Epigenetics and Human Reproduction (Epigenetics and Human Health)

Epigenetics and Human Health Series Editors Prof. Dr. Robert Feil Institute of Molecular Genetics CNRS and University ...

Author: Sophie Rousseaux | Saadi Khochbin

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Epigenetics and Human Health

Series Editors Prof. Dr. Robert Feil Institute of Molecular Genetics CNRS and University of Montpellier 1919 route de Mende 34293 Montpellier Cedex 5 France Prof. Dr. Jo¨rn Walter Universita¨t des Saarlandes Naturwissenschaftlich-Technische Fakulta¨t III FR 8.3 Biowissenschaften/Genetik/Epigenetik Postfach 151150 66041 Saarbru¨cken Germany

Associate Editor Dr. Mario-Noyer-Weidner Schwa¨bische Str. 3 10781 Berlin Germany

Sophie Rousseaux Saadi Khochbin l

Editors

Epigenetics and Human Reproduction

Editors Sophie Rousseaux and Saadi Khochbin Institut Albert Bonniot. Centre de Recherche INSERM/ Universite´ Joseph Fourier U823 Labo. Epigenetics & Cell Signalling. 38706 La Tronche CX Domaine de la Merci France sophie.rousseaux@ujf grenoble.fr; khochbin@ujf grenoble.fr

ISBN 978 3 642 14772 2 e ISBN 978 3 642 14773 9 DOI 10.1007/978 3 642 14773 9 Springer Heidelberg Dordrecht London New York # Springer Verlag Berlin Heidelberg 2011 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable to prosecution under the German Copyright Law. The use of general descriptive names, registered names, trademarks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Printed on acid free paper Springer is part of Springer Science+Business Media (www.springer.com)

Preface to the Series

“Epigenetics and Human Reproduction” is the first volume of a new Springer book series on epigenetics and human health and covers various aspects of human reproduction in relation to epigenetic reprogramming during female and male gametogenesis. Drs. Saadi Khochbin and Sophie Rousseaux, the volume Editors of “Epigenetics and Human Reproduction”, are specialists in this biomedical field. They both direct research programmes at the Albert Bonniot Institute in Grenoble (France), one of the centres in the epigenetics of reproduction in Europe. Combined, their expertise goes from fundamental studies using cells and animal models to the understanding and clinical follow-up of infertility in humans. This broad understanding of chromatin and epigenetics in reproduction was most beneficial to the balanced choice of the 15 chapters, each of which covers a different aspect of this emerging field. In fact, this comprehensively referenced book is the first one of its kind. It will be of considerable interest not only to clinicians and biomedical researchers but also to advanced medical and biology students. Many different themes are presented in the book, which is introduced by Professor Bernard Je´gou (Rennes, France) in the following pages. On the clinical side, chapters cover disorders and cancers linked to with human reproduction that are of epigenetic origin and discuss potential consequences of assisted reproduction procedures, in vitro manipulation, and somatic cell nuclear transfer technologies. This is nicely complemented with several chapters on the nuclear organisation and quality of human sperm and the epigenome of the preimplantation embryo. A major, more fundamental part of the book concerns epigenetic transitions and transcriptional regulation (including small RNAs) in developing germ cells. As concerns spermatogenesis, the highly specific chromatin remodelling process of spermiogenesis is being highlighted, also in the context of its possible consequences for the next generation. The ultimate aim of germ cell development in mammals is to produce haploid gametes, a process that requires meiosis to occur. This unique cell cycle is presented with a specific emphasis on the role of chromatin modifications. For instance, meiotic recombination, the complex crossing-over mechanism that provides the genetic diversity between the gametes, is now known to involve specific histone modifications and histone variants. v

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Preface to the Series

This then is the first volume of the new Springer Series “Epigenetics and Human Health”, edited by Professor Jo¨rn Walter (of the University of Saarbru¨cken, Germany) and myself, together with Dr. Mario Noyer-Weidner (Berlin, Germany). During the last years, Springer has become much interested in the novel discipline of Epigenetics, which has so many emerging links with human health. We are very pleased to having been involved in this exciting enterprise. Several other volumes are currently being prepared by invited Editors, to be published during the coming year, including a volume on “Epigenetics and Psychiatry” and one on “Environmental and Nutritional Epigenetics”, two other important emerging themes in medical research. It is now timely to bring together these new epigenetic discoveries of relevance to human disease and to present them in broader contexts with comprehensive referencing of the existing literature. This is the overall aim of our book series, so that it will be of lasting interest to medical and scientific readers. We hope you will enjoy this volume on human reproduction and will find it useful for your professional endeavours. September 2010

Robert Feil Mario Noyer-Weidner Jo¨rn Walter

Foreword

I am pleased and honored to have been invited to write the introduction to this volume depicting the state of the art of “Epigenetics and Human Reproduction”. As I have no particular credentials in epigenetics as such, I would guess that it is by virtue of me having been an observer and sometimes a participant in the progress in the field of reproduction over the past 30 years that I am humbly entitled to comment on the impressive work produced by the numerous contributors to this volume. For a long time, the sequencing of entire mammalian genomes, including that of humans, appeared impossible. This is probably why, when the Human Genome Project was rendered feasible by the technological revolution in the field of molecular biology in the 1980s and was officially launched in 1990, the expectations were enormous. By part of the scientific community, and then the public opinion, it was believed that mapping the whole human genome and sequencing its DNA would by itself reveal how species differ and even what makes each human being unique. However, when the human genome was finally sequenced in 2001, together with that of other selected species (e.g., the mouse in 2002), it became clear that genome sequences are so similar that they cannot per se account for the great diversity within and between species. Similarly, the illusion that for every disease genetic components could reveal its causes (and thus be used for its prevention or cure) sadly faded. Thus, by the beginning of this century, it had become clear that the extraordinary complexity of genetic regulation relies in part on changes in gene expression, without alterations in the DNA sequence itself, that is, on epigenetics. The consequence of this conclusion is that the number of publications in this field has expanded exponentially, as has the number of scientists working at one level or another in epigenetics. To the most enthusiastic analysts of the evolution of our knowledge about the regulation of genes, the epigenetic revolution is the equivalent in biology of the revolution in our understanding of the universe induced by the emergence of the new physics at the beginning of the twentieth century. Epigenetics and epigeneomics have become such strategic issues that, exactly 20 years after the start of the Human Genome Project, an International Human Epigenome Consortium (IHEC) was launched in Paris in March 2010. Its goal is to map 4,000 reference epigenomes within the next 10 years. vii

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Foreword

But how has the epigenetic revolution affected the research aimed at improving our understanding of reproduction in mammals? Most interestingly, from the beginning, the words “epigenetics” and “reproduction” have been consubstantial. Indeed, it was from cloning experiments with the mouse embryo in the 1980s that one of the primordial phenomena of epigenetics was discovered, that is, genomic imprinting. This phenomenon enables the embryo to distinguish between the maternal and paternal alleles it inherited. As a consequence, in the mammalian germ cells, the mother and the father contribute different epigenetic patterns for specific genomic loci. This finding contradicts a basic tenet of Mendelian genetics that the alleles from the father and from the mother are functionally equivalent. Understanding this phenomenon rapidly proved crucial to our comprehension of the origin of several human disorders, including the Angelman, Prader Willi, and Beckwith Wiedemann syndromes, which all result from abnormal imprinting patterns. It also raises the question of whether manipulation of gametes, such as that occurring during assisted reproduction, might induce epigenetic disruption of a chromosomal region (i.e., epimutations), an issue of great importance in humans as well as in animal models (Chaps 1 and 4). A better understanding of this problem will help greatly in evaluating the risks of artificial reproductive technologies (ART) to the conceptus and, we can hope, will increase its safety. Comprehending the etiology of infertility, which currently affects about 10 15% of the individuals in the general adult population in the developed world requires that we understand how the chemical modifications of DNA and its associated proteins via DNA methylation, histone modification, and small noncoding RNA regulation influence gene expression in both the male and the female germ lines and lead to normal embryo development (Chaps 3, 6 9, 11 15). Equally important, the outcome of epigenetic research is likely to improve our knowledge of the causes of testicular cancer, the incidence of which is increasing worldwide (Chap. 2). Furthermore, deciphering the pathways by which epigenetic modifications within the primordial germ cell control the restoration of cellular totipotency also appears crucial and will affect our perception of how the roughly 250 different types of tissues are generated in individual mammals and maintained during development (Chap. 5). Last but not the least, coping with the fast development of the technologies required to study epigenetics, and the enormous amount of data produced by the IHEC program represents a real challenge for the community of “reproepigeneticists” (Chap. 10). There is no doubt that the field of reproduction will benefit from the ongoing international research effort in epigenetics. Conversely, because the germ lineage is at the origin of life and because the cellular and molecular mechanisms that underlie the germ line are particularly complex, progress in understanding the fundamental substrate of fertility and infertility will no doubt have far-reaching consequences in numerous different domains, including systemic diseases such as cancers. Beyond the scope of this volume is another aspect that deserves to be highlighted in this introduction, related to what is now known as “transgenerational inheritance”. The question here is to understand how environmental influences

Foreword

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such as nutrition and chemical agents (solvents, metals, particles of different kinds, solvents, endocrine disruptors. . .) can alter the germ cell epigenome to make the transmission of epigenetic phenotypes possible. Although marked by a certain degree of controversy, as is normal for any emerging scientific field, an ever increasing number of animal studies progressively sustain that this phenomenon is real. The findings are strengthened by studies in human populations, showing that malnutrition during pregnancy may permanently alter chromatin at fetal stages such that epigenomic changes induced in the germ line are passed on to offspring and be at the origin of diseases such as diabetes and cancer. In conclusion, this volume provides a highly up-to-date source of information that should prove exceedingly useful to researchers, teachers, and students in human and animal reproduction, as well as to clinicians concerned with ART and with human fertility issues in general. We hope that the launching of the IHEC will also help to focus attention on the crucial importance of developing the field of “epigenetics and reproduction” and that it will contribute to convincing both IHEC and funding agencies and institutions worldwide to allocate more funds to this fundamentally important field of research. Rennes, France July 2010

Bernard Je´gou

References 1. Baccarelli A, Bollati V (2009) Epigenetics and environmental chemicals. Curr Opin Pediatr 21:243 251 2. Barton SC, Surani MA, Norris ML (1987) Role of paternal and maternal genomes in mouse development. Nature 311:374 376 3. Heijmans BT, Tobi EW, Stein AD, Putter H, Blauw GJ, Susser ES, Slagboom PE, Lumey LH (2008) Persistent epigenetic differences associated with prenatal exposure to famine in humans. Proc Natl Acad Sci USA 105(44):17046 17049 4. Kaati G, Bygren LO, Pembrey M, Sjo¨stro¨m M (2007) Transgenerational response to nutrition, early life circumstances and longevity. Eur J Hum Genet 15(7):784 790 5. Lander ES et al (2001) Initial sequencing and analysis of the human genome. Nature 409 (6822):860 921 6. Mc Grath J, Solter D (1984) Completion of mouse embryogenesis requires both the maternal and paternal genomes. Cell 37:179 183 7. Mouse Genome Sequencing Consortium et al (2002) Initial sequencing and comparative analysis of mouse genome. Nature 420(6915):520 562 8. Robaire B (2008) Is it my grandparent’s fault? Nat Med 14(11):1186 1187 9. Schiermeier Q (2010) Time for the epigenome. Nature 463(7281):587 597 10. Skinner MK, Manikan M, Guerrero Bosagna C (2010) Epigenetic transgenerational actions of environmental factors in disease etiology. Trends Endocrinol Metab 21(4):214 222 [PMID: 20074974] 11. Venter JC et al (2001) The sequence of the human genome. Science 291(5507):1304 1351

Contents

Part I Medical Aspects and Questions Raised on the Molecular Basis of Epigenome Involvement in Reproduction 1

Potential Epigenetic Consequences Associated with Assisted Reproduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Amanda Fortier and Jacquetta Trasler

2

Germ Cell Cancer, Testicular Dysgenesis Syndrome and Epigenetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 Kristian Almstrup, Olga Mlynarska, and Ewa Rajpert-De Meyts

3

Medical Implications of Sperm Nuclear Quality . . . . . . . . . . . . . . . . . . . . . . 45 Rafael Oliva and Sara de Mateo

4

Gene Expression/Phenotypic Abnormalities in Placental Tissues of Sheep Clones: Insurmountable Block in Cloning Progress? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Pasqualino Loi and Grazyna Ptak

Part II Fundamental Aspects of Genome and Epigenome Reprogramming During Gametogenesis 5

Epigenetic Reprogramming Associated with Primordial Germ Cell Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Yoshiyuki Seki

6

Epigenetic Factors and Regulation of Meiotic Recombination in Mammals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 P. Barthe`s, J. Buard, and B. de Massy

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Contents

7

Meiotic Pairing of Homologous Chromosomes and Silencing of Heterologous Regions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 157 Sam Schoenmakers and Willy M. Baarends

8

Histone Variants during Gametogenesis and Early Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 P. de Boer, M. de Vries, and S. Gochhait

9

Genome Organization by Vertebrate Sperm Nuclear Basic Proteins (SNBPs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 213 Juan Ausio´, Laurence R. Brewer, and Lindsay Frehlick

10

Epigenetics in Male Reproduction: A Practical Introduction to the Informatics of Next Generation Sequencing . . . . . . . . . . . . . . . . . . 231 Adrian E. Platts, Claudia Lalancette, and Stephen A. Krawetz

Part III

Re-organization of Nuclear Compartments During Gametogenesis

11

Organization of Chromosomes During Spermatogenesis and in Mature Sperm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261 Olga Mudrak, Irina Zalenskaya, and Andrei Zalensky

12

Nuclear Lamins in Mammalian Spermatogenesis . . . . . . . . . . . . . . . . . . . . 279 Manfred Alsheimer, Daniel Jahn, Sabine Schramm, and Ricardo Benavente

Part IV Fundamental Aspects of Gene Expression Regulation During Gametogenesis 13

Specific Transcription Regulatory Mechanisms of Male Germ Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291 Irwin Davidson

14

The Chromatoid Body: A Specialized RNA Granule of Male Germ Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 311 Ippei Nagamori, Adam Cruickshank, and Paolo Sassone-Corsi

15

Sperm RNA: Reading the Hidden Message . . . . . . . . . . . . . . . . . . . . . . . . . . . 329 David Miller

Glossary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 355 Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 369

Contributors

Kristian Almstrup Department of Growth and Reproduction, Rigshospitalet, Section GR-5064, Blegdamsvej 9, 2100 Copenhagen, Denmark, kristian@ almstrup.net Manfred Alsheimer Department of Cell and Developmental Biology, Biocenter, University of Wu¨rzburg, Wu¨rzburg 97074, Germany, [email protected] Juan Ausio´ Department of Biochemistry and Microbiology, University of Victoria, Petch Building 258a, Victoria, BC, Canada, V8W 3P6, [email protected] Willy M. Baarends Department of Reproduction and Development, Erasmus MC University Medical Centre, PO Box 2040, 3000 CA Rotterdam, The Netherlands, [email protected] P. Barthe`s Institut de Ge´ne´tique Humaine, UPR1142, CNRS, 141 rue de la Cardonille, 34396 Montpellier cedex 5, France Ricardo Benavente Department of Cell and Developmental Biology, Biocenter, University of Wu¨rzburg, Wu¨rzburg 97074 Germany, [email protected] Laurence R. Brewer Center for Reproductive Biology, Gene and Linda Voiland School of Chemical Engineering and Bioengineering, Washington State University, Pullman, WA 99164-2710, USA J. Buard Institut de Ge´ne´tique Humaine, UPR1142, CNRS, 141 rue de la Cardonille, 34396 Montpellier cedex 5, France Adam Cruickshank Department of Pharmacology, School of Medicine, University of California, Irvine, CA 92697, USA

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Contributors

Irwin Davidson Institut de Ge´ne´tique et de Biologie Mole´culaire et Cellulaire. CNRS/INSERM/UDS, 1 Rue Laurent Fries, 67404 Illkirch, France, [email protected] P. de Boer Department of Obstetrics and Gynaecology, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands, [email protected] B. de Massy Institut de Ge´ne´tique Humaine, UPR1142, CNRS, 141 rue de la Cardonille, 34396 Montpellier cedex 5, France, [email protected] Sara de Mateo Human Genetics Research Group, IDIBAPS, University of Barcelona, Casanova 143, 08036 Barcelona, Spain, Biochemistry and Molecular Genetics Service, Hospital Clı´nic i Provincial, Villarroel 170, 08036 Barcelona, Spain M. de Vries Department of Obstetrics and Gynaecology, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands Amanda Fortier Departments of Pediatrics, Human Genetics, and Pharmacology & Therapeutics, McGill University and Montreal Children’s Hospital of the McGill University Health Centre Research Institute, Montreal, QC, Canada Lindsay Frehlick Department of Biochemistry and Microbiology, University of Victoria, Petch Building 258a, Victoria, BC, Canada V8W 3P6 S. Gochhait National Centre of Applied Human Genetics, School of Life Sciences, Jawaharlal Nehru University (JNU), New Delhi, India Daniel Jahn Department of Cell and Developmental Biology, Biocenter, University of Wu¨rzburg, Wu¨rzburg 97074, Germany Stephen A. Krawetz Center for Molecular Medicine and Genetics, Wayne State University School of Medicine, Detroit, MI, USA; Department of Obstetrics and Gynecology, Wayne State University School of Medicine, Detroit, MI, USA; Institute for Scientific Computing, Wayne State University, Detroit, MI, USA; C.S. Mott Center, Wayne State University School of Medicine, 275 E Hancock, Detroit, MI 48201, USA, [email protected] Claudia Lalancette Center for Molecular Medicine and Genetics, Wayne State University School of Medicine, Detroit, MI, USA; Department of Obstetrics and Gynecology, Wayne State University School of Medicine, Detroit, MI, USA Pasqualino Loi Department of Comparative Biomedical Sciences, University of Teramo, Piazza A. Moro 45, 64100 Teramo, Italy, [email protected]

Contributors

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David Miller Reproduction and Early Development Group, Leeds Institute of Genetics and Health Therapeutics (LIGHT), University of Leeds, Clarendon Way, Leeds, West Yorkshire LS2 9JT, UK, [email protected] Olga Mlynarska Department of Growth and Reproduction, Rigshospitalet, Section GR-5064, Blegdamsvej 9, 2100 Copenhagen, Denmark Olga Mudrak The Jones Institute for Reproductive Medicine, Eastern Virginia Medical School, Norfolk, VA, USA; Institute of Cytology, Russian Academy of Sciences, St. Petersburg, Russia Ippei Nagamori Department of Pharmacology, School of Medicine, University of California, Irvine, CA 92697, USA Rafael Oliva Human Genetics Research Group, IDIBAPS, University of Barcelona, Casanova 143, 08036 Barcelona, Spain; Biochemistry and Molecular Genetics Service, Hospital Clı´nic i Provincial, Villarroel 170, 08036 Barcelona, Spain, [email protected] Adrian E. Platts Center for Molecular Medicine and Genetics, Wayne State University School of Medicine, Detroit, MI, USA; Department of Obstetrics and Gynecology, Wayne State University School of Medicine, Detroit, MI, USA Grazyna Ptak Department of Comparative Biomedical Sciences, University of Teramo, Piazza A. Moro 45, 64100 Teramo, Italy, [email protected] Ewa Rajpert-De Meyts Department of Growth and Reproduction, Rigshospitalet, Section GR-5064, Blegdamsvej 9, 2100 Copenhagen, Denmark, [email protected] Paolo Sassone-Corsi Department of Pharmacology, School of Medicine, University of California, Irvine, CA 92697, USA, [email protected] Sam Schoenmakers Department of Reproduction and Development, Erasmus MC University Medical Centre, PO Box 2040, 3000 CA Rotterdam, The Netherlands; Department of Obstetrics and Gynaecology, Erasmus MC University Medical Centre, PO Box 2040, 3000 CA Rotterdam, The Netherlands, [email protected], Sabine Schramm Department of Cell and Developmental Biology, Biocenter, University of Wu¨rzburg, Wu¨rzburg 97074, Germany Yoshiyuki Seki Laboratory of Molecular Epigenetics for Germline Development, School of Bioscience, Kwansei-Gakuin University, 2-1 Gakuen, Sanda, Hyogo 669-1337, Japan, [email protected]

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Contributors

Jacquetta Trasler Montreal Children’s Hospital Research Institute, 2300 Tupper Street, Montreal, QC, Canada H3H 1P3, [email protected] Irina Zalenskaya The Jones Institute for Reproductive Medicine, Eastern Virginia Medical School, Norfolk, VA, USA Andrei Zalensky The Jones Institute for Reproductive Medicine, Eastern Virginia Medical School, Norfolk, VA, USA, [email protected]

Part I

Medical Aspects and Questions Raised on the Molecular Basis of Epigenome Involvement in Reproduction

Chapter 1

Potential Epigenetic Consequences Associated with Assisted Reproduction Amanda Fortier and Jacquetta Trasler

Abstract Assisted Reproductive Technology (ART) includes an array of treatments designed to bypass fertility problems in both men and women in order to achieve conception. These methods vary from hormonal stimulation of follicular development and ovulation in women, to laboratory handling of both male and female gametes leading to fertilization. In the developed world 1 3% of all births are the result of ART. In recent years, concern has been raised about possible increases in the occurrence of rare genomic imprinting disorders in ART-conceived children. Studies suggest that the perturbations seen in these children could be associated with epigenetic disruption of chromosomal regions or epimutations. Genomic imprinting refers to the acquisition of a unique epigenetic profile in a small subset of genes during gametogenesis. These differential epigenetic marks in the gametes result in a parentof-origin specific expression of these imprinted genes in the offspring. An important epigenetic mark involved in the imprinting process is DNA methylation, which takes place in the male and female germlines at very specific times during the development of gametes and involves a well-orchestrated expression of enzymes. This period of DNA methylation acquisition may be susceptible to perturbations resulting from gamete handling. Furthermore, genomic imprinting established in the germline must remain unaltered following fertilization of these gametes and throughout the life of the offspring, raising another possibility for ART-induced epigenetic disturbance during the maintenance of these imprints in early embryonic life. In the present chapter we discuss the current data available on the effects of ART on genomic imprinting in humans, as well as in animal research models, and discuss the possible mechanisms involved in this epigenetic deregulation.

A. Fortier and J. Trasler (*) Departments of Pediatrics, Human Genetics, and Pharmacology & Therapeutics, McGill University and Montreal Children’s Hospital of the McGill University Health Centre Research Institute, Montreal, QC, Canada e mail: [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 1, # Springer Verlag Berlin Heidelberg 2011

3

4

A. Fortier and J. Trasler

Keywords ART Assisted Reproduction Assisted Reproductive Technologies DNA Methylation Epigenetics Genomic Imprinting Infertility Ovarian Stimulation

1.1

Introduction

A number of reports first began to appear in the literature in the early 2000s linking human-assisted reproductive technology (ART) with epigenetically based genomic imprinting disorders, in particular, Beckwith Wiedemann Syndrome (BWS) and Angelman Syndrome (AS) (reviewed in De Rycke et al. 2002; Gosden et al. 2003; Lucifero et al. 2004a). Most imprinted genes exist in clusters in the genome and their allele-specific expression is regulated by sequence elements called imprinting control regions (ICRs). Genomic imprinting is under the control of epigenetic mechanisms including DNA methylation at ICRs, also known as differentially methylated domains or regions (DMDs, DMRs). At most ICRs, methylation occurs in the female germline and is inherited from the mother. The genomic imprinting disorders seen in children conceived using ART were accompanied in many cases by a loss of maternal DNA methylation at imprinted loci. DNA methylation, one of the best studied epigenetic mechanisms, is both heritable and reversible and susceptible to perturbation during development. The fact that genomic DNA methylation patterns are acquired and dynamic during germ cell development and the preimplantation period, times that could be affected by techniques used in assisted reproduction, provided biological plausibility for the potential for ARTs to perturb genomic imprinting. However, an alternate explanation for the genomic imprinting disorders amongst ART-conceived children, the underlying infertility of the parents, was also postulated and received support from a number of studies reporting DNA methylation defects in imprinted genes in the sperm of infertile men (Filipponi and Feil 2009). Finally, it was also suggested that subfertility and ARTs might interact to predispose children to genomic imprinting disorders (Ludwig et al. 2005; Horsthemke and Ludwig 2005). The use of animal models is beginning to define which gamete and embryo developmental stages may be most susceptible to perturbation by ARTs in the absence of the complicating factor of infertility. In this chapter, the human studies linking ARTs, infertility, and genomic imprinting disorders are reviewed, and the data that have emerged from animal studies are used to suggest promising directions for future study.

1.2

ARTs and the Timing of Epigenetic Events during Gametogenesis and Embryogenesis

Epigenetic marking of the genome, including the parental germline-specific modifications of imprinted genes underlying genomic imprinting, takes place during gametogenesis and embryogenesis. The focus here is on DNA methylation as it

1

Potential Epigenetic Consequences Associated with Assisted Reproduction

5

is the epigenetic modification that has been found to be perturbed in children conceived through the use of ARTs. Other epigenetic modifications showing dynamic changes during germ cell and early embryo development, and that interact with DNA methylation, include histone modifications and expression of different kinds of non-coding RNAs (Sasaki and Matsui 2008; Ooi 2009).

1.2.1

Gamete Development

DNA methylation is found at about 20 30 million sites, predominantly within CpG dinucleotides, throughout the mammalian genome. Sequences that are methylated in the genome include retrotransposons, tandem repetitive sequences such as pericentromeric minor and major satellite DNA, genes on the inactive X chromosome, and imprinted genes (Ooi et al. 2009). This genomic methylation is erased in each reproductive cycle in the primordial germ cells and then reacquired in a sex-specific manner during gametogenesis (Trasler 2006). DNA methylation, in which a methyl group is transferred from S-adenosylmethionine (SAM) to the 50 carbon of the cytosine ring, is catalyzed by a family of DNA (cytosine-5-) methyltransferases (DNMTs) (Goll and Bestor 2005). From studies in the mouse, DNA methylation patterns are acquired at different developmental times in the male and female germlines. In the male, DNA methylation begins to be acquired before birth in gonocytes and is complete for most sequences after birth, before the pachytene phase of meiosis (Davis et al. 1999; Lees-Murdock et al. 2003; Li et al. 2004; Oakes et al. 2007; Ueda et al. 2000). Only a few imprinted genes become methylated during male germ cell development, including the ICRs controlling the H19/Igf2 and Dlk1/Dio3 domains; in contrast, ICRs that normally acquire methylation during oogenesis remain unmethylated (Bartolomei 2009). In the female germline, gametic methylation is only acquired postnatally, following pachytene and during the oocyte growth phase (Lucifero et al. 2004b; Obata and Kono 2002). Bisulfite sequencing studies, in particular, have been useful in characterizing the acquisition of DNA methylation on imprinted genes in oocytes. DNA methylation is acquired progressively during oocyte growth with some imprinted genes becoming methylated early (e.g., Snrpn) and others later (e.g., Peg1/Mest) in the oocyte growth phase (Lucifero et al. 2004b; Hiura et al. 2006). Normal DNA methylation patterns are essential for gametogenesis as evidenced by the fact that deficiency of the DNMT enzymes involved in establishing methylation patterns in male and female germ cells results in abnormal reproductive outcomes. Males deficient in DNMT3L or DNMT3a in their germ cells are infertile (Bourc’his and Bestor 2004; Kaneda et al. 2004). The offspring of females whose oocytes are deficient in DNMT3L or DNMT3a die at mid-gestation and have widespread genomic imprinting defects (Bourc’his et al. 2001; Kaneda et al. 2004). Although developmental studies are more difficult to carry out in human, there is evidence of conservation of gametic methylation, in particular for imprinted genes. For example, the paternally inherited methylation imprints at H19 and DLK1/DIO3

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are conserved in human sperm (Geuns et al. 2007b; Hamatani et al. 2001; Kerjean et al. 2000; Marques et al. 2004). In addition, imprinted genes that are normally methylated in the female germ line are unmethylated in human sperm (Kobayashi et al. 2007; Marques et al. 2008). Most studies of human oocytes have indicated that imprinted genes such as SNRPN and the chromosome 11p15 KvDMR are methylated in fully grown oocytes, emphasizing conservation with the mouse (Geuns et al. 2003, 2007a; Sato et al. 2007). In addition, similar timing of expression of the DNMTs occurs in human fetal gonads as compared to the mouse (Galetzka et al. 2007). Importantly, studies are emerging to indicate that the epigenetic patterns in human sperm (histone modifications and DNA methylation) may be important for the early stages of embryo development (Carrell and Hammoud 2010).

1.2.2

Preimplantation Embryo Development

Following fertilization, a second period of erasure occurs in the preimplantation embryo, when methylation across much of the genome is lost with the exception of imprinted genes and some repeat sequences (Lane et al. 2003; Olek and Walter 1997). Imprinted gene methylation patterns must be maintained during preimplantation development since it is only in the germline (male or female depending on the gene) that imprinted genes acquire the allele-specific methylation that will subsequently be responsible for monoallelic expression in the postimplantation embryo and adult (Bartolomei 2009). Examples of imprinted genes that become methylated in the male germline (unmethylated in oocytes) include H19, Rasgrf1, and Dlk1; examples of imprinted genes that get methylated in the female germline (unmethylated in sperm) include Snrpn, Peg3, Igf2r, and Peg/Mest1. Following the blastocyst stage, the genome is remethylated to adult levels soon after implantation, between 5.5 and 7.5 days of gestation in the mouse. Accurate reprogramming is therefore required with every reproductive cycle to ensure proper erasure, acquisition, and maintenance of methylation marks.

1.3

Assisted Reproduction and Imprinting Disorders

The two imprinting diseases linked in the initial studies to ARTs were BWS and AS. Patients with BWS show overgrowth, exomphalos, macroglossia, fetal hypoglycemia, umbilical and abdominal wall defects, and increased susceptibility to the development of childhood tumors. Five genes in the imprinted region on chromosome 11p15 that have been implicated in the pathogenesis of BWS include insulinlike growth factor 2 (IGF2) in the distal ICR1 cluster, and in the proximal ICR2 cluster, cyclin-dependent kinase inhibitor 1c (CDKN1C), potassium voltage-gated channel KQT-like subfamily member 1 (KCNQ1), and KCNQ1 overlapping transcript 1 (KCNQ1OT1, previously called LIT1) (Enklaar et al. 2006). IGF2 and KCNQ1OT1 are paternally expressed and H19, CDKN1C, and KCNQ1 are

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maternally expressed. About 50% of sporadic BWS cases result from promoter hypomethylation at ICR2 or KvDMR1, within KCNQ1OT1; loss of maternal methylation leads to biallelic expression of KCNQ1OT1 and silencing of KCNQ1 and CDKN1C. In about 2 7% of BWS patients another epigenetic defect is found, the hypermethylation of the H19 promoter and/or ICR1 region resulting in silencing of H19 and biallelic expression of IGF2. AS involves an imprinted region on chromosome 15q11 q13, results in loss of maternal gene expression of the UBE3A gene, and is characterized by mental retardation, microcephaly, movement disorders including ataxia, seizure disorders, abnormal behavior, and limited development of speech and language. In contrast to BWS, only a small proportion, about 3 5%, of AS cases are the result of methylation defects in the 15q11 q13 region (Horsthemke and Wagstaff 2008). The initial studies linking ARTs and genomic imprinting disorders suggested an increased incidence of BWS in three different countries (DeBaun et al. 2003; Gicquel et al. 2003; Maher et al. 2003, reviewed in Gosden et al. 2003). Since these initial reports were published, a number of further studies have linked ART and BWS (Bowdin et al. 2007; Doornbos et al. 2007; Halliday et al. 2004, reviewed in Owen and Segars 2009). Some reports have postulated that BWS is linked to more specific procedures such as IVF/ICSI (Sutcliffe et al. 2006) or ovarian stimulation (Chang et al. 2005). In many cases, the BWS was associated with maternal hypomethylation of KvDMR1, an imprinting centre on chromosome 11p15. Maternal hypomethylation of KvDMR1 is found in about 50 60% of cases of BWS (Weksberg et al. 2005). A report specifically examining the rates of maternal KvDMR1 hypomethylation found that for 24 out of 25 cases of BWS following ARTs, the molecular basis of the disease was maternal hypomethylation (Lim et al. 2009). The higher proportion of BWS cases associated with maternal KvDMR1 hypomethylation following ARTs supports the hypothesis that either the techniques used in ART or the underlying infertility of the parents contributes to the increased risk of BWS. Interestingly, in BWS patients with KvDMR1 hypomethylation, two groups have reported epigenetic defects at imprinted genes outside the chromosome 11p15 region (Rossignol et al. 2006; Lim et al. 2009). One of the studies found higher frequencies of loss of methylation at DMRs outside 11p15 in ART cases than in non-ART cases (Lim et al. 2009); the results suggest the presence of more extensive (multi-loci) epigenetic defects following ART. Thus, a number of studies among children with BWS have consistently reported a higher prevalence of those conceived with ARTs (DeBaun et al. 2003; Gicquel et al. 2003; Maher et al. 2003; Halliday et al. 2004; Chang et al. 2005; Sutcliffe et al. 2006). In contrast, two cohort studies, one from Sweden (Lidegaard et al. 2005) and the other from Denmark (Kallen et al. 2005), did not find an increased risk of BWS among ART-conceived children. To accurately assess the potential connection between ARTs and BWS, a large international study, comparing the prevalence of BWS among naturally conceived and ART conceived children, would be required. With an overall prevalence of BWS of about 1 in 4000 ART births, preconception counseling should be offered but screening is not warranted. While less compelling than the case for BWS, there is some evidence that AS is associated with ARTs and that the underlying molecular mechanism is demethylation

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of SNRPN (Cox et al. 2002; Orstavik et al. 2003; Sutcliffe et al. 2006; Owen and Segars 2009). Ludwig and colleagues (2005) found an increased prevalence of imprinting defects in AS patients born to subfertile couples, suggesting that underlying infertility and techniques might interact. Findings from a further study suggesting a higher risk of AS children amongst parents with fertility problems where ovulation induction was used provided further support for an interaction between infertility and specific techniques used in ART (Doornbos et al. 2007). Not all reports have been positive. For instance, a study in 92 children born after ICSI found no evidence of methylation defects at SNRPN (Manning et al. 2000); however, it is likely that much larger samples would need to be examined to detect an increased prevalence of imprinting defects. Given the rarity of AS (1/10,000 1/20,000 births) and the fact that hypomethylation is the molecular cause in only a small proportion (~6%) of cases (Williams 2005), the identification to date of five cases of AS with an epigenetic basis associated with ART suggests that further studies are warranted. One further imprinting disorder, Silver Russell Syndrome (SRS), has been linked to ARTs. The clinical features of SRS are heterogeneous but generally include severe intrauterine growth restriction, poor postnatal growth, characteristic craniofacial features, and body asymmetry. Part of the clinical heterogeneity of SRS may be attributed to the fact that several different chromosomes have been linked to the disease, suggesting that there are multiple different disorders that have been grouped into this single category. Two imprinted genes have been implicated in SRS: growth receptor bound protein 10 (GRB10) at chromosome 7p11.2 p13 and IGF2 on chromosome 11p15, which is also implicated in BWS. Epimutations at H19-IGF2 have recently been linked to SRS, following the identification of patients carrying maternal uniparental disomy for chromosome 11p15 showing characteristics of SRS. When further examination of the methylation at the H19 ICR was undertaken, hypomethylation was observed in 38 63% of cases [reviewed in (Eggermann 2009)]. If the H19 ICR is hypomethylated, reduced expression of IGF2 could occur, leading to growth defects. SRS is extremely rare, with an incidence of 1 in 100,000 live births (Perkins and Hoang-Xuan 2002). While the link between SRS and ARTs is tenuous, four cases of SRS have been described following ARTs, two in which the molecular etiology was unknown (Kallen et al. 2005; Svensson et al. 2005) and two cases with DNA methylation defects, one of which was the hypomethylation of H19 (Bliek et al. 2006; Kagami et al. 2007). Given the rarity of SRS, and the evidence of hypomethylation at H19, the association of SRS to ARTs is merely suggestive, but should be taken under advisement.

1.4

Searching for Epigenetic Abnormalities in Children Conceived Using ARTs

It has been estimated that there are about 200 imprinted genes in the human genome (Luedi et al. 2007). Imprinted genes play roles in growth and placental function and can contribute to carcinogenesis. In particular, many imprinted genes are known to

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be expressed and play a role in the normal function of the placenta (Coan et al. 2005), an important contributor to fetal development and size. Although abnormalities at only a few imprinted regions associated with ART have been identified to date, there is concern that other regions important for postnatal development or capable of contributing to cancer (e.g., growth promoters or growth suppressors) only later in life might be affected. ART-conceived children are more likely to be born small for gestational age (SGA) and are ~30% more likely to be born with a birth defect (Wilkins-Haug 2009). Investigators are beginning to search for imprinting defects at multiple loci in cohorts of children conceived using ART. One study tested the hypothesis that abnormalities in imprinting might underlie the increased proportion of SGA babies following ART (Kanber et al. 2009). The authors examined methylation patterns at six imprinted loci in DNA from buccal cells of 19 SGA babies conceived with ICSI as compared to 29 normal birth-weight naturally-conceived children. No major difference in imprinting was found between the two groups. Although one ICSI-conceived SGA child showed hypermethylation of the paternal copy of KCNQ1OT1, the significance of this finding is unclear; such a defect would be predicted to be due to a lack of erasure of the maternal methylation mark in the paternal germline, something that has been noted in male infertility (Filipponi and Feil 2009). This study argues that larger numbers of ART-conceived children should be studied with more imprinted genes targeted for analysis and additional tissues examined. Katari and colleagues (2009) used an array-based approach to examine methylation over about 1500 CpG sites in the promoters of 700 genes, including sites in all known imprinted genes. DNA methylation was compared in placenta and cord blood samples between 10 children conceived through in vitro fertilization (IVF) without ICSI and 13 children conceived naturally (in vivo). The results were used to examine gene expression profiles in a larger group of children. Overall lower levels of methylation in the placenta and higher levels in cord blood were found in the IVF group; the methylation differences correlated with altered transcription of a small number of adjacent genes. The study provided evidence that methylation abnormalities extended beyond imprinted genes although the significance of such changes for the offspring is unknown. The finding of altered methylation and expression in the placenta is notable as placental function might be affected and contribute to decreased birth weight in ART-conceived children.

1.5

Underlying Infertility as a Contributor to Imprinting Defects After ART

A number of reports have suggested a role for infertility in the aetiology of imprinting defects. The strongest evidence for such a connection comes from evidence of epigenetic defects in the sperm of infertile men. A number of studies have reported DNA methylation defects at imprinted genes in men with infertility

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and oligospermia (reviewed in Filipponi and Feil 2009). Abnormal methylation has been reported at both paternal and maternal ICRs. One of the first studies (Marques et al. 2004) reported decreased methylation of the H19 DMR in oligospermic men; interestingly, the magnitude of loss of methylation was associated with the severity of the oligospermia. In a follow-up study, Marques and colleagues (2008) reported both loss of methylation of H19 and gain of methylation of PEG1/MEST in oligospermic men. In a study of 97 oligospermic men, hypomethylation at H19 and DIO3 (also known as GTL2) was found; in addition, abnormal methylation (usually absent in sperm) was found at maternally methylated imprinted sequences (Kobayashi et al. 2007). In the latter study, methylation of LINE-1 and Alu elements was unaffected in the sperm of the oligospermic men. Houshdaran and colleagues (2007) reported more generalized effects on DNA methylation, with evidence of increases in methylation at non-imprinted sequences as well as imprinted sequences in men with poor-quality sperm. A recent study reported that abnormalities in the methylation of imprinted genes differed between men with different causes for their infertility (Hammoud et al. 2010). For all of the studies reported to date the mechanism underlying the imprinted gene defects in infertile men is unknown at present. However, the fact that male germ cell development is such a complex process and the causes of abnormalities in spermatogenesis resulting in oligospemia so numerous suggests that epigenetic defects may be secondary to the primary cause of the altered spermatogenesis. The functional significance of methylation abnormalities in imprinted genes in infertile/subfertile men, in terms of adversely affecting the offspring, is unclear. One report has approached this question by examining DNA methylation in both the father’s sperm and the resulting conceptus (Kobayashi et al. 2009). DNA methylation was assessed at seven imprinted loci, the XIST locus, and the LINE1 and Alu repeat sequences in DNA samples from ART- and non-ART-conceived aborted fetuses. Of 78 ART-conceived fetuses there were a total of 17 fetal samples that showed methylation imprint errors, with eight cases showing aberrant paternal methylation (H19, DIO3, or GTL2) and 12 cases of aberrant maternal methylation (PEG1/MEST, KCNQ1OT1, ZAC, PEG3, and XIST); the most common abnormalities were found at H19. Of the 38 non-ART-conceived conceptuses, a total of three fetal samples showed methylation imprint errors. Repeat sequence methylation was not affected in either group. Sperm DNA was examined in the fathers of the fetuses with DNA methylation abnormalities. Seven cases were found where the methylation defect in the fetus could also be identified in the father’s sperm sample, through the use of sequence polymorphisms between the maternal and paternal DNA. Interestingly, two of the seven fathers also carried alterations in the sequence of DNMT3L, providing evidence that the DNA methylation defects in the children were transmitted from the father rather than being due to the techniques used in ART. There were also four fathers’ sperm samples that had methylation abnormalities that were not found in the fetus; this finding suggests mosaicism for DNA methylation defects in sperm or the possibility of repair of defects after fertilization. In support of the former possibility, Flanagan and colleagues (2006) have reported intraindividual differences in DNA methylation in human sperm.

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Techniques Used in ART and Imprinting Defects

Limited evidence, some of which has been alluded to above (Ludwig et al. 2005; Chang et al. 2005; Sutcliffe et al. 2006), is available from human studies that techniques used in ART may result in imprinting defects. It is difficult to draw conclusions since underlying infertility may also contribute. Sato and colleagues (2007) reported that superovulation is associated with a gain of methylation at H19 and a loss of methylation at PEG1/MEST. In addition, abnormally high levels of methylation of H19 were reported in about 25% of human metaphase II oocytes matured in vitro (Bourghol et al. 2006). A further study found that the KCNQ1OT1 KvDMR1 in DNA from germinal vesicle and metaphase I oocytes was more methylated when obtained from natural cycles that from patients receiving gonadotrophin stimulation; the authors suggested that ovarian stimulation recruits immature follicles in which the methylation of imprinted genes is not yet completed (Khoueiry et al. 2008).

1.7

Assisted Reproduction and Other Potential Epigenetic Effects

Some interesting associations between ARTs and other phenomena that may be related to underlying epigenetic etiology have been reported. For instance, ARTs are associated with a 2-12 times higher rate of twinning (Aston et al. 2008). A recent meta-analysis suggested that the ART technique of blastocyst transfer is associated with a higher male female sex ratio and an increased risk of monozygotic twinning among the offspring (Chang et al. 2009). While it has been suggested that the increased risk of twins following ART is likely due to embryo manipulation or effects of culture, it may also be related to perturbation of epigenetic events during preimplantation development. BWS has been reported in naturally conceived monozygotic twins in which the twins are discordant for BWS and the BWS in the affected twin is associated with hypomethylation of KvDMR1; in most such cases the twins are female (Weksberg et al. 2002). The authors have proposed that a failure to appropriately maintain DNA methylation, through the DNMT1o DNA methyltransferase, at the “susceptible” BWS locus on chromosome 11p15, at a key time in preimplantation development, could explain the epigenetic asymmetry between the twins as well as the predominance of females (Weksberg et al. 2002; Bestor 2003). To explain the association between ART and imprinting syndromes, it is theoretically possible that ART (e.g., ovulation of poor quality or immature oocytes, embryo culture) somehow interferes with the maintenance of key DNA methylation patterns during preimplantation development resulting in imprinting defects and altered sex ratio. Such a possibility could best be tested in animal models.

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Lessons from Animal Models: Mimicking Techniques Used in Human-Assisted Reproduction

Determination of which individual techniques might perturb epigenetic events in oocytes and embryos is difficult as patients using ARTs are exposed to different procedures during their treatments. The mouse model provides an alternative, as individual techniques can be modeled in isolation. In addition, mechanisms underlying aberrant genomic imprinting can be more easily investigated in the mouse where genetic strains are available and larger numbers of oocytes and preimplantation embryos at precise stages of development can be isolated. The animal model studies have been reviewed (Huntriss and Picton 2008). Some examples will be discussed here that help provide guidance for the types of clinical studies that need to be done.

1.8.1

Superovulation and Gamete Manipulation

Sato and colleagues (2007) examined the methylation of three maternally methylated genes (Peg1, Zac, and Kcnq1ot1) and one paternally methylated gene (H19) in oocytes following superovulation. All of the maternally methylated genes had normal methylation after superovulation; however, interestingly, H19 gained methylation. The results suggested that the acquisition of maternal methylation imprints was unaffected but that oocyte quality was affected such that abnormal methylation was present on H19. In another study, Fauque et al. (2007) observed decreased levels of expression of H19 in blastocysts after superovulation; the results could be explained by an alteration in oocyte quality affecting H19 epigenetic marking or a delay in embryo development since H19 is usually first expressed at the blastocyst stage. A detailed examination of the DNA methylation status of four imprinted genes in individual blastocysts following low- and highdose superovulation regimens was reported recently. In this study a dose-dependent loss of DNA methylation at the maternally methylated genes Snrpn, Peg3, and Kcnq1ot1 and a gain of methylation of H19 were found (Market-Velker et al. 2010). The authors suggested that superovulation had two distinct effects, one to disrupt the acquisition of methylation imprints during oocyte growth and the second to impair the proper maintenance of imprints during preimplantation development. The effects of superovulation have also been examined at later times in development. A low-dose superovulation protocol was shown to alter the expression of maternally and paternally methylated imprinted genes in the midgestation mouse placenta, suggesting that trophectoderm-derived tissues may be more susceptible to disruption of imprinted genes than the embryo proper (Fortier et al. 2008). There has been concern that the use of immature gametes may be associated with imprinting defects in the offspring. From a genetic perspective, it is reassuring that

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in a transgenic mouse model, various ART techniques including IVF, ICSI, round spermatid injection (ROSI), and cell culture were not associated with an increase in the frequency or spectrum of point mutations (Caperton et al. 2007). If higher levels of methylation were induced by these techniques, they might be expected to result in mutagenesis, since methylated cytosine residues are spontaneously deaminated. Some cases of male infertility require ROSI to be performed, raising the concern for the use of immature gametes in which epigenetic patterns are not complete. Although genome-wide single base pair resolution studies have not been done, evidence to date from mouse studies suggests that DNA methylation acquisition is complete by the round spermatid stage (Trasler 2009). For the most part imprinting seems to be normal in the offspring after ROSI (Shamanski et al. 1999; Miki et al. 2004). In contrast, earlier in gestation, ROSI was associated with DNA methylation and histone methylation defects in zygotes, suggesting caution and the need for more studies (Kishigami et al. 2006; Yamazaki et al. 2007). In vitro growth (IVG) and maturation (IVM) of oocytes are additional manipulations that may be associated with altered methylation of imprinted genes. IVG of mouse oocytes has been reported to result in hypomethylation of Igf2 and Peg1/ Mest and hypermethylation of H19 (Kerjean et al. 2003). In another study using a well-established follicle culture system and varying exposure to follicle stimulating hormone, Anckaert and colleagues (2009) detected no abnormalities in the methylation of a number of imprinted genes. Since IVG and IVM are either being evaluated for or are in use clinically, further model studies are required. It will be important for future studies to use optimal culture techniques and to pursue the physiological significance of any defects detected in oocytes by examining the offspring and their placentae at later times in gestation.

1.8.2

Embryo Culture

In animal studies there have been numerous examples of culture media effects on imprinted genes. Early work linked in vitro culture media conditions during the preimplantation period with abnormal methylation patterns in the imprinted gene IGF2R and large offspring syndrome in sheep (Young et al. 2001). In the mouse model, abnormalities in imprinted gene methylation and expression have been linked to factors such as suboptimal culture media and culture additives such as serum, (e.g., Doherty et al. 2000; Khosla et al. 2001; Fernandez-Gonzalez et al. 2004; Ecker et al. 2004); some adverse effects of the culture result in abnormalities in postnatal development including behavior (Fernandez-Gonzalez et al. 2004; Ecker et al. 2004). Studies on the effects of embryo culture on genomic imprinting have shown that loss of imprinting is more pronounced in the placenta than in the embryo (Mann et al. 2004; Rivera et al. 2007). These results suggest that the placenta may be more susceptible to epigenetic disruption than the embryo; this is an important observation as abnormal placental function could in

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turn affect fetal growth. The study by Riviera and colleagues (2007) also showed that blastocyst collection and embryo transfer without culture were associated with loss of methylation on the maternal allele of the Kcnq1ot1 KvDMR1 locus, providing evidence that even relatively minor manipulations could be deleterious to the offspring.

1.9

Future Directions

Together, the importance of gametogenesis and preimplantation development for epigenetic programming, data from animal models that show that developmental DNA methylation events can be perturbed, and the human studies suggesting a link between ART and imprinting errors indicate that further studies are warranted. There is a need for well-designed prospective long-term follow-up studies to monitor ART-conceived children; it is likely that such studies will need to be continued into adulthood to determine if there is an increased risk of cancer or other adult onset diseases. Outcomes should be linked to clinical records with precise details of infertility treatments (e.g., induction protocol, and culture times and components). To begin to tease out the effects of ART versus infertility, consideration of appropriate comparison groups will be important including the children of infertile/subfertile couples who conceived naturally as well as those of couples who used ART in the absence of infertility (e.g., use of donor gametes, or in the case of preimplantation genetic diagnosis for normal embryo selection). Collecting biological samples including the placenta and DNA from the mother, father, and child will allow genomic and epigenomic studies to be performed, once the technology is available at a reasonable cost. More research is required to better understand the genetic and epigenetic causes of infertility. Due to ethical considerations and the limited access to pure populations of cells at different developmental stages, the timing and regulation of epigenetic events during human gametogenesis and embryogenesis has been difficult to study. Such studies may be easier in the future once molecular analysis of epigenetic events is possible in small numbers of cells. Mechanistic epigenetic studies will need to extend to interactions between different epigenetic pathways including DNA methylation, histone modifications, and non-coding RNAs. While counseling of prospective parents about potential epigenetic risks is important, screening for imprinting disorders amongst ART-conceived children is not currently warranted since the absolute risk of imprinting disorders is small. Acknowledgments We thank Dr. Flavia Lopes and members of the laboratory for the thought provoking discussions on this topic. Research in the Trasler laboratory is supported by grants from the Canadian Institutes of Health Research (CIHR). A.L.F. was the recipient of a CIHR Graduate Studentship. J.M.T. is a James McGill Professor of McGill University. The authors are members of the Research Institute of the McGill University Health Centre, which is supported in part by the Fonds de la recherche´ en Sante´ du Que´bec (FRSQ).

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Fauque P, Jouannet P, Lesaffre C et al (2007) Assisted reproductive technology affects develop mental kinetics, H19 imprinting control region methylation and H19 gene expression in individual mouse embryos. BMC Dev Biol 7:116 Fernandez Gonzalez R, Moreira P et al (2004) Long term effect of in vitro culture of mouse embryos with serum on mRNA expression of imprinting genes, development, and behavior. Proc Natl Acad Sci USA 101:5880 5885 Filipponi D, Feil R (2009) Perturbation of genomic imprinting in oligospermia. Epigenetics 4(1):27 30 Flanagan JM, Popendikyte V, Pozdniakovaite N et al (2006) Intra and inter individual epigenetic variation in human germ cells. Am J Hum Genet 79:67 84 Fortier AL, Lopes FL, Darricarrere N et al (2008) Superovulation alters the expression of imprinted genes in the midgestation mouse placenta. Hum Mol Genet 17:1653 1665 Galetzka D, Weis E, Tralau T et al (2007) Sex specific windows for high mRNA expression of DNA methyltransferases 1 and 3A and methyl CpG binding domain proteins 2 and 4 in human fetal gonads. Mol Reprod Dev 74:233 241 Geuns E, De Rycke M, Van Steirteghem A et al (2003) Methylation imprints of the imprint control region of the SNRPN gene in human gametes and preimplantation embryos. Hum Mol Genet 12:2873 2879 Geuns E, Hilver P, Van Steirteghem A et al (2007a) Methylation analysis of KVDMR1 in human oocytes. J Med Genet 44:144 147 Geuns E, De Temmerman N, Hilven P et al (2007b) Methylation analysis of the intergenic differentially methylated region of DLK1 GTL2 in human. Eur J Hum Genet 15:352 361 Gicquel C, Gaston V, Mandelbaum J et al (2003) In vitro fertilization may increase the risk of Beckwith Wiedemann syndrome related to the abnormal imprinting of the KCN1OT1 gene. Am J Hum Genet 72:1338 1341 Goll MG, Bestor TH (2005) Eukaryotic cytosine methyltransferases. Annu Rev Biochem 74:481 514 Gosden R, Trasler J, Lucifero D et al (2003) Rare congenital disorders, imprinted genes, and assisted reproductive technology. Lancet 361:1975 1977 Halliday J, Oke K et al (2004) Beckwith Wiedemann syndrome and IVF: a case control study. Am J Hum Genet 75:526 528 Hamatani T, Sasaki H, Ishihara K et al (2001) Epigenetic mark sequence of the H19 gene in human sperm. Biochim Biophys Acta 1518:137 144 Hammoud SS, Purwar J, Pflueger C et al (2010) Alterations in sperm DNA methylation patterns at imprinted loci in two classes of infertility. Fert Steril 94(5):1728 1733 Hiura H, Obata Y, Komiyama J et al (2006) Oocyte growth dependent progression of maternal imprinting in mice. Genes Cells 11:353 361 Horsthemke B, Ludwig M (2005) Assisted reproduction: the epigenetic perspective. Hum Reprod Update 11:473 482 Horsthemke B, Wagstaff J (2008) Mechanisms of imprinting of the Prader Willi/Angelman region. Am J Med Genet A 146A:2041 2052 Houshdaran S, Courtessis VK, Siegmund K et al (2007) Widespread epigenetic abnormalities suggest a broad DNA methylation erasure defect in abnormal human sperm. PLoS ONE 2(12): e1289 Huntriss J, Picton HM (2008) Epigenetic consequences of assisted reproduction and infertility on the human preimplantation embryo. Hum Fertil 11:85 94 Kagami M, Nagai T, Fukami M et al (2007) Silver Russell syndrome in a girl born after in vitro fertilization: partial hypermethylation at the differentially methylated region of PEG1/MEST. J Assist Reprod Genet 24:131 136 Kallen B, Finnstrom O, Nygren KG et al (2005) In vitro fertilization (IVF) in Sweden: infant outcome after different IVF fertilization methods. Fertil Steril 84:611 617 Kanber D, Buiting K, Zeschnigk M et al (2009) Low frequency of imprinting defects in ICSI children born small for gestational age. Eur J Hum Genet 17:22 29

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Kaneda M, Okano M, Hata K et al (2004) Essential role for de novo DNA methyltransferase Dnmt3a in paternal and maternal imprinting. Nature 429:900 903 Katari S, Turan N, Bibikova M et al (2009) DNA methylation and gene expression differences in children conceived in vitro or in vivo. Hum Mol Genet 18:3769 3778 Kerjean A, Dupont JM, Vasseur C et al (2000) Establishment of the paternal methylation imprint of the human H19 and MEST/PEG1 genes during spermatogenesis. Hum Mol Genet 9: 2183 2187 Kerjean A, Couvert P, Hearns T et al (2003) In vitro follicular growth affects oocyte imprinting establishment in mice. Eur J Hum Genet 11:493 496 Khosla S, Dean W, Reik W et al (2001) Culture of preimplantation mouse embryos affects fetal development and the expression of imprinted genes. Biol Reprod 64:918 926 Khoueiry R, Ibala Rhomdane S, Me´ry L et al (2008) Dynamic CpG methylation of the KCNQ1OT1 gene during maturation of human oocytes. J Med Genet 45(9):583 588 Kishigami S, Van Thuan N, Hikichi T et al (2006) Epigenetic abnormalities of the mouse paternal zygotic genome associated with microinsemination of round spermatids. Dev Biol 289: 195 205 Kobayashi H, Sato A, Otsu E et al (2007) Aberrant DNA methylation of imprinted loci in sperm from oligospermic patients. Hum Mol Genet 16:2542 2551 Kobayashi H, Hiura H, John RM et al (2009) DNA methylation errors at imprinted loci after assisted conception originate in the parental sperm. Eur J Hum Genet 17:1582 1591 Lane N, Dean W, Erhardt S et al (2003) Resistance of IAPs to methylation reprogramming may provide a mechanism for epigenetic inheritance in the mouse. Genesis 35:88 93 Lees Murdock DJ, De Felici M, Walsh CP (2003) Methylation dynamics of repetitive DNA elements in the mouse germ cell lineage. Genomics 82:230 237 Li JY, Lees Murdock DJ, Xu GL et al (2004) Timing of establishment of paternal methylation imprints in the mouse. Genomics 84:952 960 Lidegaard O, Pinborg A, Andersen AN (2005) Imprinting diseases and IVF: Danish National IVF cohort study. Hum Reprod 20:950 954 Lim D, Browdin SC, Tee L et al (2009) Clinical and molecular genetic features of Beckwith Wiedemann Syndrome associated with assisted reproductive technologies. Hum Reprod 24: 741 747 Lucifero D, Chaillet JR, Trasler JM (2004a) Potential significance of genomic imprinting defects for reproduction and assisted reproductive technology. Hum Reprod Update 10:3 18 Lucifero D, Mann MRW, Bartolomei M et al (2004b) Gene specific timing and epigenetic memory in oocyte imprinting. Hum Mol Genet 3:839 849 Ludwig M, Katalinic A, Gross S et al (2005) Increased prevalence of imprinting defects in patients with Angelman syndrome born to subfertile couples. J Med Genet 42:289 291 Luedi PP, Dietrich FS, Weidman JR et al (2007) Computational and experimental identification of novel imprinted genes. Genome Res 17:1723 1730 Maher ER, Brueton LA, Bowdin SC et al (2003) Beckwith Wiedemann syndrome and assisted reproduction technology (ART). J Med Genet 40:62 64 Mann MR, Lee SS, Doherty AS et al (2004) Selective loss of imprinting in the placenta following preimplantation development in culture. Development 131:3727 3735 Manning M, Lissens W, Bonduelle M et al (2000) Study of DNA methylation patterns at chromosome 15q11 q13 in children born after ICSI reveals no imprinting defects. Mol Hum Reprod 6:1049 1053 Market Velker BA, Zhang L, Magri LS et al (2010) Dual effects of superovulation: loss of maternal and paternal imprinted methylation in a dose dependent manner. Hum Mol Genet 19:36 51 Marques CJ, Carvalho F, Souza M et al (2004) Genomic imprinting in disruptive spermatogenesis. Lancet 363:1700 1702 Marques CJ, Costa P, Vaz B et al (2008) Abnormal methylation of imprinted genes in human sperm is associated with oligozoospermia. Mol Hum Reprod 14:67 73

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Miki H, Lee J, Inoue K et al (2004) Microinsemination with first wave round spermatids from immature male mice. J Reprod Dev 50:131 137 Oakes CC, La Salle S, Smiraglia D et al (2007) Developmental acquisition of genome wide DNA methylation occurs prior to meiosis in male germ cell development. Dev Biol 307: 368 379 Obata Y, Kono T (2002) Maternal primary imprinting is established at a specific time for each gene throughout oocyte growth. J Biol Chem 277:5285 5289 Olek A, Walter J (1997) The pre implantation ontogeny of the H19 methylation imprint. Nat Genet 17:275 276 Ooi SKT, O’Donnell AH, Bestor TH (2009) Mammalian cytosine methylation at a glance. J Cell Sci 122:2787 2791 Orstavik KH, Eiklid K, van der Hagen CB et al (2003) Another case of imprinting defect in a girl with Angelman syndrome who was conceived by intracytoplasmic semen injection. Am J Hum Genet 72:218 219 Owen CM, Segars JH Jr (2009) Imprinting disorders and assisted reproductive technology. Semin Reprod Med 27:417 428 Perkins RM, Hoang Xuan MT (2002) The Russell Silver syndrome: a case report and brief review of the literature. Pediatr Dermatol 19:546 549 Rivera RM, Stein P, Weaver JR et al (2007) Manipulations of mouse embryos prior to implantation result in aberrant expression of imprinted genes on day 9.5 of development. Hum Mol Genet 17:1 14 Rossignol S, Steunou V, Chalas C (2006) The epigenetic imprinting defect of patients with Beckwith Wiedemann syndrome born after assisted reproductive technology is not restricted to the 11p15 region. J Med Genet 43:902 907 Sasaki H, Matsui Y (2008) Epigenetic events in mammalian germ cell development: repro gramming and beyond. Nat Rev Genet 9:129 140 Sato A, Otsu E, Negishi H et al (2007) Aberrant DNA methylation of imprinted loci in super ovulated oocytes. Hum Reprod 22:26 35 Shamanski FL, Kimura Y, Lavoir MC et al (1999) Status of genomic imprinting in mouse spermatids. Hum Reprod 14:1050 1056 Sutcliffe AG, Peters CJ, Bowdin S et al (2006) Assisted reproductive therapies and imprinting disorders a preliminary British survey. Hum Reprod 21:1009 1011 Svensson J, Bj€ornstahl A, Ivarsson SA (2005) Increased risk of Silver Russell syndrome after in vitro fertilization? Acta Paediatr 94:1163 1165 Trasler JM (2006) Gamete imprinting: setting epigenetic patterns for the next generation. Reprod Fertil Dev 18:63 69 Trasler JM (2009) Epigenetics in spermatogenesis. Mol Cell Endocrinol 306:33 36 Ueda T, Abe K, Miura A et al (2000) The paternal methylation imprint of the mouse H19 locus is acquired in the gonocyte stage during foetal testis development. Genes Cells 5:649 659 Weksberg R, Shuman C, Caluseriu O et al (2002) Discordant KCNQ1OT1 imprinting in sets of monozygotic twins discordant for Beckwith Wiedemann syndrome. Hum Mol Genet 11:1317 1325 Weksberg R, Shuman C, Smith AC (2005) Beckwith Wiedemann syndrome. Am J Med Genet C Semin Med Genet 137C:12 23 Wilkins Haug L (2009) Epigenetics and assisted reproduction. Curr Opin Obstet Gynecol 21: 201 206 Williams CA (2005) Neurological aspects of the Angelman syndrome. Brain Dev 27:88 94 Yamazaki T, Yamagata K, Baba T (2007) Time lapse and retrospective analysis of DNA methy lation in mouse preimplantation embryos by live cell imaging. Dev Biol 304:409 419 Young LE, Fernandes K, McEvoy TG et al (2001) Epigenetic change in IGF2R is associated with fetal overgrowth after sheep embryo culture. Nat Genet 27:153 154

Chapter 2

Germ Cell Cancer, Testicular Dysgenesis Syndrome and Epigenetics Kristian Almstrup, Olga Mlynarska, and Ewa Rajpert-De Meyts

Abstract Testicular dysgenesis syndrome (TDS) comprises a series of disorders, which are linked by an aberrant development of the testis, including abnormal differentiation of somatic cells and delayed maturation of germ cells. Environmental factors (including endocrine disrupters) are thought to impair development of the testis. It is likewise though that epigenetic patterns can be regulated by environmental stimuli and we here review the current knowledge on epigenetic patterns in testicular dysgenesis, which to a large extend is limited to testicular cancer. Testicular Germ Cell Tumors (TGCTs) of young adults originate from a precursor cell, carcinoma in situ testis, which has features of arrested and transformed gonocytes, including high expression of embryonic pluripotency genes and low genome methylation. Overt TGCTs are capable of a marked re-programming with loss of germinal phenotype and increased somatic differentiation. These features suggest a role for epigenetic changes in the pathogenesis and progression of TGCT. In addition to chromatin modifications and gene imprinting, other regulatory mechanisms have been emerging, i.e., miRNAs, which are instrumental in the regulation of gene transcription. It is possible that epigenetic reprogramming during early development may also be a cause of genital malformations and reduced spermatogenesis, but more research is needed to identify the causative environmental factors, their precise targets and affected pathways. Keywords Epigenetics Testicular Cancer Testicular Dysgenesis Syndrome

K. Almstrup (*), O. Mlynarska, and E. Rajpert De Meyts Department of Growth and Reproduction, Rigshospitalet, Section GR 5064, Blegdamsvej 9, 2100 Copenhagen, Denmark e mail: [email protected], [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 2, # Springer Verlag Berlin Heidelberg 2011

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Introduction

Germ cell cancer in human males occurs predominantly in the testis, although in very rare instances it may also occur in extragonadal positions. Testicular germ cell tumors (TGCTs) are quite unique, as they occur predominantly in young men, and their initiation begins most likely during early development of gonads. Another unique feature of these tumors is their ability to transform into virtually any type of somatic tissue, which is strongly linked to both genomic and epigenetic changes that affect gene expression. Testicular tumors are epidemiologically and clinically linked to other disorders of male reproduction, with a common feature of developmental origins. These observations led to the proposed grouping of these disorders within the so-called Testicular Dysgenesis Syndrome (TDS). Growing evidence suggests that the majority of cases of TDS are caused predominantly by external factors, which may act directly on numerous pathways involved in testis development, and are also suspected of inflicting epigenetic changes that may be instrumental in reprogramming of germ cell differentiation. The factors remain to be identified and proven to be causal, but factors related to modern lifestyle, including endocrine disruptors have been under growing scrutiny as possible culprits. These aspects are reviewed in this chapter, with emphasis on germ cell tumors, where most information is available. The chapter focuses on human disorders, however, very little is known about the epigenetic reprogramming during human testis development and germ cell differentiation, therefore we discuss relevant findings gathered from studies in mice as well.

2.2

Epigenetics in Testicular Development: From Specification of Primordial Gem Cells to Formation of the Gonads

During embryonic development, embryonic ectoderm gives rise to somatic germ layers and primordial germ cells (PGCs) which are first observed at embryonic day E7.25 (mice) in the posterior end of the primitive streak. However already before specification of PGC’s, pre-PGC’s at E6.25 starts to express BLIMP1 (Ohinata et al. 2005), which is crucial for suppression of somatic programming of Hox genes (Ohinata et al. 2005). Around embryonic day E8 (mice), the PGCs start to migrate to the genital ridge and form a primitive gonad, tightly associated with the mesonephros, which later develops into kidney and epididymis. At the colonizationstage, the primitive gonad is still sexually bipotential and sex-determination first takes place between E10.5 and E11.5. While female germ cells subsequently enter meiosis, male germ cells, gonocytes, continue to proliferate until they differentiate to pre-spermatogonia that enter mitotic arrest. It is thought that sex-determination is determined by the expression of the male specific gene Sry in precursors of Sertoli cells. If Sry is expressed, this will trigger expression of Sox9 and several

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other masculinising genes, which are essential for Sertoli cells differentiation (recently reviewed by Koopman 2009). Sertoli cells then embed the PGC and form cords, which in mice are seen at E12. Already during specification and migration, PGCs undergo epigenetic reprogramming, including genome-wide DNA de-methylation (starting at E8), progressive erasure of H3K9me2 (starting at E7.5) and subsequently establishment of H3K27me3 (starting at E8.25) in a progressive, cell-by-cell manner, presumably depending on their developmental maturation (Seki et al. 2005, 2007). These are all modifications that repress the DNA and the PGC is thus free of repressive chromatin between E7.5 and E8.25. As the loose chromatin structure thus would allow massive transcription, in this period PGCs are kept transcriptionally quiet by transient repression of RNA polymerase II dependent transcription (Seki et al. 2007). In addition, prior to H3K9 de-methylation PGCs repress the histone methyltransferase GLP, which mediates H3K9 methylation (Tachibana et al. 2008). In migrating PGCs, BLIMP1 (expressed from E6.25) is found in the nucleus is complexed with PRMT5, which mediates symmetrical methylation of histones H2A and H4 at arginine 3 (H2AR3me2, H4R3me2). When PGCs arrive in the genital ridge this complex translocates to the cytoplasm leading to reduced levels of H2AR3me2 and H4R3me2, resulting in widespread epigenetic modifications and transcriptional repression (Ancelin et al. 2006). Translocation of the BLIMP1/ PRMT5 complex leads to down-regulation of Blimp1, which coincides with re-expression of pluripotency-associated genes like Nanog and Sox2 (Ohinata et al. 2005; Yabuta et al. 2006). When PGCs colonize the genital ridge (E11.5), extensive chromatin and epigenetic reprogramming occurs. This includes a second wave of DNA de-methylation, which includes erasure of parental imprints (Hajkova et al. 2002). Some parental imprints are already set again starting at E14.5, while others are first acquired at the newborn stage (Li et al. 2004). Linker histone H1, H3K9ac, H3K9me3, H3K27me2, DAPI chromocenters and, as mentioned above, H4/H2A R3me2, all disappear around gonadal colonization at E11.5 (Hajkova et al. 2008) as illustrated in Fig. 2.1. Using FACS sorting of dissected urogenital ridges in an Oct4-GFP transgenic mouse, Hajkova et al. (2008) were also able to show two Oct4þ (GFP) populations of PGCs at E11.5: One population of E11.5 PGC was at a more advanced stage, with low levels of H2A.Z, linker H1, H3K9ac, H3K9me3 and H3K27me3 as well as H2A/H4R3me2, when compared to the other more primitive population, which showed higher levels of these marks. The primitive population of E11.5 PGCs showed variations in the levels of DNA methylation (5-methylcytosine levels), whereas the advanced population was practically devoid of DNA methylation (Hajkova et al. 2008). It is thus evident that extensive epigenetic modifications of PGC occur at the time of gonadal colonization. Around E13, the proliferative PGCs have reached sufficient numbers, and female germ cells enter meiotic prophase while male germ cells (gonocytes) arrest in the G0/G1-phase of the cell cycle until after birth. After birth, gonocytes give rise to primitive spermatogonia (spermatogonial stem cells) that start to proliferate and develop into spermatocytes at days 10 14 post natal (Fig. 2.1).

