them discuss the role of these lncRNAs in context to the placental pathologies. More studies are necessary to establish the role of these lncRNAs in placental development and to identify their targets.

Histone modifications The DNA is packed into compact chromatin structure and nucleosomes are the building blocks of the chromatin. Within each nucleosome, there are four types of histones (H2A, H2B, H3, and H4) around which DNA is wrapped [122]. Modifications of histones by acetylation, methylation, and phosphorylation plays an important role in the regulation of gene expression by modulating chromatin structure and compaction [123]. At present, there is limited information available about the role of histone modifications in the regulation of human placental development. The most-studied histone modifications in context to placental development in either rodent models or in in-vitro systems are methylation and acetylation. The methylation can occur only on specific lysine and arginine residues whereas acetylation can occur on specific lysine residues only [122]. Depending on the position of these residues and the number of the residues (in case of methylation only), the modifications can lead to transcriptional activation or repression [124]. In a study performed on human placental samples, the levels of H3K9me3 and H3K27me3 which are associated with transcriptional repression were observed to be lower in STBin comparison toCTB. In contrary to this, STB cells were high in H4K20me3 levels, which are also repressive marks [45]. The study showed a difference in the histone marks between CTB and STB, but the effect of these modifications on trophoblast functions are not yet known. In another study, H3K4me2 (activation mark) levels were observed to be enriched in STB cells and were found to be co-localized with active RNA polymerase-II in human placental tissue [125]. The role of this histone modification in STB function has not been studied further.

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Regulation of trophoblast invasion is essential for a normal pregnancy. Matrix metalloproteinases (MMPs) and their inhibitors, known as tissue inhibitors of metalloproteinases (TIMPs), play a crucial role in invasion process and their expression level changes depending on the developmental stage of the placenta [126]. In a study, differential expression of MMPs and TIMPs was observed in three trimesters and was found to be associated withH3K9me3/H3K27me3 levels, both of which are repressive marks [127]. Another protein whose expression level varies with gestational age is Maspin, which negatively regulates migration and invasion [55]. Maspin expression has been found to be lower in first trimester compared to second and third trimester [128]. This variation in Maspin expression has been correlated with histone marks. The increased levels of Maspin in second and third trimester were correlated with increased levels of H3K9ac and H3K4me at the promoter region of this gene [129]. Pregnancy-specific glycoproteins (PSG) are secreted by STB cells of the placenta and are required for normal pregnancy [130]. The expression of these proteins has been shown to be regulated by histones in a placental-derived cell line, where inhibition of histone deacetylases (HDAC) upregulated PSG expression which was associated with an increase in H3ac marks on the promoter region of this gene [131]. Another gene regulated by the action of histone-modifying enzymes (HDAC and HAT, histone acetyltransferases) is GCMawhich plays a role in trophoblastic fusion [132]. Besides HDAC and HAT, there are other proteins like Polycomb-Group (PcG) proteins which can also modify histones. PcG genes encode for proteins that play important role in chromatin remodeling and transcriptional silencing [133]. Mutants of this PcG family members have been shown to induce placental defects in mice [134e136]. Arginine methylation is a common post-translational modification of histones and has been shown to be important for placental development in mice. Arginine methylation of H3 has been associated with the regulation of cellular potency and fate in mice [137]. The study showed the presence of higher level of H3 arginine methylation in the blastomeres that contributed toward the ICM and polar trophectoderm. This observation was also confirmed by the over-expression of CARM1-an H3-specific arginine methyltransferase-higher levels of this enzyme directed the cells to contribute predominantly toward ICM [137]. In another study, knock out of Protein arginine N-methyltransferases (PRMT1)- an H4-specific arginine methyltransferase-produced embryonic lethal phenotype associated with the failure to form an ectoplacental cavity [138]. Altogether, the above-mentioned studies dictate the importance of histone modifications in the development of the placenta.