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By E12.5 most DNA methylation is lost (Reik and Walter 2001) and de novo methylation is initiated in males at E14.5 and thereafter in females such that the mature gametes of both sexes will eventually become highly methylated. However, an additional de-methylation wave takes place in pubertal and adult male germ cells after meiosis during spermatogenesis (Bernardino et al. 2000; del Mazo et al. 1994; Norris et al. 1994), where also histones are exchanged with transition proteins and finally protamines (Fig. 2.1). In addition, remethylation and further chromatin modifications may possibly occur as the spermatozoa mature in the epididymis (Xie et al. 2002).

Fig. 2.1 Schematic drawing of major epigenetic events during male gonadal development and development of testicular germ cell tumors derived from CIS cells (modified from Rajpert De Meyts 2006). DNA methylation patterns are greatly simplified as a number of different categories of sequence specific methylation (CpG, non CpG, repeat elements, X chromosome, imprinted genes etc.) are independently regulated. Likewise, represented regulations are not exact but are intended as an overview. Data on H3K27me3 (Hajkova et al. 2008; Seki et al. 2007) is somewhat divergent and the depicted pattern is adapted from Seki et al. MSCI Meiotic sex chromosome inactivation, TER teratoma, YST Yolk sac tumor, CHC choriocarcinoma

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TDS: A Collection of Reproductive Disorders with a Possible Common Pathogenesis

TDS was proposed by Skakkebæk and co-workers (2001) as a set of developmental disorders that affect male reproduction. The concept was mainly based on testicular germ cell cancer but studies of this cancer suggested that several other disorders may have a similar aetiology and pathogenesis. These disorders (very briefly summarized below) include some forms of undescended testis, hypospadias and reduced spermatogenesis, all linked together by common risk factors that operate in utero or early infancy. The TDS concept is outlined in Fig. 2.2.

2.3.1

Testicular Cancer

The vast majority of human testicular tumors are derived from germ cells, and most of these tumors occur in young men (between 17 and 40 years of age). Germ cell tumors can also occur in infants (yolk sac tumor or teratoma) and older men (spermatocytic seminoma), but these are very rare; moreover they are thought to

Fig. 2.2 Schematic depiction of the TDS concept. Environmental factors together with genetic aberrations or polymorphisms that increase sensitivity to those factors cause testicular dysgenesis, which results in an impaired differentiation and function of somatic cells that in turn leads to a delayed germ cell maturation and decreased secretion of reproductive hormones. This cascade of events is manifested by several clinical outcomes, including impairment of testicular descent, hypospadias, neoplastic transformation of germ cells (CIS in testes or gonadoblastoma in intersex gonads) or some forms of impaired testicular function and spermatogenesis later in life. Adapted from Skakkebæk et al. (2007)

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be of purely genetic origin and not part of TDS (Oosterhuis and Looijenga 2005). Somatic cell tumors of the testis (sex cord stromal neoplasms and Leydig cell tumors) are much less frequent, have also predominantly a genetic background, and are not considered as part of TDS. Thus in this chapter we shall focus on TGCTs of young adults, which can be divided into two main types: seminoma (also called classical seminoma to distinguish it from spermatocytic seminoma) and non-seminoma (Fig. 2.1). Seminoma is a homogeneous tumor composed of mitotically dividing germ cells, whereas non-seminomas are heterogeneous tumors that may contain varying proportions of undifferentiated embryonal carcinoma (EC), partially differentiated somatic tissues (teratoma), and extra-embryonic elements such as choriocarcinoma and yolk sac tumor (Ulbright et al. 1999). A characteristic feature of TGCTs of young adults is their origin from a premalignant lesion called carcinoma in situ (CIS) testis (Skakkebaek et al. 1987), which is also known as intratubular germ cell neoplasia (ITGCN) or transepithelial intratubular neoplasia (TIN). A similar lesion, called gonadoblastoma, which consists of germ cells virtually identical to CIS cells, but surrounded by undifferentiated somatic cells in structures that do not resemble tubules, occurs in gonads of patients with more severe disorders of sexual differentiation, who most often are phenotypic female (Cools et al. 2006; Jorgensen et al. 1997). Comparative studies of protein markers (reviewed in Rajpert-De Meyts et al. 2003) and the global gene expression profile of CIS cells (Almstrup et al. 2004, 2007; Sonne et al. 2009) provided evidence that CIS cells are transformed gonocytes that failed to mature during development (Rajpert-De Meyts 2006). The transformation of delayed gonocytes to CIS cells most likely occurs during adaptation to the changed endocrine environment of the peri- and post-pubertal testis, when the cells undergo polyploidisation (Oosterhuis et al. 1989) and acquire a characteristic pattern of genomic aberrations, including gain of chromosome 12p (Ottesen et al. 2003; Rosenberg et al. 2000), often manifested as the isochromosome 12p (Atkin and Baker 1982). In analogy to PGCs and gonocytes, CIS cells highly express transcription factors associated with embryonic stem cell pluripotency and somatic tissue-specific stem cell lineages, including e.g., POU5F1/ OCT-3/4, NANOG, GDF3, KIT and AP-2g (Almstrup et al. 2004; Biermann et al. 2007; Hoei-Hansen et al. 2004, 2005; Looijenga et al. 2003; Skotheim et al. 2005; Sperger et al. 2003). Seminoma is very similar to CIS and shows expression of virtually the same genes (Almstrup et al. 2005; Gashaw et al. 2007; Hoei-Hansen 2008). By contrast, non-seminomas display a very different profile, which is characterized by the lack of expression at the protein level of the commonly used CIS/seminoma markers, e.g., PLAP, KIT, PDPN as well as germ cell-specific genes, e.g., VASA or DAZL1 (Korkola et al. 2005, 2006). This is consistent with the reprogramming of nonseminomas into a more embryonic cell type (Oosterhuis and Looijenga 2005; Rajpert-De Meyts 2006). The embryonic pluripotency genes are highly active in EC but down-regulated in differentiated components of teratomas, in line with the latter recapitulating the embryonic development (Korkola et al. 2006; Skotheim et al. 2005).

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Undescended Testis and Hypospadias

Undescended testis (cryptorchidism) and hypospadias are common urogenital abnormalities, usually diagnosed in newborn or infantile boys. These abnormalities have a broad spectrum of severity (especially hypospadias) and the most severe manifestations can also occur in combination with other malformations. Importantly, cryptorchidism is the most significant risk factor for testicular cancer (Dieckmann and Pichlmeier 2004). Testicular descent in humans first occurs trans-abdominally by enlargement of the gubernaculum and then as the inguino-scrotal phase when the testis gradually glides to its final position (Hutson and Hasthorpe 2005). In rodents, and most likely also in humans, the trans-abdominal phase is dependent on INSL3, a peptide hormone acting via its receptor, RXFP2, previously known as LGR8 (Nef and Parada 1999; Zimmermann et al. 1999). The second phase is mainly dependent on androgens; thus, different forms of androgen insufficiency (e.g., androgen receptor mutations) are often associated with undescended testes. Undoubtedly, other factors are also involved in both phases of descent, but the pathways involved are yet to be elucidated. The aetiology of hypospadias is likewise multi-factorial and not well understood, but, in the milder forms, androgen insufficiency is one of the known factors (Ferraz-de-Souza and Achermann 2007).

2.3.3

Impaired Spermatogenesis with Developmental Aetiology

Quantitative or qualitative reduction of spermatogenesis can be caused by a very large number of factors, making this disorder one of the most heterogeneous diseases, and also one of the least recognized as it is only diagnosed when it begins to affect a man’s fertility. We would like to stress that only the forms of spermatogenesis impairment that are caused by aberrations in testis development are considered part of TDS. The proportion of men that are affected is not known, because the only currently existing evidence for developmental pathogenesis is the presence of histological changes, such as distorted tubules, undifferentiated Sertoli cells, persisting gonocytes (CIS cells), patchy distribution of Sertoli-cell-only tubules or microlithiasis (Jørgensen et al. 2010). Another important point to make is that semen quality is not only defined by sperm density and total sperm count, but these variables are the easiest to establish and to correlate with clinical outcomes, so most of the existing evidence is based on the sperm production. According to several studies, male fecundity begins to be affected (usually documented as increasing waiting time to pregnancy) when sperm concentration falls down to around 40 mill/ml (Bonde et al. 1998; Guzick et al. 2001; Slama et al. 2002). However, signs of impaired development can be seen in the testis producing sperm even within the normal range.

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Endocrine Disrupters and Their Possible Influence on Epigenetic Patterns

Endocrine disrupters are chemicals that interfere with the endocrine balance in higher mammals and were at first thought to be chemicals that mimic intrinsic hormones i.e., synthetic estrogens (Sharpe and Skakkebaek 1993; Toppari et al. 1996). Later it has become clear that chemicals that interfere with all aspects of steroid metabolism i.e., aromatase inhibitors, androgen receptor blockers, can disrupt the endocrine function (Sharpe and Skakkebaek 2008). Even more importantly, the ubiquitous presence of numerous chemicals present in the environment (food, home, workplace) can lead to synergistic effects of compounds acting in a similar fashion or affecting different targets in co-operating pathways (Kortenkamp 2008). The male reproductive system appears to be especially vulnerable to endocrine disrupters because most of the known compounds in this category are known to target key pathways in testis development, i.e., anti-androgens, such as DDE, vinclozolin (Gray et al. 1994; Wilson et al. 2008) or phthalates (Mylchreest et al. 1999).

2.4.1

Evidence from Human Studies

There is very little direct evidence from human studies for a role of endocrine disrupters in testicular cancer, and only a very limited recent evidence for a role in cryptorchidism. Most of the data come from indirect epidemiological studies that are consistent with a possibility of a disruption of the early testis development by xenobiotic chemicals. The first indication of a predominant role of external factors in the aetiology of testicular cancer was the dramatic increase observed in the incidence of this cancer in the second half of the twentieth century in many countries around the world, but with marked geographic or ethnic differences, suggestive of the greatest risk among white men of Scando-Germanic origin in well developed countries (Huyghe et al. 2003; McGlynn et al. 2005; Parkin et al. 1997; Richiardi et al. 2004; Shah et al. 2007). Similar trends in other manifestations of TDS were noted first in Denmark and then in other countries (Aschim et al. 2004; Boisen et al. 2004; Carlsen et al. 1992; Hotaling and Walsh 2009; Moller et al. 1996; Paulozzi 1999), although some studies questioned the existence of such trends (Akre and Richiardi 2009). These epidemiological observations point to the primary importance in the pathogenesis of factors acting during the intrauterine or early postnatal developmental window (Rajpert-De Meyts and Skakkebaek 1993; Skakkebaek et al. 2001; Weir et al. 2000). Initially, the excess of estrogens was suspected (Henderson et al. 1979; Moss et al. 1986; Sharpe and Skakkebaek 1993; Toppari et al. 1996), later antiandrogenic effects were indicated as of primary importance (Wilson et al. 2008),

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which is in line with an increased risk of germ cell neoplasia in individuals with the androgen insensitivity syndrome. Few studies so far investigated the exposure of patients with TGCT/TDS to xenobiotics. Beside the long time interval from the fetal exposure to the disease manifestation, a big problem is the possible chemical interactions of mixtures, which are very difficult to identify in the human setting (Kortenkamp 2008). Studies of exposure to DES and other estrogens, although consistent with an increased risk of TGCT and poor semen quality in males, were not conclusive (Dieckmann and Pichlmeier 2004; Storgaard et al. 2006; Strohsnitter et al. 2001; Swan 2000). A few studies investigated exposure to other persistent endocrine disruptors, among them Hardell et al. (2003) found higher serum levels of polychlorinated biphenyls, DDE and several other persistent organic pollutants in mothers of men with testicular cancer, but no significant changes in the serum levels were found in men themselves (Hardell et al. 2003, 2006). Another study from the US demonstrated a greater burden of DDE and some chlordane-related compounds in the TGCT cases (McGlynn et al. 2008). More recently, a comparative study of these chemicals in milk samples of Danish and Finish mothers demonstrated a very different profile in the two countries, consistent with a higher risk of TDS in Denmark as compared to Finland (Krysiak-Baltyn et al. 2009). Another group of ubiquitous pollutants, which in laboratory animals cause a TDS-like syndrome are phthalates (Hutchison et al. 2008). Some studies linked phthalate exposure to under-masculinization of boys, including changes in the androgenic behavioral programming (Matsumoto et al. 2008; Swan et al. 2005, 2009). However, there is no evidence as yet from the human studies if any of the above mentioned endocrine disruptors cause epigenetic changes. Some evidence begins to emerge from animal studies, which are briefly summarized below.

2.4.2

Evidence from Studies in Animals

There are numerous toxicological studies of endocrine disrupters in animals, but few explored possible epigenetic effects. In mice exposed to DES, lesions of the rete testis and some genitourinary tumors (but not TGCTs) were observed and these effects were reported to be transmitted transgenerationally, suggesting an epigenetic mechanism (Newbold et al. 2000). Exposing rats in utero to phthalates caused a TDS-like phenotype including cryptorchidism, hypospadias, and a delay of gonocyte maturation (reviewed in Foster et al. 2001; Gray et al. 2000; Hutchison et al. 2008; Mylchreest et al. 1999), but epigenetic changes were not investigated in those studies. The most striking evidence for the involvement of epigenetic regulation in effects of some endocrine disrupters came from studies of animals exposed in utero to the anti-androgenic fungicide vinclozolin, which caused defects of spermatogenesis that persisted in a large proportion of the progeny over four generations (Anway et al. 2005). In animals that were allowed to age, numerous other disorders appeared, including kidney and prostate disease, immune abnormalities as well as an increased

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frequency of tumors, but testicular malignancies were not observed (Anway et al. 2006). Interestingly, all abnormalities were transmitted via male germ line, suggesting that in contrast to oocytes, in male germ cells exposed during embryonic sex determination some epigenetic regulation (i.e., DNA methylation) appeared to be permanently reprogrammed (Skinner 2007).

2.5

Epigenetics of Germ Cell Tumors and Testicular Dysgenesis

Normal germ cells undergo extensive chromatin changes during the process of maturation and gametogenesis. In particular the process of homologous recombination during the meiotic division, where double-strand DNA breaks are induced, requires the activation of specialized DNA repair machinery (Longhese et al. 2009). Constitutive activation of the DNA damage response process, which may be brought upon by various defects of this network, e.g., p53 mutations, is commonly found in numerous types of somatic solid tumors (Bartkova et al. 2006). By contrast, these pathways are largely silenced in TGCTs, including the pre-invasive CIS, as demonstrated by the lack of activation of the ATM kinase and phosphorylation of the downstream target histone gH2AX (Bartkova et al. 2005), or downstream effectors MDC1 and 53BP1 (Bartkova et al. 2007a) in CIS cells. This feature of TGCTs most likely reflects the physiological silencing of DNA repair pathways in foetal germ cells and may at least to some extent explain the exquisite sensitivity of TGCT (and normal germ cells) to irradiation and chemotherapy, which inflict DNA damage (Bartkova et al. 2007b). Much of the current knowledge about epigenetic changes in testicular cancer has been gained from studies on primary tissue, but to date no line representing the CIS cell has been identified. Techniques such as restriction landmark genomic scanning (RLGS) that require isolation of bulk DNA have, however, been used to study epigenetic changes in the overt tumor types derived from CIS, seminoma, and non-seminomas.

2.5.1

Chromatin Changes

Several studies have shown striking differences between seminomas and nonseminomas: there is little or no methylation in seminomas, whereas hypermethylation of specific gene promoters was observed in non-seminomas, increasing together with higher differentiation of the tumor. This was first shown by Peltomaki et al. by HpaII/MspI analysis (Peltom€aki 1991). Smiraglia et al. later confirmed this pattern by the use of RLGS and found a similar pattern, with seminomas greatly de-methylated, while the level of DNA methylation in non-seminomas was relatively high and comparable to that of solid tumors derived from somatic cells (Smiraglia et al. 2002). More recently, a matching pattern of global DNA

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methylation (low in seminoma and high in non-seminomas) was confirmed by immunohistochemistry and extended to CIS cells, which also showed global DNA de-methylation (Netto et al. 2008). The Low CpG methylation level in CIS was indeed confirmed in our laboratory as shown in Fig. 2.3, where immunohistochemical staining of CIS cells for 5-methylcytosine is shown. In line with these data, a comparative gene expression profiling of CIS, seminomas and nonseminomas revealed that the de novo DNA methyltransferase DNMT3B and the related protein DNMT3L were upregulated in non-seminomas (Almstrup et al. 2005). Interestingly, DNMT3L is known to interact directly with histone H3 when unmethylated at K4 (Ooi et al. 2007). De novo methylation consequently may be prevented at these CpG islands due to methylated H3K4 and is consistent with a strong negative correlation between DNA methylation and the presence of H3K4 methylation in several cell types (Weber et al. 2007). Typical conditions for altered epigenetic control in cancer are reduced DNA methylation of most of the genome, which may lead to genomic instability, and hypermethylation of CpG islands located in the promoter regions of tumor suppressors and tumor-related genes (Esteller 2002). CIS and TGCTs, however, represent an exception probably due to their origin from foetal germ cells, most likely from PGCs or gonocytes, the cell types in which genomes are physiologically under-methylated (Trasler 1998). This again points to the foetal origin of CIS as probably a key event in the pathogenesis of TGCT and indicates that the loss of epigenetic control may be implicated in the pathogenesis of TDS-related disorders. Poor sperm have been reported to show altered epigenetic profiles compared to normal sperm (Hammoud et al. 2009; Houshdaran et al. 2007; Poplinski et al. 2009).

Fig. 2.3 Immunohistochemical double staining for 5 methyl cytosine (5 mC, reddish brown) and placental like alkaline phosphatase (PLAP, dark blue) in carcinoma in situ (CIS) testis. PLAP is a classical cytoplasmic marker for CIS cells, which show very low 5 mC staining in the nucleus indicating hypo methylation of the genome. This is in line with a previous report showing low levels of 5 mC in CIS, varying amounts in seminoma and high levels in non seminomas (Netto et al. 2008). Note that all other normal germ cells, including spermatogonia (marked with arrows) as well as Sertoli and Leydig cells show high 5 mC levels and no PLAP signal. Bar represents 200 mm

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However, it remains to be established whether these changes observed in adult men are the result of events occurring during development, and thus can be classified as TDS. Based on the findings of transgenerational effects and late-onset diseases transmitted by sperm of animals who were exposed in utero (Anway et al. 2005), it is likely that events occurring during early differentiation of germ cells may affect the quality of sperm later in life. We are, however, in need of developing molecular markers that would corroborate this hypothesis.

2.5.2

Methylation Profiles of Specific Genes Important for Pluripotency and Tumorigenesis

A hallmark of CIS cells is expression of a range of genes known to be responsible for stemness and pluripotency (Almstrup et al. 2004; Rajpert-De Meyts et al. 2003). One of the best-known pluripotency markers is POU5F1, better known as OCT3/4. OCT3/4 is a key transcriptional factor expressed in pluripotent stem and germ cells as well as in CIS, seminomas and EC, the undifferentiated stem cell for nonseminomas. However, if EC differentiates into yolk sac tumor, choriocarcinoma or teratoma, expression of OCT3/4 becomes repressed (Looijenga et al. 2003). De Jong et al. compared the methylation pattern of the upstream region of the OCT3/4 gene in different types of TGCTs and in normal tissue. Seminoma and EC showed a hypomethylated upstream region whereas differentiated non-seminomas, teratoma and yolk sac tumor, which lack OCT3/4 expression, were found to be hypermethylated in the upstream region (De Jong et al. 2007). Methylation profiles of tumor-related genes in TGCTs differ from those in neoplasms of somatic cell origin, in particular testicular malignant lymphomas (Kawakami et al. 2003a). Methylation of E-cadherin gene (CDH1), CDKN2B, CDKN2A, BRCA1, RB1, VHL, RASSF1A, RARB, and GSTP1 was not detected in any of the investigated TGCT tissue samples, whereas tissue samples of testicular lymphoma showed hypermethylation of CDH1, RASSF1A, and RARB (Kawakami et al. 2003a). A study performed by Honorio et al. (2003) agrees with these findings, showing the absence of, or infrequent, promoter methylation at CDKN2A, GSTP1, RARB, DAPK and CDH1, in both seminomas and non-seminomas. However, in the same study, some non-seminomatous TGCTs displayed frequent methylation of RASSF1A. This disagreement may be caused by the different CpG sites of RASSF1A analysed in the two studies. In addition, studies of methylation status of the O6-Methylguanine-DNA Methyltransferase (MGMT), showed methylation in some of the non-seminomas but not in seminomas (Smith-Sørensen et al. 2002). Likewise, Zhang et al. (2005) found that the 50 ends of MAGE-A1 and MAGE-A3 were unmethylated in seminomatous tumors regardless of their expression, whereas methylation of these loci in the non-seminomas apparently prevented their expression. SYCP1 on the other hand remained unmethylated, regardless of expression in both tumor types.

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This implies that 50 upstream methylation patterns do not always dictate whether the downstream gene is expressed, which is in line with recent reports of the different usage of DNA methylation in CpG-rich and poor promoters (Weber et al. 2007). Expression of the BLIMP1/PRMT5- complex is responsible for suppression of the somatic differentiation program via Hox genes and leads to dimethylation of arginine 3 on histone H2A and H4 (H2AR3me2, H4R3me2). The BLIMP1/PRMT5 complex, as well as H4/H2A R3me2, was detected in CIS and most seminomas and, at the same time, down regulated in EC and other non-seminomas. These results led to the suggestion that H4/H2A R3me2 is a mechanism by which CIS and seminomas maintain their undifferentiated form, while loss of H4/H2A R3me2 causes differentiation of non-seminomatous tumor cells into various somatic tissue components (Eckert et al. 2008). The emerging picture of DNA methylation in TGCTs is thus an overall undermethylated CIS genome, which is retained in seminomas but becomes highly methylated in non-seminomas.

2.5.3

X-Linked Genes and XIST Expression

The possible involvement of the X chromosomes in the pathogenesis of TGCT has been implicated by several studies (Hasle et al. 1995; Lykkesfeldt et al. 1983; Rapley et al. 2000; van Echten et al. 1995). The number of X chromosomes is often increased in TGCTs (Rosenberg et al. 1999; Summersgill et al. 2001; van Echten et al. 1995), although this is most likely a reflection of their polyploidy. Moreover patients with Klinefelter’s syndrome (47 XXY) more often develop extragonadal germ cell tumors (Hasle et al. 1995). A TGCT susceptibility gene (TGCT1), also associated with cryptorchidism, was identified on Xq27 by a genome-wide linkage study, with only fragile X mental retardation 1 (FMR1) gene characterized in this region so far (Rapley et al. 2000) but this association has not been confirmed in subsequent studies (Rapley 2007). In mammalian cells with more than one X chromosome, nature has evolved a system for dosage-compensation of specific alleles by transcriptional inactivation. The X-inactive specific transcript, Xist is transcribed from inactive X chromosome (s) (Brown et al. 1991), and functions as a structural RNA covering the inactive X in a cis-like manner inactivating its “own” chromosome (Penny et al. 1996). After Xist-mediated inactivation, Xist is no longer needed to maintain the inactive state (Brown and Willard 1994), but instead additional factors, including variant histones and hypo-acetylation of histones H2A, H3 and H4 maintains X-inactivation. (Belyaev et al. 1996; Costanzi and Pehrson 1998; Hoyer-Fender et al. 2000; Jones et al. 1998; Mermoud et al. 1999; van der Vlag and Otte 1999; Wang et al. 2001). The silent non-expressed Xist allele on the active X-chromosome is methylated, while the expressed allele on the inactive X is unmethylated (Bernardino et al. 2000; Norris et al. 1994), and Xist-expression seems dependent on the methylation

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status of the X-chromosomes. However, expression of Xist is also regulated by another noncoding transcript, Tsix, that is transcribed antisense to Xist (Lee et al. 1999). Tsix has a dual function as both a repressor of the steady-state level of Xist transcription and as a regulator of the distribution of Xist transcripts within the nucleus (Lee et al. 1999; Morey et al. 2001; Stavropoulos et al. 2001). Xist is essentially undetectable in somatic male XY-cells but it is expressed in spermatocytes in the late stages of first meiotic prophase in the testis (De La Fuente et al. 1999; Farazmand et al. 2001; McCarrey and Dilworth 1992; Richler et al. 1992; Salido et al. 1992). Even though sex chromosomes are inactivated at the same stage of spermatogenesis by formation of the meiotic sex chromosome inactivation (MSCI) complex, Xist is not essential for proper MSCI (McCarrey et al. 2002; Turner et al. 2002). As reviewed by Turner (2007) MSCI is regulated by coating the large unpaired meiotic X-Y bivalent with BRCA-1 which attracts the ATR kinase to phosphorylate H2AX at serine-139 (gH2AX) leading to condensation of chromatin on the X and Y chromosomes. MSCI is further substantiated by additional chromatin modifications such as H2A ubiquitylation, deacetylation of H3 and H4, sumoylation and H3K9me2. XIST has also been found expressed in practically all TGCT subtypes including CIS and spermatocytic seminoma (Kawakami et al. 2003b; Looijenga et al. 1997). In fact, detection of the unmethylated 50 -end of XIST in plasma has been suggested as a possible diagnostic tool for the identification of germ cell tumors (Kawakami et al. 2004). The 50 -end of Xist in XY males is methylated while it is found unmethylated in TGCTs (following Xist expression) and methylation-specific PCR of this fragment on plasma samples thus represents a possible diagnostic tool. The downside is however that this is only possible for disseminated tumors where already well-established and easy measurable plasma markers as HCG, AFP and LDH exist. Kawakami et al. (2003b) showed by bisulphite genomic sequencing, that the methylation status of three X-linked genes (AR, FMR1 and GPC3), which normally do not escape X-inactivation (Allen et al. 1992; Huber et al. 1999; Kirchgessner et al. 1995), was unmethylated- and presumably active- on super-numerical X-chromosomes in both seminomas and non-seminomas, which was not influenced by XIST expression (Kawakami et al. 2003b). The role of XIST expression in TGCTs is thus probably different from its X-inactivation function observed in XX females and whether it actually has a function in chromosome inactivation in TGCTs is at present doubtful.

2.5.4

Gene Imprinting

Two genes often investigated to elucidate imprinting patterns are Insulin-like Growth Factor 2 (IGF2) and H19 both found at the 11p15 locus. While IGF2 is expressed from the paternal allele, H19 is expressed from the maternal allele (Rainier et al. 1993). The upstream region of H19 gene in human consists of

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seven CTCF (CCCTC binding factor)-binding sites (Bell and Felsenfeld 2000; Hark et al. 2000) with the sixth binding site demonstrated to be the key regulatory factor for IGF/H19 genes (Takai et al. 2001). CTCF is able to bind only unmethylated alleles. The unmethylated maternal allele is thus occupied with CTCF, which in turn blocks the IGF2 promoter and, consequently, prevents IGF2 expression. On the contrary, CTCF binding is prevented at the methylated paternal allele, which makes the enhancer region available, and IGF2 expression is stimulated (Bell and Felsenfeld 2000; Hark et al. 2000). Expression of H19 and the other imprinted genes becomes biallelic at E11.5 in developing germ cells in mice (Szabo´ and Mann 1995; Villar et al. 1995), which indicates that imprinting patterns are erased at this time of development. Biallelic expression of H19 has also been demonstrated in TGCTs (van Gurp et al. 1994). Kawakami et al. (2006) investigated methylation status of the promoter and CTCF-binding site upstream from H19 gene in TGCTs and testicular lymphomas. Both seminomatous and non-seminomatous TGCTs showed a biallelic unmethylated promoter and CTCF-binding site upstream of H19. Peripheral blood lymphocytes and testicular lymphoma on the other hand showed differential or monoallelic methylation. Similar results were reported by Sievers et al. (2005) by methylation-sensitive single nucleotide primer extension (MS-SNuPE). In other studies, hypomethylation of the sixth CTCF-binding site was detected in somatic tissue-derived cancers, such as osteosarcoma or bladder cancer (Takai et al. 2001; Ulaner et al. 2003), whereas the promoter region of H19 in osteosarcomas was hemi-methylated (Ulaner et al. 2003) and CpG regions surrounding the CTCF-binding sites in bladder cancer were significantly methylated (Takai et al. 2001). It has become clear that the methylation patterns of the promoter region and CTCF-binding site upstream of H19 are different in TGCTs than in somatic-tissue derived cancers. TGCTs exhibit thorough unmethylation of both promoter and CTCF-binding site regions, while levels of methylation in somatic neoplasms differ in those regions. Kawakami et al. (2006) suggested that the methylation status of H19 in TGCTs derives from “erasure of methylation”, whereas in somatic cancers, it is from “gain of hypomethylation”. Likewise, human foetal germ cells have a predominantly unmethylated region of H19, but in mature germ cells in adults, this region becomes significantly methylated (Kerjean et al. 2000). Taken together, these data suggested that the erasure of methylation at the promoter region and CTCF-binding site upstream of H19 in TGCTs recapitulates that of foetal germ cells or PGCs, in line with a presumably foetal origin of these tumors. Very recently, poor-quality sperm was also shown, by pyrosequencing, to have significantly different methylation profiles at the sixth CTCF binding site (Boissonnas et al. 2010). Similar altered epigenetic patterns are thus found both in poor semen and TGCTs emphasizing the suggested possible common aetiology of testicular dysgenesis (TDS). However, the methylation profile in sperm may be a reflection of de-regulation that occurred around puberty, in addition to that persisting from the foetal life. As to the aetiology, very little is known, but recently, oestrogen responsive elements have been found in the IGF2/H19 locus (Pathak et al. 2010)

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leaving a possible connection to endocrine disruption and altered methylation pattern at the IGF2/H19 locus.

2.5.5

Possible Involvement of miRNAs in TGCTs and Neoplastic Transformation

A range of studies has identified miRNAs to be abnormally expressed in TGCTs, indicating a possible mechanism in the pathogenesis of these tumors. One of the first indications of miRNAs crucial role for proper testicular development came from a targeted inactivation of protein Dicer1, which is responsible for digestion of double stranded pre-miRNAs to yield single stranded mature miRNAs. Dicer1 knockout mice showed among other things infertility due to lack of PGCs (Bernstein et al. 2003). More specific clues came later from a functional genetic screen, which identified miR-372 and miR-373 as miRNAs implicated in TGCT oncogenesis (Voorhoeve et al. 2006). In response to mitogenic signals from oncogenes, cells undergo a growth arrest. However, cells that lack functional p53 efficiently overcome this arrest, which is a prerequisite for full transformation into tumor cells. It was shown that expression miR-372 & 3 gave human primary cells the ability to circumvent the need to mutate p53 in order to drive oncogenic cell divisions. In addition, miR-372 & 3 were found highly expressed in TGCTs and in TGCT cell lines together with a normal non-mutated p53 (Voorhoeve et al. 2006) suggesting that these miRNAs might be more instrumental in the pathogenesis of TGCTs than the p53. P53 was initially suspected to play a role because of its high accumulation in a subset of CIS cells and overt tumors (Bartkova et al. 1991; Lutzker and Levine 1996), but mutations have never been convincingly demonstrated. Expression of the miR-371-373 cluster was later confirmed by expression profiling of 156 miRNAs in 69 TGCT samples (Gillis et al. 2007). This study showed, moreover, that samples clustered together depending on their maturation status, indicating involvement of altered miRNA expression in tumorigenesis (Gillis et al. 2007). One miRNA identified by Gillis et al. to be highly expressed in seminomas and EC was miR-302. Interestingly, miR-302 was recently identified as a potent suppressor of the tumor suppressor homologue p63 (Scheel et al. 2009). So far the only results concerning miRNA expression in CIS cells come from a study by Novotny et al. (2007). It was shown that expression of the transcription factor E2F1 depends on the expression levels of pri-mir-17-5p in germ cells. A similar regulation was observed in CIS cells, which express the E2F1 mRNA, but not E2F1 protein, concomitant with strong expression of the pri-miR-17-5p transcript (Fig. 2.4). As E2F1 is a key regulator of apoptosis and cell cycle progression, expression of pri-miR-17-5p could thus reduce apoptosis of CIS cells leading to sustained proliferation and ultimately neoplastic transformation. However, miR-17-5p might also be involved directly in the pathogenesis of testicular cancer as the miR-17 family recently was shown to be implicated in

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Fig. 2.4 Expression of E2F1 mRNA and protein in germ cells and CIS together with expression of the pri miR 17 5p. Adapted from Novotny et al. (2007). Serial sections of normal human testis with in situ hybridization of E2F1 mRNA (l), immunohistochemical detection of E2F1 protein (m) and in situ detection of pri miR 17 5p (n). Corner marks indicate areas magnified underneath (o, p and q). Red arrows indicate cells with expression of E2F1 protein, whereas the same cells have no or low expression of pri miR 17 5p. Blue arrows indicate examples of cells with no expression of E2F1 protein and high expression of miR 17 5p. (r, s and t) show serial sections of a tubule containing CIS cells. Bars correspond to 20 mm

regulating stem cell differentiation in mice, probably by regulating STAT3 expression (Foshay and Gallicano 2009). Still, very little is known about the complete regulatory network of miRNAs in testicular cancer and identification of many more miRNAs implicated in testicular cancer will probably be revealed in the near future. Yet questions of whether developmental mis-regulation of these miRNAs exist will also need to be elucidated. Especially remnants of embryonic miRNAs in CIS cells could be implicated in creating the neoplastic phenotype of CIS cells.

2.6

Open Questions and Perspectives

Based on the existing evidence although it is so far limited it seems that epigenetic changes are abundant in TGCTs and are likely to be involved in the pathogenesis of other symptoms of TDS. Much remains, however, unknown about

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the epigenetic status of CIS, TGCTs and TDS, if we consider the huge number of possible epigenetic modifications. In addition, the consequences of changes of epigenetic status remain to be elucidated and compared to other features of TGCTs, i.e., genomic aberrations, and phenotypic features, such as pluripotency and invasiveness. The phenotypic features are of importance for possible clinical applications of this knowledge: can the tumor type, ability to metastasise, and the response to treatment be predicted from the epigenetic profile of the precursor CIS cell? A key question concerning the aetiology of TDS and TGCT is whether endocrine disrupters indeed are responsible for aberrant epigenetic patterns, as they seem to be in some experimental animals. Here a very important avenue of research is to identify the molecular targets for each chemical, both acting alone and in mixture, and to establish the most vulnerable cell types. This research may in the future lead to the more precise preventive measures.

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Chapter 3

Medical Implications of Sperm Nuclear Quality Rafael Oliva and Sara de Mateo

Abstract The sperm nucleus is essential in the appropriate transmission of the paternal genome. Therefore, it has long been recognized that one of the key components of the sperm nuclear quality is the quality of the genomic DNA delivered by the sperm cell. In addition, it has recently been uncovered that other important components of the quality of the sperm cell nucleus are represented by its proteome and its epigenome. Sperm genome quality implies a correct number of chromosomes and the integrity of the DNA sequence. Sperm nuclear proteome quality means an appropriate composition in nuclear proteins that organize and condense the male genome, including protamines and other chromatin-associated proteins. The specific organization of the male genome, together with appropriate DNA methylation, and other components of the epigenome such as modified histones and RNA, carried by the sperm cell, constitutes the sperm epigenome. This chapter reviews the current state of the art of the normal genomic, proteomic, and epigenetic sperm cell constitution and the proven and potential medical implications of its anomalies. Keywords Chromatin DNA damage Epigenetic Genome Imprinting Nuclei Protamine Proteome Spermatozoa

R. Oliva (*) and S. de Mateo Human Genetics Research Group, IDIBAPS, University of Barcelona, Casanova 143, 08036 Barcelona, Spain Biochemistry and Molecular Genetics Service, Hospital Clı´nic i Provincial, Villarroel 170, 08036 Barcelona, Spain e mail: [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 3, # Springer Verlag Berlin Heidelberg 2011

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3.1

3.1.1

R. Oliva and S. de Mateo

Fundamental Knowledge on Sperm Genome and Proteome/Epigenome Constitution Changes in Sperm Nuclear Composition and Structure During Spermatogenesis

Spermatozoa are produced during spermatogenesis, a differentiation process involving extremely marked genetic, cellular, functional, and chromatin changes (Fig. 3.1, Baccetti and Afzelius 1976; Mezquita 1985; Poccia 1986; Oliva and Dixon 1991; Kierszenbaum and Tres 2004; Kimmins and Sassone-Corsi 2005; Rousseaux et al. 2005; Seli and Sakkas 2005; Baker et al. 2005; Oliva 2006; Shaman et al. 2007; Balhorn 2007). Spermatogonia replicate and differentiate into primary spermatocytes, which undergo genetic recombination and meiotic divisions, leading to haploid round spermatids. Round spermatids then undergo a differentiation process, called spermiogenesis, where marked cellular, epigenetic, and chromatin remodeling events take place. The nucleosomes are disassembled, and the removed histones are replaced by transition protein first and finally by the highly positively charged protamines that form a tight toroidal complex with the DNA (Figs. 3.1 and 3.2, Oliva and Dixon 1991; Oliva 2006; Balhorn 2007). In humans and in other species, it has been determined that approximately 5 15% of the DNA remains associated with histones. Finally, the spermatozoon undergoes a maturation process during its transit through the epididymis, where its chromatin is further compacted via the formation of disulfide bonds between protamines and the formation of zinc bridges, and where it acquires different specific membrane and cellular functionalities (Oliva and Dixon 1991; Baker et al. 2005; Balhorn 2007; Oliva et al. 2009; Bjorndahl and Kvist 2010). The mature sperm cell nucleus has the essential function of delivering the haploid paternal genome to the oocyte. In addition, the sperm genome is packaged with protamines, histones, and other proteins, which constitute the sperm nuclear proteome (Fig. 3.2). The association of the male genome with sperm-specific nuclear proteins and protein modifications, as well as its specific DNA methylation pattern, constitutes elements of the sperm cell epigenome (Fig. 3.2). Once in the oocyte, the male pronucleus undergoes another extremely marked chromatin remodeling process during which the nucleoprotamine structure is disassembled and a new nucleosomal and chromatin structure is assembled (Fig. 3.1). The sperm cell nuclear quality can be assessed at the genome, proteome, and epigenome levels. The most well-known and studied aspect of sperm nuclear quality is the sperm DNA integrity. More recently, the proteome and epigenome components have also been considered as important elements for fertilization and subsequent embryo development.

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Medical Implications of Sperm Nuclear Quality

Spermatogonia

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DNA replication Transcription Repair

Nucleohistone Ac

Ac

Primary Spermatocyte Secondary Spermatocyte Round Spermatid (haploid)

Incorporation of histone variants and proteins and other epigenetic modifications

Transcription Repair

Ac

Transcription Repair

Ac

Ac

Ac

Ac

Histone hyperacetylation (Ac) during spermiogenesis, incorporation of histone variants and proteins and other epigenetic modifications

Ac Ac Ac Ac Ac Ac Ac Ac Ac Ac Ac Remodelling factors

Transition proteins

Ac Ac

Ac

Binding of protamines

Histone variants

Nucleoprotamine

Epigenetic marks

Maturation (epididymis) Capacitation

Male Pronucleus

Ac

DNA in a nucleosome structure 20 to 50 Kb chromatin loops attached to nuclear matrix or chromosome scaffold

Genetic recombination Transcription Repair

Elongating Spermatid Spermatozoa

Ac

Ac

Ac

Fertilization ~15% of sperm DNA associated to histones

~85% of sperm DNA in compact nucleoprotamine toroidal structures (~50 kb of DNA each)

Ac

Sperm centriole

Remodelling factors

Ac Ac

Ac

Ac

Ac Ac

Transcription Repair DNA replication

Ac

Ac Ac

Ac

Ac

Ac

Ac

Ac

Epigenetic marks Ac Ac

Ac

Ac

Fig. 3.1 Major chromatin transitions during spermiogenesis and fertilization. The chromatin structure present in spermatogonia, spermatocytes, and round spermatids (top left) is similar to that present in all somatic cells, with the DNA being organized in nucleosomes. During spermatogen esis, histones are hyperacetylated and experience other modifications, histone variants are incorporated, nucleosomes are disassembled, and transition proteins bind the DNA. At the final stage of spermiogenesis, the transition proteins are removed and protamines progressively bind the DNA. During the sperm maturation in the epididymis, the formation of disulfide bonds in protamines further stabilizes the nucleoprotamine complex. The mature sperm nucleus consists of 85 95% of the DNA, organized into highly compact toroidal nucleoprotamine structures, and about 5 15% of the DNA organized with histones. The distribution of genes in the histone and protamine regions is not random. This organization, together with appropriate DNA methylation,

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R. Oliva and S. de Mateo

Sperm Cell Nuclear Proteome

The sperm protamines, histones, and other proteins associated with the genome constitute the sperm nuclear proteome (Fig. 3.2). Protamines are the most abundant sperm nuclear proteins organizing 85 95% of the male genome in many vertebrates and therefore represent an important component of the sperm nuclear quality (Figs. 3.2 and 3.3a, see Chap. 9 by Ausio´ et al. in the present book for a complete review of the genome organization and evolution of vertebrate sperm nuclear basic proteins). Different hypotheses are available on the function of the protamines: (1) Packaging and streamlining the sperm nucleus so that the sperm cell can swim faster, (2) Protecting the genetic message, and (3) participating in the setting up of an appropriate organization or “bar coding” of the sperm chromatin, with a potential epigenetic function (Oliva and Dixon 1991; Balhorn 2007). Protamines can be directly measured after reduction of the disulphide bonds, lysis with guanidine thiocyanate, extraction with HCl, purification and separation using acid gel electrophoresis, and visualization using Coomassie Blue staining (Fig. 3.3a, de Yebra and Oliva 1993). In addition to the relative quantities and distribution of protamines within the sperm nucleus, their crosslinking status through disulfide bond formation strongly stabilizing the nucleoprotamine complex, and also the formation of zinc bridges with protamine thiols of cysteines, should also be considered as important components of the sperm nuclear quality (Saowaros and Panyim 1979; Balhorn et al. 1992; Lewis et al. 2003; Vilfan et al. 2004; Bjorndahl and Kvist 2010). However, the recent analysis of the proteome content of the sperm cell using sensitive protein detection approaches and mass spectrometry has indicated that, in addition to the male genome complexed with protamines, the sperm cell also delivers to the oocyte an important protein “cargo” (Figs. 3.2 and 3.3b, Oliva et al. 2008, 2009). The analysis of the type of proteins identified in the different mature sperm proteomic projects has also provided some unexpected results. For example, many transcription factors, DNA binding proteins, and proteins involved in chromatin metabolism have been identified (Martı´nez-Heredia et al. 2006; Lefie`vre et al. 2007; de Mateo et al. 2007; Codrington et al. 2007; Baker et al. 2007, 2008a, b; Peddinti et al. 2008). The presence of proteins such as histone acetyltransferase and deacetylase, histone methyltransferase, DNA methyltransferase, topoisomerase, helicase, transcription factors, zinc fingers, leucine zippers, homeobox proteins, chromodomain proteins, centrosomal proteins, and telomerase in cells that are supposed to be transcriptionally inert and that have 85 95% of their DNA tightly packaged with protamine is, indeed, remarkable(Fig. 3.2 and Table 3.1, Mezquita 1985; Poccia 1986; Oliva and Dixon 1991; Rousseaux et al. 2005;

Fig. 3.1 (continued) epigenetic modification of histones and other chromatin associated proteins, appropriate chromosome positioning, and the RNA carried by the sperm cell, constitutes the sperm epigenome, which is transferred to the oocyte at fertilization (reproduced from Oliva et al. 2009 with permission from the publisher)

3 Medical Implications of Sperm Nuclear Quality

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Oliva 2006; Balhorn 2007; Oliva et al. 2008, 2009). The proportion of the nuclear proteins identified most likely represent an underestimation since they have been identified in whole sperm proteomic analysis (Martı´nez-Heredia et al. 2006; de Mateo et al. 2007; Baker et al. 2007, 2008a, b; Peddinti et al. 2008). Thus, it could be expected that an approach focusing on the mature sperm nuclei should identify

Sperm nuclei Proteome Genome RNA •DNA sequence Protamines Histones / other 5 10 3 7 MW •DNA integrity pI

• • • • • •

Repair Protection TopoII Apoptosis Necrosis ROS

100 75 50 37 25 20

•Chromosome number •DNA methylation

P1 P2

15 10

Epigenome ~5-15% of sperm DNA

~85-95% of sperm DNA » 50 Kb

Sperm histone variants and modified histones (acetylation, methylation, phosphorylation, …) DNA methylation

Nuclear matrix attachment sites

Highly packaged nucleoprotamine toroidal structures contain each about 50 kb of DNA

Distribution of genes in the protamine vs histone-associated regions. Gene / promoter –specific nucleoprotein structure

Sperm RNA

Fig. 3.2 Elements of the sperm nuclear quality. One of the key components of the sperm genome quality is the genome represented by its sequence fidelity, good DNA integrity, and correct chromosome number (top left). The sperm nuclear proteome quality also includes an appropriate protein composition (top right) and is intimately related to the condensation and organization of the male genome by protamines and the associated proteins. This organization, together with appropriate DNA methylation, epigenetic modification of histones and other chromatin associated proteins, appropriate chromosome positioning, and the RNA carried by the sperm cell, constitutes the sperm epigenome (bottom)

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additional chromatin-associated proteins. An important issue that will be important to determine in the future is whether these newly identified transcription factors and nuclear proteins are marking some regions of the male genome and have an

Fig. 3.3 Proteomic identification of sperm proteins. (a) The protamines can be analyzed after DTT reduction of the disulfide bonds, protein extraction, and polyacrylamide gel electrophoresis (left). (b) The global sperm proteome can be analyzed by two dimensional gel electrophoresis and the proteins identified by MALDI TOF (right). Alternatively, the sperm proteome can be inves tigated after tripsin digestion of the sperm proteome followed by peptide separation using liquid chromatography and peptide identification by tandem mass spectrometry (not shown). Proteomic methods are sensitive enough to allow protein identification and quantification from single sperm samples for which DNA integrity and other conventional semen and sperm parameters have also been determined. (c) The proteomic information can be correlated with other sources of information available for each sperm sample or with clinical information. For example, one of the proteins (prohibitin) identified in the proteome maps is correlated in independent infertile patients with the protamine P1/P2 ratio (left) or with the TUNEL results (right). [Modified from de Mateo et al. (2007), Oliva et al. (2008)]

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Table 3.1 Chromatin associated proteins identified in the human, bull, mouse, and rat sperm cell proteome projects (Oliva et al. 2009) Protein category Proteins detected using mass spectrometry in mature mammalian spermatozoa Abundant sperm nuclear proteins Protamine Histone H1 H2A H2B H3 H4 Histone modifying proteins Histone acetyltransferase Histone deacetylase Histone methyltransferase ADP ribosyl DNA modifying proteins DNA methyltransferase DNA helicase Topoisomerase Telomerase DNA binding proteins Chromodomain/chromobox proteins Different transcription factors Zinc finger/ZNF/ZF proteins Leucine zipper containing proteins Homeobox proteins Homeodomain proteins Oncogenes Telomere/telomeric proteins Centromere/centrosomal proteins RNA binding proteins Different RNA binding proteins

important epigenetic function or a role upon fertilization (Oliva and Dixon 1991; Oliva et al. 2008, 2009; Wu and Chu 2008). One example of sperm proteins crucial for embryo development are the proteins of the centrosome because, in humans and most mammals (with the exception of mouse), they are paternally inherited (Chatzimeletiou et al. 2008). Another example is the proteasomal proteins present in sperm (Martı´nez-Heredia et al. 2006; de Mateo et al. 2007). A role for sperm proteasomes in acrosome exocitosis, zona pellucida penetration, sperm mitochondrial sheath degradation, as well as chromatin remodeling after penetration of the spermatozoa has been described (Chakravarty et al. 2008; Rawe et al. 2008). So it is plausible that some of the newly identified sperm proteins may also have a role in the zygote. An alternative explanation for the presence of some of these proteins is that they could represent leftovers from spermiogenesis. In this case, these proteins could represent a “window” to the later stages of spermatogenesis, with potential clinical implications. Finally, some of these proteins could be involved in the mitochondrial gene expression and/or protein synthesis with mitochondrial ribosomes from nuclear-encoded mRNAs present in mature sperm (Gur and Breitbart 2008). Thus, it will be interesting to determine the role or meaning of the different sperm chromatin-related proteins detected in the different proteomic studies (Table 3.1).

Table 3.2 Sperm DNA integrity as measured by TUNEL or COMET and assisted reproduction results Method Sample size Threshold Fertilization rate Embryo quality Pregnancy according to ART proposed affected? affected? affected? TUNEL 20 ICSI 20% NS NA NA 82 IVF/50 ICSI – NA NA Yes (ICSI) 117 IVF 35% NS NS Yes 45 IVF/68 ICSI 35–55% Yes (IVF) NS Yes (ICSI) 88 IVF/234 ICSI 15% Yes (ICSI) Yes (ICSI) NS 42 ICSI 4–10% NS NS Yes 82 IVF/50 ICSI 10% Yes (IVF) Yes (ICSI) Yes (ICSI) 26 IVF/22 ICSI – NA NA NS 72 ICSI 30–40% NA NA Yes 217 IVF/86 ICSI 10% Yes NS NS 35 IVF/14 ICSI 20% NA Yes (IVFþICSI) NS 167 IVF 36.5% NS NA Yes 18 ICSI – NA NS Yes 50 IVF/54 ICSI 10%/20% Yes (ICSIþFIV) NS Yes (ICSI) 208 IVF/54 ICSI 36.5%IVF/ NS NS Yes (IVF) 24.3% ICSI 154 IUI – NA NA Yes 140 IVF – NS NS Yes 100 IVF 4% Yes NS NS 50 IVF/61 ICSI 4% Yes (IVF) NS NS 131 ICSI – Yes NS NA 143 IVF 4% Yes NS NA 30 IVF/36 ICSI 10% NS NA NA COMET 55 ICSI – NS NS NA 28 ICSI – NS NS NA 88 ICSI (TESE) – NS? NS Yes 20 IVF/40 ICSI – NS NS NA 40 IVF – NS Yes Yes NS not significant, NA not applicable or not described Daris et al. (2010) Tarozzi et al. (2009) Frydman et al. (2008) Bakos et al. (2008) Benchaib et al. (2007) Ozmen et al. (2007) Borini et al. (2006) Hammadeh et al. (2006) Hazout et al. (2006) Huang et al. (2005) Seli et al. (2004) Henkel et al. (2004) Tesarik et al. (2004) Benchaib et al. (2003) Henkel et al. (2003)

Duran et al. (2002) Tomlinson et al. (2001) Høst et al. (2000a) Høst et al. (2000b) Lopes et al. (1998) Sun et al. (1997) Sakkas et al. (1996) Abu-Hassan et al. (2006) Nasr-Esfahani et al. (2005) Thompson-Cree et al. (2003) Morris et al. (2002) Tomsu et al. (2002)

NA NA NA NA NA NA NA NA NA NA NS NA

Authors

Elevated miscarriages? NA NA Yes Yes Yes NA Yes (ICSI) NA NA NA NA NA NA NA NA

52 R. Oliva and S. de Mateo

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The determination of the sperm cell proteome also opened up the possibility to investigate the potential presence of abnormal proteins in infertile patients (de Mateo et al. 2007; Martı´nez-Heredia et al. 2008, de Aitken and Baker 2008). Using this approach, different altered proteins have already been reported, associated with different phenotypes in infertile patients (Zhao et al. 2007; de Mateo et al. 2007; Martı´nez-Heredia et al. 2008; Kriegel et al. 2009; Liao et al. 2009). Furthermore, the proteomic approach also allows to search for correlations among abundance of proteins and other aspects of the sperm nuclear quality such as the protamine content or the DNA integrity of the sperm cell (de Mateo et al. 2007). As an example, one of these proteins detected in proteomic analysis, and correlated with DNA damage and with protamine content is prohibitin (Fig. 3.3c). Two locations and functions have been described for prohibitin in somatic cells: a location in the nucleus with the function of modulation of transcriptional activity by interacting with various transcription factors, and a location in the mitochondria where it could act as a chaperone (Kurtev et al. 2004; Tatsuta et al. 2005; Gamble et al. 2007). More limited data is available on the function of prohibitin in the transcriptionally inactive mature sperm cells, although a role in the ubiquitination of prohibitin in marking of defective sperm has been proposed (Thompson et al. 2003). This example illustrates the potential of the present proteomic approaches to identifying proteins potentially involved in the pathogenic mechanisms that lead to male infertility. The detection of proteome anomalies and correlations is just now at its beginnings. Possibly, proteomics will allow the discovery of pathways of altered proteins that point towards many of the ethiologies involved in male infertility (de Mateo et al. 2007).

3.1.3

Sperm Cell Epigenome

In addition to the genome and the proteome, an important component of the sperm cell, which has been recognized more recently, is the epigenome (Fig. 3.2). A few decades ago, it was thought that the only function of the spermatozoon was to deliver a haploid complement of the paternal genetic information, encoded in the DNA sequence, into the oocyte. This idea started to change with the discovery that the male and female pronuclei were not equivalent as derived from pronuclear transfer experiments in which embryos with either two male or two female pronuclei could not develop (McGrath and Solter 1984). It was later discovered that this was due to the presence of sex-specific imprinting of genes mediated through DNA methylation differences set during gametogenesis (Reik et al. 2001). DNA methylation is one of the most well-studied aspects of the sperm epigenome and involves the enzymatic modification of cytosines from some of the CpG dinucleotides to form 5-methyl cytosine, and it is also one of the most well-studied aspects of the cellular epigenome (Klose and Bird 2006). DNA methylation plays a role in regulating genes during development and is also involved in genomic imprinting and in X-chromosome inactivation. Other proposed roles for DNA

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methylation include genome defence and blocking the expression of retrotransposons repetitive sequences in the genome (Yoder et al. 1997; Verona et al. 2003). During spermatogenesis, the genomic methylation patterns are erased and reacquired in the developing male and female gametes (Trasler 2009). The male and female germ cells experience a different reprogramming of the DNA methylation status during gametogenesis, and its complementarity in the zygote is essential for proper embryonic development (see Chaps. 1 and 5 in the present book). Because of the DNA methylation reprogramming during gametogenesis and early embryogenesis, the use of ARTs with in vitro manipulation of gametes or early human embryos has received much attention. A small but significantly increased frequency of offspring with imprinting genetic conditions has been reported as a result of assisted reproduction outcome (Cox et al. 2002; Moll et al. 2003; DeBaun et al. 2003; Manipalviratn et al. 2009; Butler 2009). A few studies have examined DNA methylation in human sperm and reported DNA methylation defects at imprinted loci in oligospermic patients (Marques et al. 2004, 2008; Kobayashi et al. 2007). Oxidative DNA damage itself has been shown to impair global sperm DNA methylation in infertile men (Tunc and Tremellen 2009). A more recent study has identified that infertile patients had significant altered methylation at six of seven imprinted loci tested, with differences observed between oligozoospermic and abnormal protamine patients (Hammoud et al. 2009b). The underlying mechanisms and significance of these DNA methylation alterations in infertile patients are not known at present, nor is it known whether they could be related to the described infants born with imprinting defects after ART, but they definitely deserve further investigation. From a more global and public health perspective, the potential detrimental effects of endocrine disruptors and other compounds potentially affecting DNA methylation or other types of imprinting deserve serious consideration. A substantial increase in male infertility and a reduction in sperm counts coupled to an increased incidence of testicular cancer and andrological disorders have been observed over the last 50 years (See Chap. 2 in the present book for a complete review; Carlsen et al. 1992; Skakkebaek et al. 2006; Sonne et al. 2008). While it is difficult to establish a causal link between contaminants and these trends, transgenerational effects could be involved (Waterland and Jirtle 2003; Anway and Skinner 2008). Altogether, the above data indicates that the DNA methylation status of the sperm cell as an important component of the sperm nuclear quality cannot be ignored, and more research is needed. Present exome genome sequencing approaches coupled to bisulphite sequencing to detect DNA methylation should offer an unprecedented tool to further clarify the role of the DNA methylation normalcy of the sperm cell. In addition to the DNA methylation, additional epigenetic components of the sperm are constituted by the sperm chromatin and gene organization, the sperm RNA, and the protein modifications. The potential importance of the sperm chromatin complement (nuclear proteome + genome) delivered by the spermatozoa was largely neglected until recently. This notion was partly due to the fact that the spermatozoon was supposed to be a transcriptionally inert cell with the DNA tightly packaged with protamines (Mezquita 1985; Oliva and Dixon 1991; Kierszenbaum

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and Tres 2004; Kimmins and Sassone-Corsi 2005; Rousseaux et al. 2005; Seli and Sakkas 2005; Oliva 2006; Shaman et al. 2007; Balhorn 2007). However, different lines of evidence indicate that, in addition to the DNA that contains the genetic information and the methylation sex specific imprints, the sperm cell also delivers to the oocyte a wealth of epigenetic information, which may be crucial for embryonic development. As indicated at the beginning of the present chapter, the sperm nuclei of many species retains about 5 15% of the sperm DNA organized by histones, many of which are sperm-specific variants (Fig. 3.1, Gatewood et al. 1987; Zalensky et al. 2002; Singleton et al. 2007). The rest of the genome (about 85 95%) is tightly packaged by protamines into toroidal structures (Fig. 3.1, Oliva and Dixon 1991; Balhorn 2007). Each of these toroidal structures contains about 50 kb of DNA, and they are thought to be attached through their linker region DNA to the sperm nuclear matrix (Shaman et al. 2007). Of potential epigenetic relevance, the distribution of genes in the genomic regions organized by protamine and in the genomic regions organized by histones is not random (Gatewood et al. 1987; Oliva and Dixon 1991; Gardiner-Garden et al. 1998; Wykes and Krawetz 2003; Li et al. 2008; Arpanahi et al. 2009; Hammoud et al. 2009a). This organization of the sperm genes into the nucleoprotamine and nucleohistone compartments has been further demonstrated recently by two independent groups, with the application of microarrays and deep genome sequencing technologies, respectively (Arpanahi et al. 2009; Hammoud et al. 2009a). In the first report, the authors used two different strategies to fractionate the human sperm and mouse chromatins into the histone and protamine regions (Arpanahi et al. 2009). One of these strategies was based on the differential extraction of the histones using 0.65 M NaCl and subsequent digestion of the “free DNA” with a combination of BamHI and EcoRI to liberate the histoneassociated genomic domains, following previously described procedures (GardinerGarden et al. 1998; Wykes and Krawetz 2003). The other strategy was based on the differential digestion of the nucleosome-associated regions of the sperm nucleus using micrococcal nuclease, also following previously described methods (Zalenskaya et al. 2000). With the different chromatin fractions, the authors then used human and mouse whole chromosome microarray CGH to determine the differential distribution of genes. The basic conclusions of this work were that regions of increased endonuclease sensitivity are closely associated with gene regulatory regions and that a similar differential packaging was observed in both mouse and man, implying the existence of epigenetic marks distinguishing gene regulatory regions in male germ with a potential role for subsequent embryonic development (Arpanahi et al. 2009). In the second study (Hammoud et al. 2009a), the authors also used the differential digestion of the nucleosome-associated regions of the sperm nucleus, using micrococcal nuclease (Zalenskaya et al. 2000). The fractionated chromatin was then analyzed by deep genome sequencing, using the Illumina GAII sequencing. In this study, DNA methylation and the differential distribution of sequences was also investigated using microarray hybridizations. The basic conclusions of this work were that retained sperm nucleosome-associated regions are significantly enriched

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at loci of developmental importance, including imprinted genes, microRNAs, HOX genes, and the promoters of developmental transcription and signaling factors (Hammoud et al. 2009a). In addition, they demonstrated that histone modifications (H3K4me2, H3K27me3) localize to particular developmental loci, and that developmental promoters are generally DNA hypomethylated in sperm, but acquire methylation during differentiation. Altogether, the results were interpreted as being indicative of epigenetic marking in sperm being extensive and correlated with developmental regulators (Hammoud et al. 2009a). In addition to the specific organization of the genes in the histones versus the protamine organized regions, it has been shown that chromosomes tend to occupy discrete regions within the sperm nucleus with telomeres located in the periphery (see Chap. 11 by Mudrak et al. in the present book; Haaf and Ward 1995; Zalensky et al. 1995, 2002; Hazzouri et al. 2000; Zalenskaya and Zalensky 2002; Mudrak et al. 2005). Furthermore, it has been shown that pericentric heterochromatin regions remain associated with specific nucleosomes and nucleosome-like structures in condensing spermatids (Rousseaux et al. 2008). In fact, it has already been demonstrated that sperm-derived histone variants contribute to zygotic chromatin in humans (van der Heijden et al. 2008; see also Chap. 8 by de Boer et al. in this book on histone variants during gametogenesis and early development). Thus, epigenetic processes implemented during spermatogenesis distinguish the paternal pronucleus in the embryo (Biermann and Steger 2007; Wu and Chu 2008). There is also some evidence that alterations in some of the proteins present in the spermatozoa may be related to subsequent embryo development. This evidence has come from the observation that the abnormal protamine complement present in some infertile patients correlates with reduced assisted reproduction outcomes (Oliva 2006; Aoki et al. 2006; Carrell et al. 2008; Carrell 2008; de Mateo et al. 2009). It has also been suggested that topoisomerase II-mediated breaks in spermatozoa may cause the specific degradation of paternal DNA in fertilized oocytes (Yamauchi et al. 2007). A final aspect of the sperm epigenome is the presence of sperm RNA. The identification of sperm RNAs and the demonstration of their transfer to the ova have provided evidence for a potential role of the sperm RNA in the oocyte (Ostermeier et al. 2004). This issue is covered in detail by Miller et al. (Chap. 15 of the present book).

3.2

3.2.1

Approaches/Tests for Exploring Sperm Nuclear Quality in Human Infertility Sperm DNA Quality and Integrity

It has long been recognized that the key role of the spermatozoa is to transmit the male genome to the offspring. Assessment of the sperm genome quality can include the evaluation of the DNA sequence, chromosome number, and sperm DNA

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integrity. Of these components, the DNA integrity is the most well-studied aspect. However, altered DNA sequence or abnormal chromosome number are also involved in male infertility and offspring disease. Genetic mutation not only allows species evolution but is also at the origin of hereditary diseases (Darwin 1859; Nei 1969; McKusick 2007). The involvement of the paternal cell line in the de novo appearance of several genetic diseases is now well-known (K€uhnert and Nieschlag 2004). Advanced age in the father has been associated with an increased frequency of diseases such as achondroplasia in the offspring (Rousseau et al. 1994; Wyrobek et al. 2006) and Apert syndrome (Yoon et al. 2009) among others. One of the proposed mechanisms for this increase of mutation rates with age is the large number of replication cycles, which spermatogonia undergo throughout life, although other mechanisms have also been proposed (Choi et al. 2008). Factors causing mutations include endogenous stochastic mutations as a consequence of replication errors and oxidative stress and exogenous factors and mutagens (Rattray and Strathern 2003). The DNA repair activity in the germ line is another important factor determining the integrity of the male genome (See Chap. 7 by Schoenmakers and Baarends in the present book; Barnes and Lindahl 2004). Predisposing factors to sperm DNA mutations may include inappropriate protection of the genetic message by protamines (Oliva 2006; Leduc et al. 2008, see also below). In addition, it is important to remember that an appropriate haploid chromosome complement in the spermatozoon is also essential. The existence of sperm cell aneuploidy rates in infertile patients and its consequences has been extensively studied (See Chap. 6 by Barthe`s et al. in the present book; Bosch et al. 2003; Sarrate et al. 2005; Martin 2005; Arnedo et al. 2006; Anton et al. 2007; Farfalli et al. 2007; Martin 2008; Rubio et al. 2009). DNA damage can be of various forms, such as DNA strand breaks, abasic sites, base modifications, such as the formation of 8-hidroxiguanine, or covalent bond formation with other DNA strands or proteins (Aitken et al. 2009). Many of these types of DNA damage also increase the chance of DNA fragmentation. DNA fragmentation can affect either one strand or both strands. DNA fragmentation affecting only one strand has the potential to be repaired during spermatogenesis or by the oocyte upon subsequent fertilization, using the other strand as a template. On the other hand, double-strand DNA fragmentation is conceptually more disastrous because of its potential in originating severe truncation of the genetic message and loss of chromosome fragments, thus resulting in miscarriage or severe unbalanced chromosomal abnormalities in the offspring. Several tests have been developed to measure DNA damage in the sperm cell. The presence of DNA strand breaks can be detected by terminal deoxynucleotidyl transferase-mediated 20 -deoxyuridine 50 -triphosphate (dUTP)-nick end-labeling (TUNEL) assay that measures the presence of free 30 hydroxyl at the breakage site (Gorczyca et al. 1993). This method can detect single- and double-strand DNA breaks (Zini et al. 2001a; Domı´nguez-Fandos et al. 2007). Assessment of DNA integrity with the TUNEL assay is undertaken either counting under a fluorescent microscope or using flow cytometry (Domı´nguez-Fandos et al. 2007; Zini and Sigman 2009). The measurements obtained either from the fluorescence

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microscopy or from flow cytometry are expressed as percentages of labeled sperm cells. Numerous studies have detected increased TUNEL scores in the sperm cells of infertile (Domı´nguez-Fandos et al. 2007). In addition, TUNEL results have been studied in relation to the assisted reproduction results (Table 3.2). Another test to measure the presence of single- or double-strand breaks in the sperm cell is the COMET assay. The basis of the COMET assay or single cell gel electrophoresis is to expose decondensed sperm cells to an electrophoresis on an agarose gel. Each spermatozoon is analyzed, and if the sperm cell has damaged DNA, the negative DNA fragments will migrate to the positive pole (anode), creating a shape similar to that of a comet (Singh et al. 1989; Aravindan et al. 1997). This assay can detect both double- or single-strand DNA breaks. COMET detects double-strand breaks under neutral conditions and double- and single-strand breaks under alkaline conditions. The DNA is stained and measured under the microscope or using a digital image processing and analysis software (Sigman et al. 2009). The parameters that can be obtained from the software are COMET head parameters and COMET tail parameters (Tomsu et al. 2002), but the most used parameter to determine the degree of DNA damage is the DNA migration with the “% tail DNA”, that is, the average percentage of DNA staining outside the area of the sperm nucleus in the electrophoresis and the tail moment (Schmid et al. 2007). COMET has been shown to correlate with other tests and semen parameters and evidenced a decrease of DNA fragmentation in infertile patients (Lewis and Agbaje 2008). COMET results have also been studied in relation to the assisted reproduction results (Table 3.2). In addition to the TUNEL and COMET assays, there are a number of tests that measure the susceptibility of damaged sperm chromatin to decondensation. One of these tests is the sperm chromatin structure assay (SCSA), which uses acridine orange (AO) staining to detect and differentiate between double- and single-strand DNA. The AO dye fluoresces green when bound to double-strand DNA and red when bound to single-strand DNA (Royere et al. 1988; Larson et al. 2000). The SCSA test assesses the sperm DNA susceptibility to denaturation in situ under acid or heat conditions (Evenson et al. 1980). This characteristic is proportional to the amount of DNA strand breaks (Bakos et al. 2008; Sigman et al. 2009). The most important measurements obtained from this test are the DNA fragmentation index (%DFI), measuring levels of DNA strand breaks with the ratio of red/(redþgreen) fluorescence, and the high DNA stainability (HDS) that measures the percentage of immature sperm cells. Each of these indexes has been used to determine the relation of the results on the SCSA test with the seminal parameters and assisted reproduction outcome. Semen parameters are correlated weakly to the SCSA results (Spano et al. 1998; Larson et al. 2000). The SCSA is a reproducible test that strongly correlates with the COMET and TUNEL assays (Aravindan et al. 1997; Zini et al. 2001a). Numerous reports have studied the relation between SCSA results and the assisted reproduction outcomes (Table 3.3). Another test that explores the DNA decondensing potential of intact and damaged chromatin is the sperm chromatin dispersion test (SCD), which is based on the assessment of the DNA dispersion and halo formation upon induced decondensation (Ferna´ndez et al. 2003). Undamaged sperm DNA releases the DNA loops

Table 3.3 Results from sperm chromatin tests measuring DNA integrity (acridine orange staining (AO), SCSA and SCD) and assisted reproduction results Method Sample size according to Threshold proposed Fertilization rate Embryo quality Pregnancy Elevated Authors ART affected? affected? affected? miscarriages? AO 26 IVF/22 ICSI – NS NA NS NA Hammadeh et al. (2006) 234 IVF 12% NS NA Yes NA Henkel et al. (2004) 50 IVFþICSI – Yes (IVF) NA NA NA Katayose et al. (2003) 183 ICSI 56% Yes Yes Yes Yes Virant-Klun et al. (2002) 154 IUI – NA NA NS NA Duran et al. (2002) 101 IVF – NS NA NA NA Nasr-Esfahani et al. (2001) SCSA 60 ICSI 15% Yes NS Yes NA Micin´ski et al. (2009) 137 IVF/86 ICSI DFI 27%/HDS 15% NS NS NS Yes Lin et al. (2008) 220 IVF/93 ICSI/197 IUI – NA NA NS NA Bungum et al. (2008) 388 IVF/223 ICSI/387 IUI DFI 30% NS NS NS NS Bungum et al. (2007) 106 ICSI 30% NA NA NS Yes Check et al. (2005) 46 IVF/54 ICSI DFI 27%/HDS 17% Yes (DFI with NA NS NA Payne et al. (2005) IVFþICSI) 60 ICSI DD 30% NS Yes NS Yes Zini et al. (2005a) 109 IVF/66 ICSI/131 IUI DFI 27%/HDS 10% NA NA Yes Yes Bungum et al. (2004) 249 IVFþICSI DFI 30%/HDS 15% Yes (HDS Yes (DFI Yes (DFI) Yes (DFI) Virro et al. (2004) with IVF) with ICSI) 55 IVF/26 ICSI DFI 27% NS NS Yes NA Larson-Cook et al. (2003) 24 (IVFþICSI) COMPa1 27% NS NS NS NA Larson et al. (2000) SCD 92 IVFþICSI – Yes (ICSI) NS NS NA Tavalaee et al. (2009) 50 ICSI – NS NA NA NA Nasr-Esfahani et al. (2008a) 622 IVFþICSI 18% Yes Yes NS NS Velez de la Calle et al. (2008) 85 IVFþICSI – Yes NS NS NA Muriel et al. (2006a) 100 IUI – NA NA NS NA Muriel et al. (2006b) NS not significant, NA not applicable or not described

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forming the halos. Damaged sperm DNA has less capacity to form halos under decondensing conditions. The halo size of the sperm cells in the SCD test is inversely correlated with the presence of DNA fragmentation (Ferna´ndez et al. 2003; Garrido et al. 2008). It has also been suggested that the SCD test measures susceptibility to form a single-stranded DNA from a native double-stranded DNA since nicked DNA denatures more easily than double-stranded DNA (Zini and Sigman 2009). The degree of fragmentation is determined with microscopy and digital image analysis as the halo size corresponding to the surface of the halo divided by the surface of the whole nucleoid (Ferna´ndez et al. 2003). A strong correlation has also been found between the SCD and SCSA tests since both tests measure susceptibility of sperm DNA to acid denaturation in vitro (Chohan et al. 2006). SCD has also been studied for its correlations with other tests and semen parameters and in relation to the assisted reproduction results in infertile patients (Table 3.3). One of the important factors involved in decreased DNA quality appears to be oxidative stress since abnormal amounts of reactive oxygen species (ROS) are detected in about half of the infertile patients (Tremellen 2008). Therefore, many different strategies have been proposed to determine the presence of free radicals and oxidative stress in the sperm cells (Tremellen 2008). One procedure to assess sperm DNA oxidative damage is the measurement of the oxidized deoxynucleoside (8-OHdG, Fraga et al. 1991; Kao et al. 2008). Chances of natural conception are inversely correlated with sperm 8-OHdG levels (Loft et al. 2003). Another common approach to detecting oxidative stress in sperm is the measurement of the effect of free radicals on membrane lipids, which results in the formation of malondialdehyde (MDA) and is measured using the thiobarbituric acid assay (Li et al. 2004; Aitken et al. 1993). MDA concentration within both seminal plasma and sperm is elevated in infertile patients (Nakamura et al. 2002; Hsieh et al. 2006). One of the effects of oxidative stress on the conventional seminal parameters is a reduction in sperm motility or the presence of asthenozoospermia (Aitken et al. 1995; Kao et al. 2008). The presence of oxidative stress itself may result in an increased susceptibility of the DNA to undergo strand breaks and therefore resulting in DNA damage detectable by other tests, such as TUNEL, COMET, SCSA, or SCD among others.

3.2.2

Measurement of the Protamine Content in Infertile Patients

A typical extraction of human protamines from mature sperm cells and its separation using electrophoresis in an acidic gel and visualization using Coomassie Blue staining is shown in Figs. 3.2 and 3.3a. The most intense protein bands that can be visualized are the protamines (Fig. 3.3a). The two major bands correspond, respectively, to the two types of protamines known to be present in mammals: the P1 protamine and the family of P2 proteins (Fig. 3.3a). The protamine P2 is formed by the P2, P3, and P4 components, and it is only present in some mammalian species including human and mouse (Gusse et al. 1986; McKay et al. 1986;

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Be´laı¨che et al. 1987; Bower et al. 1987; Bellve´ et al. 1988; Balhorn et al. 1988, Balhorn 2007; Oliva and Dixon 1991; Bianchi et al. 1992; Yoshii et al. 2005). The components of the P2 family differ only by the N-terminal extension of 1 4 residues, although the P2 component is the most abundant (Figs. 3.2 and 3.3a, Gusse et al. 1986; McKay et al. 1986; Sautie`re et al. 1988; Martinage et al. 1990; Arkhis et al. 1991; Oliva and Dixon 1991; Bianchi et al. 1992; Alimi et al. 1993; Yoshii et al. 2005). The content of protamine P1 in the human sperm nucleus is similar to the content of protamine P2 (P1/P2 ratio of approximately 1) (Ammer et al. 1986; Balhorn et al. 1988; de Yebra and Oliva 1993; de Yebra et al. 1993, 1998; Bench et al. 1996; Corzett et al. 2002; Aoki and Carrell 2003; Mengual et al. 2003; Aoki et al. 2005a). Therefore, a protamine ratio (P1/P2) around 1 has been taken as evidence for normal protamination. It should be noted, however, that the P1/P2 ratio itself does not provide information on whether there is an overall reduction of the protamine content. Anomalies in the protamine content in infertile patients were already described more than 20 years ago (Silvestroni et al. 1976; Chevaillier et al. 1987; Balhorn et al. 1988; Lescoat et al. 1988). Many subsequent studies have further confirmed the association of abnormal protamine content and abnormal seminal parameters and male infertility (Bach et al. 1990; Blanchard et al. 1990; Belokopytova et al. 1993; de Yebra et al. 1993, 1998; de Yebra and Oliva 1993; Colleu et al. 1996; Bench et al. 1998; Mengual et al. 2003; Aoki et al. 2005a; Torregrosa et al. 2006; Oliva 2006). More recently, the protamine results have also been correlated with the assisted reproduction outcomes (Table 3.4). In addition to the above studies in infertile patients, the expression of protamines has also been determined in response to thermal stress in normal testicles (Love and Kenney 1999; Evenson et al. 2000). Thermal stress in stallion testicle is associated to decreased formation of disulfide bridges in protamines (Love and Kenney 1999). This aspect has also been studied in a patient who just finished an episode of influenza, with the detection of the appearance of protamine P2 precursors and a rise in the ratio of histones to protamines between 33 and 39 days posthyperthermia (Evenson et al. 2000). The expression of the gene corresponding to the protamine P2 also has been found altered concomitant to induced thermal stress in the mouse testicle (Iuchi et al. 2003). It is also interesting to note that variation over time of protein and DNA contents in sperm from an infertile human male possessing protamine defects has been described (Bench et al. 1998). Indirect detection methods to tentatively assess the amount of protamines or measure chromatin structure based on different staining procedures or fluorochromes have also been used. For example, in situ competition between protamine and Chromomycin A3 (CMA3) indicated that CMA3 staining is inversely correlated with the protamination state of spermatozoa (Bizzaro et al. 1998). The CMA3 test has also been correlated with the extent of nicked DNA (Manicardi et al. 1995). In the evaluation of the CMA3 staining, sperm cells with bright yellow stain are CMA3-positive cells and those with dull yellow stain are CMA3-negative (Nasr-Esfahani et al. 2001). Interestingly, CMA3 staining has been shown to be increased in the sperm cells of infertile patients (Lolis et al. 1996; Franken et al.

Table 3.4 Sperm protamine and histone assessment and assisted reproduction results Method Sample size Threshold Fertilization rate Embryo quality according to proposed affected? affected? ART Direct protamine 87 ICSI/26 IVF < 1.1 and >1.5 Yes (IVF) NS measurements 124 IVF/264 < 0.8 and >1.2 Yes (IVF) NS through extraction, ICSI electrophoresis, 71 IVF/73 ICSI < 0.8 and >1.2 Yes (IVFþICSI) NS and detection (P1/P2 ratio) Indirect protamine 82 IVF/50 ICSI 29.25% Yes (IVF) NA determination 92 IVF+ICSI 40% Yes Yes (IVFþICSI) through CMA3 50 ICSI – Yes NA staining 68 ICSI 40% Yes NA 26 IVF/22 ICSI – NA? NA 28 ICSI – Yes NS 56 ICSI 30% Yes NA 45 ICSI 30% Yes NS 55 ICSI 30% Yes NA 101 IVF 30% Yes NA 140 IVF – NS NS 32 IVF/38 ICSI 30% NS NA Indirect histone 130 ICSI 75% NA NA assessment 222 Natural – NA NA through aniline conceptions blue staining 55 ICSI – NS NA 101 IVF 20% Yes NA 78 IVF 20% Yes NA 61 ICSI 29% NS NA 89 IUI – NA NA NA NS NS

NS NA NA NA NA NA NA NA NA NA NA NA NA NA NA NA NA NA NA

Yes Yes NS

Yes (IVF) NS NA NA NS NA NA NA NA NA NS NA Yes NS NA NA Yes NS NS

Razavi et al. (2003) Nasr-Esfahani et al. (2001) Hammadeh et al. (1998) Hammadeh et al. (1996) Milingos et al. (1996)

Tarozzi et al. (2009) Tavalaee et al. (2009) Nasr-Esfahani et al. (2008a) Nasr-Esfahani et al. (2008b) Hammadeh et al. (2006) Nasr-Esfahani et al. (2005) Nasr-Esfahani et al. (2004a) Nasr-Esfahani et al. (2004b) Razavi et al. (2003) Nasr-Esfahani et al. (2001) Tomlinson et al. (2001) Sakkas et al. (1996) Wong et al. (2008) Jedrzejczak et al. (2008)

Aoki et al. (2005a)

de Mateo et al. (2009) Aoki et al. (2006)

Pregnancy Elevated Authors affected? miscarriages?

62 R. Oliva and S. de Mateo

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1999; Razavi et al. 2003; Nasr-Esfahani et al. 2004a, b, 2005). CMA3 results have also been correlated to the assisted reproduction results (Table 3.4). Another indirect approach to investigate the status of the sperm chromatin has been the use of aniline blue staining to detect the presence of histones and therefore indirectly infer in the presence of lower amounts of protamines in the sperm nucleus (Table 3.4, Chevaillier et al. 1987; Colleu et al. 1988). An increase in the percentage of aniline blue cells was found in asthenozoospermic samples as compared to normozoospermic ones (Colleu et al. 1988). Acidic aniline blue was also correlated with differences in sperm nuclear morphology in sperm donors and in infertile patients (Auger et al. 1990). A decreased resistance to chromatin decondensation by treatment with sodium dodecyl sulphate (SDS) and dithiotreitol (DTT) in abnormal sperm as compared to normal sperm has also been taken as evidence for lower protamine S S stability and chromatin packaging (Bustos-Obrego´n and Leiva 1983; Le Lannou et al. 1986; Jager 1990). The accessibility for the fluorescent dye ethidium bromide to DNA has also been correlated to in vitro fertilization (IVF) outcomes (Filatov et al. 1999). In addition to the protamine content, the disulfide bonds crosslinking status between cysteines has also been studied in infertile patients. There is many data indicating that the sperm protein thiols are oxidized upon passage from caput to the cauda epididymis (Rufas et al. 1991). When comparing the thiol labeling patterns, oligospermic or infertile samples were found to have higher SH content (less disulfide bonds) as compared to the normozoospermic ones (Rufas et al. 1991; Lewis et al. 1997; Zini et al. 2001b). After thiol-specific fluorochrome monobromobimane (mBBr)-flow cytometry, spermatozoa from subfertile patients with oligo-astheno-teratozoospermia (the OAT syndrome) were characterized by a biphasic distribution reflecting both over oxidation and incomplete thiol oxidation and possibly a reduced protamine content (Ramos et al. 2008). Animal models also support a correlation between disulfide bond formation and integrity of the DNA (Shirley et al. 2004; Conrad et al. 2005; Zubkova et al. 2005; Suganuma et al. 2005).

3.2.3

Correlation Between Protamines and Sperm DNA Integrity

One of the hypotheses of the function of protamines is that they could be involved in the protection of the genetic message delivered by the spermatozoa (Oliva and Dixon 1991; Mengual et al. 2003; Oliva 2006). Incomplete protamination could render the spermatozoa more vulnerable to attack by endogenous or exogenous agents such as nucleases (Szczygiel and Ward 2002; Sotolongo et al. 2003), free radicals (Irvine et al. 2000; Alvarez et al. 2002), or mutagens. Therefore, this issue has been assessed by different groups using a variety of direct or indirect approaches. Evidence links high DNA fragmentation indexes obtained by SCSA test with lower ICSI or IVF rates (Evenson et al. 1980; Evenson and Wixon 2006). A negative significant correlation between fertilization rate and CMA3 staining or P1/P2 ratio measured directly by electrophoresis has been reported (Nasr-Esfahani

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et al. 2004b). COMET parameters also correlate with embryo cleavage score and with CMA3 staining, suggesting that DNA fragmentation is more frequent in protamine-deficient spermatozoa (Nasr-Esfahani et al. 2005). A reasonably good direct proof that DNA integrity is compromised in protamine-deficient human sperm has been obtained by direct measurement of protamines by electrophoresis (Aoki et al. 2005b). The correlation between protamines and integrity of the DNA in the sperm cells is also supported by animal models. Using transgenic knockout mice for transition proteins, it has been demonstrated that sperm fertility declines during epididymal passage, as revealed by ICSI, while genomic integrity deteriorates (Suganuma et al. 2005). This loss of genomic integrity during passage from the caput to the caudal epididymis in these mice has been related to abnormalities in the protection of the DNA by protamines (Yelick et al. 1987; Shirley et al. 2004; Suganuma et al. 2005). Furthermore, in these mice, the developmental defects appeared at implantation similarly as it has been described in clinical reports from infertile patients with decreased DNA integrity (Tesarik et al. 2004; Suganuma et al. 2005). In humans, the use of ICSI with testicular sperm has demonstrated to improve pregnancy rates in patients with poor pregnancy rates and decreased DNA integrity of ejaculated spermatozoa (Greco et al. 2005b). Thus, a reasonable explanation could be that incomplete or abnormal protamination, as has been observed in many studies, could lead to incomplete disulfide bond formation and incomplete DNA protection during epididymal passage in these patients. All the above observations have led to the proposal of a two-step hypothesis for the generation of damaged DNA (Leduc et al. 2008; Aitken et al. 2009). Abnormal protamination of the sperm cell, set during abnormal spermatogenesis, would leave the sperm genome more prone to be damaged by oxidative stress. Subsequently, free radicals would result in the attack to the sperm DNA, resulting in DNA damage. This hypothesis would explain the correlations detected between abnormal protamine content through gel electrophoresis (de Yebra et al. 1993; Mengual et al. 2003; Torregrosa et al. 2006; Domı´nguez-Fandos et al. 2007; de Mateo et al. 2009) or indirectly with CMA3 staining (Tarozzi et al. 2009; Chiamchanya et al. 2010) and decreased DNA integrity (Aoki et al. 2005b).

3.3

Sperm Nuclear Quality and Outcome of ART

A general trend, illustrated in Tables 3.2 and 3.3, is the apparent dispersion in the results of DNA integrity tests from different studies. This dispersion is likely to arise from several factors. First, the ability of the oocyte to repair DNA damage for which there is very little information and therefore is not a controlled factor in the different studies (Derijck et al. 2008; Jaroudi et al. 2009). Decreased oocyte quality associated with increased maternal age can be an additional factor (Lopes et al. 2009). Also, telomere dysfunction resulting from oxidative stress may contribute to reproductive aging-associated meiotic defects, miscarriage, and infertility

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(Keefe and Liu 2009). Another very important source of variation is the type of test used to measure DNA fragmentation. Each test (TUNEL, COMET, SCSA, SCD, measurements of oxidative stress) has its own range of normality. In addition, very small methodological variations in each test result in large variations in the result (Domı´nguez-Fandos et al. 2007; Muratori et al. 2009). The time from ejaculation and the study are also likely to have a major effect, with an 8% increase of DNA damage per hour (Gosa´lvez et al. 2009). Also, the semen recollection and processing conditions affect DNA damage (Donnelly et al. 2000; Young et al. 2003). Statistical design differences are also likely to account for much of the dispersion in the results since sample sizes fluctuate from 20 to several hundred samples in the different studies (Tables 3.2 and 3.3). Many of the studies do not control statistical Type II errors so that no conclusions can be made upon lack of detection of differences in the study groups. Finally, exogenous factors and genetic background are not yet systematically controlled at present. Altogether, the above factors are likely to result in a large dispersion of results in the different studies (Tables 3.2 and 3.3). From this perspective, larger multicentric studies and metaanalyses of the data are necessary. Some general trends in the assisted reproduction outcomes are already demonstrated from several meta-analysis studies (Zini et al. 2008). Without medical assistance, DNA damage has been related to a decrease in the probability of pregnancy and an increase in the spontaneous time to pregnancy. Using IUI, a strong odds ratio of 9.9 has been detected (Zini et al. 2008). Using FIV, a weak but significant odds ratio of 1.5 is detected (Zini et al. 2008). However, when using ICSI, DNA damage is not associated with the probability of pregnancy. A plausible interpretation for this effect is that ICSI bypasses the detrimental effects that affect spontaneous pregnancy when using IUI or FIV. However, a clear trend towards increased miscarriage has been demonstrated in a meta-analysis involving 11 studies, 1,549 assisted reproduction cycles, 640 pregnancies, and 122 abortions (Zini et al. 2008). In addition to the link between damaged DNA and ART outcomes, the effect abnormal protamine content on ART has also been studied (Table 3.4). One of the initial observations linking protamines and IVF capacity came from the observation of a limited number of patients with an altered P1/P2 ratio with a reduced fertilization index (Khara et al. 1997). Radical differences in protamine content in two siblings associated with different ICSI outcomes were also reported (Carrell et al. 1999). Also, a reduction in protamine P2 and the sperm penetration assay was reported (Carrell and Liu 2001). More recently, it has been described that spermatozoa staining with CMA3, which indirectly indicates a possible deficiency in protamines, has an IVF percentage of 36.8%, which is below the index reached (64.6%) with the negative spermatozoa after using this dye (Nasr-Esfahani et al. 2004a). Subsequent work using this approach demonstrated the presence of increased DNA fragmentation in presumably protamine-deficient spermatozoa (Nasr-Esfahani et al. 2005). This group also measured directly the protamines P1 and P2 by gel electrophoresis and found a negative significant correlation of the fertilization rate with the protamine deficiency and the P1/P2 ratio (Nasr-Esfahani et al. 2004b, 2005). The results of additional recent studies on the

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correlation of the CMA3 results with the assisted reproduction outcomes are shown in Table 3.4. The expression of the genes encoding protamines 1 and 2 in testicular spermatids of azoospermic patients biopsied and treated by ICSI has also been studied (Steger et al. 2003; Mitchell et al. 2005). Using this approach, a lower expression of the mRNA corresponding to the protamine P1 gene was found in couples who did not reach a pregnancy in comparison with couples who reached a pregnancy. At the protein level, it has been reported that a reduction of the P1/P2 ratio results in a marked reduction of the IVF index in comparison with the patients with a normal or an increased P1/P2 ratio (Table 3.4, Aoki et al. 2005a). Furthermore, the sperm P1/P2 ratios are related to IVF pregnancy rates and predictive of fertilization ability (Aoki et al. 2006). These observations have been confirmed in independent laboratories including ours (Table 3.4, de Mateo et al. 2009). In this study, a significant decrease in fertilization rate was detected in the low P1/P2 group of patients when using IVF, but not when using ICSI (Table 3.4). But even in the ICSI group, a subsequent reduction in the pregnancy rate was detected (Table 3.4). Perhaps this result could be related to the findings of a series of IVF experiments using spermatozoa injured with DTT, where the binding and penetration of the oocytes in the hamster assay was markedly reduced, except if ICSI was used, when the DTT-injured spermatozoa reached an even higher rate of pronuclear formation and decondensation of the sperm head of the spermatozoa (Ahmadi and Ng 1999a, b). It is interesting to note that most of the above studies considered only the P1/P2 ratio, but this ratio provides limited information. For instance, it does not indicate whether the abnormal ratio is due to a change in P1, in P2, or in both. It does not provide information either on the distribution of the protamines along the genome. Thus, it will be interesting in future studies to consider also the protamine to DNA ratio and the distribution of the protamines related to the assisted reproduction results. One of the important issues to consider is the clinical situations associated to a decrease of DNA integrity, as these will likely result in preventable factors. Table 3.5 lists different clinical situations and exogenous causes associated with a decrease in DNA integrity. Among all these factors, only a few have been demonstrated to result in prevention. For example, infection treatment has been shown to decrease DNA damage and improve reproduction outcome (Moskovtsev et al. 2009). Also, severe varicocele treatment through varicocelectomy has been proven to result in a beneficial effect in sperm quality (Zini et al. 2005b). DNA smoking is another preventable factor for which there is evidence that quitting smoking results in an improvement of sperm quality (Soares and Melo 2008). Altered levels of protamines present in infertile patients have been reported to improve upon patient treatment (Chen et al. 2005). However, treating or avoiding the etiological factors is not possible at present in most cases, most likely because they remain largely unknown in the infertile male clinical workup. Therefore, strategies to improve DNA integrity are being developed at present. These strategies can be divided into two groups: selective isolation of good quality sperm cells from an ejaculate and antioxidant treatment therapy (Table 3.6, Aitken et al. 2009). Selective isolation

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Table 3.5 Examples of clinical situations and exogenous causes associated with an altered sperm nuclear quality (mainly with a decrease of DNA integrity) Clinical situations and exogenous causes References Infection and inflammation. Chlamydia, mycoplasma Gallegos et al. (2008) Presence of leukocytes Erenpreiss et al. (2002) Presence of immature cells Said et al. (2005) Paternal age Schmid et al. (2007), Singh et al. (2003) Increased scrotal temperature Varicocele Smith et al. (2006) Fever episodes Banks et al. (2005) Sauna Jacuzzi Occupational exposure Evenson et al. (2000) Oncologic patients. Lymphomas, seminomas, O’Flaherty et al. (2008), Edelstein et al. testicular cancer (2008), Kobayashi et al. (2001) Chemotherapeutic agents and radiation O’Flaherty et al. (2009) Obesity Chavarro et al. (2009) Diabetes Agbaje et al. (2008) Tobacco Potts et al. (1999) Pollution Rubes et al. (2005) Cell phone radiation De Iuliis et al. (2009)

Table 3.6 Examples of therapeutic strategies used to improve sperm nuclear quality Strategy References Selective Swim up Ahmad et al. (2007), Hammadeh et al. (2001), isolation Sakkas et al. (2000) Density gradients Fariello et al. (2009), Hammoud et al. (2009c), Morrell et al. (2004), O’Connell et al. (2003), Mengual et al. (2003), Tomlinson et al. (2001), Erel et al. (2000), Sakkas et al. (2000), Larson et al. (1999), Colleu et al. (1996), Sukcharoen (1995) Filtration through glass Nani and Jeyendran (2001), Engel et al. (2001), wool columns Larson et al. (1999), Van den Bergh et al. (1997) Electrophoresis Aitken et al. (2009), Fleming et al. (2008), Ainsworth et al. (2005, 2007) Other methods Parmegiani et al. (2010), Ubaldi and Rienzi (2008), Dirican et al. (2008), Said et al. (2008), Paasch et al. (2007), Said et al. (2006), Jakab et al. (2005) Surgical recovery of Greco et al. (2005c), Hammadeh et al. (1999) testicular sperm IMSI Junca et al. (2009), Ubaldi and Rienzi (2008), Antinori et al. (2008), Hazout et al. (2006), Berkovitz et al. (2005, 2006), Bartoov et al. (2003) Antioxidant Vitamin C and E, zinc, Tunc et al. (2009), Zini et al. (2009), Tremellen et al. therapy folic acid, B carotene (2007), Greco et al. (2005a, b), Fraga et al. (1991)

pursues the selection of the sperm cells within an ejaculate that contain the least damaged genome. Several strategies have been developed based on the laboratory processing of the sperm sample. These include swim up (Sakkas et al. 2000; Hammadeh et al. 2001), density gradient centrifugation (Colleu et al. 1996;

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Hammoud et al. 2009c), and electrophoresis (Ainsworth et al. 2005, 2007; Fleming et al. 2008; Aitken et al. 2009) among others (Table 3.6). Also, the use of testicular spermatozoa can be considered a form of selective isolation (Greco et al. 2005c). More recently, a high resolution optical microscopy technique called intracytoplasmic morphologically selected sperm injection (IMSI) has been applied to select sperm cells with a morphologically normal sperm nucleus (Berkovitz et al. 2005; Antinori et al. 2008). The other strategy to improve DNA integrity is antioxidant therapy. A large number of trials have been reported to improve sperm DNA quality using an oral antioxidant therapy (Fraga et al. 1991; Greco et al. 2005a, 2005b; Tremellen et al. 2007; Tunc et al. 2009; Zini et al. 2009). However, the efficacy of antioxidant treatment in the management of male infertility is more limited, having been reported that no single antioxidant is able to enhance fertilizing capability in infertile men, whereas a combination of them could provide a better approach (Lanzafame et al. 2009; Zini et al. 2009).

3.4

Present Limitations of Sperm Nuclear Quality Assessment and Open Questions

Current sperm nuclear quality assessment in clinics is based essentially on the detection of DNA fragmentation. However, it should be noted that an important proportion of clinics do not incorporate any type of sperm nuclear quality test in the routine workout of the infertile male. One of the reasons for this could be found in the high success rates of ICSI in the treatment of male factor infertility. However, ICSI is based on the selection of motile spermatozoa at low magnification, which does not provide information of the sperm nuclear quality. However, because it is known that motile sperm tend to have intact DNA as compared to immotile sperm, this procedure provides a certain degree of sperm nuclear quality selection. However, it has been demonstrated that morphologically normal and motile spermatozoa can have fragmented DNA. Thus, DNA damage tests should be incorporated into the clinic, particularly in those cases with recurrent assisted reproduction failure upon ICSI. A quite exciting new approach is based on high magnification selection of the sperm cells prior to sperm injection (IMSI). A correlation between morphologically abnormal sperm heads under high magnification and damaged DNA has been reported. So IMSI selection has an important potential in the selective isolation of sperm cells with intact DNA. It will be interesting to also determine whether IMSI has the potential to select sperm cells with a normal protamine complement. Other important sperm selection procedures based on density centrifugation, electrophoresis, or sperm antigen-based selection should also offer practical treatment options in infertile males with decreased DNA integrity. Another limitation of the present DNA integrity tests is that most of them do not assess the full range and types of potential DNA damage present in the sperm cell. For instance, detection of single- or double-strand breaks can be just a small fraction of the damaged DNA, which may include the presence of oxidized adducts,

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abasic sites of the presence of abnormal cross-linking to chromatin proteins. While different tests are available to measure oxidized DNA, they have not been incorporated as a routine in the clinic. Finally, it should be noted that the impact of damaged DNA may also be related to the repair capacity of damaged DNA in sperm by the oocyte repair machinery, for which there is little information. In addition to the DNA integrity present in the sperm cell, it is clearly important to take into account the sperm proteome and epigenome. Protamine, proteome, and epigenome determinations are being presently applied as part of research projects. It is now clear that the sperm cell delivers an epigenome that is determined by the distribution of genes into the histone- and protamine-associated domains, the presence of histone modifications, and the specific methylation of genes. However, the impact of alterations in these factors potentially present in the infertile male remains largely unexplored. Also, the effect of the potential mutations and imprinting defects associated with infertility and passed on to the next generation with the use of ART and its long-term effect in subsequent generations remains largely unknown. However, we are now at a quite interesting moment in sperm cell genomic, proteomic, and epigenomic research. A clear proof for epigenetic effects linked to DNA methylation is beyond doubt. In addition, a proof that proteins and modified proteins are being transferred to the oocyte upon fertilization has been clearly demonstrated. The possibility of investigating the extent to which this chromatin structure, and RNA and protein “cargo” delivered by the sperm cells determine many aspects of early embryonic development is now open. It will also be interesting to further investigate the extent of involvement of sperm nuclear quality anomalies present in infertile patients in reproductive outcomes. Acknowledgments Supported by a grant from the Ministerio de Ciencia y Tecnologı´a BFU2009 07118, fondos FEDER to R.O.

References Abu Hassan D, Koester F, Shoepper B et al (2006) Comet assay of cumulus cells and spermatozoa DNA status, and the relationship to oocyte fertilization and embryo quality following ICSI. Reprod Biomed Online 12:447 452 Agbaje IM, McVicar CM, Schock BC et al (2008) Increased concentrations of the oxidative DNA adduct 7, 8 dihydro 8 oxo 2 deoxyguanosine in the germ line of men with type 1 diabetes. Reprod Biomed Online 16:401 409 Ahmad L, Jalali S, Shami SA, Akram Z (2007) Sperm preparation: DNA damage by comet assay in normo and teratozoospermics. Arch Androl 53:325 338 Ahmadi A, Ng SC (1999a) Destruction of protamine in human sperm inhibits sperm binding and penetration in the zona free hamster penetration test but increases sperm head deconden sation and male pronuclear formation in the hamster ICSI assay. J Assist Reprod Genet 16:128 132 Ahmadi A, Ng SC (1999b) Influence of sperm plasma membrane destruction on human sperm head decondensation and pronuclear formation. Arch Androl 42:1 7

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Martı´nez Heredia J, Estanyol JM, Ballesca JL, Oliva R (2006) Proteomic identification of human sperm proteins. Proteomics 6:4356 4369 Martı´nez Heredia J, de Mateo S, Vidal Taboada JM et al (2008) Identification of proteomic differences in asthenozoospermic sperm samples. Hum Reprod 23:783 791 McGrath J, Solter D (1984) Completion of mouse embryogenesis requires both the maternal and paternal genomes. Cell 37:179 183 McKay DJ, Renaux BS, Dixon GH (1986) Human sperm protamines. Amino acid sequences of two forms of protamine P2. Eur J Biochem 156:5 8 McKusick VA (2007) Mendelian inheritance in man and its online version, OMIM. Am J Hum Genet 80:588 604 Mengual L, Ballesca´ JL, Ascaso C, Oliva R (2003) Marked differences in protamine content and P1/P2 ratios in sperm cells from percoll fractions between patients and controls. J Androl 24:438 447 Mezquita C (1985) Chromatin composition, structure and function in spermatogenesis. Revis Biol Celular 5:1 124 Micin´ski P, Pawlicki K, Wielgus E et al (2009) The sperm chromatin structure assay (SCSA) as prognostic factor in IVF/ICSI program. Reprod Biol 9:65 70 Milingos S, Comhaire FH, Liapi A, Aravantinos D (1996) The value of semen characteristics and tests of sperm function in selecting couples for intra uterine insemination. Eur J Obstet Gynecol Reprod Biol 64:115 118 Mitchell V, Steger K, Marchetti C et al (2005) Cellular expression of protamine 1 and 2 transcripts in testicular spermatids from azoospermic men submitted to TESE ICSI. Mol Hum Reprod 11:373 379 Moll AC, Imhof SM, Cruysberg JR, Schouten van Meeteren AY, Boers M, van Leeuwen FE (2003) Incidence of retinoblastoma in children born after in vitro fertilization. Lancet 361:309 310 Morrell JM, Moffatt O, Sakkas D et al (2004) Reduced senescence and retained nuclear DNA integrity in human spermatozoa prepared by density gradient centrifugation. J Assist Reprod Genet 21:217 222 Morris ID, Ilott S, Dixon L, Brison DR (2002) The spectrum of DNA damage in human sperm assessed by single cell gel electrophoresis (comet assay) and its relationship to fertilization and embryo development. Hum Reprod 17:990 998 Moskovtsev SI, Lecker I, Mullen JB et al (2009) Cause specific treatment in patients with high sperm DNA damage resulted in significant DNA improvement. Syst Biol Reprod Med 55:109 115 Mudrak O, Tomilin N, Zalensky A (2005) Chromosome architecture in the decondensing human sperm nucleus. J Cell Sci 118:4541 4550 Muratori M, Tamburrino L, Tocci V et al (2009) Small variations in crucial steps of tunel assay coupled to flow cytometry greatly affect measures of sperm DNA fragmentation. J Androl. doi:10.2164/jandrol.109.008508 Muriel L, Garrido N, Ferna´ndez JL et al (2006a) Value of the sperm deoxyribonucleic acid fragmentation level, as measured by the sperm chromatin dispersion test, in the outcome of in vitro fertilization and intracytoplasmic sperm injection. Fertil Steril 85:371 383 Muriel L, Meseguer M, Ferna´ndez JL et al (2006b) Value of the sperm chromatin dispersion test in predicting pregnancy outcome in intrauterine insemination: a blind prospective study. Hum Reprod 21:738 744 Nakamura H, Kimura T, Nakajima A et al (2002) Detection of oxidative stress in seminal plasma and fractionated sperm from subfertile male patients. Eur J Obstet Gynecol Reprod Biol 105:155 160 Nani JM, Jeyendran RS (2001) Sperm processing: glass wool column filtration. Arch Androl 47:15 21 Nasr Esfahani MH, Razavi S, Mardani M (2001) Relation between different human sperm nuclear maturity tests and in vitro fertilization. J Assist Reprod Genet 18:219 225

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Chapter 4

Gene Expression/Phenotypic Abnormalities in Placental Tissues of Sheep Clones: Insurmountable Block in Cloning Progress? Pasqualino Loi and Grazyna Ptak

Abstract Although 10 years have already passed by since the production of the first mammal by nuclear transfer of somatic cells (SCNT/cloning), the overall improvement achieved so far is negligible. Even in species where SCNT is relatively successful, like cattle, the efficiency is still very low. Therefore, the development of novel strategies to improve cloning efficacy is a major challenge for reproductive biologists. The crucial question is whether procedures can be improved to minimize adverse developmental and epigenetic effects. The published data indicate that abnormalities of the placental tissues, namely in vascularization, are a main reason of the poor development of cloned conceptuses. This chapter focuses on the phenotypic and gene expression analysis of placental tissues in sheep clones and suggests possible strategies for the improvement of cloning procedures. Keywords Cloning Placenta Gene Expression Sheep

4.1

Introduction

The first animal produced by nuclear transfer of a somatic cell (SCNT/cloning) was Dolly the sheep (Wilmut et al. 1997). The breakthrough was highly emphasized by the media not only for the outstanding scientific achievement, but mainly for the ethical issues it raised. Unfortunately, the ensuing equation “animal cloning ¼ human cloning” launched by the media generated a strong aversion against SCNT. The introduction, next, of the concept of “Therapeutic Cloning,” where cloned human embryos were to be produced in order to generate embryonic stem (ES) cells for cell replacement therapy (Lanza et al. 1999), further contributed to generate a negative perception about SCNT.

P. Loi (*) and G. Ptak Department of Comparative Biomedical Sciences, University of Teramo, Piazza A. Moro 45, 64100 Teramo, Italy e mail: [email protected], [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 4, # Springer Verlag Berlin Heidelberg 2011

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Fortunately, the scientific panorama has changed. The induction of pluripotency in somatic cells through the transfection of four genes expressed in stem cells (Takahashi and Yamanaka 2006) has given an ethically impeccable alternative to therapeutic cloning for cell replacement therapies (Tweedell 2008). Furthermore, the American Food and Drugs Administration (FDA) gave its green light for foodstuff produced from cloned animals (http://www.fda.gov/bbs/topics/NEWS/ 2006/NEW01541.html, Yang et al. 2007). These two events jointly restored a positive atmosphere, confining at the same time cloning to its natural niche that is animal reproduction. SCNT allows us to make copies of a particular genotype and this has great potential applications in many fields: 1. Multiplication of elite animals (Wells 2003). As clones are the genetic copy of the cell donor, they present similar productive/reproductive performances. It should be stressed that besides quanti/qualitative animal traits, like meat or milk, current selection strategies also take into account other relevant parameters, like resistance to common pathologies (mastitis, infections and parasitic diseases) rusticity and others. Indeed, breeding out these complex traits using the traditional selection schemes might turn out to be complicated (if not impossible) and time consuming. In the next few years, sequencing systems based on nanotechnologies will be available, opening the door to very low-cost whole genome sequencing with prices decreasing to $1,000 or even to $100 per entire genome within 5 years [VisiGen Biotechnologies, Complete Genomics (http://visigenbio. com)]. The combination of SCNT with third generation DNA sequencing will allow to “copy and paste” the selected genotypes, speeding up the selection process. However, it must be stated that in all cases SCNT should complement classic breeding and reproduction technologies and be applied only strategically. 2. Production of transgenic animals (Robl et al. 2007). The nucleus to be transplanted into the enucleated egg can be genetically modified to produce cloned transgenic embryos and ultimately live animals. There are two main applications for large transgenic animals. The first is the generation of animals which express human peptides to be used in medicine. For instance, the European Medicines Agency (EMEA) has authorized the commercialization of human antithrombin III (ATyrin), the first protein purified from the milk of transgenic animals, for the treatment of haemophilic patients (Niemann and Kues 2007). The second is the generation of engineered animals (essentially pigs) for the production of organs to be used for transplantation into humans (xenotransplantation) (Niemann and Kues 2007). However, the production of transgenic pigs for xenotransplantation seems to be more complicated and a long term goal as yet. 3. Multiplication of animal species threatened by extinction (Loi et al. 2001; Holt et al. 2004). The last century has seen a dramatic reduction in animal species, mostly large mammals, mainly caused by human-related activities. The obvious consequence of such phenomenon is the progressive contraction in biodiversity worldwide. The situation has been clearly depicted in the last report published by the World Conservation Union (http://www.iucn.org/). Paradoxically, this

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problem does not only involve wild species, but also domestic ones, often local or typical breeds which are perfectly adapted to particular ecotypes, but are disappearing and are substituted by just a few more productive species. According to the Food and Agriculture Organization of the United Nations (FAO 2007), a total of 1,491 breeds world-wide (20%) are classified as being either critically endangered, critical-maintained, endangered, or endangered-maintained (http:// faolex.fao.org/docs/pdf/ber90159.pdf). Although in mammalian livestock the proportion of breeds classified at risk is lower than average (16%) in absolute terms, their number is very high (881 breeds). This is a minimum estimate, since no information is available to evaluate the risk status of a large number of breeds, mainly native of Asia and Africa. SCNT might thus represent a unique tool for the preservation/expansion of such threatened populations. Unfortunately, cloning efficiency is very low with only 1 5% of offspring delivered at the end of the process. This disappointing outcome reflects a stall in nuclear reprogramming state of the art. Following the production of the first mammal by SCNT, the best laboratories concentrated their efforts in order to understand the molecular mechanisms underlying nuclear reprogramming, at that time a completely unknown phenomenon. However, following the advent of therapeutic cloning/ES cells (Lanza et al. 1999) and the discovery of iPS cells (Takahashi and Yamanaka 2006) scientists shifted their attention to this last and more promising field, leaving SCNT at stake.

4.2

Abnormal Nuclear Reprogramming in Clones and Related Placental Phenotypes

The reversal of cellular differentiation through SCNT is an unpredictable, but crucial event. Indeed, compromised development of cloned conceptuses is due to the oocyte’s failure to restore the totipotent state in the transplanted nucleus through the process of “Nuclear Reprogramming” (Rideout et al. 2001). Nuclear Reprogramming is a very general term which describes the resetting of the cell memory established during cell commitment and differentiation. At the moment, complete reprogramming is achieved only exceptionally, while the majority of clones show epigenetic deregulation and abnormal gene expression at pre- and postimplantation stage and even after birth (Latham 2005; Ogura et al. 2002; Tsunoda and Kato 2002; Tamashiro et al. 2003; Kremenskoy et al. 2006; Morgan et al. 2006; Loi et al. 2005; Chae et al. 2009). Some possible nuclear reprogramming improvement strategies might be worth trying, but it is mandatory to establish a “reference” model to assess their effectiveness. The only proof that a complete reprogramming has been achieved is the delivery of healthy offspring after the transfer of cloned embryos into suitable foster mothers.

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Hence the question is: how can we predict that a cloned embryo is “normal” and that there will be a happy end of the story or, conversely, that such an embryo will stop developing during pregnancy, or worse, will die soon after birth due to severe malformations? In addition, the transfer of cloned embryos also threatens the life of the foster mothers, as, at least in ruminants (particularly in sheep), they often (35%) develop acute hydroallantois at the end of pregnancy which requires a cesarean section (Loi et al. 2005). Therefore, we are firmly convinced that it is unethical to carry out in vivo developmental trials unless there is a clear evidence on the efficacy of the specific reprogramming approach tested. But what might be considered as the proof of principle that a cloning strategy is better than the available state of the art? Some possible markers of nuclear reprogramming, like the expression of relevant genes involved in the maintenance of pluripotency genes, such as Pou5f1/ POU5F1 (previously known as Oct4/OCT4), Nanog, Otx2, Ifitm3, GATA6, SNRPN, etc., have been proposed (Suzuki et al. 2009; Sawai 2009; Xing et al. 2009). Some of these genes (Pou5f1/POU5F1) have also been tagged with a GFP reporter gene, thus allowing their visualization in living embryos (Wuensch et al. 2007). In addition to the analysis of a restricted panel of specific genes, large scale expression profiles might also be resolved in early cloned embryos, providing a global view on genome activity (Amarnath et al. 2007; Smith et al. 2007; Kato et al. 2007; Zhou et al. 2008). These tools are valuable and provide much information. However, the main message coming from the gene expression analyses carried out at blastocyst stage in clones is that only a minority of genes is actually deregulated (Smith et al. 2005). This finding contrasts with the large number of foetal losses and abnormalities occurring during postimplantation development. There is strong evidence that developmental errors occur immediately after implantation in mice (Jouneau et al. 2006). It might be that minor reprogramming errors, which are not detected in preimplantation embryos, are magnified later in development. Moreover, a common finding in all cloned animals is that most of the alterations described occur in extraembryonic tissues. The functional and morphological abnormalities of the placenta have been studied in detail in mice and partially in cow and sheep clones, whereas no information are available for the other cloned species. Placental hypertrophy, the common phenotype found in mice (Wakisaka-Saito et al. 2006) and in cattle (Hashizume et al. 2002), is absent in sheep (Loi et al. 2005). The main histological lesion described in mice is an enlarged spongiotrophoblast cell layer (Wakisaka-Saito et al. 2006) with concomitant enlargement of the trophoblast giant cells and disorganization of the labyrinth (Wakisaka-Saito et al. 2006). SCNT placentas in cows display fewer placentomes that are often larger than normal and irregular in size (Chavatte-Palmer et al. 2006; Constant et al. 2006; Hill et al. 2000). Histological examination revealed a hypotrophic trophoblastic epithelium and reduced vascularization (Hoffert-Goeres et al. 2007).

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A dramatic reduction in the vascular bed has been described in early cloned sheep fetuses, and the finding was associated with fetal losses (De Sousa et al. 2001). The establishment of normal vascularization and angiogenesis in placental tissues is crucial for placental and fetal development and this is certainly a bottleneck for cloned conceptuses. Therefore, we decided to focus our attention on this issue. Implantation is very different in mice and sheep. Instead of aggressing the uterine lining, as in mice, the sheep conceptus expand their extraembryonic component, quite dramatically, to get in close contact with the endometrium; then angiogenesis, confined to the chorioallantoic membrane apposed to specialized areas of the uterine epithelium (caruncolae), starts (Spencer et al. 2007). Differently from fetal angiogenesis (Kubota and Takigawa 2007), the very early phases of extraembryonic angiogenesis are not well defined in mice and data concerning sheep are essentially not available. Therefore we set out first to define the window during which placental vascularization and angiogenesis take place in sheep and found that it starts at day 20 in normal pregnancy (i.e., produced by natural mating). We next monitored the expression pattern of a suitable panel of marker genes between day 20 and 24 of pregnancy. Placental development and function is also regulated by imprinted genes, therefore, the majority of the previous studies have focused on the epigenetic deregulation of imprinted genes in cloned fetuses (Zhou et al. 2007). However, the indications given by global epigenetic marks, like DNA methylation for instance, suggest that besides imprinted genes, larger regions of the genome including nonimprinted genes must be affected in SCNT. With this in mind, we have analyzed by RT-PCR the expression of 22 imprinted and nonimprinted genes, including the main angiogenetic factors and the components of the Notch signalling pathway (Table 4.1), which are also involved in vascular development and differentiation (Gasperowicz and Otto 2008) in the extraembryonic tissues of normal conceptuses. A consistent pattern was observed for all genes analyzed, which was later compared with the expression profiles of the same genes in extraembryonic tissues of conceptuses produced by in vitro fertilization (IVF) or SCNT. As expected, significant differences for most of the imprinted genes analyzed were observed in IVF and cloned embryos (Fig. 4.1). The expression of other nonimprinted genes was also deregulated in SCNT extraembryonic tissues. Particularly, deregulation in the expression of angiogenetic and Notch, such as NOTCH 1, 2 and 4 and JAG1 (Figs. 4.2 and 4.3), suggests that reprogramming errors might undermine the Table 4.1 Panel of genes used a marker of placental vascularization/development in early post implantation sheep normal conceptuses (days 20 24) Notch pathway Angiogenetic factors Maternally Paternally imprinted imprinted NOTCH 1, 2, 4; DLL 1, 4; JAG 1, 2; VEGF; FGF 2; FGF2 R2; IGF2; H19; CDKN1C; HEY 1, 2. EPHRIN B2 ANG1; ANG2; Tie 2 MEST PHLDA2 Function Functions Vascular formation Growth factors production Vascular vein/artery differentiation Vascular proliferation Nutrients transport

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Fig. 4.1 mRNA expression of imprinted genes in chorioallantoic tissues collected on day 20 of pregnancy obtained by natural breeding (CTR), in vitro fertilization (IVF), and somatic cell nuclear transfer (SCNT). The results show a down regulated expression of IGF2, H19 and PHLDA2 in SCNT models. In addition DNMT1 expression were strongly down regulated in placentae from IVF and SCNT models. CTR natural mating; IVF in vitro fertilized; SCNT somatic cell nuclear transfer

Fig. 4.2 mRNA expression of notch pathway genes in chorioallantoic tissues collected on day 20 of pregnancy obtained by natural breeding (CTR), in vitro fertilization (IVF), and somatic cell nuclear transfer (SCNT). The results show a down regulated expression of NOTCH4 and DLL1 in IVF models. In addition placentae from SCNT embryos show an up regulation respect to those obtained by IVF for NOTCH1, NOTCH2 and JAG1. CTR natural mating; IVF in vitro fertilized; SCNT somatic cell nuclear transfer

development of clones soon after implantation by interfering with the establishment of normal vascularization. Remarkably, the expression in extraembryonic tissues of one of the master genes for the establishment of imprinting marks in the genome, the enzyme DNA Methyltransferase 1 (DNMT1) was strongly decreased in SNCT in comparison to both control and IVF conceptuses (Fig. 4.1). These results did not surprise us, although they go against the dominant view according to which abnormal reprogramming results mostly from the unbalanced

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Fig. 4.3 mRNA expression of angiogenetic factors in chorioallantoic tissues collected on day 20 of pregnancy obtained by natural breeding (CTR), in vitro fertilization (IVF), and Somatic Cell Nuclear Transfer (SCNT). The results show a down regulated expression of FGF2, ANG2 and its receptor TIE2 in IVF models respect to the controls. CTR natural mating; IVF in vitro fertilized; SCNT somatic cell nuclear transfer

expression of imprinted genes. Another important point emerging from these preliminary studies is that the genes abnormally expressed in sheep clones differ considerably from those described in mice. This is very important, as it does suggest a species-specific molecular fingerprint resulting from SCNT. Thus, having now established the reference model for the most important genes that drive placental vascularization and growth in sheep embryos, we can assess the different reprogramming strategies, at least in sheep.

4.3

Possible Directions to Improve SCNT

The many solutions tested for improving nuclear transfer have been described in exhaustive reviews, to which the reader can refer (Loi et al. 2008a; Wakayama 2007). However, none of them, so far, has produced major advancements of the current state of the art. Furthermore, the most effective of them, which is the use of ES cells as nucleus donors, can be applied only to species where bona fide ES cells are available, in other words, in none of the large animals. The second more reputed approach, the pretreatment of donor cells/reconstructed embryos with the histone deacetylase inhibitor Trichostatin A, has been validated beyond any doubt only in mice (Wakayama 2007). We recently suggested that abnormal reprogramming might result from unbalanced or asymmetric reprogramming of the somatic cell nucleus due to the fact that the reprogramming machinery of the oocyte does not seem to recognize the paternal chromosomes (Loi et al. 2008b). This hypothesis stems from the differential “treatment” reserved to the paternal and maternal sets of chromosomes early in

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development, as suggested by several studies. Instead of intermingling at syngamy, as the gross embryological observation suggests, each parental set of chromosomes is spatially separated and occupies distinct nuclear territories during early embryogenesis in mouse, suggesting a sex-dependent chromatin remodeling (Barton et al. 2001; Haaf 2001). Other indications of an “asymmetry” between parental chromosomes in early development have been detailed elsewhere (Loi et al. 2008b). Developmental programming is a complex, stepwise process that occurs separately in the male and female germ line. Oocyte chromosomes are fully reprogrammed or better “programmed” by the end of the follicular growth as a result of epigenetic modifications. Likewise, sperm cells are programmed after extensive chromatin reorganization and epigenetic formatting in the male gonads (Caron et al. 2005). When a somatic nucleus is transplanted into an enucleated oocyte is reprogrammed by the molecular mechanisms available in the oocyte, which are evolutionarily designed to recognize the maternal, not the paternal chromosomes, as those are normally postmeiotically reprogrammed in the male germ. The molecular and structural changes occurring in the chromatin during the transition from somatic to spermatozoa/specific configuration have been gradually unveiled and several key factors involved in this process have been identified. One of them, the Bromodomain Testis-Specific (BRDT) protein, has been the focus of particular attention since it possesses the unique ability to specifically induce a dramatic compaction of acetylated chromatin during spermatozoa maturation and also in somatic cells (Pivot-Pajot et al. 2003). We have recently suggested that a suitable approach for making the oocyte’s life easier during cloning might be to induce the expression of testis-specific reprogramming/remodeling molecules in somatic cells prior nuclear transfer (Loi et al. 2008b). Indeed, we speculated that reprogramming would be facilitated if we were able to make the chromatin of a somatic nucleus resemble as much as possible that of a programmed sperm cell. In a preliminary study we showed that somatic cells (skin fibroblasts) transiently transfected with DRBT were better reprogrammed, at least at the blastocyst stage, than untransfected controls (Loi, unpublished). Whereas the induced expression of only four totipotent genes in somatic cells reversed the differentiation memory and restored the multipotent status of differentiated cells (Takahashi and Yamanaka 2006), our approach goes in the opposite direction, aiming at forcing a terminally differentiated somatic cell to acquire (at least partially) the chromatin configuration of a highly specialized cell, the spermatozoa. This is not the only option available though. In case the “spermatization” of somatic cells fails, other strategies that involve the use of iPS cells, or the knowledge derived from such studies might be put forth. The IPs field is growing at a very rapid pace and new straightforward strategies are being elaborated almost every week (Lin et al. 2009). It is thus possible that insights into reprogramming might come from iPS cells as well. In particular, work carried out in iPS cells has shed new light on the role of the tumor suppressor gene p53 during nuclear reprogramming (Kawamura et al. 2009). This finding suggests that p53 silencing in somatic cells prior to nuclear transfer might result in a better reprogramming.

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In theory iPS cells should be more “reprogrammable” than differentiated cells and therefore their use as nuclei donors could in theory improve SCNT outcome, although no reports on this subject have been published as yet. A potential obstacle for the production of iPS in large animals might be the lack of knowledge on culture requirements for species-specific ES cells. Even in this regard the general picture is by far more positive that some years ago. The standardization of ES cell isolation has never been so advanced, and complete synthetic media have been developed (Ying et al. 2008). This breakthrough brings some hope for the successful isolation of farm animal ES cells, a long sought after goal.

4.4

Concluding Remarks

The state of knowledge on the mechanisms of canonically induced nuclear reprogramming (nuclear transfer) or by the newest approach (IPs) has never been so advanced. The present conjuncture is therefore highly favorable to allow an attack, perhaps definitive, on the unsolved problems in SCNT. Such attack could be ideally carried out on two fronts: 1. The “core” issue of SCNT: nuclear reprogramming 2. The “fringe” of SCNT SCNT core nuclear reprogramming must be tackled with completely innovative solutions, and what we propose goes in that direction. The “spermatization” of chromatin in a somatic cell relies on the same philosophy of iPS cells, but with the difference that in our case a differentiated cell is changed into another terminally differentiated and highly specialized cell, the male gamete. It is certainly a risky strategy, but fortunately in case of failure other options might be foreseen, not least the use of iPS and/or ES cells as nucleus donors for SCNT. The evidence that undifferentiated cells are easily reprogrammed is available since the dawn of nuclear transfer. What is important to stress is that the outcome of a winning reprogramming strategy holds the potential to be transversal, in other words, easily transferable to all “clonable” mammals. SCNT fringe SCNT is a complex multistep process, which involves the maturation of an oocyte and its enucleation, the selection and preparation of the donor cell and its electro fusion into the oocyte, the artificial activation of the recipient oocyte, in vitro culture of the cloned embryos and finally the activation of the embryo genome. The success of each step accounts for the final outcome of SCNT. Oocyte physiology, activation dynamics and preimplantation embryo metabolism differ among species, therefore, as different from the “core” issue it is unlikely that a standardized protocol will be developed for all species. A multistep approach, focusing on oocyte/embryo biology and particularly on their metabolic needs in vitro might prove to be effective. A great deal of efforts should be dedicated to the optimization of in vitro systems for mass production of fully competent recipient oocytes. Robust protocols are available for the maturation of ovine, bovine and

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pig oocytes (Galli and Lazzari 2008) but for other clonable species, like wild and rare animals, practically nothing is known. Preimplantation development of cloned embryos is predominantly carried out in vitro using culture media designed for normal embryos. SCNT embryos apparently develop better in complex media, like those employed for primary culture of somatic cells, suggesting that some of the metabolic pathways of the differentiated cells are still active after nuclear transfer (Chung et al. 2002). The development of “compromise” media (i.e., hybrid somatic cells/embryo culture media) should improve the viability of cloned embryos (Cavaleri et al. 2006; Boiani et al. 2005). Another critical factor that affects nuclear reprogramming appears to be the timing of zygotic genome activation (ZGA). Indeed, species with delayed ZGA apparently have a reprogramming advantage (more time to get a complete reprogramming). Accordingly, the mouse, in which ZGA starts during the late part of the first cell cycle, shows a much lower reprogramming efficiency than cattle, where ZGA takes place during the fourth cell cycle (Meissner and Jaenisch 2006). Unfortunately, nothing can be done to influence this feature. Thus, in the fortunate case that the core issue of SCNT will be cracked, it should not take too long to optimize the species-specific “fringes” for globally improving SCNT. Of course the definitive answer on the efficacy of any corefringe strategy will be the normal expression of crucial genes assessed immediately after implantation. A positive outcome will be represented by a match between the expression patterns of target genes in IVF and cloned conceptuses. This fortunate circumstance would give the green light for the transfer of clones into foster mothers and should provide the crucial achievement of around 30 40% live, normal offspring that will make of SCNT a valuable tool for (farm and wild) animal reproduction and breeding. Acknowledgments We are indebted to Mrs Paola Toschi and Mrs Antonella Fidanza for the expression analysis; to Drs. Marta Czernick, Federica Zacchini and Fiorella Di Egidio for the embryological work and the in vivo developmental studies. The research in our laboratory is supported by the Program “Ideas” European Research Council call identifier ERC 2007 StG 210103 angioplace; “Expression and Methylation Status of Genes Regulating Placental Angio genesis in Normal, Cloned, IVF and Monoparental Sheep Fetuses” (G PTAK); PRIN (Italian Ministry of Education) 2007 funds, n. 2007MY2M92 (G Ptak), and Teramo University funds (PL). PL acknowledges financial support from EU 7FP, NextGene project n. 244356 CP FP SICA?

References Amarnath D, Li X, Kato Y et al (2007) Gene expression in individual bovine somatic cell cloned embryos at the 8 cell and blastocyst stages of preimplantation development. J Reprod Dev 53 (6):1247 1263 Barton SC, Arney KL, Shi W et al (2001) Genome wide methylation patterns in normal and uniparental early mouse embryos. Hum Mol Genet 10:2983 2987 Boiani M, Gentile L, Gambles VV et al (2005) Variable reprogramming of the pluripotent stem cell marker Oct4 in mouse clones: distinct developmental potentials in different culture environments. Stem Cells 23:1089 1094

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Caron C, Govin J, Rousseaux S et al (2005) How to pack the genome for a safe trip. Prog Mol Subcell Biol 38:65 89 Cavaleri F, Gentile L, Scholer HR et al (2006) Recombinant human albumin supports development of somatic cell nuclear transfer embryos in mice: toward the establishment of a chemically defined cloning protocol. Cloning Stem Cells 8:24 40 Chae JI, Lee KS, Kim DJ et al (2009) Abnormal gene expression in extraembryonic tissue from cloned porcine embryos. Theriogenology 71(2):323 333 Chavatte Palmer P, de Sousa P, Laigre P et al (2006) Ultrasound fetal measurements and pregnancy associated glycoprotein secretion in early pregnancy in cattle recipients carrying somatic clones. Theriogenology 66:829 840 Chung YG, Mann MR, Bartolomei MS (2002) Nuclear cytoplasmic “tug of war” during cloning: effects of somatic cell nuclei on culture medium preferences of preimplantation cloned mouse embryos. Biol Reprod 66:1178 1184 Constant F, Guillomot M, Heyman Y et al (2006) Large offspring or large placenta syndrome? Morphometric analysis of late gestation bovine placentomes from somatic nuclear transfer pregnancies complicated by hydrallantois. Biol Reprod 75:122 130 De Sousa PA, King T, Harkness L et al (2001) Evaluation of gestational deficiencies in cloned sheep fetuses and placentae. Biol Reprod 65:23 30 Galli C, Lazzari G (2008) The manipulation of gametes and embryos in farm animals. Reprod Domest Anim 43(2):1 7 Gasperowicz M, Otto F (2008) The notch signalling pathway in the development of the mouse placenta. Placenta 29:651 659 Haaf T (2001) The battle of the sexes after fertilization: behaviour of paternal and maternal chromosomes in the early mammalian embryo. Chromosome Res 9:263 271 Hashizume K, Ishiwata H, Kizaki K et al (2002) Implantation and placental development in somatic cell clone recipient cows. Cloning Stem Cells 4:197 209 Hill JR, Burghardt BC, Jones K et al (2000) Evidence for placental abnormality as the major cause of mortality in first trimester somatic cell cloned bovine fetuses. Biol Reprod 63:1787 1794 Hoffert Goeres KA, Batchelder CA, Bertolini M et al (2007) Angiogenesis in day 30 bovine pregnancies derived from nuclear transfer. Cloning Stem Cells 9:595 607 Holt WV, Pickard AR, Prather RS (2004) Wildlife conservation and reproductive cloning. Reproduction 127:317 324 Jouneau A, Zhou Q, Camus A et al (2006) Developmental abnormalities of NT mouse embryos appear early after implantation. Development 133:597 607 Kato Y, Li X, Amarnath D et al (2007) Comparative gene expression analysis of bovine nuclear transferred embryos with different developmental potential by cDNA microarray and real time PCR to determine genes that might reflect calf normality. Cloning Stem Cells 9:495 511 Kawamura T, Suzuki J, Wang YV et al (2009) Linking the p53 tumour suppressor pathway to somatic cell reprogramming. Nature 460:1140 1144 Kremenskoy M, Kremenska Y, Suzuki M et al (2006) DNA methylation profiles of donor nuclei cells and tissues of cloned bovine fetuses. J Reprod Dev 52:259 266 Kubota S, Takigawa M (2007) CCN family proteins and angiogenesis: from embryo to adulthood. Angiogenesis 10:1 11 Lanza RP, Cibelli JB, West MD (1999) Human therapeutic cloning. Nat Med 5:975 977 Latham KE (2005) Early and delayed aspects of nuclear reprogramming during cloning. Biol Cell 97:119 132 Lin T, Ambasudhan R, Yuan X et al (2009) A chemical platform for improved induction of human iPSCs. Nat Methods 6:805 808 Loi P, Matsukawa K, Takahashi K et al (2008a) Nuclear reprogramming: what has been done and potential avenues for improvements. Cent Eur J Biol 3:11 223 Loi P, Beaujean N, Khochbin S et al (2008b) Asymmetric nuclear reprogramming in somatic cell nuclear transfer? Bioessays 30:66 74 Loi P, Clinton M, Vackova I et al (2005) Placental abnormalities associated with post natal mortality in sheep somatic cell clones. Theriogenology 65:1110 1121

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Loi P, Ptak G, Fulka J Jr et al (2001) Genetic rescue of an endangered mammal by cross species nuclear transfer using post mortem somatic cells. Nat Biotechnol 19:962 964 Meissner A, Jaenisch R (2006) Mammalian nuclear transfer. Dev Dyn 235:2460 2469 Morgan HD, Santos F, Green K et al (2006) Epigenetic reprogramming in mammals. Hum Mol Genet 14:R47 R58 Niemann H, Kues WA (2007) Transgenic farm animals an update. Reprod Fertil Dev 19:762 770 Ogura A, Inoue K, Ogonuki N et al (2002) Phenotypic effects of somatic cell cloning in the mouse. Cloning Stem Cells 4:397 405 Pivot Pajot C et al (2003) Acetylation dependent chromatin reorganization by BRDT, a testis specific bromodomain containing protein. Mol Cell Biol 23:5354 5365 Rideout WM III, Eggan K, Jaenisch R (2001) Nuclear cloning and epigenetic reprogramming of the genome. Science 293:1093 1098 Robl JM, Wang Z, Kasinathan P et al (2007) Transgenic animal production and animal biotech nology. Theriogenology 67:127 133 Sawai K (2009) Studies on gene expression in bovine embryos derived from somatic cell nuclear transfer. J Reprod Dev 55:11 16 Smith C, Berg D, Beaumont S (2007) Simultaneous gene quantitation of multiple genes in individual bovine nuclear transfer blastocysts. Reproduction 133:231 242 Smith SL, Everts RE, Tian XC et al (2005) Global gene expression profiles reveal significant nuclear reprogramming by the blastocyst stage after cloning. Proc Natl Acad Sci USA 102:17582 17587 Spencer TE, Johnson GA, Bazer FW et al (2007) Pregnancy recognition and conceptus implanta tion in domestic ruminants: roles of progesterone, interferons and endogenous retroviruses. Reprod Fertil Dev 19:65 78 Suzuki J Jr, Therrien J, Filion F et al (2009) In vitro culture and somatic cell nuclear transfer affect imprinting of SNRPN gene in pre and post implantation stages of development in cattle. BMC Dev Biol 9:9 Takahashi K, Yamanaka S (2006) Induction of pluripotent stem cells from mouse 1098 embryonic and adult fibroblast cultures by defined factors. Cell 126:663 676 Tamashiro KL, Sakai RR, Yamazaki Y et al (2003) Phenotype of cloned mice: development, behaviour, and physiology. Exp Biol Med 228:1193 1200 Tsunoda Y, Kato M (2002) Recent progress and problems in animal cloning. Differentiation 69:158 161 Tweedell KS (2008) New paths to pluripotent stem cells. Curr Stem Cell Res Ther 3:151 162 Wakayama T (2007) Production of cloned mice and ES cells from adult somatic cells by nuclear transfer: how to improve cloning efficiency? J Reprod Dev 53:13 26 Wakisaka Saito N, Kohda T, Inoue K et al (2006) Chorioallantoic placenta defects in cloned mice. Biochem Biophys Res Commun 349(1):106 114 Wells D (2003) Cloning in livestock agriculture. Reprod Suppl 61:131 150 Wilmut I, Wilmut I, Schnieke AE et al (1997) Viable offspring derived from fetal and adult mammalian cells. Nature 385:810 813 Wuensch A, Habermann FA, Kurosaka S et al (2007) Quantitative monitoring of pluripotency gene activation after somatic cloning in cattle. Biol Reprod 76(6):983 991 Xing X, Magnani L, Lee K et al (2009) Gene expression and development of early pig embryos produced by serial nuclear transfer. Mol Reprod Dev 76:555 563 Yang X, Tian XC, Kubota C et al (2007) Risk assessment of meat and milk from cloned animals. Nat Biotechnol 25:77 83 Ying QL, Wray J, Nichols J et al (2008) The ground state of embryonic stem cell self renewal. Nature 453:519 523 Zhou QY, Huang JN, Xiong YZ et al (2007) Imprinting analyses of the porcine GATM and PEG10 genes in placentas on days 75 and 90 of gestation. Genes Genet Syst 82:265 269 Zhou W, Xiang T, Walker S et al (2008) Global gene expression analysis of bovine blastocysts produced by multiple methods. Polejaeva I. Mol Reprod Dev 75:744 758

Part II

Fundamental Aspects of Genome and Epigenome Reprogramming During Gametogenesis

Chapter 5

Epigenetic Reprogramming Associated with Primordial Germ Cell Development Yoshiyuki Seki

Abstract Epigenetic modifications, including DNA methylation and histone modifications, confer variety and stability of gene expression, which ensure the generation and maintenance of numerous distinctive cell types during mammalian development and in adults. Recent studies have suggested that genome-wide changes of epigenetic modifications, referred to as “epigenetic reprogramming,” occur during the development of primordial germ cells (PGCs) in mice. This reprogramming might be critical for the reestablishment of potential totipotency in this lineage. However, the molecular mechanisms underlying these events are just beginning to emerge. We briefly summarize here our current knowledge on the development of PGCs and their epigenetic reprogramming, which may have general implications for the reprogramming of somatic cell nuclei of any kind. Keywords DNA methylation Epigenetic reprogramming Histone methylation Imprinted genes Primordial germ cells

5.1

Introduction

All cells in multicellular organisms contain identical genetic information; yet, a variety of somatic cell types are generated with different gene expression programs. These programs are usually fixed in a stable cellular function through epigenetic mechanisms, including DNA methylation (Bird 2002), histone tail modifications (Peters and Schubeler 2005), and specific nuclear architecture (Misteli 2007). Thus, each somatic cell type acquires a specific and stable epigenetic

Y. Seki Laboratory of Molecular Epigenetics for Germline Development, School of Bioscience, Kwansei Gakuin University, 2 1 Gakuen, Sanda, Hyogo 669 1337, Japan e mail: [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 5, # Springer Verlag Berlin Heidelberg 2011

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signature, referred to as “cellular memory,” which is often mitotically heritable. In contrast, the genome of the germ cell lineage, which is the sole pathway to the next generation, must be maintained in an epigenetically reprogrammable state for the new generation to continue to be created. There are essentially two modes, which are known as “preformation” and “epigenesis,” respectively, for the specification of the germ cell fate during the development of multicellular organisms (McLaren 2003). In preformation, which is seen in model organisms such as Caenorhabditis elegans and Drosophila melanogaster, localized determinants in oocytes, often referred to as the germ plasms, specify the new germline cells and segregate them from the somatic lineages at the outset of development. The germplasm set aside in precursor blastomeres for the germ cell lineage provides repressive mechanisms, including transcriptional and translational repression, which prevent the activation of the genetic programs for somatic differentiation from occurring in the germline blastomeres (Seydoux and Braun 2006). Specified germ cells in these animals in turn establish a chromatin-based silencing [e.g., low levels of histone H3 Lysine-4 di-methylation (H3K4me2)], which ensures subsequent global transcriptional quiescence (Blackwell 2004). By contrast, in the epigenesis, which is seen in mammals including mice, a potentially equivalent population of pluripotent cells at a relatively late stage of development is induced to form either germ cells or somatic mesoderm in response to signals from adjacent tissues (Lawson et al. 1999; Ohinata et al. 2005). This implies that cells recruited for the germline may have to undergo “epigenetic reprogramming” from a somatic to a potentially totipotent germline phenotype. Recent studies have demonstrated that this indeed seems to be the case (Seki et al. 2005, 2007). Here, we summarize briefly what is currently known about the epigenetic reprogramming in the PGCs of mouse embryos.