Imprinting and placental development Normal placental growth, development, physiology, and morphology are dependent on the correct establishment and maintenance of genomic imprinting [139]. Imprinted genes are those that are monoallelically expressed based on their parent of origin. These patterns of expression tend to be species and tissue-specific and in some cases also developmentally regulated [140]. The selective inactivation of one of the parental alleles can be achieved by differential DNA methylation, allele specific histone modifications, antisense ncRNA mediated silencing, and long-range chromatin interactions. An evolutionary link between imprinting and placentation has been suggested with imprinted genes in the placenta playing key roles in trophoblast differentiation and placental development [139]. Imprinted genes are typically associated with differentially methylated regions (DMRs) that are established in either the germ line as primary DMRs and maintained after fertilization

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or during embryonic development as secondary or somatic DMRs. Except the ones associated with known imprinting domains, sperm-derived methylation marks are largely lost by the blastocyst stage, there is incomplete erasure in the maternal genome such that hundreds or even thousands of imprinted DMRs are retained in the placenta [141e143]. More than 30 of placental-specific DMRs have been known to show paternal specific expression in the placenta [142] though, the function all of them is not clear. Most of these placental-specific DMRs show intermediate levels of methylation (transient imprinting) in chorion but loss of methylation in amnion, reflecting the partial trophoblast contribution to the chorion and later origin of the amniotic epithelial layer. They also show enrichment for zinc finger protein 57 (ZFP57) binding sites and for H3K9me3. In mice, di/tri-methylation of histone 3 lysine 9 (H3K9me2/3) and ZFP57 protects imprinted DMRs from demethylation during embryonic reprogramming [144]. About half of placental-specific imprinted DMRs exhibit methylation allele specific polymorphism [145]. That is, some placental samples will exhibit monoallelic methylation while the others show loss of imprinting at the given DMRs. This has been demonstrated to lead to polymorphic monoallelic/biallelic expression for a number of genes, including LIN28B, NTM, and MAGI2 [139,146]. Interestingly, some genes, such as IGF2R and the closely linked SLC22A2, show polymorphic imprinted expression that appears to be independent of DNA methylation [139].

H19-IGF2 cluster H19-IGF2 cluster is localized distally at 11p15.5 in humans and 7qF5 in mice. In both species, the structure of the locus is conserved. An ICR (Imprinting Control Region), located 3 kb from the H19 transcription start site has also been identified, with seven binding sites for the Zinc finger transcription factor CTCF. H19 is expressed exclusively form the maternal allele, while IGF2 is expressed from the paternal allele. In mice, a placental specific promoter of IGF2 was discovered. Epigenetic modification at IGF2 promoter leads to a strong decrease of placental development and placental growth [147]. By contrast, altered expression of H19, leads to placental and fetal overgrowth [148]. Among other imprinted genes that affect placental and fetal growth besides H19 and IGF2 are paternally expressed genes, generally identified in mice (PEG1, PEG3, RASGRF1, DLK1) and maternally expressed genes (IGF2R, GNAS, CDKN1C, GRB10). In mice, the receptor of IGF2, IGF2R is imprinted with a maternal specific expression. In humans, IGF2R is polymorphically imprinted since it is imprinted in few individuals [149]. Recently a polymorphism located at the IGF2/H19 locus was shown associated to placental DNA methylation and birth weight in association with Assisted Reproductive Technologies usage [150]. Deregulation of imprinted genes in the placenta is linked to placental diseases [139,151]. In a recent study, Christians and coworkers, analyzed a list of 120 imprinted genes in relation with global expression of 117 placental samples, including pre-eclampsia (PE) and Intra Uterine Growth Restriction (IUGR) cases [152] The authors identified a significant correlation between birth weight and the expression level of imprinted genes but without significant differences between paternally versus maternally expressed genes. Imprinted genes were also deregulated in preeclampsia with most changes observed in paternally expressed genes that were found to be down-regulated, while maternally expressed genes were up-regulated. The trend was similar for IUGR [139].

The pseudo malignant placental epigenome There are striking similarities between some cancers and early placentation, including growth in a lowoxygen environment, lack of cell-contact inhibition, immune privilege, invasiveness, common signaling pathways regulating these processes, and in some cases, expression of pregnancy-specific