5.1.1

From Fertilization to the Specification of the New Germ Cell Lineage

The developmental program in the mouse is initiated upon the fusion of a highly specialized female gamete, the oocyte, with a male counterpart, the sperm. One of the first events to occur in the fertilized oocyte is the reorganization of the paternal genome, which includes replacement of the protamines with maternally deposited histones and subsequent apparent genome-wide DNA demethylation (Mayer et al. 2000) (Fig. 5.1). These events take place prior to the S phase. It is notable that genome-wide DNA demethylation from the paternal genome is a conserved phenomenon across several mammalian species, which suggests the functional significance of this event (Dean et al. 2001). Apparently, this resetting of the epigenetic modifications is a well-programmed process in the initiation of embryonic development. Interestingly, it was reported that in the absence of Pgc7/stella, a small shuttling protein between the nucleus and the cytoplasm, genome-wide DNA

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Extraembryonic ectoderm (ExE) Paternal PN Active demethylation Trophectoderm (TE)

Inner cell mass (ICM)

Primordial germ cells (PGCs) Mitotic arrest

Epiblast Maternal genome Primitive endoderm (PE) Passive demethylation De novo DNA methylation X chromosome Erasure of X chromosome reactivation parental imprints inactivation

Developmental Stage Fertilization

E0.5

E3.5 Preimplantation

E6.5

Postimplantation

Paternal imprints (E15.5 17.5)

Spermatogenesis Genital ridges Meiotic entry X chromosome reactivation

E12.5

Maternal imprints (Growing Oocyte)

Oogenesis

Migrating PGCs PostmigratingPGCs

DNA methylation Genome-wide (Male) Genome-wide (Female) Imprinted genes LINE-1 (Male) LINE-1 (Female) IAP

Fig. 5.1 Dynamics of DNA methylation changes throughout germ cell development. The changes in the DNA methylation levels of the indicated categories during germ cell development are shown by the graded color codes

demethylation occurs not only from the paternal genome but also from the maternal allele (Nakamura et al. 2007), indicating that Pgc7/stella functions to prevent DNA demethylation from the maternal allele in normal development. Recent knockdown experiments of elongation factors, which contain radical SAM domain, clearly showed that the loss of elongation factors in mouse zygote impairs paternal DNA demethylation. However, the biological function of paternal DNA demethylation in mouse zygote is largely unknown (Okada et al. 2010). The resultant totipotent zygote, with key maternal factors, replicates haploid parental genomes and begins to undergo cleavage divisions, with the major zygotic transcription starting at the late 2-cell stage (Morgan et al. 2005) (Fig. 5.1). The first cell fate specification seems to occur in the 8- to 16-cell stage, when the cells located outside start to show signs of differentiation towards trophectoderm (TE) cells and the cells located inside remain undifferentiated and maintain pluripotency. Although the precise mechanisms regulating this process are unknown, reciprocal inhibition of the key lineage determinants Oct4 and Cdx2 seems to play a role in the segregation of these two lineages (Niwa et al. 2005; Morgan et al. 2005; Strumpf et al. 2005). With the formation of the blastocoel cavity, which originates at multiple locations underneath the outside cells at around the 16-cell stage and coalesces to form a single large cavity at around the 32-cell stage (refs), the developing embryos are now called blastocysts. At around embryonic day (E) 3.5, the blastocysts consist of two clearly discernable cell types, the TE and the inner cell mass (ICM) cells. More precisely, the TE can be classified into the polar TE (pTE), which contacts and covers the ICM, and the mural TE (mTE), which delineates the blastocoel cavity. Thus, blastocysts bear a polarized structure with the ICM on one side and the blastocoel cavity on the other. The ICM is the source of all the cells in the adult body, including germ cells, and is the pluripotent cell population.

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Following substantial genome-wide DNA demethylation in the preimplantation stages, embryonic DNA methylation patterns start to be imposed through lineagespecific de novo methylation that begins in the ICM of a blastocyst (Li 2002) (Fig. 5.1). Genome-wide DNA methylation levels increase rapidly, an increase that is mediated by the de novo DNA methyltransferases, Dnmt3a and Dnmt3b (Okano et al. 1999). Another repressive modification, histone H3 Lysine-9 di-methylation (H3K9me2), increases after the two-cell stage, which thus precedes de novo DNA methylation (Liu et al. 2004). The first lineage that differentiates from the ICM is the primitive endoderm (PE), which delineates the inner surface of the ICM at E4.5. The PE layer eventually grows to cover the mTE as well and is then called the parietal endoderm. The undifferentiated cells in the ICM are now called the primitive ectoderm, which maintains pluripotency. Upon implantation, pTE cells proliferate and grow into a thick column of extraembryonic ectoderm (ExE) cells. The primitive ectoderm cells, partly through apoptosis, form a cup-shaped epithelial sheet, which is called the epiblast. It is from these epiblast cells that all the somatic cells as well as the germ cells in the adult body arise. The initial patterning of embryogenesis, including anterior posterior polarity formation and the gastrulation that forms mesodermal cells, is mediated through signaling molecules from the ExE and VE that cover the epiblast. The new generation of the germ cell lineage is recruited from the proximal epiblast cells as Blimp1 (a potent transcriptional repressor with a PR domain and Zn fingers)-positive cells at E6.25 (Ohinata et al. 2005). The primordial germ cells (PGCs), the first population of the germ cell lineage and the source for both the oocyte and the sperm, form a tight cluster of alkaline-phosphatase positive cells in the extraembryonic mesoderm at E7.25 (Ginsburg et al. 1990) and migrate through the developing hindgut endoderm (Anderson et al. 2000), eventually colonizing in the genital ridges after E9.5 (Molyneaux et al. 2001), where they begin to differentiate into functional gametes through highly complicated developmental pathways. The fully differentiated oocyte and sperm can now reinitiate and recapitulate all these processes.

5.2

A Brief Overview of the Epigenetic Reprogramming During PGC Development

Blimp1 is induced in the most proximal epiblast cells at E6.25, and these cells increase gradually in number and contribute exclusively to PGCs with alkaline phosphatase activity and stella expression at E7.25 (Ohinata et al. 2005). Genomewide chromatin modifications including H3K4me2, H3K4me3, H3K9Ac, H3K9me1, H3K9me2, H3K9me3, H3K27me2, and H3K27 me3 in the Blimp1positive lineage-restricted PGC precursors (~E6.75) are indistinguishable from those of their somatic mesodermal neighbors, some of which should share common precursors with the germ cell lineage (Seki et al. 2007).

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Blimp1, Prdm14 Dppa3 Dnmt3a/b, Np95 5meCpG GLP H3K9me2 H3K27me3 H4R3me2s G2 arrest ‘Reprogramming in a progressive, cell-by-cell manner’

‘Pre-reprogramming’ Dnmt3a/b, Np95 (++), 5meCpG (++) Imprinted genes (++) GLP/H3K9me1/2 (++) H3K27me3/H4R3me2s (+)

Dnmt3a/b, Np95 (-), 5meCpG (+) Imprinted genes (++) GLP/H3K9me1/2 (-) H3K27me3/H4R3me2s (+)

Specification E6.5

Migration E7.5

E8.5

Proliferation and subsequent further ‘reprogramming’ Including imprinted erasure in the genital ridges Dnmt3a/b, Np95 (-), 5meCpG (+) Imprinted genes (+) GLP/H3K9me1/2 (-) H3K27me3/H4R3me2s (++)

Dnmt3a/b, Np95 (-), 5meCpG (-) Imprinted genes (-) GLP/H3K9me1/2 (-) H3K27me3/H4R3me2s (++)

Arrival in the genital ridges E9.5

E10.5

E11.5 E12.5 Developmental Stage

Fig. 5.2 Epigenetic reprogramming in migrating PGCs. A summary of the epigenetic reprogram ming in migrating PGCs described in the text is shown

Subsequently, from around E8.0 onwards, migrating PGCs specifically show the genome-wide reduction of the two major repressive modifications, DNA methylation and H3K9me2, whereas from around E8.5 onwards, they acquire significantly higher levels of H3K27me3 and symmetrical dimethylation of arginine-3 of H4 (H4R3me2s) (Ancelin et al. 2006; Seki et al. 2005) (Fig. 5.2). These global changes of histone modifications in migrating PGCs occur in a progressive, cell-by-cell manner, presumably depending on their developmental maturation. Prior to or concomitant with the onset of H3K9me2 demethylation, PGCs enter the G2 arrest of the cell cycle, which apparently persists until they acquire high H3K27me3 levels (Seki et al. 2007). Interestingly, PGCs exhibit global repression of RNA polymerase II-dependent transcription, which begins after the onset of H3K9me2 reduction in the G2 phase and tapers off after the acquisition of high-level H3K27me3 (Seki et al. 2007). PGCs resume rapid proliferation and migrate through the hindgut epithelium and mesentery after around E9.25, and they start migrating into the developing gonads after E9.5. Soon after PGCs enter the genital ridges, they undergo further DNA demethylation, including the erasure of parental imprints between E9.5 and E12.5 (Hajkova et al. 2002). Repetitive elements, such as IAP (Intracisternal A Particle elements, classified as LTRs, ~1,000 copies per mouse genome) and LINE1 (the autonomous long interspersed nucleotide element-like element, ~10,000 100,000 copies per genome), are highly methylated in migrating PGCs and somatic cells. Postmigrating PGCs demethylate CpG methylation on LINE1 elements, while most of the CpG cites on the IAP elements remain highly methylated (Hajkova et al. 2002; Lane et al. 2003). The female PGCs, which initially undergo random X-chromosome inactivation as in the somatic cells, reactivate the inactive X, which most likely reflect the global epigenetic reprogramming process in gonadal PGCs (Tam et al. 1994).

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The Cell Cycle State of PGCs

The precise manner of the recruitment of PGCs and their subsequent increase in number would have critical implications in the mechanism of epigenetic reprogramming in PGCs. Classically, it has been considered that once PGCs are established and have reached approximately 40 in number, they proliferate constantly with a doubling time of about 16 h (Tam et al. 1994). These assumptions were based on the counting of the number of PGCs by alkaline phosphatase staining followed by histological examination. However, it has been indicated that the AP staining often underestimates the number of PGCs, especially in early developmental stages (Lawson et al. 1999). Our study involving Blimp1-mEGFP and stella-EGFP transgenic embryos, which enable accurate counting of the PGC number from the very beginning of the germ cell lineage, has shown that PGCs do not increase their number constantly (Seki et al. 2007); from around E7.75 to E9.25, coincident with their migration in the developing hindgut endoderm, their increase in number is relatively slow compared to that after E9.5. Consistently, the BrdU incorporation experiment showed that PGCs at these stages incorporate BrdU only at low ratios, which indicates that their entry into the S-phase is less frequent. Direct calculation of the DNA contents of PGCs by flow cytometry has shown that a majority of PGCs (approximately 60%) from E7.75 to E8.75 are in the G2-phase of the cell cycle. After E9.75, PGCs exhibited a cell cycle distribution with a clear G1 peak, a broad S-phase, and a less prominent G2 peak, which is a similar cell cycle distribution to that of ES cells and is indicative of a rapidly cycling state. This study therefore indicates that a majority of PGCs migrating in the hindgut endoderm are arrested at the G2-phase of the cell cycle. The precise mechanism of the increase of the PGC number at the earliest stage is still unknown. Initially, four to eight Blimp1-positive cells, which are in the single layer of the most proximal epiblast cells in direct contact with ExE, are observed at around E6.25. These cells subsequently relocate to the posterior-proximal end of the embryo to form a tight cluster of approximately 20 cells at ~E6.75, and they further increase their number to about 40 by ~E7.25, when most of them show typical AP activity and some of them already start their migration (Ohinata et al. 2005). Since the cell cycle distribution of E7.25 Blimp1-positive cells is similar to that of PGCs after E9.75 (Seki et al. 2007), it is clear that the initial Blimp1-positive cells are proliferating. A critical issue therefore is how long the Blimp1-positive germ cell precursors continue to be recruited from the epiblast cells, the clarification of which will provide a comprehensive view of the increase of the PGC population. It is of note that, in D. melanogaster, pole cells (lineage-restricted PGCs) also become mitotically quiescencent at the G2 phase during their migrating periods (Su et al. 1998). The maternal RNA-binding protein Nanos, together with Pumilio, acts to inhibit the pole-cell division, by repressing the translation of the Cyclin B1 mRNA (Asaoka-Taguchi et al. 1999). The mouse Nanos protein, Nanos3, starts to

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be expressed in PGC as early as E7.25 (Yabuta et al. 2006) and is required for germ cell development; the homozygously null mutants for Nanos3 show abnormality in PGC development as early as E9.5 (Tsuda et al. 2003). It is of interest to examine if Nanos3 mutants show abnormal cell cycle states.

5.4

Genome-Wide DNA Demethylation in Migrating PGCs

Our immunofluorescence analysis has shown that PGCs show a significant reduction of genome-wide DNA methylation at around E8.0 (Seki et al. 2005). The genome-wide DNA methylation level declines further when PGCs enter the gonads. It is critical to determine the precise kinetics of this DNA demethylation process. Three DNA methyltransferases (Dnmts) sharing a conserved DNMT domain have been identified in mammals (Bird 2002). Dnmt1 maintains DNA methylation during replication by copying the preexisting DNA methylation of the old DNA strand onto the newly synthesized strand (Leonhardt et al. 1992). Dnmt3a and Dnmt3b are mainly responsible for de novo methylation by targeting the unmethylated CpG site (Okano et al. 1998). They also cooperate with Dnmt1 to maintain methylation patterns during cell division (Liang et al. 2002). It has been shown that Dnmt3a/b and Uhrf1 (Np95), which is essential for the recruitment of Dnmt1 into the replication foci, are actively repressed in PGCs at E6.75 to E7.5. (Seki et al. 2005; Yabuta et al. 2006), suggesting that the loss of all the Dnmts activities might contribute to the genome-wide DNA demethylation in PGCs. Active mechanisms for DNA demethylation might also exist in PGCs. Although methyltransferases that impose DNA methylation are well characterized, the molecular basis underlying active DNA demethylation is still elusive and controversial (Bhattacharya et al. 1999; Ng et al. 1999). In plants, it has been shown that DEMETER, a DNA glycosylase, functions as a DNA demethylase by excising 5-methylcytosine with subsequent DNA repair by means of a base-excision repair pathway (Gehring et al. 2006). Recently, the RNA editing enzymes Aid and Apobec1 and the growth arrest and DNA-damage-inducible protein Gadd45a have been shown in vitro to have DNA demethylation activity also through the base-excision repair pathway (Morgan et al. 2004; Barreto et al. 2007). Interestingly, Aid and Apobec1 are located in a cluster of genes with stella, growth differentiation factor 3 (Gdf3), and Nanog (Morgan et al. 2004). It is currently unknown whether Aid, Apobec1, and Gadd45 are responsible for the global DNA demethylation in migrating PGCs. A recent report showed that genome-wide erasure of DNA methylation was impaired in Aid deficient PGCs at E13.5. However, the impairment of genome-wide demethylation in Aid deficient PGCs was not remarkable, suggesting that another major genetic pathway is sure to induce genome-wide demethylation in PGCs (Popp et al. 2010).

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Genome-Wide Demethylation of H3K9me2 in Migrating PGCs

We have shown that PGCs undergo genome-wide demethylation of H3K9me2 from around E7.75 onwards. The kinetics study involving the entire PGC population in each embryo has indicated that demethylation of H3K9me2 occurs in a progressive, cell-by-cell manner, presumably depending on the developmental maturation of the PGCs. Thus, the demethylation levels of H3K9me2 in each PGC and the ratio of PGCs showing reduced H3K9me2 increase progressively from E7.75, and at E8.75, nearly all the PGCs show low H3K9me2 levels. Apparently, the levels of H3K9me2 become at least less than half of the original level during this period. As described above, a majority of PGCs are arrested at the G2-phase of the cell cycle when this demethylation is occurring. It would therefore be reasonable to consider that the preexisting H3K9me2 marks are removed by some active mechanisms. As in the case for DNA demethylation, the target sequences for H3K9me2 demethylation is currently unknown. Until recently, the presence of enzymes that demethylate methylated histones had been elusive (Bannister et al. 2002). However, three distinct classes of demethylation enzymes, PADI4 (Cuthbert et al. 2004), LSD1 (Shi et al. 2004), and Jumonji C (JmjC) domain-containing proteins (Tsukada et al. 2006), have now been identified. Among these, the JmjC domain-containing proteins constitute a large family of ~30 proteins throughout the genome and can demethylate all the methylation states (mono-, di-, and tri-) of H3K4, H3K9, and H3K36 (Klose et al. 2006). Single-cell gene expression analysis of the JmjC domain-protein family has shown that Jmdm2a (Jmjd1a), which encodes an H3K9me1 and 2 demethylase (Yamane et al. 2006), is indeed expressed in migrating PGCs. However, its expression is similarly seen in somatic neighbors (Seki et al. 2007). This suggests that PGCs may have additional mechanisms that allow genome-wide removal of H3K9me2. Our studies have found that PGCs actively repress GLP, a critical histone methyltransferase for H3K9me1 and 2, at around E7.5 (Seki et al. 2007). GLP was shown to form a complex with G9a and to be essential for the stability and the histone methyltransferase activity of this enzyme complex (Tachibana et al. 2005). These findings suggest that the repression of GLP may lead to the loss of H3K9 methyltransferase activity in PGCs, which triggers the genome-wide reduction of H3K9me2, either through a turnover of methyl groups or a replacement of the entire H3 molecule with unmodified H3 molecule. Consistently, a report involving a mass spectrometry analysis of H3K9 methyl groups differentially pulse-labeled by a heavy isotope demonstrated that H3K9me2 turns over without replication (Fodor et al. 2006). Furthermore, it was also reported that chromatinassociated and nonchromatin-associated histones are continually exchanged in Xenopus oocytes, and that the maintenance of H3K9me2 at a specific site requires the continual presence of an H3K9 histone methyltransferase (Stewart et al. 2006). Thus, repression of an essential enzyme may shift the equilibrium between methylation and demethylation toward demethylation, leading to the erasure of H3K9me2 in migrating PGCs.

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Global Transcriptional Repression in Migrating PGCs

The question arises as to what the transcriptional state of migrating PGCs would be when genome-wide reduction of two major repressive modifications, DNA methylation and H3K9me2, are occurring. Our study has demonstrated that coincident with this epigenetic reprogramming, PGCs are devoid of RNA polymerase II (RNAP II)-dependent global transcription (Seki et al. 2007). RNAP II-dependent transcription is tightly regulated by the phosphorylation on the C terminal domain (CTD) repeat sequence (YSPTSPS) (Phatnani and Greenleaf 2006). The transition from the transcription initiation to the promoter clearance is accompanied by the CTD phosphorylation predominantly on the Ser-5. For the transcription elongation, the phosphorylation site changes into the Ser-2. In migrating PGCs, phosphorylations on both Ser-5 (detected by H14 antibody) and Ser-2 (detected by H5 antibody) (Patturajan et al. 1998) become transiently absent. Precise time-course analysis showed that Blimp1-positive PGC precursors and PGCs until E7.75 were positive for Ser-2 phosphorylation, and from around E8.0 onwards, slightly after the onset of H3K9me2 demethylation, PGCs turned off Ser-2 phosphorylation progressively, in a cell-by-cell manner. At E8.5-E8.75, most PGCs lose this modification. However, at E9.25, some PGCs apparently regain this modification, and by E10.5, almost all of the PGCs become positive for this phosphorylation. It is therefore the case that during the loss of DNA methylation and the progressive removal of H3K9me2, PGCs are mainly in the G2-phase of the cell cycle and essentially stop RNAP II-dependent transcription [RNAP I-dependent transcription remains apparently intact (Seki et al. 2007)]. It is well-established that in C. elegans and D. melanogaster, RNAP II-dependent transcription is shut off at the outset of germ cell specification, which prevents the activation of somatic gene expression in their PGCs (Seydoux and Dunn 1997). This transcriptional repression is essential for the successful establishment of the germ cell lineage in these animals. In C. elegans, this transcriptional repression depends on PIE-1, a maternally inherited germplasm component (Seydoux et al. 1996). PIE-1 is a Zn finger protein with RNA binding activity and is reported to prevent CTD phosphorylation by competing with the CTD kinase, Cdk9 (Zhang et al. 2003). In D. melanogaster, this repression depends on the polar granule component (pgc), which also prevents CTD phosphorylation with an as-yet-unknown mechanism (Martinho et al. 2004). In mice, Blimp1-positive PGC precursors and PGCs are transcriptionally active with CTD phosphorylation by E7.75, which is consistent with the activation of many specific genes associated with germ cell specification (Yabuta et al. 2006). Nonetheless, PGCs exhibit global repression of RNAP II-dependent transcription, specifically during their migration period, which we assume would not be for the purpose of repressing the somatic program in PGCs, but presumably for an efficient execution of the epigenetic reprogramming. What is the mechanism for the repression of the CTD phosphorylation in PGCs in mice? An autolog for PIE-1 or pgc is not present in the mammalian genome.

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In mammals, it is known that the CTD phosphorylation mediated by PTEF B (Cdk9 and Cyclin T complex) can be inhibited by a protein called Hexim1 (Yik et al. 2003). In addition, it was reported that B2 RNA, one of the short interspersed transposable elements (SINEs), is associated with the initiation complex of RNAP II and represses global transcription by inhibiting transcriptional initiation (Espinoza et al. 2004). It would be interesting to examine whether one of these mechanisms operates in migrating PGCs, which may lead to the clarification of the functional significance of the global repression of RNAP II in PGCs.

5.7

Genome-Wide Upregulation of H3K27me3 in Migrating PGCs

Following the genome-wide loss of DNA methylation and H3K9me2, genomewide H3K27me3, another repressive modification mediated by the polycomb repressive complex 2 (PRC2), becomes upregulated in migrating PGCs from around E8.25 onwards (Seki et al. 2005, 2007). Both the percentage of PGCs with high levels of H3K27me3 and the degree of H3K27me3 upregulation in individual PGCs increase as the development proceeds, indicating that, as in the case for the demethylation of H3K9me2, H3K27me3 upregulation proceeds progressively, in a cell-by-cell manner. We are discussing the molecular mechanism underlying genome-wide upregulation of H3K27me3 in migrating PGCs. PRC2 consists of three core components, Ezh2, Eed, and Suz12. Single-cell gene expression analysis has shown that genes for these three core components are expressed at similar levels both in the PGCs and their somatic neighbors by at least E7.75 (Yabuta et al. 2006). Considering the fact that PGCs globally shut off RNAP II-dependent transcription after around E8.0, the upregulation of genome-wide H3K27me3 in PGCs after E8.25 may not depend on the de novo transcription of some specific factors. It was reported that in Suv39hdeficient cells, which essentially lack H3K9me3 at the pericentromeric heterochromatin, H3K27me3 is ectopically upregulated in this location; this suggests that there is “crosstalk” between H3K27 methylation and H3K9 methylation, which rescues the deregulated chromatin structure (Peters et al. 2003). A similar system may operate in migrating PGCs in which hypomethylated regions for H3K9me2 might be “sensed and rescued” by H3K27me3.

5.8

Arginine Methylation Mediated by Blimp1/Prmt5 Complex in Migrating PGCs

Arginine methylation is catalyzed by the PRMT class of histone methyltransferases and contributes to both active and repressive effects on chromatin function (Klose and Zhang 2007). A recent study has shown that Blimp1 forms a complex with

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Prmt5, a protein arginine (R) methyltransferase, which mediates symmetrical dimethylation of H2A and H4R3 (Ancelin et al. 2006). Blimp1 and Prmt5 are coexpressed in PGCs, and PGCs show high levels of H4R3me2 after around E8.5. A chromatin immunoprecipitation (ChIP) experiment identified putative targets of the Blimp1-Prmt5 complex, including an RNA helicase, Dhx38. The study further showed that the onset of Dhx38 expression coincides with the translocation of Blimp1-Prmt5 complex from the nucleus to the cytoplasm of PGCs at around E11.5. Furthermore, ectopic expression of Blimp1 in embryonic carcinoma (EC) cells leads to the repression of Dhx38. Thus, the Blimp1-Prmt5 complex may have essential roles in maintaining the identity of migrating PGCs. Importantly, a homolog of Prmt5 in D. melanogaster, Dart5, was shown to be critical for germ cell specification in D. melanogaster (Anne et al. 2007), which suggests that this protein has an evolutionally conserved importance in PGCs.

5.9

Potential Significance of Epigenetic Reprogramming in Migrating PGCs

What could be the potential biological significance of epigenetic reprogramming in migrating PGCs? Essentially, migrating PGCs convert their genome-wide repressive modifications from DNA methylation and H3K9me2 to H3K27me3. It has been reported that DNA methylation and H3K9me2 are critical for the stable maintenance of the repressed genes. For example, a time-course study on the DNA methylation and histone modifications on the Oct3/4 locus showed that these modifications are imposed on the locus after the gene expression is tightly shut off (Feldman et al. 2006). G9a-mediated H3K9me2, which recruits HP1 and DNA methylation, was shown to be essential for the stable maintenance of the repressed state of this gene locus. Another study showed that a CpG-free transgene undergoes transcriptional silencing to an extent similar to that of another transgene with CpG sites that becomes methylated (Feng et al. 2006). However, only the transgene with CpG methylation was shown to be resistant to reactivation. On the other hand, pluripotent ES cells bear high levels of PRC2-mediated H3K27me3, which functions to repress developmentally regulated lineage-specific genes (Bernstein et al. 2006; Lee et al. 2006). Many of these developmentally regulated genes bear bi-valent modifications with H3K27me3 and H3K4me3, which ensure that these repressed genes can be expressed quickly upon the induction of differentiation. Taken together, DNA methylation and H3K9me2 are considered to play critical roles in the stable maintenance of the repressed state of unused genes during cell fate specification, whereas H3K27me3 may be a more plastic repression of the lineage-specific genes in pluripotent cells. As PGCs bear significant levels of genome-wide H3K4me3, specific upregulation of H3K27me3 in PGCs may contribute to the creation of an ES cell-like genome organization. Indeed, it is possible to derive pluripotent stem cells from PGCs (EG cells) from E8.5 to E12.5, if they

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are cultured with basic fibroblast growth factor (bFGF), leukemia inhibitory factor (LIF), and stem cell factor (SCF) (Matsui et al. 1992). Epigenetic reprogramming and specific expression/reacquisition of the key pluripotency genes Oct3/4, Sox2, and Nanog in PGCs may thus be critical for their potential pluripotency (Yabuta et al. 2006). It might be the case that genes such as Blimp1 are important to prevent PGCs from overtly reverting to a pluripotent state in vivo. It is an essential future challenge to determine the target sequences from which DNA methylation and H3K9me2 are removed and upon which H3K27me3 is imposed.

5.10

Epigenetic Reprogramming of Imprinted Genes in Postmigrating PGCs

Genomic imprinting refers to a parent-origin-specific gene regulation present only in mammals (Delaval and Feil 2004). Imprinted genes are expressed exclusively either from maternal or paternal alleles and play essential roles in development, growth, and behavior. Currently, nearly 100 genes are known to be imprinted throughout the genome. Imprinting is mediated by epigenetic modifications including DNA methylation and histone modifications imposed on either maternal or paternal alleles. Imprinting marks of the parental germ cells generally persist in somatic lineages, whereas to maintain the correct imprinting cycle, newly formed germ cells need to erase the imprints of the previous generation and establish new ones depending on the sex of the new generation (Surani 2001). To determine when the erasure of the imprinting occurs in PGCs, several studies examined the DNA methylation states of imprinted genes in migrating and postmigrating PGCs from E9.5 to E12.5 (Lee et al. 2002; Hajkova et al. 2002; Sato et al. 2003; Yamazaki et al. 2005). Allelespecific DNA methylations in nearly all the imprinted genes were maintained in migrating PGCs until about E9.5, indicating that imprinting marks are resistant to the global DNA demethylation in migrating PGCs. Indeed, it has been shown that the erasure of imprinting commences in a part of the migrating PGCs at E9.5, progresses gradually afterward, and completes in gonadal PGCs at E12.5. The mechanism of imprint erasure is currently unknown. A study on the kinetics of the erasure of imprinting marks on several imprinted genes suggested that the erasure is an active process involving the direct removal of DNA methylation (Hajkova et al. 2002). A recent study showed that in Aid-deficient PGCs, some imprinted locus, H19 and Lit1, have partial resistance against the demethylation process (Popp et al. 2010). On the other hand, one study showed that, in the Dnmt3a and 3b double knockout ES cells with high passage numbers, several imprinted regions undergo complete demethylation, suggesting that the absence of Dnmt3a and 3b may be a critical requirement for the erasure of imprints (Chen et al. 2003). In addition, in G9a-deficient ES cells, the H3K9me2 levels of the genomic region

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for the Prader Willi syndrome imprinting center become highly reduced, together with the concomitant loss of maternal-allele-specific DNA methylation, leading to the biallelic expression of the genes in the affected region (Xin et al. 2003). This suggests the importance of H3K9me2 levels for the stable maintenance of the imprinting. Together, these findings suggest that it would be reasonable to speculate that the repression of Dnmt3a, 3b, GLP, and G9a and the genome-wide DNA and H3K9me2 demethylation in migrating PGCs provide an essential genomic background, in which the erasure of imprinting and further reprogramming of the genome proceeds efficiently.

5.11

Modification of the Repetitive Elements in PGCs

Many recently completed genome projects have revealed that large parts of the mammalian genomes consist of the repetitive sequences derived from DNA and retrotransposons [LINEs (the autonomous long interspersed nucleotide elementlike element), SINEs (the LINE-dependent, short RNA-derived short interspersed nucleotide elements), and LTRs (retrovirus-like elements with long terminal repeats)]. In humans and mice, repetitive elements cover as much as ~48 and ~44% of the genome, respectively, whereas only 1.5 and 4% of the genome encode protein-coding genes, respectively (noncoding regions comprise 50 and 52% of the genome, respectively) (Martens et al. 2005). Several studies have shown that the repetitive elements bear unique epigenetic modifications. Large-scale analysis of the genome-wide DNA methylation pattern in human somatic cells indicated that about 30% of the methylated cytosine on the whole genome is distributed on SINEs (Rollins et al. 2006). Moreover, an unbiased ChIP cloning approach of H3K9me2 in human somatic cells showed that 68% of independent clones contain SINEs (Kondo and Issa 2003), suggesting that the majority of DNA methylation and H3K9me2 targets are SINEs in the mammalian genome. Considering these findings, we can speculate that SINEs might be one of the major targets of epigenetic reprogramming in migrating PGCs. It is known that PGCs at E10.5, which is soon after their entry into the genital ridges, bear DNA methylation that is comparableto somatic cells on LINE1, IAP, and imprinted genes (Hajkova et al. 2002; Lane et al. 2003). PGCs thereafter remove methylation on LINE1 and imprinted genes, whereas the methylation of IAPs remains intact (Fig. 5.1). LINE1 is enriched on the X chromosome as compared with autosomes, and it has been proposed that it may act as way station to spread gene silencing for X chromosome inactivation (Lyon 2003). Interestingly, statistical analysis of the repeats content in the regions surrounding monoallelic and biallelic genes revealed that SINEs are excluded from the imprinted region, whereas LINE1 is more abundant in imprinted genes in human and mouse genomes (Allen et al. 2003; Greally 2002; Ke et al. 2002). These findings suggest that repeat

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elements-based epigenetic reprogramming might be involved in the erasure of parental imprints and X chromosome reactivation.

5.12

Modification of the Specific Loci in PGCs

A recent study has explored the DNA methylation states of the 16,000 promoters (between 700 bp and þ200 bp relative to the transcription start site) in human primary somatic and germline cells (sperm) (Weber et al. 2007). This study classified the promoters into three distinct entities, ones with high CpG frequency (HCP), with intermediate CpG frequency (ICP), and with low CpG frequency (LCP). It has been shown that genes bearing LCPs are mostly highly methylated irrespective of their expression states, whereas genes bearing HCP are mostly (~95%) hypomethylated, even when they are not expressed. Interestingly, this study showed that of the rarely hypermethylated HCPs in somatic cells, 17% were genes showing a testis-specific expression, including DAZL, SPO11, SOX30, BRDT, ALF, TPTE, and REC8, or testis-specific histone variants HIST1H2BA and HIST1H1T. These genes were not methylated in sperm, suggesting that these methylations are established after fertilization during somatic development, presumably by the de novo DNA methyltransferases, Dnmt3a and Dnmt3b. Somatic cells thus seem to show a systematic methylation of promoters for germline-specific genes, including strong CpG islands that are otherwise protected from DNA methylation. Consistently, another report showed that in Dnmt1-deficient embryos, germline-specific genes such as Sycp3, Mvh, and Dazl, whose promoters are methylated in somatic cells and in migrating PGCs but become unmethylated in gonadal PGCs, showed premature and ectopic expression (Maatouk et al. 2006). These findings thus provide evidence that one of the important functions of DNA methylation in mammals may be the preclusion of deleterious activation of germline-specific genes (many of which are involved in meiosis) in somatic cells. Accordingly, E2F6 has been shown to be necessary to silence several germlinespecific genes in somatic cells, and DNA methylation and H3K9me2 on the promoters of these genes becomes hypomethylated in E2F6-deficient fibroblast cells (Pohlers et al. 2005; Storre et al. 2005). E2F6 is therefore one of the candidates for the targeting of Dnmt3a/3b to the germline-specific genes in somatic cells. A recent genetic study involving Dnmt3a/3b double mutant mice demonstrated that several genes in the Rhox clusters, which are predominantly expressed in the extraembryonic tissues (mainly in the placenta), are ectopically expressed in the embryo proper in these mutants, indicating that DNA methylation imposed during postimplantation development on these genes is necessary for the silencing of these genes in the embryo proper (Oda et al. 2006). Interestingly, Rhox5, 6, and 9 are specifically reexpressed in PGCs (Pitman et al. 1998; Takasaki et al. 2000), suggesting that PGCs may undergo epigenetic reprogramming to reactivate placenta-specific genes for the next generation.

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Conclusion

Somatic nuclear transplantation experiments have demonstrated that unfertilized mature oocytes can reprogram the differentiated somatic nuclei to totipotent states (Campbell et al. 1996). However, a majority of cloned embryos die at every stage of embryonic development with a variety of abnormalities. Of those that develop to [a] later gestational stage or to term, many have placental abnormalities (Wilmut 2002). Cloned adults also show many abnormalities including short life span and obesity (Wilmut 2002). It is likely that most of the defects of the cloned animals would be caused by inappropriate epigenetic reprogramming. This notion is supported by the fact that the offspring of cloned animals do not bear any of the abnormalities mentioned above (Tamashiro et al. 2002). These findings indicate that epigenetic reprogramming through germ cell development is essential for normal embryonic development and for adult life. Therefore, in reproductive and stem cell biology as well as in regeneration medicine, we consider it a critical challenge to analyze carefully the epigenetic reprogramming that occurs at each stage of germ cell development and to identify the underlying molecular mechanisms. For this purpose, one of the essential challenges will be to develop an experimental system that allows genome-wide analysis of DNA methylation and specific histone modification sequences from a small amount of biological samples.

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Chapter 6

Epigenetic Factors and Regulation of Meiotic Recombination in Mammals P. Barthe`s, J. Buard, and B. de Massy

Abstract During meiosis, recombination is induced to promote connections between homologous chromosomes, thereby ensuring faithful chromosome segregation. These connections result from reciprocal recombination events or crossovers. Absence or defects of recombination can lead to genome instability, chromosome nondisjunction, aneuploid gametes, or reduced fertility. Recombination events are therefore highly regulated through mechanisms that are still poorly understood. The role of epigenetic modifications in meiotic recombination is suggested by several observations, particularly by the differences between male and female recombination as well as by interindividual variations in crossover rates. In this review, we describe features of meiotic recombination that might involve epigenetic controls. We then present the epigenetic marks and modifiers known to occur or to play a role during meiotic prophase when recombination takes place. Direct evidence of a functional link between histone modifications and recombination is highlighted by a recent analysis of the distribution of recombination showing that the protein PRDM9 targets chromatin modifications to specific sites in the genome where initiation of meiotic recombination takes place. Keywords Aneuploidy Crossing over Double strand breaks Genetic map Hotspot H3K4Me3 Meiosis PRDM9 Recombination

Abbreviations Chr cM CO

Chromosome Centi morgan Crossover

P. Barthe`s, J. Buard, and B. de Massy (*) Institut de Ge´ne´tique Humaine, UPR1142, CNRS, 141 rue de la Cardonille, 34396 Montpellier cedex 5, France e mail: bernard.de [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 6, # Springer Verlag Berlin Heidelberg 2011

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DNMT DSB FISH HAT HDAC HDMT Histone modifications HJ HMT Hp1 IAP JMJD1a KO LD LINE MAEL MSCI Msuc NCO PiRNA SC SEI SNP TSA

6.1

DNA (cytosine-5)-methyltransferase DNA double-strand break Fluorescent in situ hybridization Histone acetyl-transferase Histone deacetylase Histone demethylase Example H3K4Me3 is histone H3 with tri-methylated lysine 4 Holliday junction Histone methyl-transferase Heterochromatin protein 1 Intracisternal A particle Jumonji domain-containing histone demethylase 1a Knock-out Linkage disequilibrium Long interspersed element Maelstrom Meiotic sex chromosome inactivation Meiotic silencing of unsynapsed chromatin Non crossover Piwi-interacting RNA Synaptonemal complex Single end invasion Single nucleotide polymorphism Trichostatin A

Introduction

Recombination between homologous chromosomes during meiosis is one of the several steps required for production of mature gametes and thus for fertility. In the absence of recombination, meiotic cells are either eliminated or they generate nonfunctional gametes, leading to sterility. The process of homologous recombination takes place during the prophase of the first meiotic division. It is a complex and long process with multiple reactions and alternative pathways. It involves events at the DNA, chromatin, and chromosome levels that require many interactions between proteins, DNA, and RNA, only few of which are well understood. Particularly, epigenetic information, in the large sense of this term, is known to play an important role, but how it actually influences meiotic recombination is known for only a few very specific steps of this process in mammals. In this review, we first outline the molecular mechanism of meiotic recombination with the main landmarks of this process at the DNA and chromosomal levels

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(see Sect. 6.2). Several of these events have been shown to be evolutionarily conserved from yeast to mammals, although obviously distinct features and regulations have also been identified by comparative studies in different organisms. The strategies used to monitor meiotic recombination in mammals, which are also used in many other multicellular eukaryotes, are then briefly summarized (see Sect. 6.3). We then present data that support the hypothesis that epigenetic modifications might influence meiotic recombination (see Sect. 6.4). In mammals, these data come from a relatively limited number of studies on the role of heterochromatin as well as on sex-specific and interindividual properties of meiotic recombination. In addition, several works on DNA methylation and chromatin structure or histone modifications highlight specific epigenetic features of the meiotic cell during prophase, suggesting a direct or indirect role for epigenetics in recombination (see Sect. 6.5). In the last part, we present recent analyses on the control of initiation of meiotic recombination that provide direct evidence for the role of a specific histone modification in initiation (see Sect. 6.6).

6.2

Outline of the Molecular Mechanism of Meiotic Recombination

The sexual reproductive cycle requires halving of the chromosome material during gamete formation. This is achieved by the specialized meiotic cycle, which entails one DNA replication phase followed by two divisions, thereby leading to the formation of haploid gametes from a diploid cell. During the first reductional division, homologous chromosomes segregate from each other, whereas during the second division, sister chromatids segregate. The reductional division is unique to meiotic cells and requires specialized mechanisms in order to connect homologous chromosomes and ensure their proper orientation at metaphase I and faithful separation by the segregation machinery. In most eukaryotes, these connections are established by a process called crossing over (CO), which defines the reciprocal recombination events between homologous chromosomes that can be visualized cytologically as chiasmata (Petronczki et al. 2003). The mechanical role of COs implies a regulation of their frequency, with at least one CO per chromosome. In addition, CO has evolutionary consequences as it increases genome diversity (Coop and Przeworski 2007). The molecular mechanism of CO has been analyzed in detail in Saccharomyces cerevisiae and Schizosaccharomyces pombe (Hunter 2007), and several of the main properties of the CO pathway are conserved in higher eukaryotes, including mammals (Handel and Schimenti 2010). Meiotic recombination is initiated by the formation of DNA double-strand breaks (DSBs), which are catalyzed by the evolutionarily conserved SPO11 protein (Bergerat et al. 1997; Keeney et al. 1997). DNA ends are then processed and invade a chromatid from the homologous chromosome through a process called strand invasion or strand exchange, which requires several proteins,

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gH2AX

SYCP3 RAD51

SYCP3 MSH4

SYCP3 MLH1

Fig. 6.1 DNA and cytological events during meiotic prophase. Meiotic recombination is initiated by DSBs. DNA ends are processed and start strand invasion of a homologous molecule (nascent strand invasion). This intermediate can be processed in two alternative ways to finally form a crossover (CO) or not (NCO). In the CO pathway, a single end invasion intermediate (SEI) is formed, matured into a double Holliday junction (dHJ), and then resolved into a CO. In the NCO pathway, after a short extension of the 30 end, the nascent strand invasion intermediate is displaced and the DSB repaired by strand annealing giving rise to gene conversion without CO. Some DSB repair events might occur through interaction with a sister chromatid (not shown). At the cytologi cal level, these steps can be visualized by immunodetection of phosphorylated H2AX (gH2AX) (DSB formation), RAD51/DMC1 (nascent strand invasion), MSH4 (strand invasion at zygotene), and MLH1 (CO). Chiasmata are visible at the diffuse or diplotene stage. From McDougall et al. (2005)

particularly RAD51 and DMC1. Strand invasion then follows two alternative DBS repair pathways: (1) The invading DNA is displaced and DSB repair results in a gene conversion event but without CO (noncrossover: NCO); (2) The invading DNA end is extended to give rise to a double Holliday junction intermediate (single Holliday junction intermediates may also form), which is then resolved to yield a CO. This process requires many proteins, among which MLH1 (Fig. 6.1). Many questions remain open about the mechanisms and regulation of meiotic recombination. Various approaches, using genetic, cytological, or molecular strategies, have shown that CO frequency is highly regulated and that COs are not randomly distributed. A few mechanisms of CO regulation have been identified so far, but they are still poorly understood: (1) A control called “crossover assurance” ensures the formation of at least one CO per chromosome. In some species, such as mammals, the rule seems to be one CO per chromosome arm (Pardo-Manuel De Villena and Sapienza 2001); (2) The presence of one CO decreases the probability to have an adjacent CO. This property called “CO positive interference” leads to

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a more even spacing between COs than if they were randomly distributed. However, a fraction of COs (5 10% in the mouse) seems to result from a noninterfering CO pathway, which is MLH1-independent; (3) CO occurrence appears not to decrease proportionally with DSB frequency. This regulation (called “CO homeostasis”) has been shown only in S. cerevisiae (Martini et al. 2006) and has been proposed to result from the choice of DSB repair pathway (i.e., NCO or CO); (4) COs are not randomly distributed in the genome, with some regions showing higher or lower density of COs. This property might result from the non random distribution of DSBs and/or from the regulation of the choice of DSB repair pathway (Baudat and de Massy 2007b). DSB formation, the molecular landmark of initiation of meiotic recombination, is at the center of many investigations. In S. cerevisiae and S. pombe, where DSBs can be mapped at high resolution by molecular approaches, they were found not to be randomly distributed. Specific chromatin features have been shown to be associated with DSBs and have been proposed to play a role in DSB formation (see Sect. 6.6). In mammals, DSBs are not directly detectable with the current available techniques, and localization of recombination events is based in most cases on mapping CO events by various approaches (see Sect. 6.3). In humans, these approaches have shown that most COs are clustered in narrow regions (1 2 kb), called hotspots (Jeffreys et al. 2001), which are predicted to be preferred initiation sites (Buard and de Massy 2007). On the basis of population diversity analysis, over 30,000 hotspots have been identified in the human genome, spaced on average every 50 100 Kb, often outside genes and with highly variable levels of activity (McVean et al. 2004; Myers et al. 2005). In mice, based on the analysis of a 25 Mb region on chromosome (chr.) 1 (Paigen et al. 2008) and of several other chromosomal regions (de Massy 2003), initiation of meiotic recombination also appears to be clustered in small intervals according to rules similar to those described in humans. DNA sequence, chromatin structure, and/or higher order levels of chromosome organization might play a role as potential determinants for hotspot localization.

6.3

How to Measure Recombination Activity in Mammals?

Several methods, commonly used in humans and mice to measure recombination activity, are briefly presented here (see also Arnheim et al. (2007) for review)

6.3.1

Pedigree Analysis

Pedigree analysis is possible when DNA samples from families have been collected to allow the genotyping of offspring, parents, and possibly grandparents. A recombination event taking place in a meiotic cell of a parent can be inferred when alleles at consecutive loci of a child arise from the two grandparents. The frequency d of

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recombination events between two markers is converted into the genetic distance o in Morgan units (M). Pedigree-based genetic maps provide estimates of male and female recombination rates per unit of physical distance over large intervals. Indeed, accurate measurement over small intervals is limited by the number of meioses that can be screened through this approach.

6.3.2

Sperm Typing

Recombinant events can be detected from DNA of pooled sperm using Single Nucleotide Polymorphisms (SNPs) as anchors for allele-specific PCR amplification of recombinant fragments. Recombination rates are accurate down to 0.0001 cM (centi Morgan), with a ~100 bp resolution (depending on the SNP density). One limitation of this approach is the size of the amplifiable target (<15 Kb), dictated by the current PCR technology. This strategy has also been used for CO analysis in oocytes in mice (Baudat and de Massy 2007a). Molecular analysis of markers from the genomic DNA from a single sperm, also called single-sperm typing, can also be used to detect COs. In this case, parallel independent PCR amplification of two markers is performed allowing CO estimates in the range of 0.1 cM with no limitation on the size of the interval tested.

6.3.3

MLH1 Foci

MLH1 (and MLH3) are required for CO formation (Guillon et al. 2005; Svetlanov et al. 2008) and localize to CO sites at the pachytene stage of meiotic prophase (Marcon and Moens 2003). Detection of these proteins in oocyte or spermatocyte spreads by immunocytochemistry alone or combined with fluorescent in situ hybridization (FISH) allows the quantification and localization of COs per nucleus and/ or per chromosome at microscopic resolution (several Mb). Since the formation of a small subset of COs (about 5% in mice and probably a similar percentage in humans) is MLH1/3-independent, the detection and quantification of MLH1/3 foci is thus expected to slightly underestimate the total number of COs.

6.3.4

Population Diversity Analysis

A combination of alleles at consecutive loci a haplotype is the result of mutations that create new polymorphisms and of recombination events that reshuffle them. A parameter can be calculated, the linkage disequilibrium (LD) coefficient D that measures the degree of association between alleles at consecutive loci. The normalized coefficient |D’| is the most widely used measure for describing LD

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along the genome, as it varies from 0 to 1 irrespective of the allele frequencies. Coalescent-based analyses that use a probability model can predict the most likely number of recombination events needed to produce a set of current haplotypes. Since they predict the number of COs that have occurred between markers on different haplotypic backgrounds over generations by simulations, coalescent approaches allow the estimate of historical recombination rates. The resolution is determined by the number of SNPs tested. Hotspots can be identified and are then called historical or LD-based hotspots. Although the general pattern of LD is shaped by recombination, one limitation of this approach is that other factors also modulate haplotype diversity, including evolutionary forces (Stumpf and McVean 2003; Fearnhead et al. 2004) and chance (Phillips et al. 2003). Another obvious limitation of population diversity analyses is that male and female recombination activities are not assessed separately.

6.4

Indirect Evidence that Epigenetic Modifications Influence Recombination Activity in Mammals

Many types of variations of meiotic recombination activity have been reported in humans relative to gender, individuals, or populations (see also Lynn et al. (2004) for review) and may in part result from epigenetic modifications.

6.4.1

Male Versus Female Recombination

Several studies have revealed clear differences between males and females in meiotic recombination. We present below a summary of these data with examples that highlight sex-related differences in frequency and localization of recombination events in both humans and mice.

6.4.1.1

Male and Female Genetic Maps

Genetic maps in humans have been refined over the last years by using an increasing number of markers and by taking advantage of HapMap, the nucleotide polymorphism database. The development of high throughput genotyping technologies has allowed the genotyping of a large number of individuals or DNA samples, thus improving the accuracy of recombination frequencies. Recent human genetic maps have been generated from pedigree analyses based on families from various origins (for instance, the group of Icelandic families analyzed by deCODE Genetics, or the families from European ancestry collected by the CEPH, Centre d’Etude du Polymorphisme Humain) (Kong et al. 2004; Matise et al. 2007). The parental origin of each recombination event could be determined, thus establishing male and

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Table 6.1 Description of the second generation Rutgers combined linkage physical maps Chromosome No. of mapped No. of intervaled Physical Map length (cM) markers markersa length (Mb)b Sex Female Male averaged 1 1,968 326 245.3 286.2 365.5 209.8 2 1,957 245 242.5 264.5 333.7 197.7 3 1,716 287 199.1 223.8 281.0 168.7 4 1,415 258 191.1 214.7 273.4 158.2 5 1,398 342 180.3 208.5 263.0 157.1 6 1,385 252 170.6 196.0 248.0 146.3 7 1,205 199 158.5 188.1 237.1 140.7 8 1,164 138 145.7 168.7 218.8 120.8 9 995 149 139.9 167.2 199.5 136.1 10 1,149 222 134.9 175.0 215.8 136.5 11 1,252 169 134.2 161.6 197.2 127.5 12 1,145 141 132.1 175.6 214.1 139.2 13 857 144 95.6 131.8 158.0 106.2 14 799 128 86.8 125.2 144.1 107.4 15 781 89 79.8 131.8 155.7 109.4 16 744 131 88.6 133.4 158.1 111.8 17 761 221 78.5 138.9 164.8 115.5 18 758 74 75.8 129.6 150.0 110.5 19 554 108 62.7 111.1 127.8 96.4 20 600 66 62.4 114.3 124.4 105.8 21 426 40 33.2 69.1 81.1 58.5 22 360 71 34.1 80.2 90.1 71.7 X 779 153 154.0 194.9c 194.9 35.9 Total: 24,168 3,953 2,925.8 3,789.9 4,596.1 2,867.6 From Matise et al. (2007) a These markers could not be localized into a single map position and were instead localized to a larger map interval or bin b Physical length spanned by these maps; marker positions are from the NCBI Build 36.1 (March 2006 genome assembly) c The female map of the X chromosome is used when calculating the length of the entire genome

female genetic maps. These maps provide information on recombination rates (the recombination activity per physical distance in cM/Mb) over large intervals. In the combined analysis of both Icelandic and CEPH families (Matise et al. 2007), the map length (cM) for all autosomes ranged between 58.5 and 365.5 cM per chromosome, thus exceeding 50 cM, which is the length expected for a frequency of one CO per chromosome pair (Table 6.1). These data revealed several differences between male and female maps: 1. The overall CO frequency is higher in females (4,596.1 cM) than in males (2,867.6 cM) (in average 1.55-fold higher on autosomes). The CO rate is 1.6 cM/Mb in females and 1.02 cM/Mb in males. 2. All autosomes have higher CO frequencies in females.

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Fig. 6.2 Female and male recombination rates along human chromosome 1. Rates are determined over 3 Mb windows by steps of 1 Mb. Red lane female, Blue lane male. From Kong et al. (2002) with permission

3. However, along a given chromosome, CO rates can be higher in males in some areas and the difference between female and male recombination rates varies in different regions (Fig. 6.2). A striking example of higher recombination rate in males is the PAR1 region on the X chromosome, where the CO rate is around 20 cM/Mb in males, a value that is more than tenfold higher than the average recombination rate in females. This is explained by the fact that PAR1 is the main region of homology between the X and the Y chromosome, PAR1 is 2.6 Mb long, and one CO is expected to occur in this region in each meiotic cell. The other region of homology, PAR2, is only 320 kb long. 4. Near subtelomeric regions, recombination rate tends to be relatively high in males, sometimes exceeding that of females. 5. A common feature in both sexes, however, is CO repression at centromeric regions. This property is most likely due to the heterochromatic structure of these regions as shown by functional assays in other model organisms, such as D. melanogaster (Westphal and Reuter 2002) and yeast (Petes 2001). Repression of recombination adjacent to the centromeric repeated DNA might be due to spreading of constitutive heterochromatin in this region. Indeed, analysis of human chr. 21 shows a sharp transition between the proximal heterochromatic domain, which is repressed, and the distal euchromatic domain, which is active and where recombination takes place. This transition corresponds to a peak of repressive histone methylation marks (Grunau et al. 2006).

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Fig. 6.3 Female and male recombination rates along a region of mouse chromosome 1 at high resolution. Recombination rates that are off scale are shown as numbers over each interval. Red arrows hotspots that are predominantly active in females, blue arrows hotspots that are predomi nantly active in males. From Paigen et al. (2008)

In the mouse, a genotyping project using both inbred and outbred mice highlighted similar features: increased map length for each autosome in females relative to males (in average 1.3-fold higher in female) and variations of the difference between female and male recombination rates in different regions (Shifman et al. 2006; Cox et al. 2009). A higher resolution analysis (in the order of 1 100 kb depending on the interval) of a segment of mouse chr. 1, confirmed that sex differences in recombination rates are localized to specific regions. Indeed, within this chromosomal region, a 9 Mb area (169 178 Mb) showed generally higher recombination rate in females, whereas the adjacent region (178 194 Mb) had higher recombination rate in males. In addition to these regional variations, individual hotspots also can have distinct activities in males or females with differences that do not necessarily correlate with global changes (Paigen et al. 2008) (Fig. 6.3). This suggests that at least two levels of control influence recombination activity, one at the hotspot level and one at the level of chromosome domains. These sex-specific features of COs have been confirmed by independent cytological studies, where COs were quantified using MLH1 foci as markers (Tease et al. 2002). In humans, both the total number of MLH1 foci and the number of MLH1 foci per individual chromosome, determined by immuno-FISH analysis, were in average 1.4-fold higher in oocytes than in spermatocytes (Barlow and Hulten 1998). This approach also allowed to address the question of whether the difference observed was due to the genotype or to sex-specific factors arising during development of the meiotic cell, and thus to determine the genetic or epigenetic origin of this phenomenon. Ideally, one would like to know if oocytes from an XY individual would have a male or a female pattern of COs. This question has been addressed using XY* mice, which carry a Y chromosome from M. m. poschaivinus and show a sex reversed phenotype with female gonad development. In these XY* mice, the CO rate and distribution were similar to those of XX females, suggesting that sex-specific factors and not genotype influence these recombination properties

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(Lynn et al. 2005). This favors the idea that some features occurring during oocyte development might regulate meiotic recombination through an epigenetic control.

6.4.1.2

Recombination Activities in Minisatellites

Minisatellites are short DNA sequences (6 100 bp) repeated in tandem and spanning from 0.5 kb to several kilobases. Minisatellites have a high degree of polymorphism, and some have been shown to be hypermutable in the germ line, possibly due to high frequency of recombination during meiosis (Vergnaud and Denoeud 2000). Among ten unstable minisatellites, three of them, including the well-studied CEB1 minisatellite (Buard et al. 2000), are specifically unstable in the male germ line (Vergnaud et al. 1991), whereas the others do not show significant differences between males and females. At least for some minisatellites, instability has been proposed to be directly linked to the meiotic DSB repair pathways. In such cases, the difference in stability between sexes might be a consequence of the different meiotic recombination activities observed during male and female meiosis.

6.4.1.3

Regions Subjected to Parental Imprinting

Parental genomic imprinting is an epigenetic mechanism that confers distinct properties to some chromosomal regions depending on their parental origin. It is brought about by complex epigenetic modifications and some, such as DNA methylation, are laid down at different times during male and female germ line developments. Kinetic analysis of DNA methylation suggests that these marks are implemented before meiosis in the male germ line and after birth, i.e., after meiotic prophase, in the oocytes (Delaval and Feil 2004). In regions subjected to parental imprinting, one might expect epigenetic differences between male and female chromosomes to be present when meiotic recombination occurs, and consequently also differences in the process of meiotic recombination itself. Two reports have shown indeed that this may be the case (Paldi et al. 1995; Sandovici et al. 2006). By pedigree analysis, Paldi and co-workers (1995) detected a significantly higher recombination rate in males than in females in two regions containing imprinted genes, a 1.9 Mb region on chr. 15 (15q11-13) and a 1.3 Mb region on chr. 11 (11p15.5). Subsequent analysis of the 15q11-13 region could not reproduce this observation and actually found a higher recombination rate in females (Lercher and Hurst 2003). In any case, these results should be interpreted with caution as they could be due to other differences between male and female chromosomes than imprinting. For instance, recombination rates are often higher in subtelomeric regions in meiotic male germ cells. Since 11p15.5 is a subtelomeric region, its higher CO rate in males might be a consequence of its localization on the chromosome. Analysis of recombination rates in imprinted regions also revealed that recombination rates (sex averaged) are 1.4-fold higher in imprinted regions than

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in the whole genome (Lercher and Hurst 2003). This analysis was based on the deCODE map (Kong et al. 2002) with a resolution of about 1 cM, a window size considerably larger than most of the imprinted domains. Other authors, using the LD-based map, found that overall, the LD values per Mb of 17 imprinted domains (1 Mb in size in average) were significantly higher compared to those of random nonimprinted regions (Sandovici et al. 2006). A higher resolution study by pedigree analysis of a 1 Mb region of chr. 11p15.5 showed that recombination was higher in males than females, consistent with the results by Paldi et al. (1995). In a 0.5 Mb interval of this region, which contains several imprinted genes and two imprinting control centers (H19/IGF2 and KCNQ1OT1), the CO rate was about fivefold higher in males than females. Interestingly, two LD-based hotspots are located in the vicinity of these imprinting control centers (Sandovici et al. 2006).

6.4.1.4

CO Frequency and Length of the Synaptonemal Complex

During meiotic prophase, homologous chromosomes pair together and become stably associated and synapsed through a structure called the synaptonemal complex (SC). In this configuration, observed at the pachytene stage of prophase I, the protein axes of each homologous chromosome, called the lateral elements, are linked through a central structure, the central element. Recombination takes place in the context of this structure, and in particular, MLH1 foci are found on the SC. One interesting observation based on cytological analysis of chromosomes from oocytes and spermatocytes is that SC length differs between males and females. In humans and mice, a correlation between SC length and CO rate has been observed when comparing strains, individuals, or sexes (Lynn et al. 2002; Tease et al. 2002). For instance, in the mouse C57BL/6 strain, the SC length of chr. 1 is 13.2 mm in females and 10.2 mm in males and MLH1 foci frequency on chr.1 is 1.85 and 1.54 cM/Mb, respectively (de Boer et al. 2006). Thus, the intercrossover distance when measured in physical distance (mm) is in average identical in males and females. This suggests that the metric for CO control is related to the chromosome axis (Kleckner et al. 2003). Furthermore, the important role of chromosome structure and of genes coding for condensins (components of chromosome axes) in regulating CO frequency has been recently shown in the nematode C. elegans (Mets and Meyer 2009). These observations may indicate that at least some of the differences in CO frequency and/or localization during male and female meiosis may be due to differences in chromosome organization and that they are under genetic control.

6.4.2

Interindividual Variations

Many variations of meiotic recombination between individuals, species, or populations have been observed, and it has been suggested that they are due to DNA

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sequence or epigenetic modifications. However, it is rarely possible to determine whether some underlying polymorphism in the region tested is responsible for the observed effect and/or whether an epigenetic modification is involved.

6.4.2.1

Variations in Genome Wide CO Rates

The global CO activity per meiotic cell varies among individuals as measured by MLH1 foci quantification. In mice, the number of MLH1 foci was also found to differ in the strains C57BL/6, Cast/Ei, and Spret/Ei. However, the number of MLH1 foci relative to the SC length was nearly constant: 1 MLH1 focus every 7.1, 6.9, and 6.9 mm in CAST/Ei, C57BL/6, and SPRET/Ei, respectively, consistent with the SC/CO relationship (Lynn et al. 2002). In men, the number of MLH1 foci ranged from 40 to 60 per spermatocyte in 14 individuals, with a mean going from 46.2 3.3 to 52.8 4.8 (Lynn et al. 2002). In human oocytes, the intercell variation was also important with 42 102 foci per oocyte at the pachytene stage and interindividual means ranging from 52.6 to 88.3 (Lenzi et al. 2005; Cheng et al. 2009). A potential factor of variation in these studies could be the developmental stage of the cells analyzed and/or the timing of MLH1 focus formation, which could artifactually add experimental variation. Variations in the number of CO events per meiosis have also been measured genetically in humans, between parents of families used to build genetic maps (Broman et al. 1998; Kong et al. 2002; Cheung et al. 2007). Genome-wide searches identified several sequence variants associated with this variation in both males and females, including variants located within the Rnf212 gene, an ortholog of the C. elegans ZHP-3 gene involved in chiasma formation as well as four other genes with unknown functions (Kong et al. 2008; Chowdhury et al. 2009).

6.4.2.2

Variations of Hotspot Activity in Humans

High resolution analysis of CO by sperm analysis has shown major differences in recombination activities among individuals. In several cases, variations of hotspot activities measured by sperm typing have been shown to be correlated with a polymorphism located near the center of the hotspot [DNA2 (Jeffreys and Neumann 2002), NID1 (Jeffreys and Neumann 2005) and S2 (Jeffreys and Neumann 2009)], indicating a genetic control of recombination activity. In contrast, two other hotspots, MSTM1a and 1b, showed differences in activity independent of local DNA sequence variations (Neumann and Jeffreys 2006). At these two adjacent hotspots, COs were analyzed by sperm typing in 26 individuals. The MSTM1a hotspot was active in only three men, whereas the MSTM1b hotspot was active in all men tested but with recombination frequencies ranging from 1.2 to 90 10 5 (Fig. 6.4). At MSTM1a, no genetic variation specific to the three men could be identified in 100 Kb around the hotspot. For

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Fig. 6.4 Interindividual differences in CO hotspot activity. CO frequencies at two hotspots, MSTM1a and MSTM1b, were measured by PCR assays using sperm DNA from 26 men. The frequency of CO per DNA molecule is reported. From Neumann and Jeffreys (2006) with permission

MSTM1b, some men with different CO activities presented the same haplotype over 20 kb around the hotspot. Overall, these CO variations could thus be explained by the presence of a remote cis acting element (located more than 100 Kb in the case of MSTM1a), by an unlinked polymorphism with a trans effect and/or by an epigenetic modification. A genome-wide pedigree analysis in the Hutterite population, a founder population from European ancestry currently living in North America, has allowed to map CO activity in 164 individuals at the hotspot resolution (i.e., less than 30 Kb). These COs were compared to the 32996 LD-based, well-resolved hotspots, obtained using the phase II Hapmap. A high degree of interindividual variation was observed with hotspot usage ranging from 10% to nearly 100% with a mean of 60% (Coop et al. 2008). A major fraction of this interindividual variation is now explained by the regulatory activity of PRDM9 on initiation of recombination through binding to specific sequences (Baudat et al. 2010). This control involves epigenetic modifications since Prdm9 encodes for a protein with histone methyl-transferase (HMT) activity (see Sect. 6.6).

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Variations of Hotspot Activity in Mice

Recent studies in mice have analyzed in detail differences in recombination rates between hybrids. In the first study, two hybrids, which differed only by heterozygosity over a 30 Mb region on chr. 17, were shown to display genome-wide differences in CO activities, suggesting the implication of a trans acting factor. The locus responsible was mapped to a region of 6.7 Mb on chr. 17 and named Dsbc1, for DSB control 1 (Grey et al. 2009). The second study showed that the genetic origin of an overlapping 5.3 Mb region on chr. 17 influenced CO activity on chr. 1. This region was named Rcr1, for Recombination control region 1 (Parvanov et al. 2009). These studies are the first reports that identify trans-acting modifiers of recombination in mice, and the gene involved has now been mapped: it is Prdm9, the ortholog of the one identified to influence hotspot usage in humans and whose properties and function are described in Sect. 6.2.

6.5

6.5.1

Evidence for Epigenetic Modifications During Meiosis and Genes Known to Regulate such Modifications in Mammals DNA Methylation

In mammals, DNA methylation patterns are initially acquired during germ cell development. Interestingly, the timing of de novo methylation differs in male and female germ cells. In the mouse female germ line, methylation patterns are acquired postnatally during the oocyte growth phase, after completion of meiotic recombination and prophase I of meiosis (Kono et al. 1996; Walsh et al. 1998; Lucifero et al. 2002). Conversely, in the male germ cells, methylation patterns are initially acquired prenatally and further consolidated after birth during spermatogenesis (La Salle and Trasler 2006). In parallel, maintenance of DNA methylation also takes place in the context of DNA replication, in spermatogonia and preleptotene spermatocytes. These cells are therefore capable of de novo and maintenance DNA methylation. Both de novo and maintenance DNA (cytosine-5)-methyltransferases (DNMTs) work in concert to generate and propagate these genomic methylation patterns (Fig. 6.5). Currently, five DNMTs (DNMT1, DNMT2, DNMT3a, DNMT3b, and DNMT3L) have been characterized and classified according to similarities found in their C-terminal catalytic domain (Goll and Bestor 2005). DNMT1 is in charge of maintenance methylation and is the major DNMT expressed in somatic tissues. In testis, its expression is highest in type A spermatogonia and low in type B spermatogonia and preleptotene spermatocytes; it increases in leptotene/zygotene and decreases in pachytene spermatocytes to be reexpressed in round spermatids (La Salle and Trasler 2006). In contrast, DNMT3a and DNMT3b have been postulated to function primarily as de novo DNA methyltransferases (Okano et al. 1999). The

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a

Mitotic phase Spg B

Meiosis (Prophase I) PL

L

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Modifying enzymes

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Prdm9 Suv39h1 Suv39h2 G9a

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Spg A

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TH2B TH3 H3.3A H3.3B H1t

DNMT3a2 DNMT3b DNMT3a DNMT1

Fig. 6.5 Dynamics of histone modifications, histone modifying enzymes, histone variants, and DNMTs during spermatogenesis. (a) Distribution of histone modifications and expression of histone modifying enzymes in male mouse germ cells. Histone dynamics are based on immunos taining of spermatocyte spreads or testis sections. Modifying enzyme dynamics are based on immunohistochemical analysis (AOF2, G9a), immunostaining and RNA in situ hybridization on testis sections (Suv39h1/2), RT PCR amplification of testis cDNA (Prdm9), and immunostaining (JMJD1A, G9a). Spg A: spermatogonia type A, Spg B: spermatogonia type B, PL: preleptotene, L: leptotene, Z: zygotene, P: pachytene, D: diplotene, RS: round spermatids, ES: elongated spermatids, HMT: histone methyl transferase, HDMT: histone demethylase, n.d.: not determined. (1) Godmann et al. (2007), (2) Buard et al. (2009), (3) Tachibana et al. (2007), (4) Peters et al. (2001), (5) Godmann et al. (2009). (b) Chromatin remodeling and dynamics of histone variants and expression of DNA methyl transferases (DNMTs) in male mouse germ cells. Dynamics of histone variants are based on immunostaining (gH2AX), immunohistochemistry on testis sections (TH2A/B and TH3), RNase protection assay and nonradioactive in situ hybridization on testis sections (H3.3A and B), in situ hybridization and immunohistochemical techniques (H1t mRNA and protein) and real time RT PCR amplification (DNMTs). (1) Kimmins and Sassone Corsi (2005), (2) La Salle and Trasler (2006)

inactivation of Dnmt3a in germ cells causes impaired synapsis of homologous chromosomes, reactivation of retrotransposons, imprinting defects, and progressive loss of germ cells, followed by azoospermia (Kaneda et al. 2004). In the study of La Salle et al. (La Salle and Trasler 2006) the global level of Dnmt3a transcripts was

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measured and was similar to that of Dnmt1. In contrast, Dnmt3b is dispensable for spermatogenesis, and its expression is more dynamic during male germ cell development: it is highest in type A spermatogonia, decreases dramatically (by more than fourfold) in type B spermatogonia and preleptotene spermatocytes, and increases again by almost twofold in leptotene/zygotene spermatocytes. Once more, in pachytene, a steady decrease in expression is observed, while Dnmt3b transcripts are clearly detected in round spermatids (La Salle and Trasler 2006). These authors also analyzed Dnmt3a2, the second main species of mRNA transcribed from the mouse Dnmt3a gene. It presents a very dynamic expression pattern reminiscent to that of Dnmt3b. Taken together, these data indicate that, in spermatogenesis, transcription of Dnmt genes is downregulated during: (a) differentiation of spermatogonia into spermatocytes and (b) during pachytene. The authors postulate that their downregulation could be related either to the need to express testis-specific genes and/or to changes in chromatin structure. Finally, DNMT3L is closely related to DNMT3a and DNMT3b but lacks the conserved transmethylation domain. It is remarkable because Dnmt3L is exclusively expressed in germ cells and shows a strong sexual dimorphism in expression patterns. In females, it is present only in growing oocytes (Bourc’his et al. 2001). Conversely, in male germ cells, Dnmt3L expression first occurs after day 12.5 postcoitum in prospermatogonia and remains high until birth. Postnatally, Dnmt3L expression declines dramatically and is only detectable at low levels in mature germ cells (Bourc’his et al. 2001; Hata et al. 2006; Sakai et al. 2004; Webster et al. 2005; La Salle et al. 2007). It has been proposed that low DNMT3L expression is required for the continued epigenetic programming that occurs up to pachytene (Godmann et al. 2009). In fact, the phenotype observed in Dnmt3L / mice is indistinguishable from that of mice with germ cell-specific ablation of Dnmt3a (Kaneda et al. 2004). Indeed, both phenotypes are characterized by abnormal reactivation of retrotransposons, imprinting defects due to abnormal DNA methylation, impaired synapsis between homologous chromosomes, failure to progress through meiotic prophase, azoospermia, and sterility (Bourc’his et al. 2001; Bourc’his and Bestor 2004; La Salle et al. 2007; Webster et al. 2005; Hata et al. 2006). In comparison to the other DNMTs, little is known about DNMT2. It seems to have contradictory roles in both DNA and RNA methylation (Schaefer and Lyko 2009).

6.5.2

Histone Modifications

Histones are a group of evolutionarily conserved proteins that play a critical role in the packaging of DNA into chromatin. There are four classes of core histones (H2A, H2B, H3, and H4) and a linker histone (H1). The regulatory function of histones is mainly achieved through their covalent modification, primarily at amino acid residues in their N-terminal tail regions (Xu et al. 2009). Only few data are available about histone modifications during the prophase I of meiosis and their

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relationships with meiotic recombination and synapsis. The best described is the formation of the male-specific “sex body,” whereby the X and Y chromosomes undergo several specific chromatin modifications during the meiotic prophase and become transcriptionally silent (MSCI: for meiotic sex chromosome inactivation) (see below and reviewed in (Zamudio et al. 2008), and Chap. 7). To date, a limited number of studies have analyzed the spatial and temporal dynamics of histone modifications during meiotic prophase I (see Fig. 6.5a): data are available on trimethylation of H3 at lysine 4 (H3K4Me3) (Godmann et al. 2007; Buard et al. 2009), H3K4 dimethylation (H3K4Me2), H3K27 methylation (H3K27M1), H3K9 monoacetylation (H3K9Ac1), H4 acetylation (H4Ac) (Buard et al. 2009), and H3K9 mono-, di-, and trimethylation (H3K9Me1, Me2, and Me3) (Tachibana et al. 2007). Furthermore, it was recently shown that, in mouse, H3K4Me3, together with other modifications, is associated with the initiation of meiotic recombination (Buard et al. 2009), suggesting a direct implication of this modification in meiotic DSB formation, similarly to data obtained in S. cerevisiae (Borde et al. 2009). Indeed, at the active mouse Psmb9 hotspot, chromatin is enriched in H3K4Me3, H3K4Me2, H3K9Ac1, H3K27Me1, and H4Ac, whereas it is depleted in H3K9Me3 and H3K27Me3 (Buard et al. 2009). Several assays showed a specific enrichment of H3K4Me3, H3K4Me2, and H3K9Ac1 on chromatids that carry an active initiation site, presumably before DSB formation, thus suggesting that these modifications could be part of the substrate for recombination initiation at the Psmb9 hotspot. In contrast, H4Ac is increased as a consequence of DSB formation. H3K4Me3 is known to be enriched near transcription start sites of actively transcribed genes. However, no transcript could be detected at the Psmb9 hotspot, which is located in an intergenic region. Given that promoters of transcribed genes are not meiotic initiation sites, some yet unknown feature must differentiate these regions, which are both enriched for H3K4Me3. The dynamics of H3K4 methylation in spermatogenesis follows the waves of transcription and chromatin remodeling events with high levels of the modification appearing in spermatogonia up to the leptotene stage and then downregulation in pachytene cells and a brief reappearance in elongating spermatids (Fig. 6.5a; Godmann et al. 2007). In addition to these data on histone modifications, knockout (KO) mice for several genes coding for histone-modifying enzymes show impaired meiosis and thus provide insight into the role of epigenetic modifications during meiosis (see Fig. 6.5a and Table 6.2). Three types of HMT (Suv39h1-Suv39h2, G9a and Prdm9) have a critical function during meiosis in mammals. Suv39h1 (KMT1a) and Suv39h2 (KMT1b) are H3K9Me1 and H3K9Me2 trimethylases. In adult tissues, Suv39h1 is broadly expressed, whereas Suv39h2 is highly expressed in the testis, where it localizes to the nuclei of Sertoli cells and to heterochromatin of spermatocytes and round spermatids. Peters and co-workers (2001) have shown the essential function of this HMT in epigenetic programming during spermatogenesis. Whereas single KO mutants are fertile, double KO for Suv39h1 and Suv39h2 present severe spermatogenic abnormalities. Indeed, entry into meiotic prophase is delayed, homologous synapsis is affected, and apoptosis is induced during the transition from mid to late

Table 6.2 Epigenetic modifiers, adapted from Godmann et al (2009) Genes Function Substrate Phenotype of KO Dnmt1 Maintenance DNMT CpG Loss of global DNA methylation, improper expression of imprinted genes, ectopic X-inactivation, reactivation of transposable elements Dnmt3A De novo DNMT CpG Impaired synapsis of homologous chromosomes, reactivation of retrotransposons, imprinting defects ! progressive loss of germ cells followed by azoospermia Dnmt3B De novo DNMT CpG Dispensable for spermatogenesis Dnmt3L No catalytic activity CpG Impaired synapsis of homologous chromosomes, reactivation of retrotransposons, imprinting defects ! progressive loss of germ cells followed by azoospermia Kmt1a (Suv39h1) Histone H3K9Me1,2 No phenotype methyltransferase Kmt1b (Suv39h2) Histone H3K9Me1,2 No phenotype methyltransferase Suv39h1/2 Histone H3K9Me1,2 Meiotic arrest: impaired synapsis of homologous double KO methyltransferase chromosomes, missegregation of bivalents, aneuploidy, genomic instability ! infertility Kmt1c (G9a) Histone H3K9Me1,2 Meiotic arrest: Synaptonemal complex malformations, methyltransferase irregular gene expression due to loss of H3K9, silencing ! infertility Prdm9 (Meisetz) Histone H3K4Me3 Impaired DSB repair pathway, defective pairing of methyltransferase homologous chromosomes, inaccurate sex body formation, ! infertility Kdm3a (Jmjd1a) Histone demethylase H3K9Me1,2 Postmeiotic defects: impaired chromatin condensation and acrosome formation, defective heterochromatin distribution, failed expression of Tnp1, Prmt1 Hdac6 Histone deacetylase Acetylated Dispensable for spermatogenesis lysines mHr6b Ubiquitin conjugating H2Aub, Dispensable for spermatogenesis enzyme H2Bub Baarends et al. (2003), Roest et al. (1996)

Zhang et al. (2008)

Okada et al. (2007)

Hayashi et al. (2005)

Tachibana et al. (2007)

Peters et al. (2001)

Kaneda et al. (2004) Bourc’his et al. (2001), Bourc’his and Bestor (2004), La Salle et al. (2007)

Kaneda et al. (2004)

Reference Li et al. (1992), Panning and Jaenisch (1996), Walsh and Bestor (1999)

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pachytene, leading to missegregation of the bivalents, genomic instability, and infertility. These authors proposed a role of Suv39h in defining a distinct higher order chromatin structure that may be required for the initial alignment and clustering of meiotic chromosomes (Peters et al. 2001). G9a/EHMT2 (KMT1c) is a mammalian H3K9 mono- and dimethyltransferase that acts in a multiprotein complex. In contrast to Suv39h1/2, it localizes to euchromatin. In the adult testis, G9a is expressed in spermatogonial stem cells and differentiating spermatogonia until the early leptotene stage and in spermatocytes. During the female meiotic prophase, the kinetic of expression is similar to that observed in male germ cells. Germ cell-specific ablation of G9a (Tachibana et al. 2007) leads to male infertility with a dramatic reduction of germ cells in adulthood. Like in Suv39h1/2 null mice, meiosis is aborted during the early pachytene stage with synapsis defects (Tachibana et al. 2007). Interestingly, the underlying mechanisms seem to be different, as Suv39h1/2 catalyzes H3K9 methylation in heterochromatic regions (Peters et al. 2001), whereas G9a brings H3K9 methylation marks to euchromatin (Tachibana et al. 2007). G9a / mice also display altered expression of a small number of genes (Tachibana et al. 2007). It has been thus proposed that G9a deficiency could alter meiotic prophase due to modifications of chromatin structure and/or deregulation of transcription of important genes. PRDM9 (Meisetz) is a H3K4 trimethylase, which seems to be crucial for proper meiosis in both sexes (Hayashi et al. 2005). Prdm9 / spermatocytes and oocytes exhibit impaired DSB repair during the pachytene stage and undergo cell death before completing meiosis. The implication of PRDM9 in the definition of the initiation sites of recombination is discussed in the last part of this review (Sect. 6.6). These KO mouse models demonstrate that a specific association of repressive and activating histone methylation marks mediates the highly orchestrated expression of meiosis-specific genes. Some of these marks might also be important for other chromatin features linked to chromosome dynamics during meiotic prophase (pairing, recombination, and synapsis), as recently proposed for the control of the initiation of meiotic recombination by PRDM9-mediated H3K4Me3 (Baudat et al. 2010; Buard et al. 2009) Reciprocally, the controlled removal of histone methylation marks by Histone Demethylases (HDMT) is essential for spermatogenesis. First, AOF2 (Lsd1/ KDM1) specifically removes activating H3K4 mono- and dimethylation marks and, in concert with the androgen receptor, repressive H3K9 mono- and dimethylation marks (Shi et al. 2004). Noteworthy, Aof2 is expressed in a developmental- and stage-specific manner during spermatogenesis: its expression is high in spermatogonia, increased in preleptotene spermatocytes, and then dramatically decreased in pachytene spermatocytes (Fig. 6.5; Godmann et al. 2007). Moreover, its distribution pattern directly overlaps with that of H3K4 mono- and dimethylation in mouse testis. It is probably a key regulator of H3K4 methylation in spermatogenesis. Interactions between the epigenetic modifiers AOF2, HDAC1, and MBD2a and b were identified by coimmunoprecipitation of protein extracts from mouse testis,

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which suggest that these proteins play a role in chromatin reorganization. The conditional KO of Aof2 is embryonic lethal at 7.5 days postcoitum (Wang et al. 2007). On the other hand, the HDMT JMJD1a (of the family of Jumonji Domaincontaining Histone Demethylases, also called JHDM2A or KDM3a) specifically demethylates H3K9Me1 and H3K9Me2 (Yamane et al. 2006). Rat Jmjd1a was originally identified as a testis-specific gene (Hoog et al. 1991). Jmjd1a / mice exhibit postmeiotic chromatin condensation defects, and JMJD1a directly binds to and controls the expression of Transition nuclear protein 1 (Tnp1) and Protamine 1 (Prm1) (Okada et al. 2007). Analyses of Jmjd1a expression in spermatocytes suggest that the reduction of H3K9Me2/1 observed at the pachytene stage is mediated, at least in part, by JMJD1a (Tachibana et al. 2007). This hypothesis is supported by the finding that pachytene as well as leptotene/zygotene oocytes lack JMJD1a (as tested by immuno-fluorescence). Active removal of H3K9Me2/1 at the pachytene stage is thus a specific feature of male meiosis. Furthermore, histones can also be ubiquitylated on their N-terminal tail. The addition of a single ubiquitin group to H2A and H2B seems to be critical for chromatin remodeling events during postnatal germ cell development. The KO of the ubiquitin-conjugating enzyme mHr6b results in male infertility due to meiotic and postmeiotic defects (Roest et al. 1996), whereas female fertility is not affected. In addition to defects in postmeiotic chromatin remodeling, Baarends and coworkers reported a specific requirement for the ubiquitin-conjugating activity of HR6B in relation to dynamic aspects of the SC meiotic recombination in spermatocytes (Baarends et al. 2003). Increased apoptosis in Hr6b / primary spermatocytes was detected during the first wave of spermatogenesis, indicating that HR6B has a primary role during the meiotic prophase. Detailed analysis of Hr6b / pachytene nuclei showed increased SC length and number of MLH1 foci (Baarends et al. 2003). Finally, acetylation of histone tails plays a role in prophase I. Consequently, some histone acetyltransferases (HATs) and deacetylases (HDACs) must be implicated. Although almost all HDACs are ubiquitously expressed, HDAC6 is very abundant in mouse testis (Godmann et al. 2009). However, genetic ablation of Hdac6 did not affect spermatogenesis as Hdac6 / mice are viable with normal fertility (Zhang et al. 2008). The inhibition of HDACs by Trichostatin A (TSA) is an alternative approach to study the impact of histone acetylation on spermatogenesis. In TSA-treated testes, a dramatic loss of pachytene and diplotene spermatocytes was observed, whereas the number of spermatogonia was only weakly decreased and progression through metaphase I and II of meiosis was not perturbed (Fenic et al. 2004). The mechanism involved in this effect remains unclear, but this study emphasizes a critical role of HDACs in spermatogenesis. Taken together, these studies show that meiotic prophase progression must be strictly controlled by a variety of epigenetic modifiers; however, the existence of a direct or indirect functional link between histone modifications and the process of meiotic recombination remains to be evaluated.

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6.5.3

Histone Variants in Meiotic Prophase I (Fig. 6.5b)

A specific remodeling of chromatin takes place during spermatogenesis with the sequential replacement of somatic histones with histone variants at different stages of germ cell differentiation. Here, we describe the histone variants incorporated during meiotic prophase, leaving aside the changes occurring during spermiogenesis (reviewed in the Chap. 8).

6.5.3.1

The Specific Case of the XY Sex Body (Histone Modifications and Histone Variants)

The sex body is a nuclear compartment in the male germ cells where selective inactivation of sex chromosome gene expression occurs. In male meiosis, the heteromorphic X and Y chromosomes can undertake homologous pairing only in a relatively small region of homology, the pseudoautosomal region. Clustered in the sex body, they undergo the condensation process of heterochromatinization, accompanied by transcriptional silencing (MSCI). The sex body contains a number of proteins that are implicated in heterochromatinization, including the H2A variant macro-H2A1 and the Heterochromatin Protein 1 beta (HP1b) (Kimmins and Sassone-Corsi 2005). In addition, the chromatin of the sex body is characterized by increased methylation of H3K9, resulting in higher level of HP1b. Ubiquitination of H2A is also increased and phosphorylation of gH2AX is enhanced (including in Spo11 / mice indicating that this phosphorylation is not dependent on DSB formation) [reviewed in (Baarends and Grootegoed 2003)]. These phenomena are thought to result from the failure of the X and Y to synapse. In fact, a similar process called MSUC (for Meiotic Silencing of Unsynapsed Chromatin) is observed in conditions where autosomes fail to synapse (Baarends et al. 2005; Turner et al. 2005; Burgoyne et al. 2009). These chromatin modifications, which are correlated with transcription repression, might also play a role on recombination events, for instance, in allowing DSBs to be repaired between sister chromatids. On the X and Y, two important consequences of sex body formation could be first to allow the repair of DSBs formed in the nonhomologous region of the X and Y, and second to repress the expression of some genes encoded on the X and Y, and deleterious for meiotic progression.

6.5.3.2

H2A and H2B Variants

Phosphorylation of H2AX Depending on its structural organization, chromatin can generate a barrier to DSB formation and DNA damage repair. One of the roles of histone modifications is to allow recognition of and accessibility to DNA sequences where recombination (i.e. DSB formation and repair) takes place. One of the earliest recognized responses

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to DNA damage is phosphorylation of the histone variant H2AX in mammalian cells (Rogakou et al. 1998), and it is involved in the recruitment of several proteins required for DSB signaling and repair (Riches et al. 2008). H2AX is a highly conserved H2A variant, which can be differentiated from other H2A isoforms by the presence of a conserved SQ motif at the C-terminus and which is phosphorylated on the serine residue 139 in mammals (the phosphorylated form is named gH2AX). It represents 10 20% of the total H2A of chromatin (Redon et al. 2002) and it is found in large amounts in adult germ cells (Tadokoro et al. 2003; Yoshida et al. 2003). In mouse spermatocytes, gH2AX is spatially and temporally linked to meiotic DSBs and synapsis (Mahadevaiah et al. 2001). During leptotene, it is abundant, encompasses a large proportion of the chromatin, and its phosphorylation is SPO11-dependent (Baudat et al. 2000; Romanienko and Camerini-Otero 2000). As synapsis progresses, the global level of gH2AX decreases and it becomes eventually undetectable on autosomes at the end of pachytene. In spermatocytes, a high level of gH2AX is found on the chromatin of the sex body (see above). Finally, H2ax deletion is characterized, among other features, by male infertility due to a defect in sex body formation. Surprisingly, DSBs appear to be repaired properly in these mutant mice and females are fertile (Celeste et al. 2002; Fernandez-Capetillo et al. 2003).

Testis-Specific Histone Variants TH2A and TH2B Male germ cells have an unusually high number of histone variants in comparison to somatic cells. TH2A and TH2B are testis-specific variants that are expressed during meiotic prophase and present remarkable differences in their N-terminal tails. H2A and TH2A differ in eight amino acid residues in the first half of the molecules and three consecutive changes in the C-terminal region (Pradeepa and Rao 2007). TH2B differs from H2B by the addition of three potential phosphorylation sites (Ser 12, Thr 23, and Thr 34) and the repositioning of two others (Ser 5 and Ser 6), resulting in a different “phosphorylation map” of its N-terminal tail (Kimmins and Sassone-Corsi 2005). By immunohistochemistry, TH2B was detected on chromatin of early primary spermatocytes in rat but not in round spermatids (Unni et al. 1995). In humans, TH2B is present throughout spermatogenesis, from spermatogonia to mature sperm (Zalensky et al. 2002). The nucleosomal core histone of pachytene chromatin has 80% of H2B replaced by TH2B while about 10% of H2A is replaced by TH2A.

6.5.3.3

H3 Variants

H3 possesses three somatic variants, H3.1, H3.2, and H3.3 (Franklin and Zweidler 1977), but only one testis-specific variant, TH3, has been described (Trostle-Weige et al. 1984). The two variants of H3.3 (H3.3A and H3.3B) are incorporated into the germ line. H3.3A is present during male germ line development from the spermatogonia to the spermatid phase (Bramlage et al. 1997). But while H3.3A mRNA is

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present in pre- and postmeiotic cells, expression of H3.3B is essentially restricted to cells in meiotic prophase. So, by prophase I, H3 is largely replaced by H3.3A and H3.3B, which apparently associate with euchromatin, suggesting a role in the massive transcription program of spermatocytes. Indeed, in D. melanogaster, H3.3 is enriched in modifications associated with transcriptional activation and is deficient in demethylation at K9 (McKittrick et al. 2004), a modification associated with silencing and heterochromatin formation. TH3 is strongly detected in spermatogonia and similarly or even more in spermatocytes and round spermatids. TH3 is synthesized in spermatogonia/early spermatocytes but not in pachytene spermatocytes, in contrast to the other testis-specific histones TH2A, H1t, and TH2B. Therefore, TH3 may have a different role in spermatogenesis than the other testis-specific histone variants (Trostle-Weige et al. 1984). 6.5.3.4

H1 Variants

The linker histone H1 is necessary for stabilization of higher order structures of chromatin, by virtue of its association with DNA-connecting nucleosomes. Proteins of the H1 family greatly diverge in their structures, and several germ cell-specific variants have been identified. The C termini of such variants are highly specific and this feature may influence their binding to chromatin (Hendzel et al. 2004). Among these germ line-specific variants, the testis-specific H1t makes up almost 40% of the total histone H1 content in pachytene spermatocytes, and it is the only one expressed from the middle of the pachytene stage (Pradeepa and Rao 2007). The other H1 variants appear later in spermatids. It has been shown that H1t isolated from rat testis is a poor condenser of DNA and chromatin (Khadake and Rao 1995). Indeed, it lacks the 16-mer (S/TPKK) motif known to have DNA condensation properties, and the presence of H1t in pachytene chromatin seems to loosen the chromatin structure even in higher order structures (Pradeepa and Rao 2007). However, homozygous H1t / mice are fertile and show no obvious defect due to the lack of this histone (Fantz et al. 2001). One oocyte-specific H1 variant has been described (H1Foo, also called H1oo or H1X). Differently from the various male H1 variants, H1Foo is heavily polyadenylated and the protein is extremely rich in lysines, which are potential sites for methylation or acetylation. It is found from the germinal vesicle stage to the MII oocyte stage (Kimmins and Sassone-Corsi 2005). This variant is known to play a role in gene silencing (Clarke et al. 1998).

6.5.4

Noncoding RNAs During Spermatogenesis

Several classes of RNAs that are not translated into proteins (noncoding RNAs) have been identified [reviewed in (Khalil and Wahlestedt 2008)]. Noncoding RNAs have been classified based on their size and function. Among them, Piwi-interacting RNAs (piRNAs) are 26-34 nucleotide long noncoding RNAs that are exclusively

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expressed during spermatogenesis in mammals (Girard et al. 2006; Grivna et al. 2006). Different from miRNAs and siRNAs, the biogenesis of piRNAs is DICERindependent (Girard et al. 2006). The proteins of the Piwi-family, MIWI (Deng and Lin 2002), MIWI2 (Carmell et al. 2007), and MILI (Kuramochi-Miyagawa et al. 2004), are all essential for male fertility, suggesting an important role for piRNAs in mammalian spermatogenesis. Interestingly, the functions of Piwi proteins seem not to overlap since the invalidation of each gene results in defects at different stages of spermatogenesis (Khalil and Wahlestedt 2008). Miwi / mice display spermatogenic arrest at the beginning of the round spermatid stage (Deng and Lin 2002) and MIWI has been shown to interact with the translational machinery and with piRNAs to regulate spermatogenesis (Grivna et al. 2006). In contrast, Miwi2 / mice display a defect in meiotic progression during the early prophase of meiosis I and a marked and progressive loss of germ cells with age. These phenotypes may be linked to inappropriate activation of transposable elements (Carmell et al. 2007). In Mili / mice, spermatogenesis is blocked completely at the early prophase of the first meiosis, from zygotene to early pachytene, and males are sterile. However, primordial germ cell development and female germ cell production are not disturbed (Kuramochi-Miyagawa et al. 2004). These findings suggest an important role for piRNAs and their interacting proteins in the process of mammalian spermatogenesis. Genetic ablation of a murine homolog of Maelstrom (MAEL), a Drosophila protein localized in the nuage, a perinuclear germ cell hallmark (Soper et al. 2008), demonstrated that it is essential for spermatogenesis. Moreover, study of Mael / testes suggests a functional relationship between MAEL and the Piwi proteins MIWI2 and MILI. Indeed, Mael, Mili, and Miwi2 / spermatocytes fail to complete meiotic prophase and to assemble functional SC. Interestingly, Mael / and Miwi2 / spermatocytes show identical atypical morphologies (not reported for the Mili mutant) prior to their elimination by apoptosis (Carmell et al. 2007). Furthermore, Mael / and Miwi2 / testes exhibit a common pattern of activation of transposable elements (LINE, Long interspersed element, and IAP, Intracisternal A particle), whereas expression of IAP is higher in Mili / testes (Aravin et al. 2007; Carmell et al. 2007; Kuramochi-Miyagawa et al. 2008). These data suggest that MAEL plays a more prominent role in MIWI2-dependent processes, possibly at a post-piRNA production step.

6.6

6.6.1

How Could It Work: Are Meiotic Recombination Initiation Sites Defined by a Specific Combination of DNA Sequence and Chromatin Features? Methylation of DNA Inhibits the Formation of Crossovers

In the fungus Ascobolus immersus, DNA methylation of a recombination hotspot on the two homologous chromosomes has been shown to strongly inhibit CO formation compared with the unmethylated hotspot (Maloisel and Rossignol 1998). When

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only one allele is methylated, a large excess of CO events show gene conversion of the methylated region, consistent with a stronger reduction of initiation on the methylated allele than on the unmethylated one. However, CO formation is still 50fold reduced at the hemimethylated hotspot, suggesting that DNA methylation might not only interfere with initiation itself but also with the stability of recombination intermediates or other post-DSB steps (Lichten 2008). Highly recombinogenic regions are thus expected to be less methylated than low recombinogenic ones. This should lead to a higher stability of cytosines in regions of high recombination activity, because, following deamination, methyl-cytosines are converted into thymines. The positive correlation between recombination rate and CpG content (outside CpG islands) at the Mb scale observed in humans could result from such methylation effect (Buard and de Massy 2007).

6.6.2

Sequence Features and Open Chromatin at Recombination Initiation Sites

On the basis of DSB mapping in S. cerevisiae, Spo11 seems to have no or little DNA sequence-specificity at the nucleotide resolution (de Massy et al. 1995; Liu et al. 1995; Diaz et al. 2002), and the evolutionary conservation of the Spo11 DNA binding domain suggests that this property should also apply to mammalian SPO11. However, in humans, sequence comparison of more than 22,699 CO hotspots, the location of which was deduced from LD analyses, revealed that a 13 mer DNA motif CCNCCNTNNCCNC is shared by 40% of these hotspots (Myers et al. 2008). Chromatin accessibility is a crucial feature of initiation sites in S. cerevisiae and S. pombe (Lichten 2008), and DSBs occur preferentially in open chromatin (Ohta et al. 1994; Wu and Lichten 1994). H3 and H4 acetylation and H3K4Me3 have been, respectively, detected at DSB sites either by local analysis of individual sites in S. pombe (Yamada et al. 2004) or by genome-wide analysis in S. cerevisiae (Borde et al. 2009). Several mutations in HMTs and HATs have been shown to lead to decreased or increased DSB activity, suggesting a role for histone modifications in the initiation of meiotic recombination (Sollier et al. 2004; Yamashita et al. 2004; Mieczkowski et al. 2007; Hirota et al. 2008; Merker et al. 2008). Chromatin remodeling factors, such as Snf22 in S. pombe, are also involved in initiation (Yamada et al. 2004), probably by rendering the initiation sites flagged by histone modifications accessible to the recombination machinery. H3K4Me3 is a well-known mark of open chromatin, which is highly enriched at actively transcribed promoters in most eukaryotes, and the HMT SET1 is responsible for this histone modification in S. cerevisiae. Although S. cerevisiae recombination initiation sites are mostly found in promoter regions, a specific enrichment for H3K4Me3 at DSB sites that is not correlated with transcription activity has been shown (Borde et al. 2009). One important open question, however, is to understand how SET1 activity is specifically targeted to these sites. Analysis of initiation of

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meiotic recombination in S. pombe has shown that recombination hotspots could be induced by the binding of the Atf1/Pcr1 complex, a transcription factor that has a strong DNA binding specificity (a heptamer motif) and recruits the histone modifiers and chromatin remodelers described above. As previously mentioned (see Sect. 6.5.2), H3K4Me3 enrichment correlates with initiation of recombination not only in S. cerevisiae but also in the mouse (Buard et al. 2009). In contrast to yeast, CO hotspots are not found in gene promoters in mammals and no transcriptional activity is correlated with the H3K4Me3 enrichment found at active mouse hotspots (Buard et al. 2009). The recent identification of the HMT PRDM9, which is involved in defining mouse and human initiation sites, illustrates how genetic and epigenetic information can be combined for such specification. On proximal mouse chr. 17, a 4.6 Mb region has been identified that contains a genetic element controlling recombination in trans (Grey et al. 2009). According to the subspecies’ origin of this chromosomal region (M. musculus molossinus or M. musculus domesticus), the recombination activity of two individual hotspots and their H3K4Me3 enrichment are changed as well as the genome-wide distribution of recombination. This finding led to propose that two different alleles of a genetic element included in this region were able to activate distinct sets of hotspots scattered over the genome. Among the 58 genes, the eight noncoding RNAs and hundreds of piRNAs annotated in this critical 4.6 Mb region, Prdm9 (also called Meisetz) stands out as a strong candidate for regulating recombination. Indeed, PRDM9 is a meiosis-specific HMT able to trimethylate H3 at K4, and Prdm9 / mice are sterile (Hayashi et al. 2005). The predicted protein harbors three domains: the methyl transferase PR/SET domain, a KRAB domain, possibly involved in protein protein interactions, and a C2H2 zinc finger domain, known as a DNA binding module. Owing to hundreds of peptide DNA interactions tested in vitro and gathered in a database, the DNA motif that would bind to a given series of zinc fingers can be predicted (Fu et al. 2009). The zinc finger domain of PRDM9 has experienced accelerated evolution in metazoan as exemplified in closely related primate or rodent species (Myers et al. 2010; Oliver et al. 2009). For instance, extensive variations between the zinc finger tandem repeats of the M. m. molossinus and M. m. domesticus alleles have been observed: not only the number of zinc finger repeats but also the pattern of successive variant repeats differ between the two strains (Baudat et al. 2010; Mihola et al. 2009). The variations of the 84 bp zinc finger repeats consist mainly in nonsynonymous substitutions at the three codons involved in contact with DNA. Human PRDM9 presents the same features, including this tandem repeat polymorphism among different individuals (Baudat et al. 2010; Oliver et al. 2009; Parvanov et al. 2010). Strikingly, the DNA motif predicted to bind to the zinc finger domain of the two major Prdm9 human alleles found in Europeans matches the 13 mer consensus sequence shared by 40% of LD-based hotspots (Baudat et al. 2010; Myers et al. 2010). An association analysis was then conducted using a set of 82 families, in which previous genome-wide high resolution SNP genotyping permitted to determine how many CO transmitted by a given parent would fall within a LD-based hotspot (Coop et al. 2008). Three Prdm9 alleles segregated in these

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families: the two major alleles (A and B) were predicted to bind to DNA motifs matching the 13 mer consensus, whereas the third allele (I) was predicted to bind to a highly divergent motif. About 60% and 70% of the CO produced by parents of AA or AB genotype fell within LD-based hotspots, respectively. In contrast, less than 20% of the CO of AI parents did so (Baudat et al. 2010). The association between Prdm9 genotype and hotspot usage was so strong (p < 10 11) to make of Prdm9 a major player in the regulation of the genome-wide distribution of recombination in humans, most probably by defining a set of DNA sites as hotspots for each Prdm9 zinc finger allele. This idea is further strengthened by the direct demonstration of the specific interaction in vitro between the PRDM9A isoform and an oligonucleotide matching the 13 mer hotspot consensus motif and reciprocally between PRDM9I and its predicted sequence target (Baudat et al. 2010).

6.6.3

A Model: Chromatin Writers and Readers as Major Determinants of Meiotic Recombination

Altogether, the lines of evidence presented above suggest a model where a combination of DNA features and epigenetic modifications is necessary to specify initiation sites of recombination in mammals (Fig. 6.6). A given allelic isoform of PRDM9 could have thousands of potential binding sites with different affinities and accessibilities in the genome. In a given cell, at or before leptonema, PRDM9 may bind to a subset of these sites through its specific C2H2 zinc finger domain. The PR/SET domain of PRDM9 would then modify the closest nucleosomes by adding specifically a third methyl group on H3K4 only if this lysine is already dimethylated (Hayashi et al. 2005). Other chromatin modifiers (and/or other proteins), including the not yet identified HATs, would then be recruited, maybe by interacting with PRDM9 through its KRAB domain, and would add supplementary marks. A specific combination of histone modifications, including the prominent H3K4Me3, would then constitute a substrate for one (or several) protein(s) necessary for DSB formation. This putative protein may be a chromatin remodeling factor that can, on one hand, read the combination of histone modifications found specifically at the initiation site and, on another hand, recruit SPO11 to this site either directly or via the recruitment of a SPO11-interacting partner. DSBs are then recognized by the MRN complex (consisting of MRE11, RAD50 and NBS1) and by kinases, which induce H2AX phosphorylation. DSBs are then repaired while H4 is acetylated either by a HAT, such as TIP60, or by incorporation of newly synthesized (and acetylated) histones during nucleosome replacement at the repaired DSB site (Polo et al. 2006). Many unknown features of this model of chromatin changes associated with recombination remain to be tested. Major efforts are now focused on identifying the putative reader(s) of the histone modifications present before DSB formation. Other phenotypes have been reported associated with Prdm9 deletion and with Prdm9 allelic variability. PRDM9 has been first thought to be a meiosis-specific

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Fig. 6.6 Model of chromatin based specification of meiotic recombination initiation site. PRDM9 binds to a DNA motif through its zinc finger domain and adds a third methyl group onto the dimethylated K4 of histone H3 of the adjacent nucelosomes. Nucleosomes are indicated as beige cylinder and histone posttranslational modifications as red balls. Readers, or proteins with affinity to H3K4Me3, and/or to PRDM9 (KRAB/PR SET domain) are recruited, and may modify further the structure of the chromatin. Spo11, or a protein interacting with Spo11, recognizes one or several of the specific features of the DNA and chromatin at this site, binds and promotes DSB formation

transcription factor, due to its H3K4 trimethyl transferase activity and due to the observed meiotic arrest and alteration of transcription levels of at least one gene in Prdm9 / mice (Hayashi et al. 2005). This hypothesis has been challenged because the accelerated evolution of the zinc finger DNA binding module seems incompatible with binding to conserved promoters (Oliver et al. 2009). However, splicing variants of Prdm9 mRNA encoding a truncated protein with a SET domain but without the zinc finger domain have been detected (Hayashi et al. 2005) and they could have a function in the regulation of transcription. Variability of the PRDM9 zinc finger domain has been associated with hybrid sterility in the mouse, although the mechanism involved in this phenotype remains elusive (Mihola et al. 2009). The product of a hybrid sterility gene in Drosophila has been shown to bind to heterochromatic DNA only in sterile hybrid males,

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leading to heterochromatin decondensation (Bayes and Malik 2009). The same mechanism has been proposed for PRDM9: evolution of its zinc finger module towards recognition of the rapidly evolving centromeric satellite DNA would be counter-selected (Oliver et al. 2009). However, this scenario against the co-evolution of the two (DNA and protein) sequences does not explain the extraordinary variability of PRDM9 zinc fingers within a given species, as observed in humans (Baudat et al. 2010; Parvanov et al. 2010). Furthermore, this scenario does not take into account the fact that, in mouse, a responder element located on the X chromosome is also necessary for hybrid sterility to occur (Mihola et al. 2009). The characterization of this responder element might shed light on the mechanism of hybrid sterility mediated by PRDM9. PRDM9 has also been reported to be involved in human fertility with base substitutions apparently associated with infertility in cohorts of sterile men. However, the restricted number of patients genotyped in one report (Miyamoto et al. 2008) and the fact that the base substitutions suggested to be associated with infertility in the second report (Irie et al. 2009) are in fact nonsynonymous variant of repeat polymorphisms of the zinc finger array also observed in fertile men (Baudat et al. 2010), strongly suggests that larger association analyses are needed to confirm these hypotheses. In summary, the mechanisms of action of PRDM9 in specifying meiotic initiation sites and in building reproductive barriers remain to be characterized. Whether Prdm9 is subject to accelerated and positive selection in relationship to its role in speciation or, instead, whether change of hotspot usage is also a positive selection force increasing the pace of its rapid evolution remains to be addressed.

6.7

Perspectives

The recent advances in the study of initiation of meiotic recombination have generated a large number of questions directly linked to the functional relationships between epigenetic marks and recombination. What epigenetic modifications are found at recombination hotspots? How general are they and/or how many different types of hotspots can be identified based on the chromatin marks? Which of these features are specific to initiation sites? What are the enzymes involved in writing and reading these marks? Similar issues are raised by the potential roles of epigenetic marks for the repair of meiotic DSBs. In principle, one of the best ways to get some of this information would be to obtain a genome-wide description of these features. Some of the tools, such as next-generation sequencing, are already available; however, one of the limiting steps is the accessibility to suitable biological material, and thus more investment should be put into getting the required samples with the amount and purity needed. For instance, understanding male/female differences would require overcoming the scarcity of female meiotic germ cells. As can be seen from the analysis of several mouse mutants, defects in several epigenetic modifiers cause strong phenotypes, some of them specific to male germ

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cell meiosis. It would be interesting to understand better these defects, to differentiate those due to direct alteration of recombination from those where modification of gene expression indirectly alters recombination. In particular, this could help identifying genes that might be involved in pathologies of human reproduction. Acknowledgments We thank F. Baudat and C. Grey for discussions and comments on the manuscript. P. B is supported by a PhD grant from the French MENRT. This work was funded by grants from CNRS, EDF, ANR 06 BLAN 0160 01, ANR 09 BLAN 0269 01, and ARC3939 to BdM.

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Chapter 7

Meiotic Pairing of Homologous Chromosomes and Silencing of Heterologous Regions Sam Schoenmakers and Willy M. Baarends

Abstract In meiotic prophase, homologous chromosomes pair and recombine, which ensures correct segregation of the homologs at the metaphase to anaphase transition of the first meiotic division. As a first step, early in meiotic prophase, DNA double-strand breaks (DSBs) are introduced by SPO11 at hundreds of sites throughout the genome. In mammals, formation and repair of these breaks is essential for the pairing process. The sex chromosomes, X and Y, are largely heterologous and pair only in the small pseudoautosomal homologous regions. The large heterologous regions show delayed repair of DSBs and asynapsis, which triggers meiotic sex chromosome inactivation (MSCI), leading to the formation of the transcriptionally silenced XY body. Initiation of XY body formation is known to require phosphorylation of H2AX, which is also associated with damage-induced DSB repair in somatic cells. MSCI is considered as a specialized form of a general meiotic silencing mechanism that also silences autosomal unsynapsed chromatin during male and female meiotic prophase: meiotic silencing of unsynapsed chromatin (MSUC). Whereas the XY pair remains always largely unsynapsed and inactivated, autosomal nonhomologous regions frequently show heterologous synapsis and escape from inactivation. Hence, MSUC is less efficient than MSCI. However, if MSUC occurs, MSCI is incomplete, indicating that the two mechanisms are functionally interacting. Herein, we briefly review how X and Y evolved from a pair of autosomes, in relation to X-chromosomal dosage compensation in females and MSCI in males.

S. Schoenmakers Department of Reproduction and Development, Erasmus MC PO Box 2040 3000 CA Rotterdam, The Netherlands and Department of Obstetrics and Gynaecology, Erasmus MC PO Box 2040 3000 CA Rotterdam, The Netherlands e mail: [email protected] W.M. Baarends (*) Department of Reproduction and Development, Erasmus MC PO Box 2040 3000 CA Rotterdam, The Netherlands e mail: [email protected]

University Medical Centre,

University Medical Centre,

University Medical Centre,

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Next, we describe current ideas about initiation and maintenance of MSCI and MSUC. We also discuss MSCI in birds, a species with female heterogamety. Insight in the mechanism of meiotic silencing is highly relevant for our understanding of male infertility associated with translocations, particularly when these occur between an autosome and the X. Furthermore, we anticipate that dysregulation of MSCI may impact on early embryonic development. Keywords DNA repair Meiosis Meiotic Sex Chromosome Inactivation (MSCI) Meiotic Slencing of Unsynapsed Chromatin (MSUC) Synaptonemal complex formation X chromosome XY body Y chromosome

7.1

Introduction

The presence of the largely nonhomologous, or heterologous, X and Y chromosomes in the genome of mammalian males during meiosis is associated with specific problems. The Y chromosome is much smaller than the X chromosome, and carries very few genes. The human X chromosome harbors at least 1,000 genes, and many of these are essential. The X and Y share homology in the short so-called pseudoautosomal regions (PARs) that contain approximately 28 genes in humans (reviewed in Helena Mangs and Morris 2007). Despite the current differences in length and gene content, X and Y have evolved from a pair of homologous autosomes (reviewed in Graves 2006). DNA sequence analyses have revealed ancestral homology between genes on X and Y and have confirmed the evolutionary relationship between X and Y (Lahn and Page 1999; Handley et al. 2004). Below, we will address how X and Y have become so divergent, and how this has complicated regulation of X and Y pairing, their recombination, and gene expression during meiosis. Meiosis encompasses two divisions: meiosis I and II. Meiosis I is a so-called reductional division; homologous chromosomes are separated, generating two nuclei, each with a single set of replicated chromosomes. Meiosis II is more similar to mitosis; the sister chromatids are pulled apart from each other. The net result of the two successive divisions is the formation of four functional haploid nuclei in males. Meiosis is initiated continuously in adult males, but in females, all oocytes enter meiotic prophase before birth. The oocytes halt at the end of the first meiotic prophase, with an arrest in the diplotene (dictyate) stage until just prior to ovulation, when the first meiotic division generates the small first polar body and the secondary oocyte. The ovulated secondary oocyte is blocked at metaphase II. Fertilization triggers the metaphase II to anaphase II transition, and a second polar body is formed, as well as the female haploid pronucleus. The formation of small polar bodies allows the final oocyte to maintain maximal cytoplasmic volume containing many maternal factors required for the first cleavage divisions. This specific regulation

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of meiosis in females results in the formation of one single gamete and two polar bodies. The most remarkable achievement of meiosis is the separation of homologous chromosomes. This process requires that homologous chromosomes can find each other, align over their full length, and stick together until the metaphase to anaphase transition of the first meiotic division. Moreover, in order to ensure that complete genome sets are separated during the metaphase-to-anaphase I transition, pairing between nonhomologous chromosomes needs to be prevented. One important aspect of the first meiotic prophase is meiotic recombination. This involves the induction and repair of DSBs, which leads to exchange of genetic information between homologous chromosomes. Errors in this mechanism lead to an unequal number of chromosomes (aneuploidy) in the resulting gametes, which is mostly lethal to the developing embryo, but can be viable when it concerns a small chromosome (trisomy 21) or the sex chromosomes. With this review, we aim to highlight known and putative links between meiotic DNA double-strand break repair, homology recognition, synapsis, and detection and silencing of heterologous regions. Basic insight in these processes will contribute to our understanding of the causes of male infertility and may help to estimate the possible risks for the offspring, resulting from the application of assisted reproduction using sperm from males in which MSCI was impaired or MSUC was activated.

7.2

Evolution of Sex Chromosomes

Several events and mechanisms have contributed to the present heteromorphic length and gene content of the sex chromosomes. Since there is no situation in the life cycle of mammals in which two Y chromosomes are present in the same nucleus, it also never recombines with another Y chromosome. In contrast, an X chromosome can pair and recombine with another X chromosome during meiotic prophase in XX oocytes. The Y does recombine with the X chromosome in spermatocytes, but solely in the short homologous pseudoautosomal regions (PARs). In an early mammalian ancestor, the initiation of the progressive differentiation of X and Y most likely started with the evolution of the dominant male sexdetermining gene SRY (Koopman et al. 1991), on one member of a pair of autosomes, initiating the formation of a sex chromosome pair (Ohno 1967). Meiotic recombination within the region containing SRY was repressed, consolidating the new XX/XY sex-determining system, with a proto-X and proto-Y. How this is initially achieved is not known, but several mechanisms have been suggested (reviewed in Bergero and Charlesworth 2009). For example, if SRY was located in a small reversed region of the genome, this could have contributed to the repression of recombination due to meiotic pairing problems in that region. In addition, or alternatively, other male beneficial genes incorporated next to SRY might allow selection to act on this male-specific region as a whole, and this would be favored by suppression of recombination in the whole region between the proto-Y and proto-X,

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to keep SRY in one functional unit with other male beneficial genes. With time, the region of suppressed recombination has expanded and now encompasses the whole male-specific region of the Y (MSY), which is most of the Y chromosome. The complete lack of meiotic recombination for the MSY has resulted in the gradual loss of the majority of functional genes that were present on the original proto-Y chromosome through various selection mechanisms that act during evolution (reviewed in Bachtrog 2006). In addition, accumulation of repetitive sequences on Y has led to the formation of large heterochromatic regions (Lippman et al. 2004). Complete degeneration of the Y chromosome has been predicted to occur in approximately 14 million years (Graves 2006). However, recent sequence analyses of multiple single-copy Y chromosomal genes in more than 100 men of diverse genetic background has shown that there is very little variation in the protein coding regions of these genes; implying that natural selection effectively operates to preserve Y chromosome functionality in man. In addition, human Y-linked genes have been surprisingly well conserved as compared to chimpanzee Y-linked genes. Comparative sequencing showed that no degenerate gene loss in humans has occurred during the 6 MY that separates the two species (Hughes et al. 2005, 2010). Hence, the widely suggested rate of Y chromosomal decline, with 4 5 single-copy genes disappearing every one million years, does not seem applicable for the human Y chromosome. Total degeneration of the Y chromosome may also be prevented, or at least delayed, via intra-Y-homolog recombination between palindromic regions (Rozen et al. 2003), containing multicopy genes, particularly involved in spermatogenesis, although this is a vulnerable system, described as an Achilles heel (Lange et al. 2009). Stepwise fusion of the X chromosome with autosomal chromosomes, the so-called different evolutionary strata, and a series of Y-inversion events have also added to the current heteromorphic state of the sex chromosomes (Lahn and Page 1999; Graves 2006). Finally, acquirement of individual genes by autosome-to-Y and autosome-to-X transpositions also contributed to XY differentiation. Since most of these transposed genes and the remaining functional genes on Y have a testisspecific expression, the Y chromosome seems to have masculinized, and its functional regions exist mainly out of male-beneficial genes (Skaletsky et al. 2003). Mammalian XY differentiation began approximately 200 million year (MY) ago and the last major rearrangement on the human X chromosome occurred 30 50 MY ago (Lahn and Page 1999). The oldest part of the Y chromosome contains the sex-determining gene SRY (distal of Yp) and the PARs, where X and Y can still recombine. Where the Y chromosome has evolved into a male-beneficial gene-enriched chromosome, the X chromosome has become enriched for reproduction- and brain developmental-beneficial genes (Khil et al. 2004). The evolutionary changes that occurred on X and Y, generating huge differences, complicate the pairing and recombination process during male meiotic prophase. The heterologous X and Y need to associate, to ensure correct separation of X and Y during the first metaphase-to-anaphase transition.

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Homologous Chromosome Pairing in Meiotic Prophase

In meiotic prophase, all homologous chromosomes need to recognize each other before they can proceed with alignment, pairing, synapsis, and recombination. Alignment and initiation of pairing occurs following the final S phase in leptotene, pairing progresses during zygotene, and is followed by synapsis, which is completed in pachytene nuclei. Synapsis is achieved by the formation of the synaptonemal complex (SC) between the chromosomal axes of the paired homologous chromosomes. X and Y also need to recognize each other, but at the same time, pairing and recombination of the heterologous regions needs to be avoided. The exact mechanism by which homologous chromosomes recognize each other and prevent association between nonhomologous chromosomes in meiotic prophase in mammals is still enigmatic, although two important processes are known to be essential for chromosome pairing: bouquet formation and the generation and repair of meiotic DSBs.

7.3.1

Bouquet Formation

After DNA replication in preleptotene, the telomeric repeats of all chromosomes associate with the nuclear envelope in leptotene. They aggregate and cluster near the centrosome, forming the meiotic bouquet (reviewed in Ding et al. 2007; Bhalla and Dernburg 2008). Specific components of the inner and outer nuclear membrane have been shown be essential for bouquet formation and (homologous) chromosome pairing in mouse (Ding et al. 2007). Telomeric movements along the nuclear membrane enable the occurrence of transient interactions between chromosomes (Ding et al. 2004), possibly contributing to homology recognition and pairing. Also, specific unique (DNA) sequences or chromatin components such as yet unknown pairing centers and centromeres may contribute to the initial homologous interactions (Bhalla and Dernburg 2008).

7.3.2

Meiotic DNA Double-Strand Break Formation

Simultaneously with formation of the bouquet configuration of chromosomes at leptotene, several hundreds of meiotic DSBs are induced by the topoisomeraselike enzyme SPO11 throughout the mammalian genome (Keeney et al. 1997). The presence of DSBs leads to the activation of the checkpoint kinase ATM, which phosphorylates histone H2AX (forming gH2AX) in the regions surrounding the breaks (Rogakou et al. 1999; Mahadevaiah et al. 2001; Bellani et al. 2005). In somatic cells, gH2AX also marks damage-induced DSBs (Rogakou et al.

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1999). Formation and repair of meiotic DSBs is essential to achieve complete and correct chromosome pairing (Baudat et al. 2000; Romanienko and CameriniOtero 2000).

7.3.3

Synaptonemal Complex Formation

During formation and repair of meiotic DSBs, a proteinaceous structure is assembled between the homologs. This structure, named the SC, physically connects the homologous chromosomes, facilitates and stabilizes synapsis, and enables meiotic recombination (reviewed in Heyting 1996). The assembly of the SC starts with the formation of the axial elements (AE) at leptotene, when SYCP2 and SYCP3 proteins form filaments along the chromosomal axes of the connected sister chromatids (reviewed in Costa and Cooke 2007). During this phase, specific sites on the chromosomes, such as “sticky” centromeric and pericentric heterochromatic regions in respectively budding yeast and Drosophila (reviewed in Bhalla and Dernburg 2008), and pairing initiating regions in Caenorhabditis elegans (MacQueen et al. 2005), could establish transient interactions and stabilize the aligning chromosomes (reviewed in Bhalla and Dernburg 2008). The sites of meiotic DSB repair (see below) seem to form the first physical connections between homologous chromosomes and could function as initiation sites of synapsis, at least in budding yeast (Bhalla and Dernburg 2008). However, the clustered and paired telomeres at the bouquet stage could also function as starting points of synapsis. This is suggested by the observation that synapsis during meiosis in humans appears to initiate from both ends of the SC, proceeding to the middle of the aligned chromosomal axes (Brown et al. 2005). Apart from SYCP2 and SYCP3, the chromosomal axes also contain cohesin, the protein complex that holds sister chromatids together. The meiosis-specific cohesin proteins REC8 and SMC1b colocalize with the AE component SYCP3 (Eijpe et al. 2003). Analysis of mice deficient for genes encoding meiosis-specific components of the cohesin complex have shown that they are required not only to keep the sister chromatids connected during the first metaphase-to-anaphase transition but also to regulate the length of the SC and synapsis (Revenkova et al. 2004; Xu et al. 2005). Upon homologous chromosome alignment, the transverse element (TE) protein SYPC1, in cooperation with the central element (CE) components SYCE1, SYCE2, and TEX12, connects the homologous AEs (reviewed in Costa and Cooke 2007). The appearance of SYCP1 defines the moment of synapsis, and the AEs are now termed lateral elements (LEs) in the completed SC. Meiotic chromosome pairs in spermatocytes from Sycp1 / , Syce1 / , Syce2 / , and Tex12 / mice show normal alignment, but do not synapse and fail to complete repair of meiotic DSBs. In addition, the X and Y chromosomes do not pair in these mutants (de Vries et al. 2005; Bolcun-Filas et al. 2007; Hamer et al. 2008; Wojtasz et al. 2009). Also in SYCP2 and SYCP3 deficient mice, chromosome pairing is severely affected (Yuan et al. 2000; Yang et al. 2006). It is important to note that the severity of the

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meiotic phenotype that is observed upon mutation of genes involved in meiotic recombination or synaptonemal complex formation is frequently different between males and females (Hunt and Hassold 2002; Morelli and Cohen 2005). For example, In Sycp3 knockout females, meiosis proceeds up to the diplotene stage, but many oocytes are lost during the first week of postnatal development (Yuan et al. 2002). Some oocytes even survive until adulthood, and embryos obtained from such oocytes are frequently aneuploid (Yuan et al. 2002). This and other sex-differences observed for the functions of genes involved in meiosis are thought to be due at least in part to a sex-differential regulation of checkpoints (Wang and Hoog 2006) (see also 7.3.5).

7.3.4

Meiotic DNA Double-Strand Break Repair

In mitotic cells, DSBs can be repaired via nonhomologous end-joining (NHEJ) or via homologous recombination (HR) pathways. For the latter process, an intact homologous template is required, for which after S phase, the sister chromatid is usually available. In meiotic prophase, expression of proteins involved in NHEJ is repressed (Goedecke et al. 1999), and all meiotic DSBs thus need to be repaired by the more accurate HR pathway. The meiotic DSB repair process is somewhat different from homologous recombination repair of damage-induced DSBs in somatic cells. First, meiotic breaks are not induced by damage, or stalled replication forks, but generated by SPO11, an enzyme that remains covalently attached to the cleavage site (Neale et al. 2005). To allow DNA repair through HR, endonucleolytic cleavage of these protein-linked DSBs is necessary to remove the attached SPO11 together with some oligonucleotides (Neale et al. 2005). In fission yeast, subunits of the MRN complex (MRE11, RAD50 and NSB1) in combination with the endonuclease Ctp1 are required for meiotic DSB processing, including cleavage and subsequent resection of the ends (Milman et al. 2009; Williams et al. 2009), yielding 30 ended single-stranded DNA (ssDNA) overhangs (Acharya et al. 2008). The MRN complex is also involved in sensing the presence of (meiotic) DSBs and aids in the localization of the kinase ATM to damage sites (reviewed in Borde and Cobb 2009). Apart from the different origin and initial processing of SPO11-induced breaks compared to damage-induced DSBs in somatic cells, homologous recombination has also been adapted for meiosis. The most important meiotic specialization of this repair pathway involves the inhibition of the use of the sister chromatid as a template for repair, thereby stimulating the search for the homologous chromosome. Several meiosis-specific DNA repair proteins and the SC contribute to the so-called interhomolog bias of the meiotic DSB-repair machinery. After the 30 ssDNA overhangs on each end of the DSB have been formed, the RecA-like strand exchange protein RAD51, essential for strand invasion of the homologous DNA,

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forms filaments on these overhangs in mitotic cells (Haaf et al. 1995). Using immunocytology, the formation of RAD51 filaments can be observed as foci. In meiotic cells, not only RAD51 but also its meiosis-specific paralog DMC1 assembles on the processed ssDNA ends (Tarsounas et al. 1999), and these RAD51/ DMC1 filaments initiate the search for homology (Kinebuchi et al. 2004, reviewed in Neale and Keeney 2006). From these two repair proteins, DMC1 is believed to be the one that promotes repair via the homolog (Bishop et al. 1992; Schwacha and Kleckner 1994). When the ssDNA filament invades the dsDNA of the homolog, this brings them in close proximity to each other. The search for homology is most likely facilitated by the bouquet configuration and the active movements of the chromosomes. This whole process, occurring on several sites of DSB repair on each chromosome pair, most likely mediates correct alignment and pairing of homologous chromosomes. The importance of the generation and in particular the repair of meiotic DSBs for chromosome pairing is illustrated also by the infertility phenotypes of Dmc1 and Spo11 knockout mice (Pittman et al. 1998; Yoshida et al. 1998; Baudat et al. 2000; Romanienko and Camerini-Otero 2000). Once homology has been detected, the ssDNA with the RAD51/DMC1 filaments invades the homologous template and a brief phase of DNA synthesis occurs (reviewed in Neale and Keeney 2006). Subsequently, crossovers (actual exchange of chromosome arms) and noncrossovers (gene conversion events) are formed through separate pathways that are strictly regulated; the number of crossovers is rather constant per species and sex and highly outnumbered by the number of noncrossovers. Special mechanisms reduce the likelihood of two crossovers occurring in close proximity of each other, but they ensure that each chromosome pair, including X and Y, contains at least one obligate crossover.

7.3.5

Meiotic Checkpoint Regulation During Male Meiotic Prophase

To maintain genomic integrity, the repair process and the associated pairing of homologous chromosomes need to be tightly controlled. In budding yeast, two main meiotic cell cycle checkpoints have been identified: the replication or doublestrand-break checkpoint, coupling the completion of premeiotic DNA replication to the introduction of SPO11-induced meiotic DSBs, and the pachytene checkpoint, monitoring correct (homologous) DSB repair, meiotic recombination, and synapsis (reviewed in Hochwagen and Amon 2006). Careful analysis of many meiotic mutants in mouse have revealed that incomplete meiotic DSB repair and synapsis trigger apoptosis around midpachytene, corresponding to Stage IV of the cycle of the seminiferous epithelium in mouse (Ashley et al. 2004a, b), indicating that a checkpoint operates at this stage. It has also been proposed that the loss of spermatocytes at Stage IV is actually not due to

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checkpoint activation, but caused by failure to inactivate the X and Y chromosome (meiotic sex chromosome inactivation, see below) (Burgoyne et al. 2007, 2009). This attractive hypothesis also provides a possible cause for the sex-difference in meiotic phenotypes of many mutants, because the two X chromosomes are active during normal female oogenesis. The two related kinases ATM and ATR perform pivotal functions in the activation of cell cycle checkpoints in somatic cells. The SPO11-mediated induction of meiotic DSBs activates ATM, which enables the first meiotic wave of phosphorylation of H2AX (Bellani et al. 2005). ATR is expressed somewhat later and is required for gH2AX formation on XY body chromatin (see also below) (Bellani et al. 2005, Turner et al. 2004). Atm / spermatocytes show a failure in meiotic DSB repair and chromosome synapsis, and apoptosis is induced at the mid-pachytene checkpoint, indicating that ATM is not required for this checkpoint (Barchi et al. 2005). Spo11 heterozygosity allows progression of the meiotic prophase of Atm / spermatocytes until the first metaphase, when increased spermatogenetic apoptosis is still observed, indicating that metaphase I also functions as a checkpoint moment (Barchi et al. 2005; Di Giacomo et al. 2005). In these Spo+/ Atm / mice, reduced or delayed formation of meiotic DSBs may help ATR to compensate for the loss of ATM (Barchi et al. 2005; Di Giacomo et al. 2005). In many organisms, cell cycle checkpoint proteins (such as ATR and ATM) not only function as surveillance proteins, but are also indispensable for completion of DSB repair and meiotic recombination. Phosphorylation of the yeast meiosisspecific HORMA-domain protein Hop1 by ATR and ATM homologs (Mec1 and Tel1) is required for meiotic prophase checkpoint activation as well as for interhomolog-directed repair (Carballo et al. 2008). Recently, two mammalian homologs of the yeast Hop1 protein, named HORMAD1 and HORMAD2, have been identified (Wojtasz et al. 2009). Early in meiotic prophase, HORMAD1 and 2 accumulate on unsynapsed chromosomal axes. They are removed when the SC forms between homologous chromosomes (Wojtasz et al. 2009). As expected, HORMAD1 and 2 both remain present on the unsynapsed parts of X and Y (Wojtasz et al. 2009).

7.4

7.4.1

Sex Chromosomal Behavior During Mammalian Spermatogenesis Pairing of X and Y in Meiotic Prophase

During zygotene, synapsis between X and Y is initiated in the pseudoautosomal regions (PARs) and appears to continue along most of the length of the Y chromosome at early pachytene in mouse [Fig. 7.1 þ (Tres 1977)]. Hereafter, X and Y desynapse gradually until they only remain connected at their tips in late pachytene

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Fig. 7.1 Extensive (heterologous) synapsis between X and Y at early pachytene is followed by desynapsis and subsequent end to end association (a) overview of the different XY associations during pachytene in mouse spermatocytes. Spermatocyte spread nuclei immunostained for SYCP3 (red). The position of the XY pair is indicated by a box, and schematically depicted in the lower right corner of each image. Bar represents 10 mm. (b) Enlargement of the XY pair. Bar represents 5 mm. (c) Human spermatocyte in pachytene stained for SYCP3 (red) and gH2AX (green). Bar represents 10 mm. The higher magnifications of XY pair are shown on the right. Bar represents 5 mm

and diplotene (Tres 1977). Throughout pachytene and diplotene, X and Y reside in a specific region adjacent to the nuclear membrane, named the XY body. In human pachytene spermatocytes, the morphology of the X and Y chromosomal axes is more complicated as compared to mouse (Fig. 7.1), although extensive XY synapsis in early pachytene has also been described (Chandley et al. 1984). Partial XY synapsis is visible in immunofluorescent analyses of early pachytene nuclei, but the XY chromatin condenses rapidly thereafter, and the SC structure has a disorganized appearance (Tres 1977; de Boer et al. 2004).

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Table 7.1 Proteins involved in DNA repair and checkpoint activation enriched on the XY body Protein Function Pattern on XY Reference ATR Protein kinase involved in Axial elements Keegan et al. (1996) DNA replication, DNA and damage response, and chromatin checkpoint activation XY body Walpita et al. (1999) BLM Member of the RecQ chromatin helicase family, functions in DSB repair BRCA1 Functions in homologous Axial elements Scully et al. (1997) recombination repair of DSBs Ubiquitin ligase activity, checkpoint mediator protein BRCA2 Functions in homologous Axial elements Chen et al. (1998) recombination repair of DSBs, stimulates RAD51 filament formation DMC1 Meiosis specific paralog of Focal (X) along Tarsounas et al. (1999) RAD51 axial elements gH2AX H2AX phosporylated at XY body Tarsounas et al. (1999), S129, mediated by ATR chromatin Mahadevaiah et al. (2001), at the XY body Bellani et al. (2005) Baarends et al. (1999) H2AK119ub1 H2A ubiquitylated at K119 XY body silences chromatin and chromatin recruits DSB repair proteins HORMAD1/2 Meiosis specific HORMA Axial elements Fukuda et al. (2009), Wojtasz et al. (2009) domain containing proteins involved in DSB repair, interhomolog bias, and checkpoint activation XY body van der Laan et al. (2004) HR6B Functions in DSB and chromatin postreplication repair and is required for H2B ubiquitylation. Ubiquitin conjugating enzyme activity XY body Goedecke et al. (1999) Ku70 Involved in chromatin nonhomologous end joining pathway XY body Ahmed et al. (2007) MDC1 Early responder to DSBs facilitates ATM and chromatin MRE11 complex recruitment MRE11 Component of the MRE11 XY body Goedecke et al. (1999) complex involved in chromatin sensing and repair (continued)

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Table 7.1 (continued) Protein Function of DSBs and checkpoint signaling. Endonuclease activity RAD1 Component of the 9 1 1 complex involved in cell cycle checkpoint activation RAD18 Functions in DSB and postreplication repair pathways. Ubiquitin ligase activity. Ubiquitylates PCNA RAD51 Catalyzes homologous recombination reaction using ATP dependent DNA binding activity and a DNA dependent ATPase TOPBP1 Functions in DNA replication and DNA damage response, activates ATR 53BP1 Role in the recruitment of proteins to double stranded breaks in DNA, checkpoint mediator protein

7.4.2

Pattern on XY

Reference

Axial elements Freire et al. (1998)

XY body chromatin

van der Laan et al. (2004)

Focal (X) along Moens et al. (1997) axial elements

Axial elements Perera et al. (2004), Reini et al. (2004) and chromatin XY body chromatin

Ahmed et al. (2007)

Meiotic DSB Repair at X and Y

During the zygotene-to-pachytene transition, the XY pair becomes easily recognizable because it is the only chromosome pair with incomplete synapsis. At this stage, the number of RAD51/DMC1 foci in spermatocyte nuclei has already decreased dramatically compared to leptotene/zygotene (Moens et al. 2002). Few foci still persist on the autosomes in early pachytene, but the unsynapsed part of the X chromosome is always marked by several persistent RAD51/DMC1 foci (Moens et al. 1997, 2002). Concomitant with the gradual decrease in the number of RAD51/ DMC1 foci, the overall level of gH2AX also decreases. However, a second wave of gH2AX formation occurs specifically on the XY body and this phosphorylation is mediated by ATR (Turner et al. 2004; Bellani et al. 2005). ATR first localizes the unsynapsed parts of the XY pair, after which it also spreads to the synapsed part (Keegan et al. 1996; Moens et al. 1999; Baart et al. 2000). The unsynapsed part of the X chromosome is also marked by increased levels of persistent foci of DSB repair and checkpoint proteins (Table 7.1) until late pachytene/early diplotene,

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indicating that repair of DSBs along the X chromosome is severely delayed. Since persistent RAD51 foci on the unsynapsed region of Y in pachytene are hardly ever noticed (Moens et al. 1997), it is not known whether SPO11 does not induce breaks on this part of Y in mouse, or whether these breaks are somehow more rapidly repaired on the Y compared to the X chromosome. Due to the repetitive nature of the Y chromosome, such repair might occur via intrachromosomal recombination during zygotene, simultaneously with homologous recombination repair on autosomes (Skaletsky et al. 2003; Lange et al. 2009). In addition to several DSB repair and checkpoint proteins, the postreplication DNA repair pathway proteins HR6A, HR6B, and RAD18 also accumulate on the X and Y chromosomes during pachytene (van der Laan et al. 2004). HR6A and HR6B encode two very similar mammalian homologs of the ubiquitin-conjugating enzyme RAD6 in yeast (Koken et al. 1991) and form a complex with the ubiquitin ligase RAD18 (Xin et al. 2000). In yeast, the RAD6-RAD18 complex is required for the ubiquitylation of the sliding clamp protein PCNA (Hoege et al. 2002), which forms a trimeric ring around the DNA and is indispensable for DNA replication. HR6A and HR6B are also involved in the ubiquitylation of the histone H2B in yeast and mammalian cells (Sung et al. 1988; Kim et al. 2009). Although HR6A and HR6B perform redundant essential functions in somatic cells (Roest et al. 2004), HR6B is specifically required during meiotic and postmeiotic male germ cell development (Roest et al. 1996). Around mid-diplotene, the DNA repair proteins and gH2AX have disappeared from the XY body, indicating that the breaks have been repaired. At this stage, repair via the sister-chromatid may no longer be blocked. Alternatively, or in addition, NHEJ may have been reactivated at this stage and participate in repair of the persistent meiotic DSBs (Ahmed et al. 2010). In contrast to delayed DSB repair on the unsynapsed parts of X and Y, repair of DSBs that localize to the pseudoautosomal region of the XY pair appears to occur with normal or perhaps even accelerated timing (Anderson et al. 1999). RAD51 foci do not persist in this region of X and Y, and the single obligate crossover appears to be formed slightly ahead of the crossovers on the autosomes (Anderson et al. 1999),

7.4.3

Meiotic Sex Chromosome Inactivation

During the consecutive stages of spermatogenesis, X- and Y-linked genes show three different expression profiles. In mitotically dividing spermatogonia, genes from the X and Y chromosomes are actively transcribed, and male germ-cell specific genes have been found to be overrepresented on the X chromosome in these cell types (Wang et al. 2001). However, Namekawa et al. (2006) reported that in type A and B spermatogonia, the Y chromosome appears to be at least partially inactive, based on results from RNA FISH analyses.

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Upon entrance into meiotic prophase, transcription of genes located on the X chromosome starts to decrease (Khil et al. 2004). Concomitant with completion of synapsis between all autosomes around the zygotene-to-pachytene transition, MSCI is initiated and the largely unsynapsed X and Y form the inactivated XY body (or sex body) in the periphery of the nucleus (Solari 1974). The XY body remains visible as a more or less separate DAPI domain until the end of meiotic prophase. Immunocytochemical analyses have shown that the XY body is visibly depleted for RNA polymerase II from midpachytene onwards (Namekawa et al. 2006; Turner et al. 2006). In addition, microarray analyses have shown that the average RNA expression level of X chromosomal genes is very low in spermatocytes as compared to spermatogonia fractions (Namekawa et al. 2006). However, many X-linked miRNA genes escape MSCI since they remain actively transcribed by RNA polymerase II throughout meiosis (Song et al. 2009). How the transcription of miRNAs is achieved in the XY body is unclear; perhaps specific factors that bind to miRNA gene promoters allow RNA polymerase to transcribe these genes. The amount of RNA polymerase II that is actually involved in this process may be low, explaining why the whole XY body appears to be devoid of RNA polymerase II. The X chromosome contains many genes that perform functions that are essential for cell viability. For several of these genes, retroposed intronless copies have emerged on autosomes during the course of evolution, to compensate for the loss of transcription from the X-encoded variant because of MSCI, and these autosomal genes mostly show testis-specific expression (Wang 2004). The ATR-mediated phosphorylation of H2AX is the earliest known histone modification on the XY body and this modification has been shown to be required for the initiation of MSCI (Fernandez-Capetillo et al. 2003). In addition, unknown chromatin components become sumoylated around late zygotene (Vigodner and Morris 2005), and this is followed by ubiquitylation of H2A on the XY (Baarends et al. 1999) and deposition of histone macroH2A1.2 (Fernandez-Capetillo et al. 2003) around early pachytene. In pachytene, the X and Y chromosomes also undergo a series of histone modifications, such as deacetylation of H3 and H4 (Khalil and Driscoll 2006), most likely contributing to transcriptional silencing of the XY body. In addition, a global nucleosome replacement occurs around midpachytene (van der Heijden et al. 2007). The exact functions of these modifications and proteins are not known, but they could play a role in the maintenance of MSCI. So far, a few knockout mouse models have been described in which initiation of MSCI is perturbed. In the complete absence of formation of meiotic DSBs, as occurs in Spo11 / mice, chromosome pairing in spermatocytes is severely affected; X and Y hardly ever associate in these cells and are not silenced. This indicates that DSB formation could be essential for “true” XY body formation. However, one or two subnuclear regions containing part of the asynapsed chromatin but mostly not overlapping with X and Y still show enhanced accumulation of ATR and gH2AX (Bellani et al. 2005), apparently independent of the presence of meiotic DSBs. These so-called pseudo-XY body regions are also transcriptionally silenced (Mahadevaiah et al. 2008), similar to the silencing of the XY body in wild types. Thus, it seems that in the absence of meiotic DSBs, there is still some

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unknown activity on (part of the) asynapsed axes that nucleates the formation of a silenced chromatin area; the pseudo-XY body. When meiotic DSB repair is blocked, such as in Dmc1 knockout mice, the XY body or a pseudo-XY body are not formed (Pittman et al. 1998; Yoshida et al. 1998), and in such nuclei, RAD51 and gH2AX persist at many sites in the nucleus. This indicates that components of the DSB-repair machinery and the XY silencing machinery may be shared and limiting in amount. In addition, such nuclei also show impaired homologous chromosome pairing. Based upon the fact that in Spo11 mutants, the pseudo-XY body forms on only a small part of the asynapsed axes, it appears that the presence of asynapsed axes by itself is not sufficient to trigger meiotic silencing. However, the pseudo-XY body formation in the Spo11 knockout also argues against an essential role for the meiotic DSBs itself in the initiation of MSCI. The presence of DSB-repair associated proteins on the XY body and the inhibition of XY body formation when DSB-repair is blocked still point towards a certain overlap between the DSB-repair machinery and the XY silencing machinery. Apart from Spo11 and meiotic-DSB repair genes, the poly(ADP-ribose) polymerase 2 (PARP2) enzyme also appears to be involved in MSCI. Parp2 / spermatocytes show a general decrease in crossover frequency, defective SC formation between X and Y chromosomes, and also an impairment in MSCI. PARP2 controls chromatin structure and genomic integrity in response to DNA damage, again providing a link between DNA damage response pathways and MSCI (Dantzer et al. 2006). Taken together, it appears likely that the detection of heterologous regions is tightly coupled to the process of homology recognition. Both processes appear to have DSB-dependent and DSB-independent aspects. In addition, checkpoint proteins, such as ATR and perhaps also the newly identified HORMAD proteins, play essential roles in marking unsynapsed regions (often containing persistent meiotic DSBs) for meiotic silencing.