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proteins [153e155]. Cancers and placenta also share several molecular pathways leading to invasion of TGF-b signaling and the expression of human telomerase. Placenta growth factor [156] and placenta proteins such as pregnancy specific b-1 Glycoprotein 1 were found in non-trophoblastic malignant tumors [157]. A direct link between aberrant methylation and tumorigenesis is now well established [158e160] and the similarity between the placenta and invasive cancers is reflected in the epigenome. The global hypomethylation seen in cancer is similar to that of normal placental tissue [49,161]. Recent studies have shown that around 37% of the placental genome shows low to intermediate gene methylation; the same regions are often hypermethylated in somatic cells. These partially methylated domains are also characteristically found in malignant cells and in vitro cultures, [162,163]. Although the functions of these partially methylated domains are incompletely understood, it is thought that they contribute to hypomethylation in tumor cells. The potential consequence of global hypomethylation is the loss of imprinting (LOI), a common feature seen in both cancers and placenta. Some studies have demonstrated biallelic expression of imprinted genes in the placenta, suggesting LOI occurs in those genes [164]. As this most often occurs during the first trimester, LOI is suggested to function in regulating placental invasion and establishment [165]. However, the extent to which genomic imprinting affects placental function and fetal development is still poorly understood.

Disturbed placental epigenetics Placental epigenetics in relation to placental-related pathologies Preeclampsia Preeclampsia is a pregnancy-related multifunction disorder characterized by maternal hypertension, diagnosed after 20th week of gestation [166]. It is a leading cause of maternal and neonatal morbidity and mortality worldwide [167], affecting worldwide almost 10% of pregnancies. Currently, there is no effective treatment or biomarkers available for preeclampsia and the best possible option is discontinuation of pregnancy. The defective placenta is believed to be the causative organ behind preeclampsia development, hence removal of placenta and delivery of infant is the only option available in severe cases [168]. The pathological cause of preeclampsia is attributed to defective and shallow extravillous trophoblast invasion, resulting in inadequately remodeled spiral arteries, impairing maternal blood flow to the placenta [92]. This consequently produces stress in trophoblasts and releases cell fragments, microparticles, and extracellular vesicles into the maternal circulation [169], deteriorating maternal as well as fetal endothelial function [170,171]. These exosomes release miRs that modify maternal gene expression. Recent studies using microRNA-omic analysis have highlighted the importance of these miRs in preeclampsia pathogenesis [172]. Tumor suppressor genes and oncogenes regulate the invasive potential of trophoblasts. Abnormal promoter methylation of such regulatory genes amounting to their aberrant expression contribute to the pathogenesis of preeclampsia [63,173]. Studies have also suggested an imbalance between vasodilator and vasoconstrictor molecules behind endothelial dysfunction in preeclampsia. Dysregulation of VEGF/VEGFRs imbalance acts as a hallmark of endothelial dysfunction in preeclamptic women and their children [174]. Similarly, abnormal placental methylation of various angiogenic genes such as fms-related tyrosine kinase 1, kinase insert domain receptor, VEGF and EGFR has been associated with pathogenesis of

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preeclampsia [175,176]. Lower function of a proliferative type of endothelial progenitor cells known as fetal endothelial colony-forming cells (ECFC), in preeclampsia has also been related to abnormal methylation of fetal ECFC, which promotes atypical cellular interaction and Wnt signaling [177]. Many genome-wide studies have correlated aberrant methylation with preeclampsia. Different epigenetic patterns distinguish between early- and late-onset preeclampsia. Early-onset preeclampsia is associated with higher expression of DNMT1 resulting in higher LINE1 methylation, while such aberrant methylation is not found in late-onset preeclampsia [178]. Similarly, Yuen et al. has reported 34 loci versus 4 loci with lower methylation in early-onset preeclampsia versus late-onset preeclampsia when compared to control pregnancies [179]. The methylation at DMR regions have exhibited difference between preterm versus term preeclampsia, showing more DMRs especially related to cell adhesion altered in preterm preeclampsia [180]. Abnormal methylation at DMRs of imprinting genes has been also reported in preeclampsia [181]. Certain animal model studies have found fetal sex-specific differences in the DNA methylation profiles of placenta, reporting significant DNA methylation changes in female fetus bearing placentas [182]. Two of the LncRNAs MALAT-1 and SPRY4-IT1 having role in placental development have been shown to have an aberrant expression in PE as discussed in Long non-coding RNAs (lncRNAs) section.

Intra uterine growth restriction (IUGR) Another common placental disorder is IUGR, characterized by smaller placentas with abnormal vascularization similar to preeclamptic placentas. However, both these disorders show distinct molecular mechanisms. IUGR is associated with reduced fetal growth due to placental insufficiency with a birthweight of