7.4.4

Meiotic Silencing of Unsynapsed Chromatin

MSCI appears to be a specialized form of a general meiotic silencing mechanism that silences unsynapsed (autosomal) heterologous chromatin in spermatocytes and oocytes (Baarends et al. 2005; Turner et al. 2005), termed MSUC (Schimenti 2005). This mechanism is activated, for example, when translocations or inversions interfere with normal chromosome pairing and on the single X chromosome in oocytes of XO mice (Baarends et al. 2005; Turner et al. 2005). In these unsynapsed, silenced, heterologous chromatin regions, persistence of meiotic DSB repair proteins is also observed, similar to what has been reported for the unsynapsed part of the X chromosome in the XY body. The X and Y chromosomes in spermatocytes are always subjected to meiotic silencing, but autosomal chromosomes with

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nonhomologous regions or the single X chromosomes in XO oocytes can sometimes evade MSUC by an adaptive process called synaptic adjustment, which results in complete heterologous synapsis. The axial/lateral elements of the SC can be considered as a basal axis that connects the loops of chromatin. The length of these loops determines axis length, and during synaptic adjustment, chromatin loop length is adjusted to equalize the axis length of the two chromosomal regions that are heterologous (Moses and Poorman 1981). It is evident that heterologous synapsis should be generally avoided because it interferes with homologous chromosome pairing. However, heterologous synapsis of small regions might be beneficial, to offer escape from a possible synapsis checkpoint, or to prevent silencing of essential meiotic genes by MSUC, hereby allowing spermatocytes with a minor pairing problem to survive and continue with their meiotic progression. Certain heterologous regions remain unsynapsed in a fraction of nuclei, whereas they synapse heterologously in the rest of the nuclei (Moses and Poorman 1981; van der Laan et al. 2004). When heterologous regions remain unsynapsed, they accumulate the DNA repair- and MSCI-linked proteins DMC1/RAD51. BRCA1, ATR, HR6B, and RAD18, and the phosphorylated form of H2AX, in human and mouse spermatocytes and oocytes (Baarends et al. 2005; Turner et al. 2005; Garcia-Cruz et al. 2009), and this is then always associated with transcriptional silencing of these unsynapsed regions. Conversely, when heterologous synapsis occurs, DNA repair proteins are not observed in these synapsed regions, and the chromatin is not silenced. Thus, there is a tight coupling between persistence of DSBs and unsynapsed silenced chromatin. In combination with the well-known link between meiotic DSB repair and homologous chromosome pairing, this prompted us to investigate whether the presence of persistent DSBs in nonhomologous regions might be instrumental in the prevention of synaptic adjustment and/or in the initiation of transcriptional silencing of this region. If this is the normal sequence of events in nonhomologous regions, induction of extra meiotic DSBs would lead to more, or more frequent, occurrences of DSBs in the nonhomologous region, which could then lead to more frequent detection of nonhomology and/or stronger inhibition of synapsis followed by induction of silencing. Such an effect was indeed observed for a specific small translocation bivalent in mouse spermatocytes. However, a much larger bivalent that contained the same nonhomologous region synapsed heterologously in virtually all nuclei, irrespective of the presence of extra meiotic DSBs (Schoenmakers et al. 2008). This indicates that with increased length of the homologously synapsed part of the chromosome, heterologous synapsis of the rest of the chromosome may become more likely. Above a certain threshold length, this effect may become dominant over the inhibitory signal that is elicited from the presence of unrepaired DSBs. Once heterologous synapsis has occurred, interhomolog bias may be relieved, allowing repair via the sister chromatid. In spermatocytes in which MSUC has been activated, MSCI seems to be impaired (Homolka et al. 2007). This appears somewhat similar to what is observed in mouse mutants with impaired meiotic DSB repair as mentioned in Sects. 7.3.5 and 7.4.3. In such mutants, persistent DSB sites also interfere with

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MSCI (Pittman et al. 1998; Yoshida et al. 1998; Fernandez-Capetillo et al. 2003; Ashley et al. 2004a, b; Barchi et al. 2005; Bellani et al. 2005; Di Giacomo et al. 2005; Mahadevaiah et al. 2008). These findings indicate that functional components of the DNA repair machinery and the MSCI/MSUC machinery are shared and limiting in their amount; for example, the amount of ATM/ATR that has to phosphorylate H2AX may be limiting. Unsynapsed autosomal chromosomes often localize adjacent to the X and Y chromosomes, and the formation of such a single silenced chromatin region may help to minimize the competition for the necessary shared factors that mediate MSCI and MSUC in the spermatocyte nucleus.

7.4.5

Postmeiotic Silencing of Sex Chromosomes

Inactivation of the majority of X- and Y- chromosomal genes continues beyond meiotic prophase, in spermatids, and persists until the end of spermatogenesis, when the whole genome is shut down (Namekawa et al. 2006; Turner et al. 2006). Around late pachytene, the inactivated XY pair acquires di- and trimethylated H3K9, which is associated with transcriptional repression (Khalil and Driscoll 2006). H3K9me3 mediates the recruitment of the Chromobox protein 1 (CBX1), a well-known heterochromatin marker, to the XY chromatin (Lachner et al. 2001). The histone methylation marks and CBX1 remain present on X and Y until the histone-to-protamine transition in late spermiogenesis (Namekawa et al. 2006; Turner et al. 2006), indicating continued repression of X and Y. Recently, it has been shown that the multicopy Y-chromosomal gene Sly plays a pivotal role in X- and Y-chromosome silencing in spermatids (Cocquet et al. 2009). Through inhibition of Sly expression with small RNAs expressed from a transgene, it was shown that Sly is involved in the maintenance of CBX1 and H3K9me3 on X and Y in postmeiotic nuclei (Cocquet et al. 2009). In addition, deficiency for SLY leads to a global postmeiotic derepression of X and Y genes, resulting in sperm defects (Ellis et al. 2005; Cocquet et al. 2009). How can the expression of a Y chromosomal gene function in the maintenance of repression of Y chromosomal gene transcription, if it is repressed by MSCI itself? This can be explained by the fact that, although most X- and Y-encoded genes remain repressed in postmeiotic cells (Namekawa et al. 2006; Turner et al. 2006), the majority of X- and Y-linked multicopy genes including Sly show postmeiotic reexpression (Mueller et al. 2008). In addition, a small number of single-copy genes are reexpressed (Hendriksen et al. 1995; Mueller et al. 2008). In spermatocytes and spermatids from Hr6b knockout males, the maintenance of XY body silencing is also impaired, in association with increased methylation of H3K4 on XY chromatin in late spermatocytes and round spermatids (Baarends et al. 2007). The exact molecular function of HR6B in the regulation of the X and Y chromosomes during meiosis and postmeiotic germ cell development is not known.

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Silencing of X and Y During Spermatogenesis and X Chromosome Inactivation in Female Somatic Cells Are Independent Mechanisms

It can be imagined that MSCI and postmeiotic repression of X and Y would leave some epigenetic mark that could affect the regulation of gene expression from the sex chromosomes in early embryos. This idea is relevant in the context of the fact that species with heterologous sex chromosomes not only have to deal with pairing problems in meiotic prophase of the heterogametic sex but also have to compensate for the dosage difference in X-linked genes between males and females (Huynh and Lee 2003). Female mammals achieve dosage compensation through inactivation of one of their X chromosomes during early development (reviewed in Payer and Lee 2008). In addition, the active X shows a twofold upregulation in somatic tissues, to equalize the average X chromosomal gene expression level to that of autosomes (Nguyen and Disteche 2006). Male mammals also double the expression of their single X chromosome (Lin et al. 2007). From the 2-cell stage onwards, the paternal X chromosome is specifically inactivated in female mouse embryos. This imprinted form of XCI persists in the extraembryonic tissues but is reversed and replaced by random X-inactivation in the embryo proper. The process of mammalian X chromosome inactivation is thought to have spread gradually over those regions of the X that had lost homology with Y (reviewed Graves 2006), most likely paralleling the occurrence of the different strata. The fact that most of the human X-linked genes without a Y homolog that still escape XCI are located on the younger strata (Carrel and Willard 2005) supports this idea and indicates that XCI may still be expanding. In order for XCI to occur, the presence of the X inactivation center (Xic) is necessary (Rastan 1983; Brown et al. 1991). Several noncoding RNA genes are located in the Xic, including the Xist en Tsix genes. (Penny et al. 1996; Marahrens et al. 1997; Kalz-Fuller et al. 1999). Xist is specifically expressed from the inactivated X chromosome (Xi) and is required for XCI to occur in cis. The spreading of Xist RNA over the X chromosome from which it is transcribed induces its heterochromatinization (Plath et al. 2003; Silva et al. 2003). Tsix is transcribed antisense to Xist, and both genes overlap in mice. Tsix negatively regulates expression of Xist. Recently, it was described that human female preimplantation embryos also show progressive Xist RNA accumulation on one of the two X chromosomes (van den Berg et al. 2009). It is not known whether this early XCI occurs at random or only on the paternal X, as in mouse. Althought Xist and Tsix gene are not required for MSCI and postmeiotic silencing of sex chromosomes (PMSC) (Marahrens et al. 1997; Turner et al. 2002), it has been suggested that the two mechanisms (MSCI and XCI) might be functionally connected: as described above, the inactivation of X and Y during meiosis may leave an epigenetic imprint on the X chromosome, and this might play a role in the imprinted paternal X chromosome inactivation (XCI) in the early female mouse embryo from the 2-cell stage onward. To test the hypothesis that

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MSCI facilitates paternal X-inactivation in the early mouse embryo, Okamoto et al. (2005), analyzed the expression of an autosomally localized transgenic copy of Xist in germ cells and early embryos. They found that the transgene was not silenced by MSUC during meiotic prophase and that the transgenic Xist gene still showed expression only when present on the paternal copy in the zygote. In addition, they showed that the paternal X chromosome is active in two-cell embryos, excluding the inheritance of the X in a “pre-inactive state” (Okamoto et al. 2005). The Xist gene is absent from the marsupial genome, and imprinted inactivation of the paternal X in this species occurs in all cells by an unknown mechanism. MSCI does occur in the male germline of marsupials, but similar to mouse and man the paternal X is active in early embryos (Cocquet et al. 2009), again arguing against a link between MSCI and imprinted X-inactivation, even in the absence of Xist (Cocquet et al. 2009). In contrast to the theory that proposes a preinactive paternal X, it has also been suggested that the protamine-to-histone transition occurring directly after fertilization in the male pronucleus of the zygote may facilitate rapid activation of the paternal X chromosome, thereby enhancing the chance of Xist transcription from the paternal X. This hypothesis has not yet been tested.

7.6

Meiotic Silencing of Sex Chromosomes in a Species with Female Heterogamety

In contrast to the XY/XX male/female sex chromosome system in mammals, birds have a ZW female and ZZ male sex chromosome constitution. Recently, it was discovered that sex in birds is determined by the dosage of the Z-linked DMRT1 gene. The higher DMRT1 dose in ZZ chickens drives the bipotential gonads to become testes (Milman et al. 2009). Similar to the mammalian male Y chromosome, the avian female W chromosome carries few genes; so far only four genes have been identified (International Chicken Genome Sequencing Consortium 2004). In addition, the gene content of the Z chromosome appears to have masculinized, showing an overrepresentation of male-biased genes (2004). This is comparable to the overrepresentation of female-biased genes on the mammalian X chromosome (Khil et al. 2005). The heterologous mammalian X and Y chromosomes remain largely unsynapsed during the male meiotic prophase. However, despite the fact that the avian Z and W are also largely nonhomologous, they synapse completely during the female meiotic prophase in chicken oocytes. MSCI and MSUC in mammals are always associated with asynapsis, and it has been suggested that the heterologous synapsis of Z and W in bird oocytes needs to occur to escape from such a silencing mechanism, since the presence of a silenced Z chromosome would be incompatible with oocyte development during the lengthy (arrested) meiotic prophase in females (Jablonka and Lamb 1988). We performed a detailed analysis of the behavior of the Z and W chromosome during meiotic prophase in chicken oocytes. Surprisingly, we observed markers of transcriptional inactivation associated with the synapsed ZW

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sex chromosomes during pachytene. In addition, real-time RT-PCR analyses showed low levels of mRNAs from selected Z- and W-linked genes in pachytene oocytes (Schoenmakers et al. 2009). However, in diplotene, the Z chromosome was reactivated. The complete heterologous synapsis of Z and W is difficult to understand; how does the cell allow heterologous synapsis between Z and W and simultaneously prevent synapsis between nonhomologous autosomal chromosomes? Upon entrance in meiotic prophase, the W chromosome is already heterochromatic and inactivated, while Z is indistinguishable from the autosomes. It is apparent that during homologous chromosome pairing, in zygotene, the Z and W are the last pair to synapse. RAD51 foci and gH2AX, indicative of the presence of (unrepaired) DSBs, persist on the asynapsed axes, but disappear where heterologous synapsis has taken place. When Z and W start to desynapse in late pachytene, RAD51 foci and gH2AX reappear, indicating that repair of meiotic DSBs on Z is suppressed during synapsis, perhaps to avoid recombination between heterologous parts of Z and W. Upon desynapsis, in late pachytene and diplotene, this suppression is apparently relieved, and repair (via HR with the sister chromatid or through NHEJ) can occur. This appears similar to the timing of meiotic DSBs repair on the unsynapsed arm of the X chromosome in mouse diplotene spermatocytes. The obligatory crossover in the pseudoautosomal region of Z and W is formed with normal timing, as a single MLH1 focus can be observed on the ZW of most pachytene oocytes. This crossover is essential to ensure correct separation of Z and W during the first meiotic division. Markers associated with silencing, such as ubiquitylated H2A and H3K9me3, appear first on W at leptotene, and around midpachytene, when Z and W are completely synapsed, these inactivation markers mark the whole ZW body. We have suggested that the temporary inactivation of the avian sex chromosomes during pachytene is a consequence of spreading of heterochromatin from W onto Z, facilitated by the heterologous synapsis between the Z and W chromosomes (Schoenmakers et al. 2009). As mentioned above, the second wave of gH2AX formation by the kinase ATR initiates MSCI in late zygotene in mammals. In chickens, silencing of Z and W is achieved in the absence of gH2AX, but a second wave of H2AX phosphorylation associated with the postponed repair of meiotic DSBs could contribute to the maintenance of Z chromosome silencing from late pachytene until early diplotene. In conclusion, meiotic inactivation of sex chromosomes occurs also for Z and W in birds, but this mechanism, which is independent from the final achievement of synapsis, may have evolved independently from MSCI in mammals.

7.7

What Drives Meiotic Silencing in Mouse and Man?

Our analysis of MSCI in the avian germ line has shown that silencing of heterologous sex chromosomes during meiosis is widely conserved among species but does not always depend on the presence of asynapsed axes. However, as indicated above,

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the molecular mechanisms of initiation of meiotic silencing may differ between mammalian and avian species. For the initiation of MSCI in mammals, two mechanisms appear to come together: The mechanism that regulates progression of synapsis along aligned and paired chromosomes and the mechanism of meiotic-DSB repair. These two mechanisms are tightly coupled; in DSB-repair mutants, synapsis cannot proceed, in mutants that lack components of the SC, DSB repair is impaired, and in both cases, true XY body formation does not take place. The formation of the pseudo-XY body in Spo11 knockout mice supports the hypothesis that formation of a silenced asynapsed region is independent of meiotic DSBs. However, the lack of silencing associated with the majority of unsynapsed axes in these mutants, and the presence of many DNA repair-related molecules in the pseudo-XY body, indicate that asynapsis alone does not do the job and implicate functions of DSB-repair proteins in forming the final inactive chromosome structure. The chromosomal pairing dance of X and Y starts with telomere attachment to the nuclear envelope, followed by active movements along the membrane, aided by cytoskeletal fibers, until they meet and cluster together. The sex chromosomal telomeres most likely still share homology, and the largely nonhomologous regions outside of the telomeric regions may remain undetected during these early stages of chromosome pairing. At the end of the dynamic bouquet stage, when homologous telomeres have paired and clustered, the chromosomes keep moving around and most likely scan and assess homology during transient states of interaction. Around this time, when the chromosomes are already actively being pulled around, SC assembly starts with dedeposition of SYCP3 along the axes, SPO11 introduces DSBs throughout the genome, and ATM mediates the first wave of gH2AX formation in regions surrounding the meiotic DSBs. The movement of chromosomes gives the ssDNA strands coated with RAD51/DMC1 the opportunity to invade homologous dsDNA, and temporary recombination intermediates are formed. If homology is not established, recombination intermediates are destabilized and the search continues. If a homologous template is encountered, the repair process can continue and is completed to form crossovers and noncrossovers and gH2AX and RAD51 foci disappear concomitant with the formation of the complete SC. The X and Y chromosomes initiate synapsis from the PARs, but by the end of zygotene, the X chromosomal arm is still left with numerous RAD51/DMC1 foci. A second wave of gH2AX formation is then induced and covers the complete X and Y chromosomes, marking the formation of the XY body and initiation of MSCI. The persistence of DNA repair proteins on meiotic DSBs may be used as an indicator for the absence of a homologous partner. Alternatively, or in addition, persistence of DSB repair proteins may inhibit progression of synapsis. The recruitment of ATR, to replace ATM at such sites, appears to be the crucial event in the initiation of meiotic silencing, and it remains to be established if this is normally initially triggered by the presence of persistent DSBs, or by the presence of some component that specifically associates with asynapsed axes, or depends on both features associated with nonhomologous chromatin. If ATR is initially recruited by the persistent DSBs, it may subsequently spread along the chromosomal axes and into the chromosome loops until the end of the chromosomes is

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a

b homolog-directed repair paternal homolog / sister

meiotic recombination

maternal homolog / sister

= DSB + repair proteins = search homologous repair template

paternal homolog / sister

maternal homolog / sister

= SYCP3 = γH2AX

= SYCP3 = crossover

= SYCP1

= gH2AX

c homolog-directed repair Y chromosome / sister

X chromosome / sister

meiotic recombination Y chromosome / sister

X chromosome / sister

X

X X

X X

pseudoautosomal region

= accumulation DNA repair proteins along asynapsed regions

Fig. 7.2 Model for the process of homology recognition, linked to meiotic DSB repair and MSCI. (a) In early leptotene, all telomeres become attached to the nuclear membrane, and cytoskeletal fibers originating in the cytoplasm pull the telomeres along on the membrane. Simultaneously, hundreds of meiotic DSBs are induced (yellow dots) throughout the genome, and a search for a

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reached. This then results in the massive phosphorylation of H2AX, and incorporation of macroH2A, chromatin sumoylation, H2A ubiquitylation, and other histone modifications, occur downstream. Together, this results in the formation of the silenced XY body. This putative sequence of events is schematically summarized in Fig. 7.2.

7.8

Clinical Relevance of Meiotic Silencing

Interference with MSCI appears to be incompatible with fertility in mouse and man (Yoshida et al. 1998; Fernandez-Capetillo et al. 2003; Dantzer et al. 2006; Leng et al. 2009). Inappropriate expression of X- and Y-linked genes during meiotic prophase may trigger apoptosis of pachytene spermatocytes due to the toxic effect of expression of one or more of these genes (Burgoyne 2007, 2009). In addition, activation of MSUC due to the presence of chromosomal aberrations that interfere with normal chromosome pairing may also trigger germ cell death due to inappropriate inactivation of genes essential for meiosis. For example, X-autosome translocations are frequently associated with male infertility (Madan 1983; Kalz-Fuller et al. 1999; Ma et al. 2003), which is most likely due to spreading of the meiotic silencing signal from the X to the (asynapsed) autosomal part of the translocation chromosome (Turner et al. 2005). In addition, autosome-to-autosome translocations ä Fig. 7.2 (continued) homologous repair template is initiated. The chromosomal movements facilitate numerous transient interchromosomal interactions, allowing an efficient homology scan by the DSB repair machinery. If homology is confirmed, stable interactions can be formed, and strand invasions associated with DNA repair follow. Pairing regions and “sticky heterochro matic” regions (colored rectangled block) may also contribute to homology recognition and aid in the stabilization of homologous pairing. (b) Each individual autosomal homologous chromosome pair acquires many DSBs and simultaneously assembles the lateral element, SYCP3, of the synaptonemal complex. DSB repair occurs in association with the SC, and the intimate contacts that occur during strand invasion allow SYCP1 and other central elements of the SC to assemble between the SYCP3 proteins on both homologs. Some of the DSBs (at least one per chromosome pair) are converted to crossovers, and these are required for proper segregation during the first metaphase to anaphase transition. Following completion of the DNA repair process, H2AX becomes dephosphorylated. (c) The X and Y chromosomes undergo the same processes as the autosomes: induction of DSBs and assembly of SYCP3. However, apart from the pseudoautoso mal region in which the obligatory crossover will be formed, no regions of homology are present. This interferes with the homologous recombination repair process of meiotic DSBs on the X chromosome due to the absence of a homologous repair template. SYCP1 deposition and SC formation initiates from the pseudoautosomal region but does not proceed, or is unstable, in the heterologous regions. The delayed repair of DSBs could cause continuous accumulation of DNA repair proteins, and above a certain threshold ATR may be recruited. Alternatively, or in addition, accumulation of (DNA repair) proteins that specifically associate with the unsynapsed chromo somal axes may recruit ATR. This leads to spreading of ATR and other (DNA repair) proteins along the XY chromosomal axes. Subsequently, phosphorylation of H2AX spreads into the surrounding chromatin, mediating recruitment of downstream factors, including the factors that mediate transcriptional silencing in the XY body

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can trigger MSUC and are associated with (sub)fertility in mouse (Peters et al. 1997) and man (Leng et al. 2009). It cannot be excluded that inappropriate regulation of chromosome regions during spermatogenesis due to defective MSCI and/or activation of MSUC affects regulation of gene expression in the early embryo via transmission of epigenetic marks. Therefore, in the context of the field of assisted human reproduction techniques (MESA, TESA, IVF, and ICSI), it is of utmost importance to understand the basics of the meiotic silencing mechanisms to estimate the possible risks of producing zygotes from gametes in which MSCI was incomplete and/or MSUC was activated.

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Chapter 8

Histone Variants during Gametogenesis and Early Development P. de Boer, M. de Vries, and S. Gochhait

Abstract The objective of this chapter is to give an overview of the role of histone variants mainly in mammalian spermatogenesis. Two aspects emerge: their roles in the process of spermatogenesis itself and second, are histones including their variants instrumental in passing on information to the next generation? As to the first topic, enough experimental data have been assembled to be certain about the role of histone variants, accommodating chromatin regulation especially for meiosis and spermatid nucleus elongation. As to the second question, there is only nonconclusive experimental evidence for a role of histone variants per se, although the concept of male-transmitted epigenetic inheritance and effects has been convincingly proven for the mouse. The biological value in such a concept is that adaptation to newly arising circumstances does not have to rely on mutation and selection only, but could be accommodated on a shorter time axis. One particular area where some of the variation in transmission, putatively evoked by sperm-retained histones, could be studied is in the sphere of human artificial reproduction because male gametes, that in a free competition would never lead to offspring, are now called to action and are capable of producing offspring of normal phenotype. An overview will be given on histones in meiosis, during spermiogenesis and preserved in sperm. Subsequently, mouse data on the principle of male transgenerational inheritance and effects are provided. The chapter is closed by formulations on future directions of research.

P. de Boer (*) and M. de Vries Department of Obstetrics and Gynaecology, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands e mail: [email protected] S. Gochhait National Centre of Applied Human Genetics, School of Life Sciences, Jawaharlal Nehru University (JNU), New Delhi, India

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Keywords Embryonic development Histone variants spermatogenesis Sperm Spermiogenesis

8.1

Introduction

In this chapter, we will primarily deal with histone variants in the germline of mammals. When reviewing the literature that recently has been published on this theme, it is apparent that the focus of study is on male chromatin, probably because female chromatin is assumed to largely adhere to the principles of somatic chromatin (Boussouar et al. 2008; Churikov et al. 2004a, b; Godde and Ura 2009; Orsi et al. 2009; Rousseaux et al. 2008; Santenard and Torres-Padilla 2009; Ooi and Henikoff 2007). In the area of male reproduction, our knowledge on transmission of histones is growing. The explanation of embryonic development from female oocyte chromatin does not show much advance (Albert and Peters 2009). Epigenetic asymmetry of zygote pronuclei (Santos et al. 2005; van der Heijden et al. 2005) that over subsequent cleavage divisions is resolved by equalizing male to female H3 and H4 N-tail posttranslational modifications (PTMs) (Puschendorf et al. 2008) shows the dominant position of female chromatin in this respect. This epigenetic asymmetry is also found in the human (van der Heijden et al. 2009). Histone variants are mainly characterized as to their use in replication-coupled nucleosome formation including DNA repair (Polo et al. 2006) versus incorporation into chromatin outside the context of DNA synthesis. The first ones are called canonical replication dependent, the second group noncanonical replication independent. Apart from this distinction, canonical and noncanonical histone variants can be tissue or cell type specific or being expressed in a variety of ic in all tissues. The general significance of histone variants as an extra layer of information as to status and function of chromatin has been reviewed by Bernstein and Hake (2006). Because of the increased requirement of canonical histones in proliferating cells, the replication-dependent genes are present in the genome as clusters, offering another classification tool. The genes for germline-specific variants (except for H1t, TH2A/TH2B and TH3) and the common noncanonical variants of the core histones are found in isolation and differ from those present in clusters in the presence of polyadenylated mRNA, later expression in spermatogenesis and a bigger chance on introns. The informational complexity of chromatin is further enhanced by histone PTMs. These will only be touched upon in this chapter when they add to understanding. Table 8.1 gives a list of histone variants in spermatogenesis, incorporating data from rat, mouse, and human. The prime focus of interest in the transmission of paternal nucleosomes is in their possible role as transgenerational carriers of information for postfertilization development. Experiments mainly on mice demonstrate transgenerational male epigenetic inheritance and effects to exist. This evidence will be presented in part V. Progress in this area of research is curtailed by the need for cell purification methods, lack of in vitro systems, necessity for genetic manipulation of the genes concerned,

Hils1 intronless

10q31

8q22

2q44

HILS1

H2A.X

H2A.Z

H2afz 5 exons

H2afx intronless

H1fnt intronless

HANP1/H1T2 7q36

Hist1h1t intronless

17p11

H1t

Hist1h1a intronless

Hist1h1c intronless

17p11

Gene

Rat (location) Mouse

H1c

H1a

Variants

3G3

9A5.2

11D

15F1

13A3.1

13A3.1

13A3.1

Location

57–59%

88–90%

15–24%

17–19%

46%

61–84%

62–68%

Conserved

H2AFZ 2 exons

H2AFX intronless

HILS1 two exons

H1FNT intronless

HIST1H1C intronless HIST1H1T intronless

HIST1H1A intronless

Gene

Human

17–19%

20–25%

50%

64–87%

66–69%

Conserved

4q23

58–59%

11q23.2–q23.3 90%

17q21.33

12q13.11

6p22.2

6p22.2

6p22.2

Location

Table 8.1 Details of the histone variants enriched or exclusively expressed in the male germline

Male sterility

Artificially fertile

Fertile

Fertile

Fertile

Null phenotype

Proposed function

In mouse from type A Somatic linker and B histone enriched spermatogonia on, in germ cells protein decreases in post-meiotic stages. In humans is found post-meiotically Type A and B spermatogonia Evident in 21-day old Facilitates histone rats and found from replacement by pachytene opening up spermatocyte to chromatin round spermatid (Step 8) Expressed in round and Chromatin elongating remodeling, spermatids (Steps establishes 4–13) polarity by presence in the apical nuclear region Present in elongating Chromatin and elongated remodeling spermatids (Steps 9–16) g-H2AX appears in Sex body formation, spermatogonia, meiotic mostly restricted to telomere the XY body in the function pachytene diplotene spermatocyte Pachytene Suggested to replaces spermatocytes till macroH2A1.2 spermatozoa in sex chromosomes post meiosis

Expression pattern

(continued)

West and Bonner (1980), Greaves et al. (2006)

Trostle-Weige et al. (1982), FernandezCapetillo et al. (2003a, b)

Iguchi et al. (2003), Yan et al. (2003)

Tanaka et al. (2005) and Martianov et al. (2005)

Meistrich et al. (1985), Fan et al. 2001 Meistrich et al. 1985; Lin et al. (2000, 2004)

Meistrich et al. (1985), Franke et al. (1998), Lin et al. (2004), Burfeind et al. (1994)

References

Hist1h2aa intronless

TH2A

H2BL1

H2BFWT

TH2B

17p11

H2bl1

Gm14920, XE1-2 LOC100046339, H2afb3, EG624153, LOC100039277 mTH2B (Hist1h2ba) 13A3.1 intronless

H2A.Bbd

13D1

XA1.2

1700054O13Rik

H2AL3

XA1.1 2A3

H2al1 intronless H2al2 intronless

13A2-A3

13B1

Location

H2AL1 H2AL2

17p11

H2afy 10 exons

Gene

Rat (location) Mouse

macroH2A1.2 17p14

Variants

Table 8.1 (continued)

38%

88%

23–34%

24%

30% 41%

86–87%

58–60%

Conserved

6p22.2

5q31.1

Location

H2BFWT, 3 exons

Xq22.2

hTSH2B 6p22.1 (HIST1H2BA) intronless

H2AFB2 intronless Xq28

HIST1H2AA intronless

H2AFY 10 exons

Gene

Human

43%

85%

48%

90–91%

59–60%

Conserved Fertile

Null phenotype

Proposed function

References

Chromatin Spermatogonia, first Meistrich et al. 1985; meiotic prophase remodeling van Roijen et al, spermatids and 1998; Zalensky sperm in human. et al. 2002; Unni Same for rat up to et al. 1995 meiosis, likely beyond Nuclear periphery of the Configurates Churikov et al. round spermatids telomeric (2004a, b) and mature sperm nucleosomes in human sperm Condensing spermatids Remodeling of Govin et al. (2007) and mature heterochromatin spermatozoa in elongating spermatids?

Peaks in pachytene Transcriptional Pehrson and Fried spermatocyte, silencing of sex (1992), localized to body, higher Changolkar et al. chromocenter in chromatin (2007) spermatids and organization present in spermatozoa Appears in 16 day rat Trostle-Weige et al. coinciding with (1982) pachytene spermatocyte till early spermatids. Condensing spermatids Chromatin Govin et al. (2007) and mature remodeling of spermatozoa but the pericentric disappears after regions along fertilization with TH2B Protein not detected in germ cells Elongated spermatids in Chromatin Ishibashi et al. (2010), mouse, present in remodeling Chadwick and human spermatozoa Willard (2001)

Expression pattern

5B1

11B1.3

41%

97.0%

1q42.13

1q42.12 17q25.1

CENP-A Isoform a 2p23.3 (6 exons), Isoform b (4 exons)

HIST3H3 (H3FT) intron less

97% within 119 Nterminal amino acids and unique Cterminal 12 amino acids 96% H3F3A, H3F3B, 4 exons

31%

46%

96.0%

96.0%

Embryonic lethal

Impaired fertility

H3.3B is restricted to Is associated with meiotic prophase transcription while H3.3A is and general observed chromatin throughout as basal remodeling expression In rat it is present from spermatogonia to spermatids but transcribed only in spermatogonia. In humans transcription is restricted to pachytene spermatocytes Observed as discrete foci Centromere in the mature sperm functioning (bull)

Protein not detected in germ cells Spermatid specific, highest in late round stages

Palmer et al. (1990), Howman et al. (2000)

Trostle-Weige et al. (1984), Witt et al. (1996), Govin et al. (2007)

Bramlage et al. (1997), Couldrey et al. (1999)

Moss et al. (1989), Unni et al. (1995)

The names, locations, and orthology of the genes have been assigned following the UCSC (http://genome.ucsc.edu), Rat Genome (http://rgd.mcw.edu/), and Mouse genome database (http://www.informatics.jax.org). The protein sequence homologies, when not available from the literature, were established by comparing to the major somatic histone variants using the ClustalW2 protein sequence alignment tool (http://www.ebi.ac.uk/Tools) with the default parameters

Cenpa 5 exons

CENP-A

6q14

LOC382523

TH3

H3f3a, H3f3b 4 exons 1D2.3, 11E2

13q26, 10q32.3

H3.3

13A3.1

LOC328201, two exons

ssH2B

XF1

H2bl2

H2BL2

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and the general complexity of chromatin. The reason for obtaining insight in this area, apart from its central position in the propagation of life, also stems from the fact that in the world of artificial reproduction, application always is ahead of insight in biological significance, leading to a situation in which we are our own experimental model as to the effects of sperm with variable amounts of histones.

8.2

Meiotic Chromatin

By microscopy, male germline chromatin is different from the first meiotic division onwards. This was already known by older generation cytogeneticists. For instance, spermatogonial mitotic chromosomes are well-delineated, chromatin making a compact expression with well-aligned sister chromatids. Chromatin of diakinesis metaphase I bivalents is much more fuzzy, to which process the loss of sister chromatid cohesion around cross-over sites at the chiasmata will contribute (Hodges et al. 2005; Revenkova et al. 2004). Chromatin of spermatocyte bivalents is not suitable for G-banding to recognize individual bivalents, but shows excellent C-banding representing centric constitutive heterochromatin in the mouse that we now know to be marked with H3K9me3 and H4K20me3 PTMs (Tachibana et al. 2007; van der Heijden et al. 2007). Sister chromatid differentiating chromosome staining by S-phase BrdU incorporation, which was developed to visualize sister chromatid exchange (SCE), does not show reliable meiotic differentiation with the exception of the sex chromosomes, where these exchanges can be easily seen (private communication JM Veltmaat, AH Peters). This is an indication for the different chromatin organization of autosomes and gonosomes during the first meiotic division. This difference is maintained in the metaphase chromosomes of secondary spermatocytes.

8.2.1

Histone Variants

The group of Meistrich (Trostle-Weige et al. 1982, 1984; Meistrich et al. 1985) was the first to quantitatively report on the testis-specific H1 linker histone (H1t, Table 8.1) that becomes abundant and constitutes around 50% of the H1 population, when transcription is upregulated in mouse stages of the seminiferous epithelium IV V (Oakberg 1956a, b). In histological and acid spread preparations of primary spermatocytes beyond these stages, the nuclear diameter increases indicating more open chromatin. With our current knowledge, that enables us to translate the presence of testisspecific H1 linker histones (H1t) into decreased chromatin compaction (see in Godde and Ura (2009), Oakberg (1956a, b)), the specific meiotic chromatin morphology could in theory be related to the changing histone variants composition especially during male meiosis. However, single knock outs (ko) for H1t do not show any phenotype as to male fertility (Lin et al. 2000; Drabent et al. 2000; Fantz

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et al. 2001). Cytogenetics of the first meiotic division in mice double ko for H1t and the somatic H1a (the type that had previously been found to replace H1t in its absence (Lin et al. 2000)) showed no phenotype after okadaic acid-induced bivalent condensation, that always produces over contraction, thereby possibly hiding any deviant chromatin morphology in the mutant (Lin et al. 2004). Despite changes in H1 nucleosome stoichiometry and a modest change in germ cell gene expression, fertility was normal in the double ko mice. Spermatogenesis is characterized by testis specific variants for H2A, H2B, and H3 that were for the rat first described by Meistrich and co-workers (Trostle-Weige et al. 1982, 1984; Meistrich et al. 1985) and denoted by the prefix T (Table 8.1). TH2A and TH2B are prominently present in early primary spermatocytes. In rat, pachytene spermatocytes TH2A accounts for about 25% of total H2A (TrostleWeige et al. 1982) and H2B is largely replaced by TH2B (Meistrich et al. 1985), supposedly leading to a more open chromatin structure (Churikov et al. 2004b). However, their impetus on nucleosome stability and possible interaction with PTMs have not been further researched. TH2A and TH2B are already expressed in late spermatogonial stages. Expression of TH2A does not extend beyond meiosis (Trostle-Weige et al. 1982). In a later study, despite its overall decline in rat elongating spermatids, immunodetection of TH2B by the N-terminal region improves (using the anti tyrosine hydroxylase antibody) (Unni et al. 1995). This may indicate that at this stage, TH2B is more weakly bound to the DNA due to displacement by transition proteins. TH2B is uniformly present throughout human spermatogenesis and, as judged by immunohistochemistry with the same tyrosine hydroxylase antibody, most abundant in round spermatids (van Roijen et al. 1998). From the discussion in Meistrich et al. (1985), it can be deduced that testis-specific forms of H2A and H2B are not uniformly present among mammals. Most H3 variants are not germline specific (see Table 8.1). However, we have listed H3.3 in this table because of the growing interest of this variant with respect to cell memory (see part VI). Canonical H3 variants are H3.1 and H3.2 that only differ in one amino acid. The noncanonical replication-independent variant H3.3 differs in only four amino acids, of which the H3.3 serine in position 31 is at the transition from N-tail to globular domain. One of the findings that makes this variant important is its enrichment in PTMs associated with transcription activation (Bernstein and Hake 2006). Mouse and human H3.3 is coded by two genes, H3.3A and H3.3B (Table 8.1), that give the same product. H3.3B is only expressed during first meiotic prophase, whereas expression of H3.3A extends throughout spermatogenesis (Bramlage et al. 1997). By Lac Z integration, Couldrey et al. (1999) ablated the H3.3A gene. Homozygosity for the mutation resulted in a few pregnancies after copulation and reduced postnatal survival. Spermatogenesis was not further characterized. The specific importance of H3.3 has recently been demonstrated in a Drosophila genetic study (Sakai et al. 2009). A simple ko of the gene resulted into male sterility. At cytological inspection, bivalent contraction and segregation were hampered. Also, a cytological indication for a chromatin remodeling problem at spermatid nuclear elongation was obtained.

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Using a transgenic mouse line carrying copies of H3.3 marked with a V5 tag, it was noted that expression of this construct commenced in around stage II III primary spermatocytes (Oakberg 1956a, b), visible in the X chromosome first (van der Heijden et al. 2007). Likely as a consequence of increased general transcription beyond stage IV, the autosomal H3.3 signal clearly increased in late pachytene, diplotene spermatocytes. Recently, it has been shown in Drosophila that at transcription start sites, a higher H3.3 concentration connects with a lower H1 concentration, supposedly to open up chromatin (Braunschweig et al. 2009). This fact, in concert with H1t, TH2B, and H3.1/3.2 nucleosome eviction in favor of H3.3, could assist in rendering meiotic prophase chromatin less compact. In the work of Meistrich et al. (1985), it was shown that in late pachytene spermatocytes, the summed presence of H3.3 and TH2B was increased. H2A.Z (Table 8.1) in combination with H3.3 functions to open chromatin, either by decreased nucleosome stability (Jin and Felsenfeld 2007) or due to the altered linker histone binding of H2A.Z and increased nucleosome sliding (Thakar et al. 2009). This phenomenon could also be a factor in the open chromatin configuration at late first meiotic prophase as H2A.Z is increasing in this phase (Meistrich et al. 1985; Greaves et al. 2006). Macro-H2A1.2 (Table 8.1) that is generally associated with gene silencing, both at the level of single genes and at the chromatin domain level such as the inactive X chromosome (see in Bernstein and Hake (2006)), is at the beginning of first meiotic prophase present in sex body chromatin. The term sex body applies to the male meiotic sex chromosomes bivalent that from its formation on separates as a chromatin domain, often at the periphery of the nucleus. From mid pachytene on, the immunofluorescence (IF) signal for macro-H2A1.2 comes up over autosomal chromatin, but for the sex chromosomes now is only above this level for the centromeric regions and for the pseudo-autosomal region (Hoyer-Fender et al. 2004). After meiosis in round spermatids, H2A.Z comes up over the sex chromosomes in the mouse and is suggested to replace macro-H2A1.2 (Greaves et al. 2006). Gonosomal chromatin is allocyclic early condensing toward the first meiotic division, and as indicated before, much more on a par with classical cytogenetical staining methods than autosomal meiotic chromatin. Most likely, the absence of transcription for the sex chromosomes in first meiotic prophase (meiotic sex chromosome inactivation MSCI (Turner 2007)), respectively, its presence in autosomal chromatin, is one aspect of the fuzzy microscopical manifestation of autosomal bivalents at all stages of bivalent contraction as could already be concluded from the patterns of histone variants at late first meiotic prophase. Primary spermatocytes are relatively rich in the replication-independent variant H2A.X (Table 8.1) (Meistrich et al. 1985; Trostle-Weige et al. 1982). The concentration of this histone variant is high in spermatogonia and dropped slightly in round spermatids (Trostle-Weige et al. 1982). The presence of H2A.X is a prerequisite for a mature DNA damage response to develop, especially for double-strand (ds)DNA repair (Rogakou et al. 1998). Phosphorylation of serine at position 139 is achieved by one of the three phosphoinositide 3-kinase related kinases (PIKKs), ATM,

8

Histone Variants during Gametogenesis and Early Development

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ATR, and DNA-PKcs (Lempiainen and Halazonetis 2009), the product being coined gH2A.X. By immunohistochemistry, it was found that in the mouse, gH2A.X was uniformly distributed in all In(termediate) and B spermatogonia. Less intense signals were found in A spermatogonia. Leptotene and zygote nuclei stained, whereas beyond stage XII at the onset of pachytene, meiocytes were only positive for the sex chromosomes in the sex body (Hamer et al. 2003). Irradiation, by the induction of dsDNA breaks, induced gH2A.X foci in all stages concerned (Hamer et al. 2003; Ahmed et al. 2007). There is no apparent reason why gH2A. X is high in In and B spermatogonia except for their relatively long S-phase. No apparent autosomal meiotic role for H2A.X is known apart from its function at leptotene zygotene, whereby ATM-dependent phosphorylation of S139, it signals SPO11 ds break induction (Mahadevaiah et al. 2001) though not with the intensity of, for instance, an irradiation-induced ds DNA break (Chicheportiche et al. 2007). The ATR-dependent marking of unsynapsed chromatin can be observed from late zygotene (Turner et al. 2004) (Chap. 7). In a normal dsDNA break repair process, the gH2A.X chromatin reaction is taken to continue until repair has been completed or has definitely failed (Srivastava et al. 2009). In dsDNA break processing in the context of reciprocal exchange or gene conversion, H2A.X usually (but not always) is dephosphorylated (Chicheportiche et al. 2007; Mahadevaiah et al. 2001), once a recombination intermediate is formed in an early recombination nodule positioned at the central element of the synaptonemal complex. These recombination nodules can be conveniently marked by IF of MSH4, RPA, RAD51, and DMC1. At this stage, that commences at zygotene, the process of recombination is ongoing until MLH1 and MLH3 (Baker et al. 1996; Svetlanov et al. 2008) signify the presence of a mature crossing over product at mouse stages V VI, the onset of late pachytene.

8.2.2

More Sex Chromosome Aspects

The meiotic X and Y chromosomes are one of the current examples of a role for canonical to noncanonical (replication-dependent to replication-independent) H3 substitution. Although not yet fully researched, gonosomal nucleosome eviction of at least those that contain H3.1/H3.2 in the mouse takes place from early pachytene on, first detectable about 16 h after the zygotene to pachytene transition, and continues till stages IV/V at the onset of the transcription intensifying late pachytene stage (van der Heijden et al. 2007). These authors have linked this substitution to the phenomenon of meiotic sex chromosome inactivation (MSCI, Chap. 7). So in this case, a H3.3-rich environment is compatible with lack of transcription, in line with the presence of the inactivation PMTs H3K9me2,3. Addition (to the sex body) of an asynaptic autosomal segment constituting G bands 1A5 1C1.1 (approximately 40 Mb, see Schoenmakers et al. (2008)) from the reciprocal translocation T(1; 13)1Wa (de Boer et al. 1986) resulted in remodeling of this segment as well, according to the general principle of meiotic silencing of unsynapsed chromatin

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(MSUC, Chap. 7). In about 40% of nuclei, exchange in the total area undergoing nucleosome eviction was incomplete as judged by a residue of fluorescent H3.1/ H3.2 (van der Heijden et al. 2007). Generally, in small translocation marker chromosome trisomy and segmental asynaptic chromosome mutants, the extra chromosome or asynaptic segment congregates with the largely asynaptic X and Y (de Boer and Branje 1979; Forejt and Gregorova 1977; Burgoyne et al. 2009; de Boer et al. 1986; van der Heijden et al. 2007; Peters et al. 1997). Variegation as to nuclear elongation in the later spermatid stages always is the case in these chromosome mutants (de Boer and de Jong 1989) (Fertility and Chromosome pairing: recent studies in plants and animals ed CR Gillies, CRC Press, Boca Raton, Florida, (1989), Gillies (1989), (de Boer, unpublished results). Hence, spermiogenesis is abnormal, by morphology resembling the oligoasthenoteratoozoospermic condition (OAT) condition as it is known in the human. A disturbance of X, Y control of gene expression, extending into the round spermatid stage and gene dosage effects due to silencing of unsynapsed autosomal chromatin will both contribute to abnormal spermiogenesis (Burgoyne et al. 2009; Homolka et al. 2007). The fact that different autosomal inclusions lead to the same spermiogram next to the more than random involvement of gonosomal genes with reproduction (Delbridge and Graves 2007; Krausz and Degl’Innocenti 2006; Jobling 2008) would opt for the importance of sex chromatin regulation. In the mouse, a specific role in spermatogenesis for H2A.X has been demonstrated as the ko mouse model does not make a sex body despite normal autosomal synapsis in early pachytene spermatocytes, before death at around stages IV, V follows (Fernandez-Capetillo et al. 2003b). H2A.X was also found to have a role in formation of the bouquet stage at the onset of zygotene, explaining the effect of the kinase ATM on this process (Fernandez-Capetillo et al. 2003a, b). As indicated before, from late zygotene on, just before sex body formation, both X and Y chromosomes are marked by S139 phosphorylation of H2A.X, the signal disappearing at dissolution of the sex body in late diplotene (Mahadevaiah et al. 2001) when the sex bivalent chromosomes visualize. As the mouse Y is largely devoid of RAD51-signaled SPO11-induced ds DNA breaks (Moens et al. 1997), phosphorylation is indicative for chromatin remodeling alone, the marker appearing 1 2 days before H3.1/3.2 nucleosome eviction starts. For the proximal part of the Y and for the X, management of meiotic ds DNA breaks likely is equally involved in H2A.X S139 phosphorylation. Whether gH2A.X is a prerequisite for the next stage of H3.1 nucleosome eviction cannot be answered at this stage.

8.3

Spermiogenesis and Mature Sperm

In mouse round spermatids, the conspicuous AT-rich, hence DAPI-intense pericentric constitutive heterochromatin aggregates in one chromocentre, with the sex chromatin adjacent to it (Namekawa et al. 2006; van der Heijden et al. 2007). This

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configuration does not change upon entering the Golgi and capping phases of acrosome assembly in round spermatids. According to expectation, in the mouse, the gonosomal absence of H3.1/3.2 is maintained during the round spermatid stage (van der Heijden et al. 2007). As mentioned before, the combination of H3.3 with H2A.Z that is present in mouse spermatid sex chromatin (Greaves et al. 2006) is claimed to destabilize the nucleosome (Jin and Felsenfeld 2007) or alter linker histone adherence/facilitating chromatin remodeling (Braunschweig et al. 2009; Thakar et al. 2009). This would seem to be counterproductive for strict control of postmeiotic gonosomal gene expression (Mueller et al. 2008; Homolka et al. 2007; Namekawa et al. 2006), the escape of which seems associated with spermiogenesis mischief (Homolka et al. 2007; Reynard and Turner 2009; Burgoyne et al. 2009). However, the functioning of H2A.Z is context determined as judged by its different roles such as for heterochromatin formation and in the context of gene function insulation (Zlatanova and Thakar 2008). Chromatin remodeling and formation of the compact sperm nucleus has been the subject of a number of recent reviews (Pradeepa and Rao 2007; Ward 2010; Rousseaux et al. 2005; Miller et al. 2010; Eirin-Lopez and Ausio 2009; Gaucher et al. 2010). Histone variants, notably those for H1, H2A, and H2B, do play a role in this process (Table 8.1). Round spermatid development very early confers asymmetry to the cell. Acrosome and developing flagellum plus centrosomes are clearly visible at opposite sides of the nucleus from mouse/rat stage VII on. The acrosome contacts the nucleus at stage IV in these species, about halfway the round spermatid stage that lasts till stage VIII. The fact that the nucleus positions eccentrically at stage VIII at the side of the acrosome that now touches the cell membrane is a further consequence of this developmental cellular asymmetry. Nuclear elongation is not the automatic consequence of chromatin remodeling as the acrosome (Yao et al. 2002; Kang-Decker et al. 2001), perinuclear theca with acroplaxome formation, and the Sertoli cell F-actin hoops at the side of docking all play a role (Kierszenbaum et al. 2007). The asymmetry can be made visible by the histone variant H1T2 (Table 8.1) (Martianov et al. 2005; Tanaka et al. 2005) and the HMG4 chromatin protein (Catena et al. 2009) that are mutually needed to confer polarity. When H1T2 is lost, so is the posterior localization of HMG4. The anterior localization of H1T2 is independent from acrosome formation, but is destroyed by factors that help to organize chromatin such as the transcription factor TRF2 and HMGB2 (Catena et al. 2006).This chromatin gradient is subsequently maintained and later also visible in the direction of protamine cysteine cross linking that in the testicular late spermatids starts apically, during epididymal transit moving toward the tail in wavelike fashion. When staining mouse sperm with CMA3, that in human in vitro fertilization (IVF) laboratories is advocated to show more open histone-rich chromatin, caput epididymis sperm stains in the posterior part of the nucleus. As chromatin remodeling is completed before transfer to the epididymis, this staining pattern most likely reflects protamine thiol cross linking moving from anterior to posterior. The direction of chromatin remodeling upon fertilization follows the

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reverse direction, starting at the posterior side (for an illustration, see van der Heijden et al. (2005)). Homozygosity for H1T2 ablation results in subfertile males with aberrant sperm due to incomplete chromatin packaging at spermatid step 11 and the presence of cytoplasm at step 13. Apart from H1T2 that has a human homologue (Martianov et al. 2005) that has not yet been tested for a comparable role, more histone variants are connected with chromatin remodeling in elongating spermatids (see Table 8.1). Putatively involved with chromatin remodeling is the H1 variant HILS1 (Yan et al. 2003). HILS binds to nucleosomes with a lower affinity than H1a. The expression of H1t ends when HILS1 (and also H1T2) appears which is at the onset of nuclear elongation at IX, suggesting a role in chromatin remodeling (for an excellent review on linker histones, see Godde and Ura (2009)). The phosphorylation pattern of H1t at this stage suggests reduction of binging strength to DNA, which fits the concept (Rose et al. 2008). In the human, indirect evidence has been obtained for a role of HILS insufficiency in asthenozoospermia as RNA levels of HILS and the TNP1 and TNP2 transition proteins were lower in sperm cells from these patients (Jedrzejczak et al. 2007). H2AL1 and 2 are strongly enriched in step 12 16 mouse late elongating spermatids and are involved in the formation of the pericentric heterochromatin sperm chromocentre (Govin et al. 2007). The experimental evidence indicates that TH2B prefers H2AL1 and 2 in an unusual nucleosome lacking H3 and H4 (Govin et al. 2007). Incorporation of H2AL2 into the nucleosome shortens the bound DNA to 130 instead of 146 bp, leads to a more open configuration and also changed the interaction with chromatin remodeling complexes (Syed et al. 2009). The nontestisspecific H2A.Bdb variant was recently discovered during mouse spermiogenesis and in human sperm (Ishibashi et al. 2010) where it has not yet been visualized. These localizations suggest a role in preferential nucleosome occupation of sperm DNA. H4K8ac and H4K12ac are found in constitutive-centric heterochromatin in elongating spermatids and fertilizing sperm (Godde and Ura 2009; van der Heijden et al. 2006). Hence, a variation in the nucleosome composition in the mouse sperm chromocentre is likely. The first report pointing at nucleosome enrichment for centromeric regions made use of CENP-A IF in bull sperm (Palmer et al. 1990). This procedure was repeated for the human with the same result (Zalensky et al. 1993). However, Wykes and Krawetz, using DNA fractionation of human sperm, based on the fact that in a condition of increasing salt, histones are released earlier than protamines, could not find an enrichment for histones relative to protamines for centromeric DNA (Wykes and Krawetz 2003). After gamete fusion, an antibody recognizing both H2AL1 and H2AL2 was unable to detect these proteins in male mouse pronuclei (Wu et al. 2008). This observation does not preclude a function for H2AL1/2 in paternal chromosome structure and/or chromatid segregation in early development, but makes this less likely. Two H2B variants play a role in spermiogenesis among which is TH2B with its reported nonuniform presence in a human sperm population (van Roijen et al. 1998) using the anti tyrosine hydroxylase monoclonal antobody. When somatic nucleosomes containing H2A, H3, and H4 were reconstituted using TH2B, decreased

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stability was suggested (Li et al. 2005). The surprising fact is that in mature sperm, this histone is present in 20 40% of nuclei. More recently, Zalensky and co-workers have tried to answer if the presence of TH2B is a positive or negative selection marker. In a Xenopus egg extract sperm nuclear expansion essay, TH2B positive sperm were found to form pronuclei more readily, which could be interpreted to be a positive sign (Singleton et al. 2007a). However, in a double staining with CMA3 (see above), a positive correlation was found between CMA3 staining for nuclear immaturity and a positive IF signal for TH2B after sequential heparin induced nuclear decondensation (Singleton et al. 2007b). In a comparative investigation into nuclear characteristics of normospermic and oligospermic sperm samples, Ramos et al. (2008) found a higher presence of TH2B in the normospermic sample. In a subsequent project, TH2B correlated with higher density centrifugation selected sperm samples for both normospermics and oligospermic probands (de Mateo and Ramos, personal communication). In short, TH2B might be a positive nuclear marker for mature sperm. The X-linked H2BFWT variant (Table 8.1) does not change the stability of the nucleosome but resists chromatin contraction (Boulard et al. 2006) and most likely has a function in telomere maintenance in human sperm (Boulard et al. 2006; Churikov et al. 2004b; Gineitis et al. 2000; Zalenskaya et al. 2000). From its promoter CREM binding site, expression is expected in round spermatids where the protein is found (Churikov et al. 2004b). No studies are known that relate to a proposed function, apart from preserving molecular memory for a key element of chromosome structure and function. One could speculate on a role in telomere maintenance at the early cleavage divisions when telomeres are known to increase in length in the mouse (Liu et al. 2007) and in the transfer of a telomeric chromatin state to paternal human blastocyst chromosomes when telomerase activity is sharply increased (Wright et al. 2001). It cannot be excluded that the histone variants that are part of the nucleosomal make up of mature sperm did function in the chromatin remodeling process per se without any consequence for further embryonic development. This could explain the prominent presence of H2A.X in western blots of human sperm samples (Banerjee et al. 1995; Gatewood et al. 1990). Here, H2A.X likely underlines the importance of DNA repair pathways in chromatin remodeling at elongation (Leduc et al. 2008a, b). Double-strand DNA breaks needed to alleviate nucleosomeinvolved supercoiling are generated by TopoisomeraseIIB activity (Laberge and Boissonneault 2005), the enzyme being able to religate the break. Chromatin in elongating spermatids bears the marks of DNA repair linked chromatin remodeling as in mouse and rat, elongating spermatids are positive for gH2A.X (rat (MeyerFicca et al. 2005), mouse (Leduc et al. 2008a; Blanco-Rodriguez 2009)). The polyribosylation product PAR is at these stages found in the rat (Meyer-Ficca et al. 2005) and the inducing enzyme PARP1 in the mouse (Ahmed et al. 2010). In human elongating spermatids, polyribosylation has been found to be active (Maymon et al. 2006) and the presence of PARP proteins can be shown in sperm fractions, correlating with maturity (Jha et al. 2009). Another indication for some active DNA metabolism at spermatid elongation is found along a genetic route by

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studying triplet expansion in a mouse Huntington’s disease model. Triplets were shown to increase in number at the elongating spermatid stage (Kovtun and McMurray 2001). Pivotal to the theme of this chapter is if histone variants are only instrumental in chromatin remodeling or, when maintained in the mature gamete, gene action after fertilization is influenced, touching upon the landscape of transgenerational epigenetic effects. The absence of H1 from mature sperm (Gatewood et al. 1990) would for this histone type argue for a role during chromatin remodeling only. The presence of histones in sperm is already known for a long period (Gatewood et al. 1987, 1990) and has recently been explored in the mouse (Govin et al. 2007). H2A, H2A.X, H2A.Z, H2AL1, H2AL2, H2B, TH2B, H3.1, H3.3, CenP-A, and H4 have been shown to be present. The first insight that coverage of the genome in sperm by nucleosomes is nonuniform at the level of the gene came from the demonstration of local histone enrichment in the b globin gene cluster (GardinerGarden et al. 1998) and the PRM1, PRM2, TNP2 gene cluster (Wykes and Krawetz 2003). Indications for histone enrichment in chromosomal areas with specialized functions such as telomeres and centromeres have been obtained, as reported above. Maintenance of histones is likely mandatory for chromosome remodeling postsperm oocyte fusion, giving the oocyte a male chromatin environment for instance guiding histone dimer deposition (de Boer et al. 2010; Ward 2010; van der Heijden et al. 2006). Models for chromatin assembly within the sperm nucleus assume matrix-associated linker regions to be more accessible for nucleases, hence to be enriched for nucleosomes at a periodicity of around 50 kb (for a recent review, see Ward (2010)). Also, this periodicity could aid in reconstructing DNA replication competent chromosomes for embryonic development (de Boer et al. 2010).

8.4

Significance for Nucleosomes, Histone Variants for Embryonic Development

Absolute proof for a contribution of paternal histones to embryonic development has not yet been obtained, but there is some strong circumstantial evidence. First, van der Heijden et al. (2006) could by microscopic observation follow paternal nucleosomes at sperm nucleus expansion upon gamete fusion. Nucleosomes were followed by IF for H4K8ac and H4K12ac that are enriched in the mouse sperm chromocenter. The signal for H4K12ac was readily overwhelmed by H3H4 dimer deposition as H4K12ac is a marker for this process (Sobel et al. 1995). Evidence for persistence of paternal nucleosomes up to zygotic S-phase was obtained by use of an antibody against H3.1/3.2 (van der Heijden et al. 2005) that is present in sperm (Gatewood et al. 1990), and is not obscured by de novo nucleosome formation that uses H3.3 H4 dimers (Torres-Padilla et al. 2006; van der Heijden et al. 2005). Both heterologous ICSI with mouse oocytes and tripronuclear human zygotes were used to arrive at this conclusion, the signal being more dispersed in the heterologous

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situation (Ramos et al. 2008). Centric heterochromatin is predicted to be enriched for H3.1/H3.2 (Bernstein and Hake 2006) which could fit the observations in the human zygotes. Research into a function for nucleosomal DNA in sperm was only recently possible by the advent of the development of methods that allow isolation of either nonprotaminated DNA (to contrast with the protamine bound fraction) and/or the nucleosomes from the ultra compact sperm nucleus for use in comparative genome hybridization annex CHIP on chip and chip seq methodologies (Arpanahi et al. 2009; Hammoud et al. 2009; Brykczynska et al. 2010). Comparative genome hybridization by an Agilent’s whole genome tiling array, with a median probe spacing of 44 K, showed the soluble nucleosomal DNA to be enriched in gene-dense regions. Further analysis revealed that promoter sequences were overrepresented (Arpanahi et al. 2009), particularly those that were in proximity of CCCTC-binding factor (CTCF) binding sites ((Phillips and Corces 2009), transcripts of this gene hardly being present in testicular germ cells (http://biogps. gnf.org)). The conclusion is that a motive that helps in three-dimensional structuring of somatic chromosomes is preferably in a nucleosomal context in sperm. The CHIP on chip and chip seq studies of Hammoud and co-workers extended on these conclusions (Hammoud et al. 2009). By GO ontology analysis, nucleosomal promoters were found to be especially enriched for transcription factors and signaling pathways in embryonic development. Western blots showed sperm chromatin to contain H3K4me3 and to a lesser extend H3K4me2 and H3K27me3. At closer inspection, H3K4me3 was enriched for genes, previously activated because of their roles in spermatogenesis. The H3K27 mark was found on genes that have to be inactive in ES cells to prevent differentiation. The histone variants TH2B and H2A.Z were by these authors also tested for functional enrichment. Nothing was found for H2A.Z. TH2B was involved with genes that are important in sperm biology, capacitation, and fertilization. This would fit the indications presented earlier that TH2B is a positive marker for human sperm cells. MicroRNA gene clusters showed some enrichment for nucleosomes. In a recent recapitulation of this research theme, general conclusions were substantiated and extended to the mouse. This points to an evolutionary conserved pattern of histone occupancy in the male gamete (Brykczynska et al. 2010). Imprinted gene clusters (Hammoud et al. 2009) were especially interesting, in that nucleosomal enrichment was found at both male- and female-expressed genes implicating the histone code, but not specifically a histone variant to be involved with transmission of monoallelic expression to the next generation. Paternally expressed genes were characterized by H3K4me3, paternally silenced genes by H3K9me3. The fact that for imprinted genes in the mouse, subsequent gene activity can be read from the histone PTMs during spermiogenesis up to stage XII when compaction starts, was found by Delaval et al. (2007). Here the analysis was not extended beyond the elongated spermatid stage XII as without pretreatment, micrococcal nuclease digestion will not work on the compacted chromatin, present from this stage on.

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As introduced before, at paternal chromatin remodeling that follows immediately upon gamete fusion, dimers of H3.3 and H4 replace protamines (Torres-Padilla et al. 2006; van der Heijden et al. 2005). One specific chaperone for this dimer, HIRA, colocalizes with male chromatin from the onset of sperm nucleus expansion and chromatin remodeling (van der Heijden et al. 2005). At the first S-phase, semiconservative DNA replication and use of H3.1/3.2 histones from the oocyte will dilute the contribution of H3.3 to not only paternal but also maternal nucleosomes. The paradigm that H3.3 stands for active chromatin (Hake and Allis 2006) is not kept during the first cell cycle as the maternal to zygote transition in the mouse takes place at the two-cell stage when the embryonic genome awakens (Zeng et al. 2004). Some leaky transcription in the mouse is from the male pronucleus (Aoki et al. 1997) that might be regarded as an expression of epigenetic asymmetry. The male pronucleus is low in H3,H4 repressive PTMs (Santos et al. 2002; van der Heijden et al. 2005) (but rich in Polycomb repressive marks (Puschendorf et al. 2008)). There are a number of reasons why separate analysis of sperm nucleosomes for those that carry H3.1/H3.2, TH3 versus those that carry H3.3 is worthwhile. By transplantation of endodermal nuclei in Xenopus oocytes, Ng and Gurdon (2008) detected that transcriptional memory for many cell generations was conferred to H3K4 methylation of H3.3. More recently, Katz et al. (2009) discovered that germline memory likely among other factors depends on the proper H3K4 methylation status. When using hypomorphic alleles of a H3K4 demethylase over many generations of propagation in C elegans, reproductive catastrophe sets in for spermatogenesis. As in male infertility, incomplete chromatin remodeling at gamete formation is indicated (Ramos et al. 2008; Zhang et al. 2006), it would be of interest to know at what level of H3K4 methylation for both replication-dependent and replication-independent H3 variants, genes/promoters have to be passed on to the next generations to achieve a balance between maintenance of transcriptional memory and its eradication to prevent accumulation of epigenetic factors that in the end would lead to male sterility (Katz et al. 2009).

8.5

Male Transgenerational Inheritance and Effects

As is clear from the previous section, the challenge of studying histone variants in mammalian transmission is to find handles for transgenerational epigenetic inheritance (Ooi and Henikoff 2007). The prime example of epigenetic inheritance in mammals is genetic or genomic imprinting (Bartolomei 2009; Ideraabdullah et al. 2008). Following the definitions introduced by Youngson and Whitelaw (2008), the term transgenerational epigenetic inheritance is in place when the mechanism of inheritance is known as with genomic imprinting. If a phenotype is transmitted but the mechanism not proven, the term transgenerational epigenetic effect is coined (Youngson and Whitelaw 2008). Interruption of female parthenogenetic development at around the phase of organogenesis close to day 10 postconception (McGrath and Solter 1984; Surani

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et al. 1984) is the reflection of incorrect dosage of the paternal active copies of maternally imprinted genes and vice versa the maternal active copies of paternally imprinted genes (Cattanach et al. 2006; Bartolomei 2009). By ingenious oocyte manipulation, that yields digynic embryos, it was discovered that if the two genomes were from nongrown primary oocytes, prenatal development was extended by 3 days (Kono et al. 1996) which can be explained by the fact that imprinting marks are laid down in the growing primary oocytes of growing follicles (Lucifero et al. 2007). By genetic engineering, one imprinting control region (ICR) including the H19 transcriptional unit on mouse chromosome 7 was deleted, and a nongrowing primary oocyte of this genotype was fused with a meiosis-competent enucleated oocyte. The derived meiotic metaphase II spindle could be transplanted into a normal secondary oocyte leading to maternal diploidy. Limited numbers of live offspring were obtained (2/371 oocytes) (Kono et al. 2004). So when only one of the three male germline-methylated ICRs was deleted in the female, some prenatal development became possible. When this principle was expanded to include a second male imprinted ICR (Dlk1 on mouse chromosome 12), the efficiency of reproduction was markedly increased as suggesting there was no further male role from the epigenetic point of view (42 pups were developing to term from 286 blastocysts transferred) (Kawahara et al. 2007). Data for prenatal survival of a technical control with paternal and maternal genomes were not supplied. Postnatal survival was comparable to IVF controls (Kawahara et al. 2007). The above evidence would suggest that apart from classical imprinting, a male epigenetic transgenerational contribution does not exist, at least not for one generation. Multigeneration transmission experiments, however, could suggest otherwise. Examples of epigenetic male inheritance in mice mainly concern insertional retroviral intracysternal A particle (IAP) mutagenesis with the Avy and Axinfu alleles of the agouti (A, hair color pattern) and Axin (regulates embryonic axis formation via Wnt signaling pathway) genes as standing examples (Morgan et al. 1999; Rakyan et al. 2003). For the A viable yellow (vy) allele, the epigenetic aspect of inheritance usually is shown in maternal transmission. In certain maternal backgrounds (129P4/RrRk), this can also be paternal (Rakyan et al. 2003). When the female is heterozygous for the Mei2 member of the polycomb repressive complex 1 (PRC1), male epigenetic inheritance of Avy can be observed in a C57BL/6 background (Blewitt et al. 2006). A paternal allele of Avy can also be modified through in vitro culture from the zygote to the blastocyst stage (Morgan et al. 2008). Epigenetic inheritance is read by the CpG methylation status of the long terminal repeats (LTR) of the IAP that if unmethylated, assumes promoter activity, leading to the continuous expression of yellow hair pigment (Morgan et al. 1999). When variably methylated, a continuum of darker phenotypes is produced, culminating into pseudo-agouti. Further examples of male multigeneration transmission of an epigenotype, including one that has irregular replication of a long expanded simple tandem repeat (ESTR) as the readout, have been found in the area of irradiation mutagenesis (Barber et al. 2002; Barber and Dubrova 2006; Dubrova et al. 2000). This

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field of male irradiation stress-induced multigenerational effects has been the subject of a recent review, that includes speculation as to the mechanism(s) of transmission involved (de Boer et al. 2010). Also sperm RNA has been proven to be of significance for male transgenerational epigenetic inheritance (Rassoulzadegan et al. 2006). Sensitivity of the embryonic primordial germ cell stage (when normally there is erasure of DNA CmpG methylation and many histone PTMs) for male transgenerational epigenetic perturbation has recently been demonstrated by Lee et al. (2009) when they studied offspring after an in vitro culture phase of these developing gonocytes. Analysis into the F4 generation was suggestive for an expanding imbalance as measured through mainly the methylation status of the ICR of the male-imprinted H19 gene. A normal H19 epigenotype could after transmission be changed into an abnormal one and vice versa. In this study, transmission was mainly via the male germline with some female germline passages also. The authors examined histone posttranslational modifications such as H3K9me2, H3K9me3, H3K27me1, H3K27me3 of the H19 ICR in the embryonic germ cell culture and found deviations from normal. It was known for a long time that the human sperm nucleus is relatively rich in nucleosomal chromatin, assessments ranging from 4 to 15% of DNA (Gatewood et al. 1987, 1990; Hammoud et al. 2009). In sperm samples from men with oligoasthenoteratozoospermia (OAT) that are used in the IVF clinic, values on average are higher (Ramos et al. 2008; Zhang et al. 2006) which poses questions as to the information content of paternal nucleosomal DNA for embryonic development and beyond (van der Heijden et al. 2006, 2008). Also via a genetic approach, there is some evidence that the chromatin environment during gametogenesis can affect the variation in expression of (in this case) the maternal Avy gene. Chong et al. (2007) found that when the male was heterozygous for a hypomorphic allele of Smarca5 ¼ Snf2h, member of the ISWI family of chromatin remodeling protein complexes, in the Smarca5 wildtype offspring this caused a shift to lighter, more yellow coat colors, as if the male gamete could influence the CpG methylation content of the LTR promoter of the maternal Avy allele. The authors postulate it to be the protein itself that was transmitted via the sperm cytoplasm. More likely is a route in which embryonic expression of paternal target genes is changed, for instance by their histone occupancy/promoter code during spermatogenesis, which could fit the fact that Snf2h is highly expressed in the male germline.

8.6

Future Directions

Interest into not strictly genetic aspects of information transfer between generations is growing. In the exploration of the theme that the male contribution is bigger than just DNA, several aspects have been touched upon, such as RNA content, the nonrandom distribution of nucleosomes, the meaning of histone PTMs, and the

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possible value of histone variants. Also other aspects of sperm nuclear structure such as loop domain organization are worth studying. With interest arising in these softer aspects of the paternal contribution, chromatin research of female chromatin has not yet received the attention it should, collection of material likely being a factor why this area is not yet well explored. Further studies in both male and female chromatin at transmission are needed as they may reveal a better definition of the conception of gamete quality. This expression has been used for a long time to define those gametes that contribute to the next generation in an efficient way. As by artificial reproductive techniques the spectrum of gametes included is growing, a better reference to “normality” is needed. Composition of chromatin may well offer a tool for the characterization of gametes. As this, up to now, offers too big a target, and also for conceptual reasons, imprinted gene regions have been selected for more detailed study. This field first has to advance by the application of genetic methods to evaluate the contribution of genes that are involved in chromatin maintenance and chromatin modification. In this context, multigeneration experiments are needed. As chromatin remodeling is so prominent in the male germline, spermatogenesis is expected to be a sensitive readout in such systems. However, male and female gametogenesis and contribution to the next generation have to be evaluated both separately and interactively.

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Chapter 9

Genome Organization by Vertebrate Sperm Nuclear Basic Proteins (SNBPs) Juan Ausio´, Laurence R. Brewer, and Lindsay Frehlick

Abstract DNA is tightly packed in the sperm nucleus through its association with specific chromosomal proteins that are heterogeneous in size and known as sperm nuclear basic proteins (SNBPs). Despite their structural diversity, SNBPs are evolutionarily related and can be classified into three major groups or types: histone (H-type); protamine (P-type), and protamine-like (PL-type). The three types are widespread amongst vertebrates. During spermiogenesis these proteins replace the somatic histones that are present at the onset of spermatogenesis. Mammals exhibit an increased level of complexity as transition proteins (TPs) temporarily bind to DNA before being replaced by protamines in the mature sperm. The proper sequential chromatin remodeling is dependent on global post-translational modifications (PTMs) of the chromosomal proteins involved, including acetylation and phosphorylation. A transient 20 nm chromatin fiber that is independent of the SNBP type is often formed. The temporally and spatially organized compaction of chromatin during spermiogenesis may be important for the understanding of post-meiotic events such as gene expression, Huntington’s disease CAG expansion, and the remnant histones that are present in the mature sperm of certain mammals, including humans. Alterations in SNBP composition result in DNA damage and infertility, underscoring the importance of these proteins for male germline genome integrity.

The name SNBP refers to the basic proteins that are associated with DNA in the tightly packed chromatin that is found in the nucleus of mature sperm. This name was informally proposed in the course of the Seventh International Symposium on Spermatology at Cairns, Australia, in 1994. The meeting was attended by several of the researchers working with these proteins at that time including: Dominic Poccia, Norman Hecht, Marvin Meistrich, Harold Kasinsky, Lluis Cornudella, and Juan Ausio´. It provides a description following a decreasing order from the cell type (sperm) organelle (nucleus) to the molecular aspects (basic) of the proteins. J. Ausio´ (*) and L. Frehlick Department of Biochemistry and Microbiology, University of Victoria, Petch Building 258a, Victoria, BC, Canada V8W 3P6, e mail: [email protected] L.R. Brewer Center for Reproductive Biology, Gene and Linda Voiland School of Chemical Engineering and Bioengineering, Washington State University, Pullman, WA 99164 2710, USA

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 9, # Springer Verlag Berlin Heidelberg 2011

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Keywords Chromatin Histone Protamine Transition protein

9.1

Packing the Paternal Genome with Small and Large Sperm-Specific Nuclear Basic Proteins

In contrast to somatic chromatin where the DNA is found associated mainly with histones, the tight chromatin organization that is observed in the sperm nucleus results from the interaction of DNA with sperm nuclear basic proteins (SNBPs) (Eirin-Lopez and Ausio 2009). The SNBPs constitute a family of chromosomal proteins that is very heterogeneous in size. Protein members range in size from the small protamines of salmonid fish (30 40 amino acids) to the large protamine-like protein of the winter flounder Pseudopleuronectes americanus (approximately 1000 amino acids) (Kennedy and Davies 1982). Experimental evidence accumulated during the last two decades has shown that SNBPs can be classified into three main types: histone (H-type), protamine (P-type), and an intermediate group that has been referred to as protamine-like (PL-type) (Ausio´ 1999; Eirin-Lopez and Ausio 2009; Eirı´n-Lo´pez et al. 2006a). The term protamine was first used by Miescher in 1874 to refer to an organic base extracted from salmon sperm nuclei (Miescher 1874). The word histone was coined by Kossel 10 years later. He combined the terms histos (tissue) and peptone (as proteins had not been discovered yet) to describe the nature of the extracts obtained from the nucleated erythrocytes of goose (Kossel 1884). The histone H-type SNBPs involve a group of proteins that are compositionally and structurally similar to the core (H2A, H2B, H3, and H4) and linker (H1/H5) histones that are found associated with the chromatin of the somatic tissues. Hence, somatic histones and SNBPs of this type exhibit a similar electrophoretic behavior (see Fig. 9.1, lane 1). They are enriched in both lysine and arginine but their overall basic amino acid composition never exceeds 25 30% mol/mol (Fig. 9.1b). In contrast, the P-type consists of proteins with over 30% arginine (Fig. 9.1b) that are often smaller than histones (30 100 amino acids) and exhibit a higher electrophoretic mobility (Fig. 9.1, lane 3). In addition to arginine, the members of this type can also contain substantial amounts of cysteine, an amino acid seldom present in chromosomal proteins. Cysteine-rich protamines are present in chondrichthyan fish (Kasinsky 1989) and eutherian mammals (Ausio et al. 2007). The term protaminelike was first used by Subirana to refer to the intermediate compositional nature (arginine þ lysine content usually amounts to at least 35 50% mol/mol) (Fig. 9.1b) between histones and protamines of the members of this group (Subirana et al. 1973). These proteins exhibit a wide range of molecular masses ranging from 5000 to 100,000, and thus exhibit a broad range of electrophoretic mobility. They are all structurally and evolutionarily related to a histone H1 ancestor (Eirin-Lopez and Ausio 2009).

9

Genome Organization by Vertebrate Sperm Nuclear Basic Proteins (SNBPs)

a

b Ictalurus H1x CM

H1 H5 H3 H2B H2A

1

2

3

215

(H)

1

11

AAAPVQPSQP

AAKKKGPASK AKPA SAEKKN

41

51

61

SAQAEETAPE

21

31

71

KKKKGKGPGK

YSQLVINAIQ

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Fig. 9.1 The three main types of vertebrate (fish) SNBPs. (a) Electrophoretic analysis of proteins representative of the three SNBP types in fish. Lane 1: Catfish (Ictalurus punctatus); Lane 2: Red mullet (Mullus barbatus); Lane 3: Salmon (Onchorrhynchus keta). The electrophoresis was carried out in polyacrylamide gels (PAGEs) consisting of 2.5 M urea and 5% acetic acid (Ausio´ 1992). (b) Amino acid sequence of several of the SNBPs shown in (a). Ictalurus histone H1x [from I. punctatus, accession number: AAQ99138] (Evans et al. 2005); Mullus PL I [from M. surmule tus, accession number: Q08GK9] (Saperas et al. 2006); and protamines (Salmine) from O. keta (Hoffmann et al. 1990). The boxes highlight the sequence corresponding to the trypsin resistant winged helix domain that is characteristic of linker histones (Eirin Lopez and Ausio 2009). CM chicken erythrocyte histone marker

9.2

Packing Sperm Chromatin. The Roles of Acetylation and Phosphorylation

The transition from nucleosomally organized somatic chromatin at the onset of spermatogenesis to the highly compacted chromatin state found in mature sperm involves one of the most remarkable examples of global chromatin remodeling (Fig. 9.2a b). It is assisted by post-translational modifications (acetylation, phosphorylation, and ubiquitination) of both the chromosomal proteins and the chromatinremodeling complexes through processes that are not yet well understood (Gaucher et al. 2010). In many vertebrate organisms, the final chromatin conformation in the mature sperm, as a result of the replacement/displacement of histones by

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Fig. 9.2 Chromatin transitions during spermiogenesis in organisms containing different SNBP types. (a) At the onset of spermatogenesis, somatic chromatin is organized into fibers of repetitive nucleosome complexes. These polynucleosome arrangements may adopt an extended fibrillar conformation, which is characteristic of transcriptionally active euchromatic (Euc) domains, or ˚ fibers, which are usually present in inactive heterochromatin (Het). (b) The further fold into 300 A

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protamines or protamine-like SNBPs, is completely different than that of somatic cells (Fig. 9.2c). Before discussing what is known about the chromatin organization and transitions in sperm, it is important to briefly describe the structures of the SNBP members. SNBPs of the H-type, like somatic histones, can be classified into two major categories: core histones (H2A, H2B, H3, and H4) and linker histones. The former contain characteristic histone-fold domains that are flanked by N- and C-terminal intrinsically disordered domains that can adopt a certain extent of secondary structure upon interacting with DNA (Baneres et al. 1997). The histone fold (Arents and Moudrianakis 1995) is a dimerizing domain that allows these proteins to form an octameric protein complex that serves as a core around which approximately 146 bp of DNA are wrapped forming the basic structural subunit called the nucleosome. Linker histones contain wing helix domains that are also flanked by intrinsically disordered N- and C-terminal domains (Roque et al. 2005; Vila et al. 2001) (Fig. 9.2c). PL-type proteins can be grouped into large PL (PL-I) proteins with a similar structural organization to that of linker histones and small PL proteins (<100 amino acids) with structural similarity to either the N- or C-terminal domains of linker histones (Eirı´n-Lo´pez et al. 2006b). Like the intrinsically disordered domains of linker histones and PL proteins, protamines (P-type) (Fig. 9.2c) take on a secondary structure upon binding to DNA (Hud et al. 1994). In contrast to core histones the SNBPs all bind to linear DNA, like the histone H1 family which binds to linker DNA, and hence it is not surprising that they share a common phylogenetic origin (Eirin-Lopez and Ausio 2009). The occurrence of these three SNBP types in different vertebrates is shown in Fig. 9.2c. Despite the presence of three structurally distinct SNBP types, it has been shown in certain organisms with each type that the chromatin fiber is reorganized into ˚ granules at the onset of spermiogenesis (Kurtz et al. 2009; Martinez-Soler 200 A et al. 2007a; Ribes et al. 2001; Worawittayawong et al. 2008) (Fig. 9.2b). This fiber contains acetylated H4 and H3 and it has been proposed to fulfill the minimum functional requirement representing the evolutionary ancestral chromatin conformation that precedes sperm chromatin condensation in animals (Kurtz et al. 2009). ä Fig. 9.2 (Continued) initiation of spermiogenesis in many organisms involves the acetylation of histones H3 and H4 (represented here by darker nucleosomes) and the formation of a more ˚ in diameter, homogeneous chromatin organization consisting of granules of approximately 200 A regardless of the SNBP type of spermiogenesis (Kurtz et al. 2009). This precedes (c) the replacement of somatic like histones by specialized SNBP of the H, PL, or P types that result in ˚ cross sectional diameters the densely packed fibrillar or granular structures with 300 400 A (Eirin Lopez and Ausio 2009). In vertebrates, the three types of SNBPs are represented across the phylogenetic groups (Ausio´ 1999; Eirin Lopez and Ausio 2009). In mammals the replacement of histones by protamines is preceded by a transient replacement of the former by transition proteins (TPs) (see Fig. 9.4). A schematic representation of the protein structures of different SNBPs is also shown. HC, histone core, HF, histone fold, and WHD, wing helix domain. The asterisk indicates that in H type spermiogenesis, it is not clear whether or not the histones that are present in the mature sperm correspond to distinct histones that are expressed from spermiogenesis specific genes

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Confirmation of the universal nature of this fiber in mammals is still needed. Histone H4 has been shown to be acetylated at multiple sites at this stage in fish with P-type (Oliva et al. 1987, 1990) and H-type SNBPs (Kurtz et al. 2009), in birds (Oliva and Mezquita 1986), and in mammals (Hazzouri et al. 2000; Lahn et al. 2002). This acetylation takes place at a time when mammalian histone H1t is extensively phosphorylated (Rose et al. 2008; Sarg et al. 2009) (Fig. 9.3) and histones start being replaced by protamines (Fig. 9.3). Phosphorylation of histone H1 increases its dynamic exchange with chromatin (Dou et al. 2002) and mimics H1 removal (Dou et al. 1999), and core histone acetylation opens the chromatin ˚ structures (Fig. 9.2b) are highly reminiscent of the relaxed fiber such that the 200 A acetylated chromatin in the absence of H1 (Garcia Ramirez et al. 1995). This relaxation is likely to play a critical role in facilitating the subsequent formation ˚ fibrogranular structures consisting of SNBPs (Fig. 9.2c). The of the 300 400 A molecular details involved in the transition are still not well understood and likely involve chromatin-remodeling factors such as the mammalian bromodomaincontaining protein Brdt (bromodomain testis-specific). Brdt co-localizes with hyperacetylated histone H4 in post-meiotic spermatids and is able to condense

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Fig. 9.3 Steps of spermiogenesis in the mouse. The spermatid type as well as a schematic representation of the spermatid shape and degree of chromatin condensation are indicated in the upper part of the figure. Some of the chromosomal protein transitions undergone during this process, including the replacement of the histones, the presence of histone H1t, the hyperacetyla tion of histone H4, the ubiquitination of H2A, the deposition of transition proteins (TP1, TP2), and the deposition of protamines (P1, P2), are indicated in the lower part of the figure. The extent of phosphorylation is shown in red. pP2 refers to the preprotamine 2 (see Fig. 9.3). Figure modified and adapted from (Zhao et al. 2004b)

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chromatin both in vitro and in vivo (Moriniere et al. 2009; Pivot-Pajot et al. 2003). The first bromodomain of Brdt was shown to cooperatively bind acetylated lysine 5 and 8 of histone H4, two marks that along with acetylated lysine 12 increase in postmeiotic elongating spermatids (Moriniere et al. 2009). Mice with a deletion of the first bromodomain of Brdt are sterile (Shang et al. 2007). Hence, the role of histone acetylation certainly goes beyond the mere displacement of histones by protamines initially proposed (Oliva and Dixon 1991), which is not surprising considering that ˚ fiber has also been visualized in vertebrates acetylation of histone H4 and the 200 A ˚ that contain H-type SNBPs in their mature sperm. In this latter group the 200 A fiber may allow for the incorporation of sperm-specific histone H1s. Despite the varied complexity of the interactions of DNA with the different SNBP types, the ˚ fibrilar organization (Fig 9.2c) resulting complexes all seem to exhibit a 300 400 A (Casas et al. 1981). This organization seems to be an intrinsic property of the chromatin protein DNA complexes (Subirana 1992). Phosphorylation of PLs and protamines is also important for the chromatin transitions described above and has been shown to facilitate the proper deposition of these proteins onto DNA (Lewis et al. 2003). Phosphorylation of transition proteins (TPs) in eutherian mammals appears to work in a similar fashion (Pradeepa and Rao 2007). It is likely that this PTM also plays a role in the formation of the manifold supramolecular chromatin structures of the lamellar (Martens et al. 2009), granular (Gusse and Chevaillier 1978; Saperas et al. 1993), and toroidal (Balhorn et al. 1999; Brewer et al. 1999; Ward and Coffey 1991) types that follow the initial chromatin transitions shown in Fig. 9.2c and that ultimately result in the high condensed chromatin state that is observed in the mature sperm nuclei (Wiesel and Schultz 1981).

9.3

Mammalian Transition Proteins

In mammals, the histone displacement from nucleosomes coincides with the deposition and binding of the TPs to genomic DNA (Figs. 9.2b and 9.3). These proteins are subsequently replaced by highly basic, arginine-rich protamines (P-type) that further condense and inactivate the spermatid genome. The primary mammalian TPs, TP1 and TP2, undergo deposition in the nucleus at the start of the elongation of the spermatid cell (Fig. 9.3). They have been observed in many eutherian mammals including the mouse, rat, boar, bull, ram, and man (Wouters-Tyrou et al. 1998). TP1 is a 6 kDa protein and TP2 is a 13 kDa protein and their sequences have been determined for the mammals listed above (see Fig. 9.4). Additional TPs, TP3 and TP4, have also been identified for these same species, but they are present in much lower quantities (Dadoune 2003; Wouters-Tyrou et al. 1998). The exact function of the TPs, particularly their role in histone displacement during mammalian spermiogenesis remains unclear. TPs exhibit an amino acid composition that is similar to that of the PL-type SNBPs. They contain both lysine and arginine residues in addition to histidine

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Fig. 9.4 Mammalian (mouse) SNBPs. (a) Electrophoretic analysis of mouse (Mus musculus) testicular (MT) proteins in comparison to a chicken erythrocyte histone marker (CM). The approximate site of migration of the mouse transition proteins (TPs) in this type of gel is indicated by the broken arrows. The proteins were obtained from whole testicular tissue upon pyridylethyla tion in the presence of urea followed by extraction with 0.4 N HCl. (b) Primary structure of the mouse TP and protamines [mouse TP1/TP2 accession numbers: AAB21244 and NP 038722, respectively]; [mouse P1/P2 accession numbers: P02319 and NP 032959, respectively]. The arrow heads in (a) and (b) point to the sites of post translational cleavage of the TP2 precursor (Debarle et al. 1995). The box highlights the mature TP2 product that is found in the mature sperm

(ArgþLysþHis ¼ 25 40% mol/mol), and like mammalian protamines they may also contain cysteine. Their size ranges from 50 to 120 amino acids (see Fig. 9.4). Their compositional similarity to small PLs led to the proposition that the ontogenic SNBP changes observed during mammalian spermatogenesis may recapitulate the phylogenetic transition from the H-type to the P-type that is observed in the course of metazoan SNBP evolution (Ausio´ 1999). The presence of histidine and cysteine in the highly evolved mammalian SNBPs is intriguing. In TPs these residues allow for the formation of zinc-fingers (Baskaran and Rao 1991) and in protamines they contribute to the formation of inter-fiber chromatin links increasing the chromatin compaction. In both instances they may also enhance DNA-binding ability (Brewer et al. 2002; Vilfan et al. 2004). A number of groups have studied the ability of the TPs, TP1 and TP2, to condense DNA in vitro (Caron et al. 2001; Kundu and Rao 1995, 1996; Levesque et al. 1998;

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Meetei et al. 2002). It was shown that TP2 condenses DNA more effectively than TP1 (Baskaran and Rao 1990, 1991) and preferentially binds to GC-rich DNA in a zinc-dependent way (Kundu and Rao 1995). It was demonstrated that the phosphorylation of TP2 prevents it from condensing DNA (Levesque et al. 1998; Meetei et al. 2002). This result supports the hypothesis that dephosphorylation of the spermatid nuclear basic proteins regulates the timing of the initiation of genome condensation. Brewer and Balhorn developed a technique which permitted the study of the kinetics of binding and condensation of TP1 and TP2 on single DNA molecules, avoiding the complications due to inter-molecular aggregation found in bulk studies (Brewer et al. 2002).This methodology also allowed the direct measurement of the equilibrium constants for the DNA-binding reaction for each transition protein. In contrast to the previous measurements (Kundu and Rao 1995, 1996; Meetei et al. 2002), it was found that both TP1 and TP2 condensed DNA with rates similar to the protamines, P1 and P2. It was also shown that the C-terminal domain of TP2 is responsible for condensing DNA in a way that was not affected by the presence or absence of zinc. At the beginning of mammalian spermiogenesis histones are nearly globally displaced from nucleosomes. The number of nucleosomes retained in humans is 15% and in the mouse is 4%. During the displacement process, each nucleosome releases approximately two negatively wound supercoils of DNA. However, in the genome of mature sperm there is an absence of DNA supercoils (Ausio´ 1995; Risley et al. 1986; Ward et al. 1989). Although the actual mechanism of supercoil release during spermiogenesis is not yet understood, using the TUNEL (terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling) assay, it has been established that global DNA-strand breaks transiently exist between steps 9 and 11 in Fig. 9.3 (Marcon and Boissonneault 2004). The origin of the strand breaks is unknown but has been attributed to type I topoisomerases, which have recently been identified in mature human sperm (Har-Vardi et al. 2007). A type I topoisomerase would temporarily cut a single strand of the DNA helix to relieve the superhelical tension of the DNA surrounding histone octamers, thus assisting in histone displacement. TP1 has been demonstrated to stimulate the ligation of DNA singlestrand breaks in vitro. Accordingly, a model has been proposed in which TP1 binds across the single-strand breaks optimally positioning the ends for ligation after the displacement of histones, in preparation for the condensation of genomic DNA (Caron et al. 2001). The timing of the deposition of the TPs during spermiogenesis suggests that they may be responsible for displacing histones during spermiogenesis. Although histone displacement was still observed to occur in knockout mice that were null in both TP1 and TP2, this does not indicate that they are incapable of displacing histones, only that another spermatid nuclear basic protein, such as P1, could assume that role in their absence, as it does during spermiogenesis in other vertebrates that lack TPs. Functional redundancy among the spermatid nuclear basic proteins has already been observed between TP1 and TP2 in knockout mice and may similarly occur between the TPs and protamines in knockout mice during histone displacement.

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Protamines. Packing Chromatin in an Orderly Fashion

Exhaustive reviews on protamines have recently been published (Balhorn 2007; Oliva 2006a). Therefore, in this section, we will mainly focus on mammalian protamines and a few of the poorly understood aspects of their deposition onto the genomic DNA during spermiogenesis. Protamines have a widespread distribution in vertebrates. Although they are all extremely arginine-rich (as stated earlier), in general their length has gradually increased in the course of vertebrate evolution with fish containing the shortest versions (Fig. 9.1b). As is the case in other Amniota (reptiles and birds), the sperm of mammals exclusively contains SNBPs of the P-type (see Fig. 9.2c). In the transition from metatherian to eutherian (placental) mammals protamines acquired cysteine, an amino acid that is otherwise quite rare in chromosomal proteins. This increase in compositional complexity was accompanied by the appearance of two distinctive P1 and P2 protamines (Fig. 9.4), the latter one being expressed only in some of the organisms within this group, including humans (Lewis et al. 2003). The biological significance of P2 is very intriguing and this protein undergoes a complex pattern of post-translational cleavage at its N-terminus (Fig. 9.4b) similarly to the N-terminal post-translational processing that has also been observed in some invertebrate protamines (Lewis et al. 2003) during spermiogenesis (Fig. 9.3). The functional relevance of the processing of P2 in mammals has been related to the protein half-life and plays an important role in the protamine deposition process (Hecht 1989). All of this results in a highly condensed packing of the mammalian male genome that presumably is optimized by the inter- and intra-chromatin disulfide bridges of the cysteine-containing protamines and by the presence of P2. An interesting phenomenon that has often been overlooked in the past is the orderly packing of chromatin associated with protamine deposition. Observations of this have been made in invertebrates, as for instance during maturation of sperm in Sepia (cuttlefish) (Martinez-Soler et al. 2007b). However, a similar situation in other vertebrates is not unlikely considering the non-random organization of chromosomes that is present in the sperm nucleus of birds and mammals (Foster et al. 2005; Tsend-Ayush et al. 2009), including humans (Zalensky and Zalenskaya 2007). Whether this is modulated to a certain extent by perinuclear microtubule structures, as is the case in Sepia (Martinez-Soler et al. 2007b), or by the nuclear matrix (Ward and Zalensky 1996), it is highly likely that the process of protamine deposition is a tightly regulated process rather than a mere uncontrolled diffusion of protamines throughout the spermatid nucleoplasm. This process could account for the unexpected nuclear metabolic activities, detected in the post-meiotic spermatids, which have recently received a lot of attention. It was initially believed that all transcription ceased during spermiogenesis (Steger 2001) at a time when protamines start being replacing histones. However, there is now plenty of evidence in support of post-meiotic gene expression in invertebrates such as in Drosophila (Barreau et al. 2008; Vibranovski et al. 2009) and in different vertebrates including fish (Schulz et al. 2010) and mammals

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(Chalmel et al. 2007). Also, Huntington’s disease CAG expansion in humans has been shown to occur late in male germ cell development as spermatids are entering the epididymis (McMurray and Kortun 2003). How these dynamic processes can take place in the midst of the tightly packed chromatin structure resulting from the protamine DNA interactions remains unclear but could be explained by a progressive temporal and spatial regular deposition of protamines in an orderly fashion. The orderly packing of sperm chromatin may help explain how some histones remain at the mature sperm of certain mammals (described earlier). From a biophysical point of view it is difficult to explain how the nucleosomes of these remnant histones are spared replacement by protamines. These nucleosomes even contain specific variants such as H2A.X, H3.3 (Gatewood et al. 1990), and H2ABbd (Ishibashi et al. 2010), which have been shown to exhibit an unusual instability in different settings (Eirin-Lopez et al. 2008; Heo et al. 2008; Jin and Felsenfeld 2007). In addition, the chromatin region encompassed does not affect a random region of the genome (Arpanahi et al. 2009) but appears to involve a set of defined groups of genes that presumably play a critical role in gene expression after fertilization (Hammoud et al. 2009). The disorderly accumulation of an excessive amount of histones in the sperm nuclei has been linked to infertility, which will be described next.

9.5

Mammalian SNBPs and Infertility. The Intriguing Role of TPs and Protamines in DNA Integrity

The most commonly cited cause of infertility in couples is defective sperm, accounting for 40% of clinically reported infertility cases (Larson-Cook et al. 2003). In the United States alone, it has been reported that 2 3 million couples seek the aid of assisted reproductive technologies (ARTs) (Begley 1995; Mosher and Pratt 1990) to achieve fertilization. Intra-cytoplasmic sperm injection (ICSI), introduced in 1992 and used today in 50% of in vitro fertilizations (IVFs), transformed the treatment of male infertility factors such as poor sperm motility, low sperm concentration, and abnormal shape (Larson-Cook et al. 2003). However, in a large percentage of cases, where significant sperm DNA fragmentation was present, ICSI did not work (Larson-Cook et al. 2003). It is now known that sperm DNA fragmentation affects pregnancies that are initiated both naturally, and via ART, resulting in abnormal embryo development and failed pregnancies (Larson-Cook et al. 2003; Tomlinson et al. 2001). SNBPs appear to play a role in the sperm DNA integrity. There are several ways in which SNBPs can lead to infertility: altered histone: protamine ratio, altered TP expression, altered P1/P2 ratio, and mutations of P1 and P2 (Ausio et al. 2007; Oliva 2006a). Numerous studies have hypothesized that the DNA-strand breaks in mature sperm are due to the abnormal replacement of histones or TPs by protamines (Becker et al. 2008; Sakkas et al. 2002, 2003;

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Spano et al. 2000). Recent studies have added further evidence to support this hypothesis: not only is there an increased number of DNA-strand breaks in the sperm of infertile men but also a greater number of retained histones and a reduction in the amount of protamine bound to genomic DNA (Zhang et al. 2006; Zini et al. 2007, 2008). Reduced sperm protamine content has also been linked to abnormal sperm head morphology, an increase in the number of DNA-strand breaks (Aoki et al. 2005; Gazquez et al. 2008; Torregrosa et al. 2006) and male infertility (Aoki et al. 2005, 2006; Oliva 2006b). All of this underscores the functional importance of the proper SNBP transitions. Knockout experiments (Meistrich et al. 2003; Zhao et al. 2001, 2004a, b) carried out with mice that were either null in TP1 or TP2 indicated that histones were displaced during spermiogenesis and some sperm were fertile. However, chromatin condensation and the processing of the protamine P2 to its cleaved form were incomplete and the genomic DNA exhibited an increased incidence of DNA-strand breaks. These measurements suggested that TP1 and TP2 compensate for each other when one is absent and that they may have functionally redundant roles (Meistrich et al. 2003; Zhao et al. 2001). A second set of experiments used mice that were null in both TP1 and TP2 (Zhao et al. 2004a, b). These measurements showed that while histones were still displaced, chromatin condensation was poor, DNA-strand breaks were elevated, and over 80% of the epididymal sperm were dead. These measurements showed that both TPs are required for normal chromatin condensation, for the protection of genome integrity, and for fertile sperm. In contrast to the effect of TPs, haploinsufficiency of either P1 or P2 protamines caused by knockouts in one of the alleles for either one of the genes encoding these proteins caused sterility in mice (Cho et al. 2001, 2003). However, similarly to TPs, depletion of P2 was found to result in sperm DNA damage and reduced chromatin compaction. Whereas it is not difficult to envisage how the highly compacted chromatin organization resulting from the interaction of DNA with SNBPs protect against DNA damage during the sperm transit in the female tract or in an external environment, the role of TPs and protamines in maintaining DNA integrity during spermiogenesis is still poorly understood and remains intriguing.

9.6

Concluding Remarks

Some important progress has been made in recent years in our understanding of the organization of the male genome by SNBPs and in the evolution of these proteins (Eirin-Lopez and Ausio 2009). However, in comparison to other areas of research, progress has been slow and several important questions remain unanswered. For instance, what prevents the sperm histones that remain associated with sperm chromatin in some organisms, such as mice and humans, from being replaced? If the remaining histones in these organisms are so important for development (Hammoud et al. 2009) how do other mammals manage without them? If this bestows these organisms with a reproductive edge, what kind of an edge? Similarly,

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if the P1/P2 ratio is so important for fertility in certain eutherian mammals, how can those organisms that only express P1 be as reproductively successful? What edge do TPs confer to eutherian mammals that the rest of mammals can dispense with? In cases where infertility arises from a higher histone to protamine ratio, do the regions covered by the additional histones encompass random portions of the genome? Obviously some of the answers to these questions will be related to the increase in organism complexity, yet a detailed mechanistic explanation is still missing. Acknowledgments This work was supported by a Natural Sciences and Engineering Research Council of Canada (NSERC) OGP 46399 07 Grant (to J.A.) and Washington State University (start up funds to L.R.B.).

References Aoki VW, Moskovtsev SI, Willis J, Liu L, Mullen JB, Carrell DT (2005) DNA integrity is compromised in protamine deficient human sperm. J Androl 26:741 748 Aoki VW, Emery BR, Liu L, Carrell DT (2006) Protamine levels vary between individual sperm cells of infertile human males and correlate with viability and DNA integrity. J Androl 27: 890 898 Arents G, Moudrianakis EN (1995) The histone fold: a ubiquitous architectural motif utilized in DNA compaction and protein dimerization. Proc Natl Acad Sci USA 92:11170 11174 Arpanahi A, Brinkworth M, Iles D, Krawetz SA, Paradowska A, Platts AE, Saida M, Steger K, Tedder P, Miller D (2009) Endonuclease sensitive regions of human spermatozoal chromatin are highly enriched in promoter and CTCF binding sequences. Genome Res 19:1338 1349 Ausio´ J (1992) Presence of a highly specific histone H1 like protein in the chromatin of the sperm of the bivalve mollusks. Mol Cell Biochem 115:163 172 Ausio´ J (1995) Histone H1 and the evolution of the nuclear sperm specific proteins. Advances in Spermatozoal Phylogeny and Taxonomy. In: Jamieson BGM, Ausio´ J, Justine JL (eds) Memoires de Museum National d’Histoire Naturelle, Paris, Vol 166, pp 501 514 Ausio´ J (1999) Histone H1 and evolution of sperm nuclear basic proteins. J Biol Chem 274: 31115 31118 Ausio J, Eirin Lopez JM, Frehlick LJ (2007) Evolution of vertebrate chromosomal sperm proteins: implications for fertility and sperm competition. Soc Reprod Fertil Suppl 65:63 79 Balhorn R (2007) The protamine family of sperm nuclear proteins. Genome Biol 8:227 Balhorn R, Cosman M, Thornton K, Krishnan V.V, Corzett M, Bench G, Kramer C, Lee IV J, Hud NV, Allen M, Prieto M, Meyer Ilse W, Brown TJ, Kirz J, Zhang X, Bradbury EM, Maki G, Braun RE, Breed W (1999) Protamine mediated condensation of DNA in mammalian sperm. The Male Gamete: from Basic to Clinical Applications. In: Gagnon C (ed) Cache River Press, Vienna, IL, pp 58 70 Baneres JL, Martin A, Parello J (1997) The N tails of histones H3 and H4 adopt a highly structured conformation in the nucleosome. J Mol Biol 273:503 508 Barreau C, Benson E, Gudmannsdottir E, Newton F, White Cooper H (2008) Post meiotic transcription in Drosophila testes. Development 135:1897 1902 Baskaran R, Rao MR (1990) Interaction of spermatid specific protein TP2 with nucleic acids, in vitro. A comparative study with TP1. J Biol Chem 265:21039 21047 Baskaran R, Rao MR (1991) Mammalian spermatid specific protein, TP2, is a zinc metalloprotein with two finger motifs. Biochem Biophys Res Commun 179:1491 1499

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Becker S, Soffer Y, Lewin LM, Yogev L, Shochat L, Golan R (2008) Spermiogenesis defects in human: detection of transition proteins in semen from some infertile men. Andrologia 40: 203 208 Begley S (1995) The baby myth. Newsweek 126(10):38 47 Brewer LR, Corzett M, Balhorn R (1999) Protamine induced condensation and decondensation of the same DNA molecule. Science 286:120 123 Brewer L, Corzett M, Balhorn R (2002) Condensation of DNA by spermatid basic nuclear proteins. J Biol Chem 277:38895 38900 Caron N, Veilleux S, Boissonneault G (2001) Stimulation of DNA repair by the spermatidal TP1 protein. Mol Reprod Dev 58:437 443 Casas MT, Munoz Guerra S, Subirana JA (1981) Preliminary report on the ultrastructure of chromatin in the histone containing spermatozoa of a teleost fish. Biol Cell 40:87 92 Chalmel F, Rolland AD, Niederhauser Wiederkehr C, Chung SS, Demougin P, Gattiker A, Moore J, Patard JJ, Wolgemuth DJ, Jegou B, Primig M (2007) The conserved transcriptome in human and rodent male gametogenesis. Proc Natl Acad Sci USA 104:8346 8351 Cho C, Willis WD, Goulding EH, Jung Ha H, Choi YC, Hecht NB, Eddy EM (2001) Haploinsuf ficiency of protamine 1 or 2 causes infertility in mice. Nat Genet 28:82 86 Cho C, Jung Ha H, Willis WD, Goulding EH, Stein P, Xu Z, Schultz RM, Hecht NB, Eddy EM (2003) Protamine 2 deficiency leads to sperm DNA damage and embryo death in mice. Biol Reprod 69:211 217 Dadoune JP (2003) Expression of mammalian spermatozoal nucleoproteins. Microsc Res Tech 61:56 75 Debarle M, Martinage A, Sautiere P, Chevaillier P (1995) Persistence of protamine precursors in mature sperm nuclei of the mouse. Mol Reprod Dev 40:84 90 Dou Y, Mizzen CA, Abrams M, Allis CD, Gorovsky MA (1999) Phosphorylation of linker histone H1 regulates gene expression in vivo by mimicking H1 removal. Mol Cell 4:641 647 Dou Y, Bowen J, Liu Y, Gorovsky MA (2002) Phosphorylation and an ATP dependent process increase the dynamic exchange of H1 in chromatin. J Cell Biol 158:1161 1170 Eirin Lopez JM, Ausio J (2009) Origin and evolution of chromosomal sperm proteins. Bioessays 31:1062 1070 Eirin Lopez JM, Ishibashi T, Ausio J (2008) H2A.Bbd: a quickly evolving hypervariable mam malian histone that destabilizes nucleosomes in an acetylation independent way. FASEB J 22:316 326 Eirı´n Lo´pez JM, Frehlick LJ, Ausio´ J (2006a) Protamines, in the footsteps of linker histone evolution. J Biol Chem 281:1 4 Eirı´n Lo´pez JM, Lewis JD, Howe LA, Ausio J (2006b) Common phylogenetic origin of protamine like (PL) proteins and histone H1: evidence from bivalve PL genes. Mol Biol Evol 23:1304 1317 Evans DL, Kaur H, Leary J 3rd, Praveen K, Jaso Friedmann L (2005) Molecular characterization of a novel pattern recognition protein from nonspecific cytotoxic cells: sequence analysis, phylogenetic comparisons and anti microbial activity of a recombinant homologue. Dev Comp Immunol 29:1049 1064 Foster HA, Abeydeera LR, Griffin DK, Bridger JM (2005) Non random chromosome positioning in mammalian sperm nuclei, with migration of the sex chromosomes during late spermatogen esis. J Cell Sci 118:1811 1820 Garcia Ramirez M, Rocchini C, Ausio´ J (1995) Modulation of chromatin folding by histone acetylation. J Biol Chem 270:17923 17928 Gatewood JM, Cook GR, Balhorn R, Schmid CW, Bradbury EM (1990) Isolation of four core histones from human sperm chromatin representing a minor subset of somatic histones. J Biol Chem 265:20662 20666 Gaucher J, Reynoird N, Montellier E, Boussouar F, Rousseaux S, Khochbin S (2010) From meiosis to postmeiotic events: the secrets of histone disappearance. FEBS J 277(3):599 604

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Marcon L, Boissonneault G (2004) Transient DNA strand breaks during mouse and human spermiogenesis new insights in stage specificity and link to chromatin remodeling. Biol Reprod 70:910 918 Martens G, Humphrey EC, Harrison LG, Silva Moreno B, Ausio J, Kasinsky HE (2009) High pressure freezing of spermiogenic nuclei supports a dynamic chromatin model for the histone to protamine transition. J Cell Biochem 108:1399 1409 Martinez Soler F, Kurtz K, Ausio J, Chiva M (2007a) Transition of nuclear proteins and chromatin structure in spermiogenesis of Sepia officinalis. Mol Reprod Dev 74:360 370 Martinez Soler F, Kurtz K, Chiva M (2007b) Sperm nucleomorphogenesis in the cephalopod Sepia officinalis. Tissue Cell 39:99 108 McMurray CT, Kortun IV (2003) Repair in haploid male germ cells occurs late in differentiation as chromatin is condensing. Chromosoma 111:505 508 Meetei AR, Ullas KS, Vasupradha V, Rao MR (2002) Involvement of protein kinase A in the phosphorylation of spermatidal protein TP2 and its effect on DNA condensation. Biochemistry 41:185 195 Meistrich ML, Mohapatra B, Shirley CR, Zhao M (2003) Roles of transition nuclear proteins in spermiogenesis. Chromosoma 111:483 488 Miescher F (1874) Das Protamin eine neue organische Base aus den Samenf€aden des Rhein lachses. Ber Deut Chem Ges 7:376 379 Moriniere J, Rousseaux S, Steuerwald U, Soler Lopez M, Curtet S, Vitte AL, Govin J, Gaucher J, Sadoul K, Hart DJ, Krijgsveld J, Khochbin S, Muller CW, Petosa C (2009) Cooperative binding of two acetylation marks on a histone tail by a single bromodomain. Nature 461: 664 668 Mosher WD, Pratt WF (1990) Fecundity and infertility in the United States, 1965 1988. Advance data from vital and health statistics, No. 192. National Center for Health Statistics, Hyattsville, MD Oliva R (2006) Protamines and male infertility. Hum Reprod Update 12(4):417 435 Oliva R, Dixon GH (1991) Vertebrate protamine genes and the histone to protamine replacement reaction. Prog Nucleic Acid Res Mol Biol 40:25 94 Oliva R, Mezquita C (1986) Marked differences in the ability of distinct protamines to disassemble nucleosomal core particles in vitro. Biochemistry 25:6508 6511 Oliva R, Bazett Jones D, Mezquita C, Dixon GH (1987) Factors affecting nucleosome disassem bly by protamines in vitro. Histone hyperacetylation and chromatin structure, time dependence, and the size of the sperm nuclear proteins. J Biol Chem 262:17016 17025 Oliva R, Bazett Jones DP, Locklear L, Dixon GH (1990) Histone hyperacetylation can induce unfolding of the nucleosome core particle. Nucleic Acids Res 18:2739 2747 Pivot Pajot C, Caron C, Govin J, Vion A, Rousseaux S, Khochbin S (2003) Acetylation dependent chromatin reorganization by BRDT, a testis specific bromodomain containing protein. Mol Cell Biol 23:5354 5365 Pradeepa MM, Rao MR (2007) Chromatin remodeling during mammalian spermatogenesis: role of testis specific histone variants and transition proteins. Soc Reprod Fertil Suppl 63:1 10 Ribes E, Sanchez De Romero LD, Kasinsky HE, del Valle L, Gimenez Bonafe P, Chiva M (2001) Chromatin reorganization during spermiogenesis of the mollusc Thais hemostoma (Murici dae): implications for sperm nuclear morphogenesis in cenogastropods. J Exp Zool 289: 304 316 Risley MS, Einheber S, Bumcrot DA (1986) Changes in DNA topology during spermatogenesis. Chromosoma 94:217 227 Roque A, Iloro I, Ponte I, Arrondo JL, Suau P (2005) DNA induced secondary structure of the carboxyl terminal domain of histone H1. J Biol Chem 280:32141 32147 Rose KL, Li A, Zalenskaya I, Zhang Y, Unni E, Hodgson KC, Yu Y, Shabanowitz J, Meistrich ML, Hunt DF, Ausio J (2008) C terminal phosphorylation of murine testis specific histone H1t in elongating spermatids. J Proteome Res 7:4070 4078

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Sakkas D, Moffatt O, Manicardi GC, Mariethoz E, Tarozzi N, Bizzaro D (2002) Nature of DNA damage in ejaculated human spermatozoa and the possible involvement of apoptosis. Biol Reprod 66:1061 1067 Sakkas D, Seli E, Bizzaro D, Tarozzi N, Manicardi GC (2003) Abnormal spermatozoa in the ejaculate: abortive apoptosis and faulty nuclear remodelling during spermatogenesis. Reprod Biomed Online 7:428 432 Saperas N, Ribes E, Buesa C, Garcia Hegart F, Chiva M (1993) Differences in chromatin condensation during spermiogenesis in two species of fish with distinct protamines. J Exp Zool 265:185 194 Saperas N, Chiva M, Casas MT, Campos JL, Eirin Lopez JM, Frehlick LJ, Prieto C, Subirana JA, Ausio J (2006) A unique vertebrate histone H1 related protamine like protein results in an unusual sperm chromatin organization. FEBS J 273:4548 4561 Sarg B, Chwatal S, Talasz H, Lindner HH (2009) Testis specific linker histone H1t is multiply phosphorylated during spermatogenesis. Identification of phosphorylation sites. J Biol Chem 284:3610 3618 Schulz RW, de Franca LR, Lareyre JJ, Legac F, Chiarini Garcia H, Nobrega RH, Miura T (2010) Spermatogenesis in fish. Gen Comp Endocrinol 165(3):390 411 Shang E, Nickerson HD, Wen D, Wang X, Wolgemuth DJ (2007) The first bromodomain of Brdt, a testis specific member of the BET sub family of double bromodomain containing proteins, is essential for male germ cell differentiation. Development 134:3507 3515 Spano M, Bonde JP, Hjollund HI, Kolstad HA, Cordelli E, Leter G (2000) Sperm chromatin damage impairs human fertility. The Danish First Pregnancy Planner Study Team. Fertil Steril 73:43 50 Steger K (2001) Haploid spermatids exhibit translationally repressed mRNAs. Anat Embryol (Berl) 203:323 334 Subirana JA (1992) Order and disorder in 30 nm chromatin fibers. FEBS Lett 302:105 107 Subirana JA, Cozcolluel C, Palau J, Unzeta M (1973) Protamines and other basic proteins from spermatozoa of molluscs. Biochim Biophys Acta 317:364 379 Tomlinson MJ, Moffatt O, Manicardi GC, Bizzaro D, Afnan M, Sakkas D (2001) Interrela tionships between seminal parameters and sperm nuclear DNA damage before and after density gradient centrifugation: implications for assisted conception. Hum Reprod 16: 2160 2165 Torregrosa N, Dominguez Fandos D, Camejo MI, Shirley CR, Meistrich ML, Ballesca JL, Oliva R (2006) Protamine 2 precursors, protamine 1/protamine 2 ratio, DNA integrity and other sperm parameters in infertile patients. Hum Reprod 21:2084 2089 Tsend Ayush E, Dodge N, Mohr J, Casey A, Himmelbauer H, Kremitzki CL, Schatzkamer K, Graves T, Warren WC, Grutzner F (2009) Higher order genome organization in platypus and chicken sperm and repositioning of sex chromosomes during mammalian evolution. Chromo soma 118:53 69 Vibranovski MD, Lopes HF, Karr TL, Long M (2009) Stage specific expression profiling of Drosophila spermatogenesis suggests that meiotic sex chromosome inactivation drives geno mic relocation of testis expressed genes. PLoS Genet 5:e1000731 Vila R, Ponte I, Collado M, Arrondo JL, Jimenez MA, Rico M, Suau P (2001) DNA induced alpha helical structure in the NH2 terminal domain of histone H1. J Biol Chem 276: 46429 46435 Vilfan ID, Conwell CC, Hud NV (2004) Formation of native like mammalian sperm cell chroma tin with folded bull protamine. J Biol Chem 279:20088 20095 Ward WS, Coffey DS (1991) DNA packaging and organization in mammalian spermatozoa: comparison with somatic cells. Biol Reprod 44:569 574 Ward WS, Zalensky AO (1996) The unique, complex organization of the transcriptionally silent sperm chromatin. Crit Rev Eukaryot Gene Expr 6:139 147 Ward WS, Partin AW, Coffey DS (1989) DNA loop domains in mammalian spermatozoa. Chromosoma 98:153 159

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Wiesel S, Schultz GA (1981) Factors which may affect removal of protamine from sperm DNA during fertilization in the rabbit. Gamete Res 4:25 34 Worawittayawong P, Leigh C, Weerachatyanukul W, Manochantr S, Sobhon P, Breed WG, Sretarugsa P (2008) Changes in distribution of basic nuclear proteins and chromatin organiza tion during spermiogenesis in the greater bandicoot rat, Bandicota indica. Cell Tissue Res 334:135 144 Wouters Tyrou D, Martinage A, Chevaillier P, Sautiere P (1998) Nuclear basic proteins in spermiogenesis. Biochimie 80:117 128 Zalensky A, Zalenskaya I (2007) Organization of chromosomes in spermatozoa: an additional layer of epigenetic information? Biochem Soc Trans 35:609 611 Zhang X, San Gabriel M, Zini A (2006) Sperm nuclear histone to protamine ratio in fertile and infertile men: evidence of heterogeneous subpopulations of spermatozoa in the ejaculate. J Androl 27:414 420 Zhao M, Shirley CR, Yu YE, Mohapatra B, Zhang Y, Unni E, Deng JM, Arango NA, Terry NH, Weil MM, Russell LD, Behringer RR, Meistrich ML (2001) Targeted disruption of the transition protein 2 gene affects sperm chromatin structure and reduces fertility in mice. Mol Cell Biol 21:7243 7255 Zhao M, Shirley CR, Hayashi S, Marcon L, Mohapatra B, Suganuma R, Behringer RR, Boisson neault G, Yanagimachi R, Meistrich ML (2004a) Transition nuclear proteins are required for normal chromatin condensation and functional sperm development. Genesis 38:200 213 Zhao M, Shirley CR, Mounsey S, Meistrich ML (2004b) Nucleoprotein transitions during sper miogenesis in mice with transition nuclear protein Tnp1 and Tnp2 mutations. Biol Reprod 71:1016 1025 Zini A, Gabriel MS, Zhang X (2007) The histone to protamine ratio in human spermatozoa: comparative study of whole and processed semen. Fertil Steril 87:217 219 Zini A, Zhang X, San Gabriel M (2008) Sperm nuclear histone H2B: correlation with sperm DNA denaturation and DNA stainability. Asian J Androl 10:865 871

Chapter 10

Epigenetics in Male Reproduction: A Practical Introduction to the Informatics of Next Generation Sequencing Adrian E. Platts, Claudia Lalancette, and Stephen A. Krawetz

Abstract At fertilization, the male germ cell conveys a richly layered genetic landscape consisting of both DNA and its associated epigenetic information. A systems level understanding of these forms of information could reveal some of the origins of idiopathic male infertility. Characterizing the genetic and epigenetic contributions to fertilization could also offer insight into the root causes of aberrant development. Perhaps some of these elements reflect the fetal origins of adult disease. As a host of new tools and techniques emerge, we have the opportunity to reassess our models of gametogenesis in the male. The challenge is no longer to construct biological models from sparse data but to assimilate a wealth of data being generated by high throughput technologies. By aggregating data from multiple high throughput and targeted experiments, bioinformatics offers potential insight into how genetic and epigenetic information are utilized in the sperm-oocyte system. In this chapter, we will review online resources that can aid in conducting an epigenetic investigation as well as describing approaches to managing second and third generation deep sequencing data. Keywords Bioinformatics Epigenetics Imprinting Male gamete Micro RNA NGS-Next generation sequencing RNA

A.E. Platts and C. Lalancette Center for Molecular Medicine and Genetics, Wayne State University School of Medicine, Detroit, MI, USA Department of Obstetrics and Gynecology, Wayne State University School of Medicine, Detroit, MI, USA S.A. Krawetz (*) Center for Molecular Medicine and Genetics, Wayne State University School of Medicine, Detroit, MI, USA Department of Obstetrics and Gynecology, Wayne State University School of Medicine, Detroit, MI, USA Institute for Scientific Computing, Wayne State University, Detroit, MI, USA e mail: [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 10, # Springer Verlag Berlin Heidelberg 2011

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Abbreviations CDC CpG DGE DoE gDNA HAT HDAC HMT ICR JGI miRNA NGS piRNA rasiRNA sncRNA snoRNA SNP TByte WGS

10.1

Centers for Disease Control and Prevention CG dinucleotides (linked by a phosphor diester bond hence CpG) Differential gene expression Department of Energy Genomic DNA Histone acetyltransferase Histone deacetylase Histone methyltransferase Imprinting control region Joint genome initiative Micro RNA Next generation sequencing, also referred to as second generation sequencing or “now generation” sequencing (Illumina) Piwi-interacting RNA Repeat associated small interacting RNAs Small non coding RNA Small nucleolar RNA Single nucleotide polymorphism Tera Byte, 1012 bytes, 1,000 gigabytes or 1,000,000 megabytes Whole genome sequencing

Introduction

The definition of what comprises epigenetic information continues to evolve. The term was invoked in 1940 by Conrad H. Waddington to describe interactions between the genes of an embryo and its environment (Waddington 1940). This use was refined to refer to heritable nonnucleotide changes in DNA, notably cytosine methylation. Methylation of cytosine bases is the most widely accepted epigenetic mechanism of gene regulation since it is a readily detectable, mitotically heritable property of DNA that can be directly correlated with the expression of proximate genes. While evidence for histone modifications in the nucleosomes of vertebrate testis emerged in 1975 (Honda et al. 1975), it was not until the mid-1990s when the mechanism of X-chromosome inactivation by Xist RNA that epigenetics began to be appreciated as encompassing a wider range of processes. Recent advances have further broadened the term and it is now used to refer to the several layers of information that may be used to regulate cellular expression. The broad definition of epigenetics as layers of heritable regulatory information not coded by the nucleotide sequence allows a useful aggregation of these many elements. Several multinational consortia are actively exploring the different layers of

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Table 10.1 Web resourcesa wRef1 wRef2 wRef3 wRef4 wRef5 wRef6 wRef7 wRef8 wRef9 wRef10 wRef11 wRef12 wRef13 wRef14 wRef15 wRef16 wRef17 wRef18 wRef19 wRef20 wRef21 wRef22 wRef23 wRef24 wRef25 wRef26 wRef27 wRef28 wRef29 wRef30 wRef31 wRef32 wRef33 wRef34 wRef35 wRef36 wRef37 wRef38 wRef39 wRef40 wRef41 wRef42 wRef43 wRef44 wRef45 wRef46 wRef47 wRef48 wRef49 wRef50 wRef51 wRef52

http://www.epigenome.org/ http://nihroadmap.nih.gov/epigenomics/ http://epigenome.eu/ http://www.epigenie.com/ http://www.epigenome noe.net/ http://www.ebi.ac.uk/Tools/emboss/cpgplot/index.html http://www.cpgislands.com/ http://www.cdc.gov/ExposureReport/ http://www.methdb.de/ http://www.har.mrc.ac.uk/research/genomic imprinting/ http://igc.otago.ac.nz/home.html http://histonehits.org http://genome.nhgri.nih.gov/histones/ http://bioinfo.hrbmu.edu.cn/hhmd http://www.bioinformatics2.wsu.edu/cgi bin/ChromatinDB/cgi/visualize select.pl http://www.jgi.doe.gov/ http://www.illumina.com solid.appliedbiosystems.com http://www.polonator.org/ http://www.helicosbio.com/ http://www.pacificbiosciences.com http://www.iontorrents.com http://www.ibm.com http://www.illumina.com/downloads/DeNovoAssembly TechNote.pdf http://bozeman.mbt.washington.edu/consed/consed.html http://soap.genomics.org.cn http://www.ebi.ac.uk/~zerbino/velvet/ http://www.bcgsc.ca/platform/bioinfo/software/abyss http://www.dnastar.com/ http://www.clcbio.com http://sph.umd.edu/epib/faculty/mltlee/web front r.html http://bowtie bio.sourceforge.net/index.shtml http://maq.sourceforge.net/bwa man.shtml http://maq.sourceforge.net/ http://www.novocraft.com/ http://rulai.cshl.edu/rmap/ http://apps.sourceforge.net/mediawiki/cloudburst bio/index.php?title CloudBurst http://compbio.cs.toronto.edu/shrimp/ http://www.genomatix.de http://www.sanger.ac.uk/Users/lh3/NGSalign.shtml http://www.sph.umich.edu/csg/qin/HPeak/index.html http://www.1000genomes.org http://www.mirbase.org/ http://pirnabank.ibab.ac.in/ http://genome.ucsc.edu/cgi bin/hgTables http://mfold.bioinfo.rpi.edu/ http://bioinformatics.psb.ugent.be/software/details/Randfold http://web.bioinformatics.cicbiogune.es/microRNA/miRanalyser.php http://www.bioinformatics.org/mirfinder/ http://cbcsrv.watson.ibm.com/rna22.html http://www.noncode.org/ http://genome.ucsc.edu/ (continued)

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Table 10.1 (continued) wRef53 http://galaxy.psu.edu/ wRef54 http://www.clcbio.com/index.php?id 28 wRef55 http://genome.ucsc.edu/cgi bin/hgLiftOver wRef56 http://nar.oxfordjournals.org wRef57 http://www.pictar.org/ wRef58 http://www.microRNA.org wRef59 http://research.imb.uq.edu.au/rnadb/ wRef60 http://www.illumina.com/technology/mate pair sequencing assay.ilmn a This chapter contains many references to online resources all of which were openly available and accessible at the time of writing. However, internet resources are dynamic by their nature and are rapidly subsumed by later resources. If a resource cannot be found at the time of reading, internet archives such as http://www.archive.org queried with October 2009 will frequently provide a snapshot of the resource as it was available at the time of writing. Any mention of any commercial product is not an endorsement

epigenetic information. These include the international human epigenome project (wRef1), the NIH Roadmap Epigenome project (wRef2), and the European Union’s Epigenomic network (wRef3). These and other website references (wrefs) are listed in Table 10.1.

10.1.1

Heritability, Disease, and Model Systems

Epigenetically transmitted information may provide a mechanism for one generation to communicate environmental information to the next, for example, nutrition status. As a result, it is also susceptible to assaults from the environment, particularly at vulnerable developmental periods such as embryogenesis and early childhood (Barker 1997, 2004). The epigenetic state is more malleable than the genetic code and can be modified in response to environmental factors (Jirtle and Skinner 2007). This was recently shown in the rat model where vinclozolin, an endocrine disruptor that impacts male reproduction, induced transgenerational effects without any evidence of changes to the nucleotide sequence (Guerrero-Bosagna and Skinner 2009). Clearly, the transgenerational patterns of inheritance are clinically significant. Mendelian principles are of limited utility without in depth consideration of the mechanisms by which each form of epigenetic information is communicated (Nadeau 2009). Generalizations across species can also be problematic since different organisms appear to utilize epigenetic pathways in very different ways. For example, while honey bee and fruit fly are both insecta, bees are able to methylate DNA while fruit flies are essentially devoid of DNA methylation (Wang et al. 2006). Even in mammals such as mouse and man, the extent of chromatin remodeling brought about during spermiogenesis through the replacement of histones by protamines may be very different, but potentially impacts the prepotentiation of early developmental genes (Hammoud et al. 2009). Hence,

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bioinformatic approaches based on conservation between species must be applied with caution. An understanding of the basic processes of epigenetic heritability and marking should underlie the analytical strategy and these are reviewed in conjunction with relevant database projects and online resources.

10.2

Approaches and Online Resources for Epigenetic Investigation

Human forms of epigenetic information can be divided into at least four broad layers: methylation of DNA at CpG islands and surrounding CpG sites, changes to the constituent proteins and the 10 nm chromatin fiber, conjoner modification at specific histone sites, and the action of noncoding-RNAs during and after transcription. As recently demonstrated, each layer can be examined in isolation or in a broader more integrated systems level approach (Lister et al. 2009; Lu et al. 2009). A substantial set of online data resources are available to aid in both forms of investigation. For example, both Epigenie (wRef4) and the Epigenome Network of Excellence (wRef5) provide comprehensive protocol archives.

10.2.1 DNA Methylation and Imprinting Cytosine methylation occurs in many but not all higher eukaryotes. Typically in conjunction with histone modifications, a segment of the genome will undergo a series of conformational changes rendering it less accessible to transcription initiation (Bird 2002). This provides one mechanism for reducing gene expression or generating facultatively accessible heterochromatin. It can also be directed to silence genes during differentiation or as a defensive mechanism to suppress transcription from retrotransposons. Methylation of the human genome is primarily determined through the action of de novo methyltransferases DNMT3a and DNMT3b. The pattern is subsequently maintained over mitotic divisions by the maintenance methyltransferases, e.g., DNMT1. Candidate CpG rich island sites susceptible to methylation can be bioinformatically identified using what are now standard base profile windowing strategies. Tools include both generic bioinformatic suites such as Emboss (wRef6) (Rice et al. 2000) as well as specific tools such as the CpG Island Searcher (wRef7) (Takai and Jones 2003). Male sexual maturation has been suggested to be impacted by high levels of exposure to certain endocrine disruptive agents such as Phthalates (DBP) and Bisphenols that may interfere with DNA methylation (Mylchreest et al. 1999; Kawai et al. 2003; Kang and Lee 2005). Data on environmental levels and population exposure to potentially disruptive agents in the United States are maintained by the CDC (wRef8). The DNA methylation database (wRef9) (Amoreira et al. 2003)

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archives many resources, protocols, and tools useful for characterization. In addition, this database also provides an archive of methylation sequence data from numerous methylation state studies of the human epivariome. While DNA methylation likely plays a role in orchestrating the intricate processes of male sexual maturation, the spermatozoal genome exists in a somewhat hypomethylated state (Martorell et al. 1997). Interestingly, global patterns of methylation may not be fully reestablished until early embryogenesis. Genomic imprinting is typically reflected by parent of origin allele-specific methylation that yields monoallelic expression (Bartolomei 2009). To date, approximately 100 imprinted genes have been characterized in rodents and humans, with several other candidates awaiting confirmation. One of the best characterized systems is the IGF2/H19 locus (Edwards and Ferguson-Smith 2007) where differential methylation is mediated through the ICR, imprinting control region. A series of web-based databases including those at the Medical Research Council Harwell Research Center (wRef10), and the parent of origin effects database at the University of Otago (wRef11) (Morison et al. 2001) maintain lists of human and mouse imprinted genes.

10.2.2 Histone Substitution and Higher Order Chromatin Structure The testis contains a rich suite of histone variants, many of which are testisspecific (Grimes et al. 1997; van Roijen et al. 1998). During spermatogenesis, the somatic histones are replaced by a steadily varying set of stage specific isoforms. Several reviews and databases of histone variants (Lindner et al. 2003) and their associated properties are available (wRef12) (Morison et al. 2001). Other proteins such as the Polycomb group proteins may also associate with DNA to change the chromatin structure, acting as a transcriptional repressor through chromatin condensation or as a potentiator through decondensation (reviewed in Martins and Krawetz 2005).

10.2.3 Histone Modification Compared to the H1 linker, the core histones H2A, H2B, H3, and H4 are largely inaccessible to modification (Mersfelder and Parthun 2006), with the exception of the site-specific modification directed to their N-terminal tails. Several databases that describe the distribution of these modifications are available (Sullivan et al. 2002; Zhang et al. 2010) and continue to grow as new modifications are reported (wRef13, wRef14, wRef15). The number of residues that can be modified

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combined with the number of possible modifications makes the combinatorics of the histone code substantial. Modifications include lysine acetylation catalyzed by Histone Acetyltransferases, HATs, their removal catalyzed by Histone Deacetylases, HDACs, and methylation to varying degrees (mono, di or tri methylation) achieved by HMTs. Serine residues may be phosphorylated by the RSK-2 kinase and components of the ubiquitin pathway ubiquitinylate or through a like pathway sumoylate various functional groups. Each modification can, but does not always, preclude another modification to the same residue. Unlike DNA methylation, histone methylation may mark both activation and suppression depending upon which lysine is targeted and the degree to which it is modified. Hence mono, di and tri methylation typically convey different messages. The mechanism(s) by which these patterns are inherited is being actively pursued. In humans, some are paternally transmitted at fertilization as the paternally bound histone regions are incorporated into the embryonic genome (Hammoud et al. 2009). In addition to the database resources (wRef15), many excellent reviews that catalog the increasing number of histone modifications, their functions (Marmorstein 2001), and activities are available (Schones and Zhao 2008).

10.2.4 Transcriptional Modulation by Small RNAs Like histone modifications and DNA methylation, small noncoding RNAs (sncRNAs) including “epi-miRNAs” (Valeri et al. 2009) may also regulate gene expression. Recent studies suggest that feedback may take place between the small RNA and DNA methylation processes when both epigenetic systems interact (Han et al. 2007; Sinkkonen et al. 2008; Grandjean et al. 2009). The three main classes of sncRNAs, i.e., micro-RNAs (miRNAs), small interacting RNAs (siRNAs), and piwi-interacting RNAs (piRNAs) (reviewed in Moazed 2009), are primarily associated with silencing pathways. Both siRNAs and miRNAs are processed from longer precursor RNAs yielding small RNAs between 19 and 24 nucleotides in length. They both function through the dicer-dependent RNA silencing pathway that is essential for mouse spermatogenesis. Dysregulation of methylation in dicer-1 deficient mouse embryonic stem cells (Benetti et al. 2008) supports the view that the interaction between the sncRNA pathways and centromeric and telomeric DNA methylation are perhaps linked. In comparison, piRNAs are a subset of those that, like Xist, interact directly with the DNA. As orthologs of the Drosophila repeat associated small interacting rasi-RNAs, they appear to protect the genome against retrotransposition. Several other classes of sncRNAs, e.g., the promoter associated small RNAs (Taft et al. 2009), likely function in male reproduction. The possible interactions between small RNAs and DNMTs as part of the mechanism of RNA-mediated paramutation in mouse is beginning to be addressed (Grandjean et al. 2009). The experimentally identified set of small RNAs are increasing along with the number of small RNA analysis tools and databases. These include MIRBase

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(wRef43) (Griffiths-Jones et al. 2008), MicroRNA.org (wRef58) (Betel et al. 2008), PIRNABank (wRef44) (Sai Lakshmi and Agrawal 2008), the NonCode Database (wRef51) (He et al. 2008), and RNAdb (wRef59) (Pang et al. 2005). Utilities such as PicTar (wRef57) may in turn identify small RNA-binding targets. A wealth of newly developed resources is published in the annual nucleic acids research special editions for databases and web tools (wRef56).

10.3

Deep Sequencing Approaches for Epigenetic Investigation

The current clinical evaluation of male reproductive fitness primarily relies on the characteristics of cellular morphology, proteomic panels, cytogenetic and molecular genetic assays, e.g., those undertaken to detect Y-chromosome micro-deletions. The adoption of highly parallel microarray hybridization platforms by the research community (Ostermeier et al. 2002a, b, 2005) and subsequently a range of deep sequencing technologies (Hammoud et al. 2009; Han et al. 2009) have largely enabled a paradigm shift to sequence-level whole genome analysis. At present, whole genome sequencing (WGS) is at present the de facto standard for a wide range of genetic and epigenetic investigations. This relies heavily on using next generation sequencing technologies (NGS) to address many questions in the reproductive sciences.

10.3.1

Scale of Sequencing Experiments

NGS represents a major leap forward from the earlier Maxam Gilbert and Sanger sequencing processes that were the basis of the first generation of sequencers. They reached their potential as a parallel ensemble of capillaries (384), each capable of resolving up to around 800 nucleotides. Over several hours of sequencing, this typically yielded a total of 300,000 nucleotides. In contrast, a single run of second generation sequencing may yield in excess of 180,000,000,000 bases (180 gigabases) of processed sequence. This is approximately the same nine times the amount of data as the 2002 release of GenBank (Benson et al. 2003) a year after the first complete drafts of the human genome were obtained (Lander et al. 2001; Venter et al. 2001). A single experiment may now generate more sequence data than was generated during the course of the first multinational decade-long human genome sequencing project and more raw data than was previously held on the computer systems of an entire academic institution. Consequently, we are faced with a shift in magnitude of scale of the data that sequencing experiments generate. Frequently large sequencing centers such as the US DoE’s JGI (wRef16) have replaced their first generation sequencers with an equal number of NGS instruments, permitting a vast increase in the number of genomes or cell-types that can be sequenced in parallel. For example, in 2010 MIT’s Broad Institute housed over 100 second generation sequencers. But several implementations of NGS

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technologies have emerged, with each platform differentiated by throughput, flexibility, read length, accuracy, and/or cost.

10.3.2

NGS Technologies Overview

In 2010 there were three primary platforms currently in general use. Their characteristics (Illumina, Roche, and ABI) along with other platforms are summarized in Table 10.2. The Illumina sequencing by synthesis technology (wRef17) is now in its third iteration as the Genome Analyzer (GA/GAIIx/HiSeq) platform. This is currently the most widely adopted short-read (15 150 nt) platform. The Roche 454 bead-based platform was among the first to be commercialized utilizing pyrosequencing that offered long-range sequencing capabilities (300 500 nt). The Applied Biosystems SOLiD platform (wRef18) is also a bead-based platform but uses a dual-base (four color) coding system capable of highly accurate sequencing by redundantly reading each base. The Polonator platform (Shendure et al. 2005) commercialized by Dover Systems is a low-cost very short paired-read (13 nt) sequencer (wRef19). NGS solutions are evolving at a rapid pace with the emergence of a third generation of higher throughput technologies. At present, the third generation Helicos platform (wRef20) sequences short-individual molecules (~35 nt) rather than amplified aggregates. It is anticipated that sequencing capabilities exceeding 2 GBase/h with very extended read lengths will be realized when other third generation instruments are introduced from Biotage (Varshney et al. 2009), Nanogen (Varshney et al. 2009), Visigen (Varshney et al. 2009), and Pacific Biosciences (wRef21). Sequencing systems are expected to continue to evolve as Table 10.2 Overview of sequencing platform properties Platform Yield/run Run time Read length (up to) Illumina GAII/ 12 GBase 4 days 1 105 GAIIx (8 SE lanes) 10 days bases 90 GBase (8 PE 2 250 lanes) bases Roche 454 400 MBase 10 h Median 600 bases Median 30 Helicos 28 GBase (2 flow 8 days bases heliscope cells in parallel) Dover systems 4 GBase (8 lane) 2 13 polonator bases ABI SOLiD 30 GBase 7 days 1 50 (8 SE) 14 days bases 60 GBase 2 50 (8 PE) bases

Raw Technology error rate ~0.5% Four color sequencing by synthesis (SBS) ~0.1% Pyrosequencing <5% ~2.5% ~0.06%

Single color SBS, single molecule sequencing Polony Sequencing Two base/four color coding

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the solid state nanopore DNA-transistor technology proposed by IBM (wRef23) is developed with others like IonTorrent (wRef22) also releasing technologies.

10.3.3

Principles of NGS Sequencing

The current platforms such as ABI’s SOLiD, Illumina’s GAII, and Roche’s 454 all use solid state or clonal amplification reactions to generate large numbers of similar sequences from sets of input material. The spatially constrained clusters of sequences are then sequenced in a massively parallel process. The raw output is typically a set of high resolution photomicrographs of the resulting clusters with one set generated for every sequencing cycle. When the thousands of imaged regions are combined, a typical sequencing run may generate in excess of 500,000 images each of approximately 5 10 Mbytes in size. While the sequencers tend to work in similar ways and generate a similar output, the differences are important to consider. This is highlighted when one compares the processes undertaken to sequence DNA/RNA on two of the most widely adopted platforms Illumina’s GAII and Roche’s 454 FLX discussed below.

10.3.4

Illumina GAII and Roche 454 FLX

The 454 and GAII platforms differ over a range of metrics, most obviously in the length of their reads and in their error characteristics. Where a choice of sequencers is available, these different characteristics should be considered to match the sequencing approach to experimental design. Several distinct methodologies are available for Illumina GAII sequencing. Tag-based counting may be used for small RNA sequencing, ChIP-seq, and oligo dT-primed mRNA expression studies. Libraries are created from short cDNA or DNA fragments and sequenced. To increase processing efficiency, identical ~18 bp tags are aggregated together and their numbers recorded. Only the distinct sequences, rather than all sequences, are then aligned to a genome. This contrasts the sequencing of full length RNAs (sequenced as cDNAs) or DNA that is fragmented to a set size range prior to sequencing. Each sequence is then individually aligned back to a reference genome or used for de novo genome assembly. Sequencing can be carried out in single end, paired end, or mate pair mode where in the former only one end and, in the latter, opposite ends of the same molecule are sequenced. Currently, GAII sequencing can generate read lengths of up to ~150 bases per end. Accordingly, single-ended sequencing is typically used for sequencing templates that are slightly longer than the read; paired end sequencing for sequences up to 800 bases or mate pair sequencing for elements that are separated by as much as 20kbase (GAII) or 40 Bbase (454) (wRef60)). Genomic sequencing may be undertaken for a wide range of genetic and epigenetic investigations. These include genome wide SNP polymorphism analysis,

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Fig. 10.1 Typical quality metric data from an Illumina GAII experiment. Cumulative base error rates following alignment to a reference genome (ELAND) are shown. Data is shown for a single ended run (left) and paired end run (right)

DNA methylation status (following a bisulfite treatment), de novo genome assembly, or for the identification of structural elements (ChIP-seq). The latter is typically carried out to identify the genomic regions associated with specific histone variants or other transfactors (Fullwood and Ruan 2009). A variant of 3C chromatin conformational capture termed HiC has been developed to map the regions of DNA that are in contact with each other yet may originate on distinct chromosomes (Lieberman-Aiden et al. 2009). Characteristic of several NGS platforms, the base error rate on the the GAII sequencer steadily increases as each of the bases is sequenced. The initial parts of reads are generally the most accurate (Fig. 10.1). GAII errors are almost always miscalls a base where base insertion deletion errors are rare. The Roche 454 FLX platforms generate reads that are generally in excess of 350 bases or with conversion to the late 2010 reagents up to a routine length of ~500 bases. Error rates for nucleotide miscalls may be as low as 0.1% and are not as dependent on position as they are on the GAII sequencer. However, ~39% of 454 read errors are often associated with the length of long homopolymer tracks (Huse et al. 2007). This error is rarely observed when using the GAII sequencer. Other differences between the platforms include the amount of template required and the time required to complete each sequencing run. For example, the 454 sequencer typically requires five- to tenfold more starting template, but sequencing is complete within 10 h. In comparison, a GAII sequencer run may take as long as 10 days to complete.

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10.3.5 Applications of Deep Sequencing Unlike other high throughput technologies such as microarrays, each data point (read) generated by a sequencer may originate from virtually any location in the input sequence(s) unless a specific anchoring protocol is used. This is not to suggest that experimental design considerations can be relaxed. While sequencing data may seem more objectively derived, caution needs to be exercised to avoid unintentional bias or subjectivity during the analysis. The data analysis is typically staged and carried out through an analysis pipeline in which a series of discreet steps are taken to align to a reference genome, remove noise, identify features of note, and finally infer biological meaning. Each of these steps requires careful consideration in order to both avoid the common pitfalls of NGS analysis while also matching the most appropriate analysis tools to the biological question(s).

10.3.6 Types of Sequencing Many different sequencing strategies have been developed and applied in a range of protocols across the different NGS platforms. These include but are by no means limited to Single End, Paired End, Mate Pair, mRNA, Small RNA, sequence barcoding, ChIP-Seq, HiC-seq, and metagenomic (Williamson et al. 2005) sequencing. Several of these require specific postprocessing routines within the downstream analysis pipeline. Platform vendors generally provide a customized protocol for each. An overview of several of the approaches that we have utilized for our GAII sequencing projects is outlined below. For Single End (SE) sequencing, a library is first created by fragmenting DNA/ cDNA to a size somewhat longer than the proposed sequence reads, ligating adapters to the ends of the fragments, followed by clustering (solid-state amplification) and asymmetric sequencing. Failure to create libraries of sufficient length will generally require postprocessing to remove the adapter sequence from the reads. The length of the SE library contrasts Paired End (PE) sequencing libraries that are designed to be of a larger size. PE and the related mate pair sequencing permits one to construct scaffolds that span highly repetitive regions that are difficult to align or assemble. The PE library is flanked by asymmetric adapters with sequencing initiating at the 50 end before the sequence is “flipped” and the sequencing is reinitiated from the complimentary strand. RNA Sequencing can use either SE or PE approaches with most platforms starting with fragmented cDNA. Postprocessing for RNA sequencing is generally required to remove residual rRNAs/tRNAs and their pseudogenes as well as to process sequences that contain exon junctions and thus cannot be directly aligned to a genome (typically alignment is made to a prespliced genome representing all known mature RNAs). Caution needs to be exercised when applying PE to RNA-seq since antisense RNAs are of interest, but not all paired end RNA-seq protocols are strand-sensitive.

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Small RNAs form a poorly characterized but potentially rich source of novel epigenetic agents and modifiers. In principle, the creation of sequencing libraries uses the same approach as those used to generate longer RNAs. The primary differences reflect the type of adaptors used and the size selection prior to, or subsequent to, library construction. When sequencing small RNAs, attention should be paid to remove 30 adapter sequences as well as any residual primer sequences that frequently contaminate these libraries. Sequence barcoding has been developed to increase throughput by simultaneously sequencing multiple samples. Before sequencing begins, libraries are ligated with a short sequence that is unique to an originating sample. Sequences from different samples are then combined and sequenced together. Postprocessing is used to disaggregate the sequences by sample using the unique sequence barcode assigned to that library.

10.3.7

Sequencing Depth

One of the first considerations in designing an experiment is the amount of sequencing required to answer a given biological question. For example, how much sequencing would be required to assemble the genome of a species? If de novo assembly is needed, a considerable depth of sequencing will be required to construct a reference set of scaffolds. Other types of sequencing that necessitate considerable depth of coverage include SNP resolution. In this case, high levels of coverage may be needed to resolve with confidence the zygosity of each SNP even in the presence of a reference sequence. These considerations become further involved when barcoding strategies are employed (Church and Kieffer-Higgins 1988; Meyer et al. 2007) since the likely variance in sequencing depth between the different samples sequenced in parallel must also be estimated. When estimating the required depth of coverage to attain preset limits of sensitivity and specificity, factors that include read type, read length, and typical read error characteristics of the platform must be considered. In all cases, sequencing depth may be moderated by the need to sequence repetitive/highly repetitive regions of a genome. As a general rule of thumb, ~20 30 reads per base should be considered for de novo assembly for a de novo assembly with most of these reads derived from paired end libraries and a smaller number derived from longer insert mate pair libraries. This is far greater than the sevenfold coverage that was initially sought in the 1990s in order to ensure that every base was sequenced at least once (Singh and Krawetz 1995) or the read depth sought for alignment against a reference genome, e.g., sequencing the methylome (Lister et al. 2009). Consider a 3 GBase genome sequenced by PE as 2 75 bp nonoverlapping reads. Four flow cells or 32 lanes of GAII sequencing should generate up to ~106 GBases of sequence or 35-fold coverage. This should be sufficient to construct an initial set of contigs. Contigs generated from this type of sequencing (Table 10.3) might be expected to exceed an adjusted median size of (N50) 15 20 kb in length. Assembly error rates tend to rise

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Table 10.3 The size of contig outputs from three popular ab initio genome assembly algorithms Assembler (a) Coverage of (b) Raw base error rate N50 contig Maximum contig genome introduced (%) size (bp) size (bp) Velvet 5 0 6,104 54,331 0.5 3,728 36,873 1 2,084 33,892 10 0 42,856 338,003 0.5 12,957 176,366 1 2,213 91,254 30 0 59,430 465,907 0.5 1 50 0 61,894 750,171 1 a SOAPdenovo 5 0 1,313 20,620 0.5 618 12,250 1 227 5,013 Scaffold Scaffold size Scaffold size 0 34,739 354,068 0.5 16,876 209,162 1 3,364 40,512 10 0 1,620 30,692 0.5 354 9,395 1 120 409 Scaffold Scaffold size Scaffold size 0 125,616 bp 1,259,752 0.5 63,407 831,725 1 5,641 60,718 30 0 1,628 33,594 0.5 44 592 1 40 429 Scaffold Scaffold size Scaffold size 0 130,783 1,336,945 0.5 8,168 63,079 1 5,157 21,372 (32,727) Abyss 5 0 2,381 31,590 1 3,506 9,898 10 0 4,129 73,343 1 333 6,931 a SOAPdenovo can generate both contigs and scaffolds Each was presented with artificially generated PE fragments from a 100 Mbase genome with (a) different levels of genome coverage and (b) different levels of sequence error. Fragment sizes and separations were typical of a Roche 454 sequencing run with a 3 kbase library protocol. Base error generated reflected the type of errors found on the 454 platform. Where no data is presented, the analysis exceeded the available 96 GBytes of RAM on the analysis server.

with contig length and the assembly of many short contigs into scaffolds may be preferable. In some instances, a combination of technologies may be the optimal way to assemble a genome, by either mixing mate pair with paired end sequencing or combining long read data (e.g., 454) with paired end short read data (e.g., Illumina).

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While this depth of sequencing may result in scaffolds of up to several megabases, it is unlikely to be more than 80% complete relative to a typical vertebrate genome. To build a complete de novo genome, that is to characterize most repetitive regions and generate chromosome size assemblies, a sequencing depth typically in excess of 100 reads per base may be anticipated (Diguistini et al. 2009). This would seem to generate much more data than was even required to construct the initial builds of the human genome. However, short read de novo assembly does not have the advantage of even a restriction map from which to confirm assembly. Consequently, while assembling small genomes is now a computationally simple task (wRef24), the assembly of vertebrate and plant genomes remains a computational challenge. For example, a 3 GBase genome sequenced with 90 GBase of short reads generates billions of possible overlaps between reads, the combination of each of which needs to be assessed. Phred/Phrap/Consed (wRef25) (Ewing et al. 1998; Gordon et al. 1998) were some of the tools developed for the human genome sequencing project. The Celera project in turn created a whole genome shotgun (WGS) assembler that is now being developed as a community sourceforge project. Commercial or proprietary sequence assemblers include the Roche GS Assembler or Newbler, SeqMan from DNAStar (wRef29), and CLC Genomics’ assembler (wRef30) amongst others. New classes of fast sequence assemblers such as SOAPdenovo (wRef26) (Li et al. 2008a, b) are becoming available alongside open source solutions such as Velvet (wRef27) (Zerbino and Birney 2008) and Abyss (wRef28) (Simpson et al. 2009). These have established themselves as tools of choice for the assembly of larger genomes. Many employ variants of de Bruijn graph strategies to assemble contigs, where input reads are treated as groups of shorter k-mers (typically 13 56 nts). The association between the k-mers is then used to populate a graph that determines an assembled path between the reads. The underlying characteristics of these sequence assemblers vary with some taking more time and memory to seek their more optimal alignments while others are faster but may produce less accurate contigs (Table 10.3). SOAPdenovo, for example, produces short contigs and long weaker scaffolds, while Velvet and Abyss generate contigs that may be considered as an input for SOAPdenovo scaffolds. Each assembly pipeline benefits from modeling to optimize parameters and minimize aberrant assembly. Assembly error rates and types can be readily estimated by random in silico fragmentation followed by the reassembly of a reference chromosome.

10.3.8

Alignment Tools

The majority of epigenetic sequence data is collected with the expectation that it will be aligned against a reference genome. The choice of alignment algorithm is key to interpretation since algorithms differ in their error tolerance, alignment speeds, and flexibility of input format. It is frequently desirable to align the

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sequences using several algorithms that are well matched to the data source and biological question. This helps to ensure that the maximal amount of data has been retrieved and alignment errors minimized. The genesis of next generation alignment tools has been fueled by the need to align millions of sequences in the most expedient manner so that the time to align is much less than that taken to generate the sequence. This ensures that at least to the data analysis stage the pipeline does not reach a bottleneck. Both commercial and open source solutions are available to process generic (multi-platform) sequencing data. These contrast the platform specific alignment solutions that have been developed with characteristics tailored to the platform’s technology and base error characteristics. For example, the Illumina ELAND (Efficient Large scale Alignment of Nucleotide Databases) algorithm was developed to use the early 16 32 nt GAII sequencer reads as alignment seeds. In conjunction with other components of the Illumina pipeline, it is tailored to the read types and characteristics of the GAII platform. This specification contrasts the generic commercial solutions that frequently include a range of added value end-toend analysis systems designed to work with must current platforms. Examples of these systems include; NovoAlign (wRef35), a comprehensive utility with many useful postprocessing options; CLC bioinformatics’ workbench aligner and the Genomatix mapping station (GMS) (wRef39). The GMS solution is notable since it can align the output from the major sequencing platforms discussed in this chapter and has the options to identify SNPs, splice variation, and gene expression during the alignment process. Several generic open source alignment solutions are now available for NGS. These include: Bowtie (wRef32) (Langmead et al. 2009), an ultra fast alignment tool; BWA (wRef33) (Li and Durbin 2009), a tool similar to Bowtie; MAQ (wRef34) (Li et al. 2008a, b), a tool that uses base call quality when determining optimal alignments; SHRiMP (wRef38) (Rumble et al. 2009), a tool capable of using both FASTA and ABI color space sequences; RMAP (wRef36) (Smith et al. 2009), a fast solution that supports PE and bi-sulfite sequencing data, and CloudBurst (wRef37) (Schatz 2009), a derivative of RMAP that functions in a cloud computing environment. The generic alignment algorithms frequently accept a range of input formats such as FASTA, FASTQ (FASTA files with quality information), and platform-specific sequence formats, e.g., Illumina’s QSEQ and ABI’s color space variant of the FASTA format (CFASTA). The extent to which these algorithms are able to use the per-base quality score data in determining an optimal alignment is an important consideration. Alignment tools can be classified by how they approach the task. Whether they index the genome, index the input sequences, or transform the genome/sequences prior to processing. Each strategy scales differently as a function of genome size and the number of input reads. For example, a simple approach is the use of lookup tables (hash tables) to index all short segments of a reference genome. The input read would then be segmented into similarly sized sequences and all possible locations of the sub sequences in the genome determined. Simple Boolean logic could then suggest the most likely subset of locations in a reference sequence. The

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edit distance or quality metric, i.e., the smallest number of base changes required to transform the input sequence into the sequence in the reference genome, could also be calculated to provide an alignment confidence score. Where an alignment algorithm uses the quality scores from the sequencer, the confidence of the base calls in conjunction with the mismatches in the genome can also be considered. When sequencing large genomes, approaches that index the reads rather than the genome may demonstrate scaling advantages [see Bateman and Quackenbush (2009), Horner et al. (2010) and wRef40]. Conversely, when the number of reads exceeds a threshold, genome indexing solutions may prevail. Alignment rates may vary considerably from less than one million to greater than 50 million sequence reads processed per hour. But the fastest alignment approaches may not always be the best. The process of Burrows Wheeler indexing of the reference and query sequences has allowed some approaches such as BOWTIE (Langmead et al. 2009) and BWA (Li and Durbin 2009) to achieve remarkable efficiencies in both time and system memory space. However, in the case of BWA and BOWTIE, the mathematically optimal alignment may not always be achieved and alignments to multiple regions of the genome cannot be precluded. Thus, while extensions are available [e.g., crossbow (Thomas 2009)], these approaches may not be those best suited to precise applications such as detecting SNP zygosity. In this case, both MAQ and the Genomatix solution may be better suited as they seek optimal alignments in conjunction with base quality.

10.3.9 Identification of Small RNAs While mRNAs are typically aligned readily to either putative or refseq gene models, the task of analyzing small RNA data may be more challenging. Following sequence alignment, the first stage in a small RNA analysis pipeline is frequently the association of reads with families of known small RNAs. Genomic locations obtained from the alignment can be quickly mapped against files corresponding to known classes of small RNAs, e.g., miRNAs and piRNAs. For this task, the genomic locations of most of the non-coding RNA genes can be obtained from the UCSC genome tables in a bed format file (wRef45). Current mappings of miRNAs and piRNAs can be obtained from the Sanger MIR database (wRef43) and the piRNAbank (wRef44), respectively. A database of other small RNAs, including piRNAs, siRNAs and miRNAs, is available from Noncode (wRef51). A few applications are available for de novo small RNA discovery. These currently include MirDeep (Friedlander et al. 2008) that focuses exclusively on microRNAs and Novoalign (wRef35) that provides a useful set of postprocessing options for small RNAs. The identification of miRNA candidates relies on the identification of stable pre-miRNA hairpin-loop structures. This can be accomplished by extending each end of a cluster of small RNA reads by ~100 nucleotides. The stability of the folded structure for each potential hairpin can then be calculated under biological conditions by using, for example, RNAfold (wRef46) (Zuker

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2003). It is noteworthy that when working with small human testis RNAs, the temperature at which stable hairpins should form will be several degrees centigrade lower than the core human body temperature. Automated tools for miRNA discovery in deep sequencing include miRDeep (Friedlander et al. 2008) that uses RandFold (wRef47) (Bonnet et al. 2004) to detect hairpin loops, miRanalyser (wRef48) (Hackenberg et al. 2009), MirFinder (wRef49) (Huang et al. 2007), and RNA22 (wRef50) (Miranda et al. 2006). Other classes of small RNAs like the endogenous siRNAs can also be identified by their ability to form hairpin structures. Phylogenetic comparison is another approach that can be usefully explored for sncRNA discovery between species that share classes of small RNAs.

10.3.10

Sources of Error in Alignment and Interpretation

Some sources of NGS error are not related to any given sequencing technology or alignment solution. Rather they may reflect the maturity of the reference assembly, i.e., the number and the nature of the gaps and other general properties of the target. This has given rise to concepts such as mapability, a continuous measure of how easily reads of a given size may be mapped to any given region of a reference genome. Failure to consider mapability may result in errors of omission leading to both false positive but more often to false negative conclusions. Reads that align to repeat rich or information poor sequence are inherently difficult to map because they may have arisen from many different sites. If these reads are simply rejected, then a bias against certain regions of the genome will have been applied. Several strategies have been developed to mitigate this effect (Rozowsky et al. 2009). In silico genome sequence sampling can be used to create a reference track of mapability (Fig. 10.2). This is typically accomplished by sampling data, e.g., gathering 36 bp sequences every 5 bp through a reference genome then aligning each sequence to the genome. Those regions that have few or no sequences aligned have inherently low mapability. Comparing the observed mapping of sequence reads relative to a “mapability track” can help to distinguish between regions that are poorly represented because they were not sequenced from those to which few reads have been aligned due to inherently low mapability. The opposite, errors of commission, may occur where too many reads are aligned to a genomic location. For example, this occurs when regions of the reference sequence are so repetitive that they have not been characterized. These may generate spurious alignments to shorter similar locations that have been characterized. For example, the centromeres and telomeres of many organisms have not yet been completely assembled in comparison to their pericentromeric satellite repeat rich counterparts. Despite being repeat rich, the pericentromeric regions are typically poorly constrained and hence rich in SNPs appearing somewhat distant from their canonical repeat sequences. Consequently, these regions appear to the alignment algorithm to be relatively unique (Fig. 10.2) even when sequenced with short reads. This frequently results in sequences being readily

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Fig. 10.2 Mapability of human chromosome 6. Mapability is (brown track) shown as a custom track in the UCSC browser. Also shown are SNP density (lower grey track) and read density (upper black track) in a region proximate to the centromere, demonstrating a typical pericentro meric peak

assigned to the small ~10 kb pericentromeric regions even though their true origin may not be known but likely includes the large centromeric regions. When viewed in a genome browser, a set of large peaks frequently appear around specific centromeres as a consequence not of the biology but of misassignment of reads that originate from a sequencing background. Normalization should be considered when comparing sequencing data. This has been reviewed for both small RNA and mRNA studies (Landgraf et al. 2007; Mortazavi et al. 2008; Wilhelm and Landry 2009). The utility of normalization is readily apparent when using RNA-seq density to compare the level of transcripts and their splice-isoforms. When comparing isoforms, it is useful to specifically adjust the read count relative to the length of each exon considered thereby creating a read density per exon. This reflects the anticipation that longer mature RNA regions are more likely to generate reads when the RNA is fragmented for sequencing than are shorter mature RNAs. When comparing similar elements across different sequencing experiments, the total reads from a single region needs to be considered as a function of the total number of reads from each sequencing experiment. This can be calculated and expressed as the number of reads/million reads generated from that run. While length and total read count can be calculated with ease, it should be noted that the distribution is not entirely standardized since the tails will be longer for the deeper sequencing runs.

10.3.11

Peak Detection

In many NGS applications, it is necessary to distinguish with a given level of confidence a peak or cluster of reads from a continuously varying background of

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Fig. 10.3 Epigenetic data viewed in two different genome browsers. Histone associated DNA in spermatozoa sequenced by Illumina GAII (Hammoud et al. 2009) viewed in (bottom) the ENSEMBL browser (top) the UCSC browser

reads (Fig. 10.3). The background can be the result of a deficient fractionation and/ or inappropriate base calls that generate spurious alignments. Several strategies have been developed to identify a cluster of sequence reads, including the differential analysis between a test and control state. If a control state is not available, algorithms such as the Genomatix RegionMiner can be used. Sequence reads are assigned to discrete bins and the distribution of reads within the bins established. A Poisson distribution is then modeled and a background and signal threshold determined for a given p-value. This has the advantage of being a fast and readily applied technique; however, it is not capable of modeling the highly variable background levels that are often observed as a function of sequence mapability. Several alternative approaches are available to develop local models of read distribution and expected peak characteristics and assign a level of confidence within the constraints of a read cluster. These solutions are available in the R/Bioconductor suite, HPeak a markov model-based approach to peak detection (wRef41) as well as the model based analysis of ChIP-seq data (MACS) (Zhang et al. 2008).

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Estimating False Discovery

When NGS sequencing is being used to determine the level of RNA within a tissue or cell it may also be important to select sufficient biological replicates to resolve differences with high confidence. Approaches to estimating false discovery rates may be analogous to those used in microarray experiments to estimate power and sample size (Page et al. 2006). The literature on NGS power analysis is in its infancy [see for example Wang et al. (2009)]. The replicate signal dispersion characteristics for each sequencing platform needs to be determined as they may be very different (wRef31) (Lee and Whitmore 2002). All high throughput platforms have an inherent risk of false positive assignment of significance (failure to reject the null hypothesis H0) due both to the large number of measurements and the large number of questions that may be asked of the data. For example, a question often asked of ChIP-seq data is whether there exists a preferential association between the peaks found in a sequencing study and sets of transfactor binding sites. While this appears as a single question, it actually corresponds to several hundred questions, the exact number being determined by the number of different transfactor binding families that are considered. In these cases, it may be difficult to estimate degrees of freedom and the number of parallel tests to set a significance threshold with a reasonable false discovery rate. Even where these can be readily estimated, the choice of correction that best controls for multiple testing needs to be considered. These range from strong Bonferroni-type corrections (Lee and Whitmore 2002) that will limit false positive error, but necessarily increase false negative error to weaker correction such as the Holm step down adjustment (Osier et al. 2004). Alternatively, the likely false discovery rate can be determined by resampling the data (Ge et al. 2003). In this approach, the false discovery rate is estimated by randomly reassigning the data to different locations in the genome. The association test is then repeated with a selected p-value limit. After repeating this randomization a number of times, an average false discovery rate for a given p-value can be determined.

10.3.13

Data Visualization

Visualizing large amounts of sequencing data in a meaningful manner is challenging. The ability to review the data features at a number of levels is essential. It needs to be flexible to enable genomic features to be included or excluded to render a meaningful display at varying levels of resolution. At the level of the gene or locus, small scale features such as SNPs or small deletions can be presented in the context of the surrounding intron exon structure. When peak detection has been used for ChIP data or small RNA data, a parallel track that shows peak significance, i.e., q-value (Kall et al. 2009), relative to background is useful. As the perspective is expanded to the megabase scale, features such as periodicity and larger deletions can be noted. However, not all rearrangements are visible in high throughput

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sequencing unless paired end sequencing is used to note pair alignment pairs [see breakdancer Chen et al. (2009)]. At the scale of the chromosome or genome, trends related to cytoband, gene density, and di-nucleotide frequency are revealed. These features are broad appearing in many different types of experiments. Care must be taken when conclusions are drawn as typically these features are removed by normalization or considered as a locally variable background. Generic web-based visualization solutions include the UCSC Genome Browser (Kent et al. 2002), ATLAS, and Ensembl (Hubbard et al. 2009) (wRef52, wRef53). It may also be useful to install local mirrors to avoid excessive time uploading data to the viewers. While some web viewers such as the NCBI’s mapviewer (Wolfsberg 2007) allow more detailed gene models to be explored, including user defined (NCBI model maker), the user may be limited in the freedom to incorporate their own sequencing data. Many other solutions that run locally are available from commercial vendors. These include Genome Studio from Illumina and the CLC workbench (wRef54) from CLC biosciences. Frequently, the sequencing or probe alignment will be made relative to a recent genome build such as human 37.1, but it is key to ensure that the genomic features used to contextualize data are aligned relative to the same build. One should be aware that the features may shift by distances as large as hundreds of kilobases if, for example, the features were aligned relative to an earlier build such as human 36.3; Coordinate translation solutions such as UCSC’s liftover tool (wRef55) to synchronize the feature information with the data are imperfect but necessary.

10.3.14

Sequencing Systems and Epigenetics

The construction of a systems level model of epigenetic information is intricate as one must consider two aspects. First, sequencing is generally used to measure properties from large aggregations of cells, where the heterogeneity of the cell population is unknown. How does the property of a cell aggregate differ from subpopulations or individual cells? Second, the measurements are generally taken as biological snapshots of highly dynamic systems responding to cell environmental interactions. Our understanding must include both spatial and temporal terms. The key to spatial assessment is to select the level of consistency that needs to be isolated to address the hypothesis being tested. For example, if we are considering spermatozoa as a means to understand idiopathic infertility, the general population trends within semen sample taken at multiple times from the infertile individual could inform the origins of the infertility. However, if the investigation concerns how spermatozoa interact with their environment as reflected by their ability to incorporate a molecule, then the level of investigation is more appropriately conducted closer to the level of the single gamete. The second general issue is the difference between the snapshot generated by most high throughput sequencing technologies and the very dynamic environment

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in which a cell is functionally operational. Cells may themselves exhibit a finer gradation of changes such as circadian clock-driven cycles of activity, seasonally and hormonally driven cycles over the broader range, and a range of very fine homeostatic interactions specific to the individual cell rather than population. It is an experimental design that will discern the key effects. While we may wish to dramatically alter the level at which we seek coherence based on the question asked, the technologies used for genetic analyses are typically less flexible in their ability to make these measurements. Recent innovations in single cell genetics have helped us move in this direction. The question becomes: when will we have the capability to simultaneously measure each cell without impacting that cell? At present, the task must be to assess when the variance within each level of the system may affect our ability to understand the system.

Suggested Reading and Environmental Epigenetic Effects Bioinformatics of DNA methylation 1. Smith ZD, Gu H, Bock C, Gnirke A, Meissner A (2009) High-throughput bisulfite sequencing in mammalian genomes. Methods 48(3):226 232 [Epub 2009 May 12.PMID: 19442738] 2. Bock C, Walter J, Paulsen M, Lengauer T (2008) Inter-individual variation of DNA methylation and its implications for large-scale epigenome mapping. Nucleic Acids Res 36(10):e55 [Epub 2008 Apr 15.PMID: 18413340] Bioinformatics of histone modifications 1. Zang C, Schones DE, Zeng C, Cui K, Zhao K, Peng W. A clustering approach for identification of enriched domains from histone modification ChIP-Seq data (2009) Bioinformatics. 25(15):1952 1958 [Epub 2009 Jun 8. PMID: 19505939] 2. Park PJ. ChIP-seq: advantages and challenges of a maturing technology (2009) Nat Rev Genet 10(10):669 680 [Epub 2009 Sep 8.PMID: 19736561] 3. Taslim C, Wu J, Yan P, Singer G, Parvin J, Huang T, Lin S, Huang K (2009) Comparative study on ChIP-seq data: normalization and binding pattern characterization. Bioinformatics 25(18):2334 2340 [Epub 2009 Jun 26.PMID: 19561022] 4. Shin H, Liu T, Manrai AK, Liu XS (2009) CEAS: Cis-regulatory element annotation system. Bioinformatics 25(19):2605 2606 [PMID: 19689956] 5. Auerbach RK, Euskirchen G, Rozowsky J, Lamarre-Vincent N, Moqtaderi Z, Lefranc¸ois P, Struhl K, Gerstein M, Snyder M (2009) Mapping accessible chromatin regions using Sono-Seq. Proc Natl Acad Sci USA 106(35): 14926 14931 [PMID: 19706456] 6. Choi H, Nesvizhskii AI, Ghosh D, Qin ZS (2009) Hierarchical hidden Markov model with application to joint analysis of ChIP-chip and ChIP-seq data. Bioinformatics 25(14):1715 1721 [Epub 2009 May 14.PMID: 19447789]

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7. Nix DA, Courdy SJ, Boucher KM (2008) Empirical methods for controlling false positives and estimating confidence in ChIP-Seq peaks. BMC Bioinformatics. 9:523 [PMID: 19061503] Histone modifications on testis-specific histones 1. Lu S, Xie YM, Li X, Luo J, Shi XQ, Hong X, Pan YH, Ma X (2009) Mass spectrometry analysis of dynamic post-translational modifications of TH2B during spermatogenesis. Mol Hum Reprod 15(6):373 378 [Epub 2009 Apr 3. PMID: 19346237] Bioinformatics of Nucleosomal organization and chromatin structure 1. Zhang Y, Shin H, Song JS, Lei Y, Liu XS (2008) Identifying positioned nucleosomes with epigenetic marks in human from ChIP-Seq. BMC Genomics 9:537 [PMID: 19014516] 2. Jiang C, Pugh BF (2009) Nucleosome positioning and gene regulation: advances through genomics. Nat Rev Genet 10(3):161 172 [Review. PMID: 19204718] Acknowledgments This work is supported in part by the NIH grant HD36512, the Presidential Research Enhancement Program in Computational Biology and the Charlotte B. Failing Profes sorship to SAK. We gratefully acknowledge the use of the UCSC genome browser (http://genome. ucsc.edu/) and the Ensembl genome browser (http://www.ensembl.org) that were used in the creation of some of the illustrations.

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Part III

Re-organization of Nuclear Compartments During Gametogenesis

Chapter 11

Organization of Chromosomes During Spermatogenesis and in Mature Sperm Olga Mudrak, Irina Zalenskaya, and Andrei Zalensky

Abstract One of the prominent features of male germ cell differentiation is a profound sequential, structural, and spatial reorganization of the chromosomes, which manifests in diverse events, such as chromatin remodeling, DNA recombination and repair, dynamic movements of chromosomes, and chromosome domains (centromeres and telomeres) within the nucleus. Divergent states of male germ cell chromosomes during proliferation, meiosis, and final differentiation to sperm are accompanied and supported by unprecedented in-scale changes in the chromatin protein composition. It is known that in interphase somatic cell nuclei, each chromosome occupies a defined chromosomal territory (CT), whose position is nonrandom. Despite dramatic changes during spermatogenesis, the principle of nonrandom CT positioning is preserved in sperm. A specific feature of sperm chromosomes is their extraordinary compactness and hairpin structure. A distinctive feature of sperm chromatin is its tripartite organization supported by protamines and two pools of residual histones. We propose that the combination of these unique features plays a role during male pronucleus formation after fertilization. Keywords Spermatogenesis Spermatozoa Chromosome structure Chromosome positioning

O. Mudrak The Jones Institute for Reproductive Medicine, Eastern Virginia Medical School, Norfolk, VA, USA Institute of Cytology, Russian Academy of Sciences, St. Petersburg, Russia I. Zalenskaya and A. Zalensky (*) The Jones Institute for Reproductive Medicine, Eastern Virginia Medical School, Norfolk, VA, USA e mail: [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 11, # Springer Verlag Berlin Heidelberg 2011

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Introduction

Spermatogenesis is a complex and highly coordinated process of differentiation from adult spermatogonial stem cells to mature spermatozoa. One of the principal attributes of male germ cell differentiation is a profound sequential, structural, and spatial reorganization of the chromosomes, which manifests in diverse events, such as chromatin remodeling, DNA recombination and repair, dynamic movements of chromosomes, and chromosome domains (centromeres and telomeres) within the nucleus. Divergent states of male germ cells chromosomes during proliferation, meiosis, and final differentiation to sperm are accompanied and supported by unprecedented in-scale changes in the chromatin protein composition. Testisand sperm-specific histone variants and histone modifications (Boussouar et al. 2008; Churikov et al. 2004; Delaval et al. 2007; Govin et al. 2007; Meistrich et al. 2003) that emerge at different stages are followed by histone to transitional proteins to protamine transition during spermiogenesis (Eirin-Lopez and Ausio 2009; Meistrich et al. 2003). In some mammals (more considerably in humans) protamines are neighboring with the residual histones, which might have an important consequence for fertilization, and we will briefly discuss the current state of knowledge in this area. In this review, we will focus on the chromosomes in differentiating male germ cells of mammals (mostly humans). We will use the term “chromosome organization” to describe both chromosome higher order structure and their intranuclear positioning. The same characteristics have been called “higher order chromatin arrangement” (Koehler et al. 2009), “genome architecture” (Zalensky 1998), and they are the immanent constituents of a more broad concept of “nuclear architecture” (Nunez et al. 2009). We will review experimental data on changes in chromosome higher-order structures, specific intranuclear localization of chromosomes with a special attention to their telomeres and centromeres heterochromatic regions that play a leading role in formation of genome architecture. We suggest that the well-defined chromosome organization in spermatozoa that contributes to sperm epigenome is a combination of features essential for successful fertilization.

11.2

Chromosome Organization in Somatic Interphase: Brief Outlook

It is now well-established that spatial distribution and structural organization of chromosomes within eukaryotic nuclei at various stages of cell differentiation and cell cycle are dynamic, nonrandom, and tightly connected with gene regulation (Spector 2003).

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Numerous studies of the last decades exploited DNA probes specific to individual chromosomes or selected chromosomal domains, to examine three-dimensional (3-D) genome organization using constantly improving fluorescence in situ hybridization (FISH) and microscopic techniques, for more details, see reviews by Dernburg and Sedat (Dernburg and Sedat 1998; Fraser and Bickmore 2007; Walter et al. 2006). It has been demonstrated that in the interphase nucleus, each individual chromosome occupies a defined space referred to as a chromosome territory (CT) (Cremer et al. 1993, 2004; Foster and Bridger 2005; Parada et al. 2004; Ramirez and Surralles 2008). Importantly, individual CTs are characterized by their preferred intranuclear positions (Cremer et al. 2004; Fedorova and Zink 2009; Fraser and Bickmore 2007; Misteli 2004; Verschure 2004). Thus, Bickmore and her colleagues found that in diploid lymphoblasts and primary fibroblasts, gene-rich chromosomes are preferentially located at more internal regions, while the gene-poor chromosomes concentrate towards the nuclear periphery (Gilbert et al. 2005). The existence of the radial positioning suggests its functional relevance, most probably, to the control of gene expression. Further details about 3-D chromosome structure and specific interchromosomal contacts are being acquired using conformation capture technique (Sexton et al. 2009), where chromatin fragments in close proximity are captured by fixation, restrictionenzyme digestion, and intramolecular ligation (Fraser and Bickmore 2007). The mechanisms by which genomes are spatially organized are unknown and several possibilities have been proposed: activity-driven self-organization, specific interactions with nuclear matrix/membrane, and associations between heterochromatic domains (Parada et al. 2004). In addition, involvement of hypothetical protein complexes including nuclear actin, myosin, lamin, and emerin was suggested (Mehta et al. 2008).

11.3

From Spermatogonia to Early Spermatids

11.3.1 Proliferation The Spermatogonia cell population is being constantly renewed by mitotic division. A subgroup of spermatogonia that is committed to further differentiation undergoes a limited number of cell divisions and enters meiosis. Experimental data on the chromosomal organization in the spermatogonia are scarce. It has been shown using FISH with painting probes that the majority of spermatogonia displays compact CTs, which develop into long chromatin threads at the onset of meiotic prophase (Scherthan et al. 1996). Telomeres and centromeres are found to be dispersed throughout the nuclei of the mouse, bovine, and human spermatogonia (Scherthan et al. 1996; Tanemura et al. 2005; Zalensky et al. 1997), (Fig. 11.1a). A similar pattern of telomere distribution is observed in somatic cells during G0/G1 and S-phases (Luderus et al. 1996; Ramirez and Surralles 2008).

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a

b

d late preleptotene

e zygotene

pachytene

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c mid preleptotene

centromeres telomeres

Fig. 11.1 Telomeres at early stages of spermatogenesis. Schematic presentation of telomere (open circles) and centromere (filled circles) relocation during early human and mouse spermato genesis (based on data of Scherthan et al. (1996) and Zalensky et al. (1997)). CT territories and chromosomes paths are shown in grey. Modified from Zalenskaya and Zalensky (2002), with permission

It can be suggested that the general principles of CT structures and behavior in the spermatogonia would be similar to those of proliferating somatic cells.

11.3.2

Stages of Meiosis I

Spermatogonia that cease continuous proliferation and move on to the spermatocyte stage undergo two successive divisions, meiosis I and meiosis II. As a result, each spermatogonium gives rise to four spermatids, haploid germ line cells that transform to mature sperm during spermiogenesis. The longest phase of meiosis is the first stage of meiosis I prophase I that is subdivided into preleptotene, leptotene, zygotene, pachytene, diplotene, and diakinesis stages. During prophase I, homologous chromosomes align side by side, forming a structure called bivalent. Nonsister chromatids pair and may exchange their parts by crossing over at points called chiasmata. Genetic recombination at prophase I and random assortment of the homologous chromosomes during cell divisions provide diverse combinations of parental genetic material. Specific and remarkable structural organization of meiotic chromosomes and participating proteins have been intensively studied during last decades, and results were summarized in numerous excellent review papers, for example,

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(Hernandez-Hernandez et al. 2009; Page and Hawley 2003; Scherthan 2007; Stack and Anderson 2001).

11.3.2.1

Movements and Interactions of Telomeres and Centromeres During Prophase I of Meiosis

During the extended meiotic Prophase I, pairing, synapsis, and recombination of homologous chromosomes are accompanied by significant changes in the chromosome morphology and localization. Characteristic of these processes are unprecedented movement and intranuclear repositioning of the key structural domains of meiotic chromosomes centromeres and telomeres. Telomere and centromere dynamics in meiosis have been studied in detail in mouse (Scherthan et al. 1996; Tanemura et al. 2005), and human (Scherthan et al. 1996; Zalensky et al. 1997) using FISH and immunolocalization of proteins specific for these domains (i.e., centromere-specific histone H3, CENP-A). In preleptotene, centromeres move to the nuclear periphery and form small clusters (Scherthan et al. 1996; Zalensky et al. 1997), Fig. 11.1b. This is the earliest movement, which has been registered both in mouse and man (Scherthan et al. 1996). Most of the telomeres remain dispersed throughout the nucleus. In the late preleptotene/leptotene, mouse centromeres form clusters at the nuclear envelope, while in humans, they move back to the nucleus interior, Fig. 11.1c. In both cases, chromosomes expand and unravel into cord-like CTs, with the ends attached to the inner nuclear membrane (Scherthan et al. 1996). By zygotene, telomeres move along the nuclear membrane and form a transient cluster located on one side of the nuclear envelope, the bouquet-like formation (Pfeifer et al. 2001; Scherthan et al. 1996), Fig. 11.1d. The driving forces behind such movement in mammals are not known; involvement of cytoskeleton motors has been suggested (Scherthan 2001). Parallel visualization of the telomeres and proteins of the axial element and transverse filament demonstrated that the telomere clustering is connected with the onset of chromosome pairing (Pfeifer et al. 2001; Scherthan et al. 1996). In pachytene, chromosome homologs are tightly paired and are seen as a single tract. Telomeres remain anchored to the inner surface of the nuclear envelope (Pfeifer et al. 2001; Scherthan 2001) and their pairing remains prominent, Fig. 11.1e. In diplotene, when synaptonemal complex is dissolved, telomere pairing is still observed, suggesting its possible additional function (Liebe et al. 2004).

11.3.2.2

Proteins Participating in Telomere Movement

Data are accumulating that demonstrate that the dynamic behavior of telomeres is the major factor directing proceeding of meiosis I. What are the molecular mechanisms mediating telomere movements from nucleoplasm to nuclear membrane, their attachment to and sliding along the nuclear envelope? In recent years, a number of proteins have been reported that may be related to the telomere movement. Their

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description is beyond the scope of the current review; therefore, only a short listing of the potential players is provided. Mammalian telomere-binding proteins TRF1 and TRF2 and Rap1 were found at the telomeres in mice spermatocytes, and their involvement in the telomere tethering was suggested (Liebe et al. 2004). Sun1, an integral protein of the inner nuclear membrane (Dreger et al. 2001), is associated with the telomeres from the leptotene to diplotene stages of meiotic prophase I (Ding et al. 2007). Disruption of Sun1 in mice prevents telomere attachment to the nuclear envelope, efficient homolog pairing, and synapsis formation in meiosis (Ding et al. 2007). Yet another transmembrane protein, Sun2, has been localized to the telomeric attachment sites on the nuclear envelope during meiosis (Schmitt et al. 2007). EM analysis has demonstrated that Sun2 is a part of membranespanning fibrillar complex that interconnects attached telomeres with cytoplasmic structures (Schmitt et al. 2007).

11.3.3 Sex Chromosomes During pachytene, the X- and Y-chromosomes demonstrate a distinct behavior. They migrate to the nuclear periphery and form there a specialized nuclear compartment called the sex- or XY-body. Unlike the autosomal chromosomal bivalents, where homologous recombination and transcription occur, X- and Y-chromosomes within the sex body are condensed and transcriptionally repressed. Recently, it has been demonstrated that in contrast to the autosomes, X- and Y-chromosomes have no homology between them except for a small area known as a pseudoautosomal region (PAR). While the autosomal homologs synapse completely, synapsis between X- and Y-chromosomes occurs only at PAR, leaving the largest part of the XY chromosomal bivalents unsynapsed. The unsynapsed regions somehow get recognized and this triggers the process of meiotic sex chromosome inactivation (MSCI) marked by the large-scale remodeling, condensation, and transcriptional silencing of sex chromosomes, for review, see (Turner 2007). The probable function of MSCI is the prevention of recombination between nonhomologous regions of X- and Y-chromosomes and, hence, protection against aneuploidy in the subsequent generations. A number of specific proteins associated with XY bivalents have been identified such as BRCA1, ATR, gH2AX ; (Baarends et al. 2005; Turner et al. 2004; van der Heijden et al. 2007). FISH analysis of pachytene nuclei of human, chimpanzee, and mouse has revealed that within the sex body X and Y chromosomes are organized into two separate nonentangled domains; each chromosome is bended so that their four telomeres are clustered at the nuclear envelope (Metzler-Guillemain et al. 2000). In primary spermatocytes, X and Y chromosomes are located at the nuclear edge (Foster et al. 2005; Handel 2004). But after the first meiotic division (in the secondary spermatocytes), they migrate to the nuclear center and stay there all the way through the spermiogenesis and in mature spermatozoa (Foster et al. 2005).

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267

Spermiogenesis

In mammals, the complex process of differentiation of haploid spermatids into mature spermatozoa is supported by massive and radical reorganization of chromatin. Most of the histones are replaced by protamines (Govin et al. 2004; Meistrich et al. 1978, 2003), which results in extreme compaction of DNA within the sperm nucleus to a volume of about 5% of that in somatic cells. Using FISH in the study of chromosome arrangement during rat spermiogenesis, Meyer-Ficca et al. (1998) find that in the rat round spermatids of steps 1 9 both pericentromeric and telomeric signals are randomly dispersed throughout the nucleoplasm. During steps 9 12, the heterochromatic domains start to associate: some pericentromeres are clustered into compact chromocenters, some are seen as oblong linear arrays, supposedly representing partially dispersed chromocenters. Initially, most telomeres assemble around the nucleolus, pronounced telomere telomere paring being markedly increased with the further progression of spermiogenesis. Similar patterns of telomere and centromere dynamics are observed in the mouse spermatids using immunolocalization of domain-specific proteins CENP-B and TRF2 (Dolnik et al. 2007). A clearly distinct chromocenter forms in the nucleus of the mouse stages 3 4 spermatids. Interestingly, some cells reveal two spatially separated linear arrays. These data support an earlier hypothesis that the pairwise association of the centromeres is the universal principle of centromere arrangement at the postmeiotic stages (Haaf et al. 1990).

11.5

Spermatozoa

The concept of nonrandom chromosome architecture in the nuclei of mature human spermatozoa has been developed based on earlier FISH and protein immunodetection studies (Haaf and Ward 1995; Zalensky et al. 1993, 1995, 1997). Hybridization with pan-centromeric probes and immunolocalization of CENP-A demonstrated that centromeres of all nonhomologous chromosomes are collected into the chromocenter that was localized, using confocal microscopy, to the nuclear interior (Zalensky et al. 1993). Upon controlled sperm decondensation, the chromocenter stretches into two V-shaped linear arrays of individual centromeres (Zalensky et al. 1993). All telomeres are positioned at the nuclear periphery where they interact with each other to form dimers and tetramers (Zalensky et al. 1997). Later it was found, using FISH with arm-specific subtelomere probes, that telomere pairs are formed through specific interactions between the two ends of the same chromosome (Solov’eva et al. 2004). The model of the human sperm nuclear architecture has been proposed according to which CTs are stretched between the chromocenter located centrally and telomeres occupying nuclear periphery (Zalenskaya and Zalensky 2002; Zalensky

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et al. 1995, 1997), Fig. 11.2a. According to this model, human sperm chromosomes adopt a looped, hairpin conformation. Similar location and interactions were demonstrated for sperm telomeres of other mammals: rat (Meyer-Ficca et al. 1998; Zalensky et al. 1997), mouse, bull, stallion, boar, and opossum (Zalensky et al. 1997, unpublished data). The sperm chromocenters were found in bull (Powell et al. 1990; Zalensky unpublished), mouse (Dolnik et al. 2007), and rat (Meyer-Ficca et al. 1998). Rat and mouse chromocenters are build up of the centromere arrays that are apparently similar to the human ones (Dolnik et al. 2007; Meyer-Ficca et al. 1998). It can be suggested that general principles of the sperm nuclear architecture maintained by dynamic telomere and centromere interactions are preserved through mammalian evolution. As a result of substitution of histones by protamines, sperm chromatin structure is fundamentally different from that of somatic cells. Nevertheless, chromosome arrangement into distinct CTs has been shown to be preserved in the sperm of humans (Manvelyan et al. 2008; Mudrak et al. 2005; Zalenskaya and Zalensky 2004; Zalensky et al. 1995), rat (Meyer-Ficca et al. 1998), boar (Foster et al. 2005), and distant monotremes (Greaves et al. 2003). Although direct comparison between CT dimensions in somatic and sperm cells is impeded due to the cell-specific variations in the FISH protocols, nevertheless, based on the observed difference in the DNA compaction, it can be postulated that the sperm CTs are much tighter than the somatic ones. Internal organization of the sperm CTs was further studied in sperm cells subjected to controlled decondensation using treatment with increasing concentration of Heparin in the presence of DTT (Zalensky et al. 1993). This treatment causes partial unraveling of compact CTs, thus allowing visualization of the inside structures (Fig. 11.2b).

a

b Pk

qk pl ql

Fig. 11.2 Human sperm chromosomes (a) Model of chromosome organization in human sperm. Centromeres (red circles) are collected into chromocenter positioned in the nucleus interior, telomeres (green circles) interact in dimers at the nuclear periphery. Schematic paths of two CT are shown. Based on data of Zalensky et al. (1993, 1995, 1997); and Haaf and Ward (1995). (b) Unpacking of HSA1. Sperm cells at two stages of decondensation induced by Heparin/DTT treatment. FISH using arm specific painting probes (q arm green, p arm red), total DNA is counterstained with DAPI

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Using this approach, Mudrak et al. (2005) performed FISH with arm-specific DNA probes for HSA1, HSA2, and HSA5 to picture a relative arrangement of the arm domains in human sperm nuclei (Fig. 11.3). The extended territories of chromosomes were found to be oriented parallel to the long nuclear axis. Similar extended territories were shown for rat spermatozoa (Meyer-Ficca et al. 1998). Examination of FISH data demonstrates that the CTs are formed by closely positioned p- and q-arms, either aligned or intertwisted; chromosome arms within CTs displaying a distinct individuality. The chromosomal threads sharply bend in the vicinity of centromeres, whereas the tips of p- and q-arms are located close to each other (Fig. 11.3a, c). This observation is in line with the earlier discovery of specific interactions between p- and q-subtelomeres (Solov’eva et al. 2004) and endorse the earlier proposed model of CT hairpin conformation in human spermatozoa. Further decondensation of the sperm nuclei allows for more detailed examination of the chromosome arms. It has been found that chromosome arms consist of 1,000 nm width fibril, composed of chromatin globules 500 nm in diameter, often forming two strings (shown by arrows in Fig. 11.3b). Globules are interconnected by thinner chromatin filaments (Fig. 11.3b). It is speculated that each globule corresponds to a collection of 100 200 nucleoprotamine toroids, while the thinner chromatin threads may represent the nucleohistone. Earlier Haaf and Ward, using FISH on sperm chromatin spreads, observed beaded fibers with basic package units of 180, 360, and 600 nm (Haaf and Ward 1995). The model of hierarchical organization of chromosome territory (Mudrak et al. 2005) is shown in Fig. 11.3c.

11.6

Chromosome Positioning

Numerous studies of 3-D genome organization in somatic interphase proved that CTs have favorite intranuclear positions (Sect. 11.2). Concentration of gene-rich chromosomes at internal regions and gene-poor chromosomes towards the nuclear periphery suggests functional relevance, most probably to the control of gene expression. Although, in contrast to somatic cells, spermatozoa are genetically inactive, about 10 years ago, evidence of nonrandom spatial positioning of CTs in human sperm started to emerge (Gurevitch et al. 2001; Hazzouri et al. 2000; Sbracia et al. 2002; Tilgen et al. 2001). CT localization was performed using either chromosomespecific near-centromere (Hazzouri et al. 2000; Zalenskaya and Zalensky 2004) or whole-chromosome painting probes (Mudrak et al. 2005; Zalenskaya and Zalensky 2004; Manvelyan et al. 2008). There are some discrepancies between the existing results, which are most likely due to variation in procedures used for sperm decondensation prior to FISH. It could be possible that in some cases, harsh treatment with alkali or high concentration salts may cause an artificial

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a

p arm

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+ DAPI

HSA1

HSA2

HSA5

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HSA1 q arm

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Fig. 11.3 Imaging of chromosomes in human sperm using FISH. (a) Simultaneous visualization of p and q arms of HSA 1, 2, and 5. (b) Internal organization of chromosome arm fiber. The chromatin globules of 500 nm are indicated by arrows. Arrowheads point to the double rows of

Organization of Chromosomes During Spermatogenesis and in Mature Sperm Localization on periphery (% of CHR)

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chr.18

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chr.21

70 chr.Y

chr.20

60 y = 82,8696-2,606*x

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40 chr.17

30 chr.22

20 10

chr.19

0

2

4

6 8 10 12 14 16 18 20 22 24 26 gene density (mbp genes per CHR)

Fig. 11.4 Correlation of radial chromosomes distribution in human sperm with gene density (Manvelyan 2008)

chromosome movement. Nevertheless, all the studies indicate that a preferred intranuclear position of chromosomes exists. Chromosomes in human sperm display radial positioning for example, locations of HSA7 and HSA6 are mostly peripheral, while those of HSA16 and HSAX are more internal (Zalenskaya and Zalensky 2004). Analogous to somatic cells, radial intranuclear positioning seems to be driven by gene density (Manvelyan et al. 2008). Of eight small chromosomes, the gene-reach ones occupy more central positions while gene-poor chromosomes more peripheral (Fig. 11.4). Also in this study, correlation between the radial positioning and the chromosome size has been reported. In contrast, no correspondence has been found between CT size and distribution in boar sperm (Foster et al. 2005). Size-independent positioning is also observed for human chromosomes as determined from their centromere localization (Zalenskaya and Zalensky 2004). Since sperm nuclei in many mammals have a polarized extended shape with two poles, apical (acrosomal) and basal (tail attachment), it is possible to access the longitude positioning of CT. This has been done for human (Manvelyan et al. 2008; Mudrak et al. 2005; Zalenskaya and Zalensky 2004) and boar (Foster et al. 2005) spermatozoa. In human spermatozoa, at some chromosomes, HSA1, HSAX, and HSA7 were located closer to apical pole, whereas HSA5 and HSA6 closer to basal pole (Mudrak et al. 2005; Zalenskaya and Zalensky 2004). Existence of longitude ä Fig. 11.3 (continued) globules that form fiber of chromosome arm. (c) Model of hierarchical organization of chromosome in human sperm nuclei. From left to right: compact CT, p and q arms are tightly merged; extended chromosome hairpin, antiparallel arms are intertwined or stacked; 1,000 nm fiber of an individual arm; two rows of 500 nm nodular structures that form 1,000 nm fiber. From (Mudrak et al. 2005) with permission

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positioning may be a factor important in process of artificial fertilization using ICSI technique. Another potentially important characteristic of CT positioning is their relative localization within sperm nuclei. Few existing data demonstrate that in sperm some HSA CT have a tendency to be located in proximity (Tilgen et al. 2001; Zalenskaya and Zalensky 2004). Recently, contacts between different loci on different chromosomes have been documented, and it has been proposed they may play important roles in the regulation of gene expression (Cavalli 2007). Likewise, specific neighboring of sperm chromosomes may participate in programmed activation of male pronuclei during fertilization. Molecular mechanisms responsible for specific intranuclear positioning of sperm are unknown.

11.7

Tripartite Organization of Human Sperm Chromatin

The tight organization of the sperm chromosomes is provided by interactions of the sperm DNA with protamines highly basic proteins that almost completely replace histones during spermiogenesis. Nucleoprotamine is arranged into large toroidal subunits, each containing approximately 50 kbp of DNA (Balhorn 2007). Histones remain associated with not more than 1 3% of the sperm DNA in all mammalian species studied, except humans. In human sperm, histones constitute 10 15% of the total nuclear protein (Tanphaichitr et al. 1978). Function(s) of the residual histones remains unclear. A model has been suggested according to which histones are located in the linker regions separating nucleoprotamine toroids (Ward 2010), Fig. 11.5a. Important questions to be addressed in this connection are: (1) what

a

b CEN

TEL

DNA nucleoprotamine toroids gene containing nucleosomes pericentromeric and telomeric nucleosomes

Fig. 11.5 Distribution of histones and protamines along chromosomes in human sperm. (a) Short stretches of nucleosomes are sandwiched between nucleoprotamine toroids, authors interpretation of model by Ward (2010)

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are DNA sequences that are associated with the histones? (2) How are the nucleohistone and nucleoprotamine regions distributed along the genome? It was shown using chromatin digestion approach that in human sperm, part of the DNA is organized by histones into nucleosomes (Banerjee et al. 1995; Nazarov et al. 2008; Zalenskaya et al. 2000). The particles obtained by a mild digestion with micrococcal nuclease contain telomere-specific DNA sequences (Zalenskaya et al. 2000). Recently, it was also demonstrated using high-resolution genomic examination that the fraction resulted from the mild micrococcal digestion is enriched with gene sequences allegedly important for embryonic development (Arpanahi et al. 2009; Hammoud et al. 2009). In an alternative approach, an extensive combined action of restriction and endogenous nucleases was used. The chromatin solubilized by this method also displayed a nucleosomal periodicity; and presence of histones (Nazarov et al. 2008). As revealed by atomic force microscopy, nucleosomes are associated with large supranucleosomal globular particles, which in turn form compact fiber arrays (Nazarov et al. 2008). This portion of chromatin is enriched with pericentromeric sequences, which has not seen in the mildly digested chromatin. This is in line with the reports that the histones that have not been removed during chromatin reorganization in mouse spermiogenesis are associated with pericentromeric regions (Govin et al. 2007). Such data fit into the model where histones are located in the linker regions separating nucloprotamine toroids (Fig. 11.5a) proposed by Ward (2010). In summary, protamines are the key proteins that ensure tight packing of sperm chromosomes, while residual histones bind the DNA sequences that could be important during and/or immediately after fertilization. Those sequences could be genes or structural domains, such as telomeres and centromeres. The latter might be among the structures that are required at the initial stages of the male pronucleus development and chromosome movements. A schematic model is suggested that shows protamines and two types of histone localization (1) spread in patches between the nucleoprotamine toroids throughout the length of a chromosome and (2) concentrated at heterochromatic regions (Fig. 11.5b).

11.8

Concluding Remarks

Dynamic behavior of chromosomes in differentiating mammalian male germ cells results in their highly specific organization in mature sperm. Among many others, there are some interesting questions to be answered: What are the molecular mechanisms that organize sperm chromosomes into hairpins? Do the short acrocentric chromosomes in humans have the same higher-order looped structure? What supports a preferred intranuclear positioning of the sperm chromosomes? Are there chromosome positioning mechanisms that are common to sperm and somatic cells? Hypothetically, multiple levels of the unique chromosome organization in sperm have all features characteristic of epigenetic code that would be deciphered

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in the descending cells (Zalensky and Zalenskaya 2007). We anticipate that comprehensive analysis of the unpacking and activation of male genome during the early stages of pronuclei formation will be the leading topic of the studies of the near furure. Acknowledgments This work has been supported by the Jones Foundation grant and in part by National Institutes of Health Grant HD 042748 to A. Z.

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Meyer Ficca M, Muller Navia J, Scherthan H (1998) Clustering of pericentromeres initiates in step 9 of spermiogenesis of the rat (Rattus norvegicus) and contributes to a well defined genome architecture in the sperm nucleus. J Cell Sci 111(Pt 10):1363 1370 Misteli T (2004) Spatial positioning; a new dimension in genome function. Cell 119:153 156 Mudrak O, Tomilin N, Zalensky A (2005) Chromosome architecture in the decondensing human sperm nucleus. J Cell Sci 118:4541 4550 Nazarov IB, Shlyakhtenko LS, Lyubchenko YL, Zalenskaya IA, Zalensky AO (2008) Sperm chromatin released by nucleases. Syst Biol Reprod Med 54:37 46 Nunez E, Fu XD, Rosenfeld MG (2009) Nuclear organization in the 3D space of the nucleus cause or consequence? Curr Opin Genet Dev 19:424 436 Page SL, Hawley RS (2003) Chromosome choreography: the meiotic ballet. Science 301: 785 789 Parada LA, Sotiriou S, Misteli T (2004) Spatial genome organization. Exp Cell Res 296:64 70 Pfeifer C, Thomsen PD, Scherthan H (2001) Centromere and telomere redistribution precedes homologue pairing and terminal synapsis initiation during prophase I of cattle spermatogenesis. Cytogenet Cell Genet 93:304 314 Powell D, Cran DG, Jennings C, Jones R (1990) Spatial organization of repetitive DNA sequences in the bovine sperm nucleus. J Cell Sci 97(Pt 1):185 191 Ramirez MJ, Surralles J (2008) Laser confocal microscopy analysis of human interphase nuclei by three dimensional FISH reveals dynamic perinucleolar clustering of telomeres. Cytogenet Genome Res 122:237 242 Sbracia M, Baldi M, Cao D, Sandrelli A, Chiandetti A, Poverini R, Aragona C (2002) Preferential location of sex chromosomes, their aneuploidy in human sperm, and their role in determining sex chromosome aneuploidy in embryos after ICSI. Hum Reprod 17:320 324 Scherthan H (2001) A bouquet makes ends meet. Nat Rev Mol Cell Biol 2:621 627 Scherthan H (2007) Telomere attachment and clustering during meiosis. Cell Mol Life Sci 64:117 124 Scherthan H, Weich S, Schwegler H, Heyting C, Harle M, Cremer T (1996) Centromere and telomere movements during early meiotic prophase of mouse and man are associated with the onset of chromosome pairing. J Cell Biol 134:1109 1125 Schmitt J, Benavente R, Hodzic D, Hoog C, Stewart CL, Alsheimer M (2007) Transmembrane protein Sun2 is involved in tethering mammalian meiotic telomeres to the nuclear envelope. Proc Natl Acad Sci USA 104:7426 7431 Sexton T, Bantignies F, Cavalli G (2009) Genomic interactions: chromatin loops and gene meeting points in transcriptional regulation. Semin Cell Dev Biol 20:849 855 Solov’eva L, Svetlova M, Bodinski D, Zalensky AO (2004) Nature of telomere dimers and chromosome looping in human spermatozoa. Chromosome Res 12:817 823 Spector DL (2003) The dynamics of chromosome organization and gene regulation. Annu Rev Biochem 72:573 608 Stack SM, Anderson LK (2001) A model for chromosome structure during the mitotic and meiotic cell cycles. Chromosome Res 9:175 198 Tanemura K, Ogura A, Cheong C, Gotoh H, Matsumoto K, Sato E, Hayashi Y, Lee HW, Kondo T (2005) Dynamic rearrangement of telomeres during spermatogenesis in mice. Dev Biol 281: 196 207 Tanphaichitr N, Sobhon P, Taluppeth N, Chalermisarachai P (1978) Basic nuclear proteins in testicular cells and ejaculated spermatozoa in man. Exp Cell Res 117:347 356 Tilgen N, Guttenbach M, Schmid M (2001) Heterochromatin is not an adequate explanation for close proximity of interphase chromosomes 1 Y, 9 Y, and 16 Y in human spermatozoa. Exp Cell Res 265:283 287 Turner JM (2007) Meiotic sex chromosome inactivation. Development 134:1823 1831 Turner JM, Aprelikova O, Xu X, Wang R, Kim S, Chandramouli GV, Barrett JC, Burgoyne PS, Deng CX (2004) BRCA1, histone H2AX phosphorylation, and male meiotic sex chromosome inactivation. Curr Biol 14:2135 2142

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Chapter 12

Nuclear Lamins in Mammalian Spermatogenesis Manfred Alsheimer, Daniel Jahn, Sabine Schramm, and Ricardo Benavente

Abstract Nuclear lamins are important structural protein components of the nuclear envelope. The composition and properties of the nuclear lamina of spermatogenic cells differ significantly from those of somatic cells of the same species. Mammalian spermatogenic cells selectively express short lamin isoforms showing several peculiarities. In this chapter, we summarize what is known about germ linespecific lamins and discuss their possible relevance for germ cell differentiation and fertility. Keywords Lamins Mammals Meiosis Spermatogenesis Spermiogenesis

12.1

The Nuclear Lamina

The nuclear envelope of eukaryotic cells separates the nuclear content from the cytoplasm. It is a complex structure that consists of four components: (1) the outer nuclear membrane that is interconnected with the endoplasmic reticulum, (2) the inner nuclear membrane, which contains a variety of specific integral membrane proteins, (3) the nuclear pore complexes, which selectively regulate the exchange of molecules between the nucleus and the cytoplasm, and (4) the nuclear lamina, a filamentous protein meshwork that confers mechanical stability to the nuclear envelope. It is tightly associated with the inner nuclear membrane and fixed to it by several integral membrane proteins. The nuclear lamina is mainly composed of the lamins, which are intranuclear members of the intermediate filament protein family (McKeon et al. 1986). Lamins are fibrillar proteins that are able to polymerize into higher order filament arrays (Aebi et al. 1986); for review, see Stuurman et al. (1998) and Krohne (1998).

M. Alsheimer (*), D. Jahn, S. Schramm and R. Benavente (*) Department of Cell and Developmental Biology, Biocenter, University of W€ urzburg, W€ urzburg 97074, Germany e mail: [email protected] wuerzburg.de, [email protected] wuerzburg.de

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 12, # Springer Verlag Berlin Heidelberg 2011

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Because of their biochemical properties, their behavior during the cell cycle, and their expression pattern, two groups of nuclear lamins can be distinguished: the ubiquitously expressed B-type lamins and the A-type lamins, which are developmentally regulated and not present in undifferentiated cells. Differentiated mammalian somatic cells contain at least four different lamin variants: two B-type lamins, B1 and B2 that are encoded by two different genes (LMNB1 and LMNB2), and two A-type lamins, A and C, which are different splice forms of the lamin A gene (LMNA; for review, see Herrmann and Aebi (2004) and Dechat et al. (2008)). Like other members of the intermediate filament protein family, somatic lamins contain a large central a-helical coiled coil-forming rod domain flanked by non-ahelical head and tail domains (Fig. 12.1; Stuurman et al. 1998; Krohne 1998; Herrmann and Aebi 2004). In addition, they contain a nuclear localization signal and a CaaX motif at the very end. This motif represents a target site for postranslational farnesylation that is essential for nuclear envelope association of the molecules. One exception concerns lamin C that lacks a CaaX motif and requires other lamins for its nuclear envelope association (for review, see Krohne (1998) and Stuurman et al. (1998)). Although it is currently not known how lamin polymers are assembled in vivo, in vitro studies indicate that the initial step for lamin polymerization is a formation of parallel coiled coil in register of two lamin molecules via their a-helical rod domains. These dimers further assemble into higher order filaments by head-to-tail and lateral interactions, whereby both rod ends and the immediate flanking amino acids within the head and tail domains seem to play a crucial role (Fig. 12.1; Stuurman et al. 1998). Besides the commonly accepted architectural function of the lamins, that is, forming a mechanically stable lamina network, many additional functions have

nuclear envelope association

longitudinal and lateral association

lamin subtypes

farnesylation

dimer formation

lamin B1 lamin B2 lamin A

NLS 1B

1A

2A

2B

CaaX

lamin C 120

lamin C 2 GNAEGR

myristoylation

nuclear envelope association 207

lamin B3

CaaX 84 AA

Fig. 12.1 Schematic representation of the molecular organization of somatic and germ line specific lamins of mammals. Domains in somatic lamins essential for dimer formation and longitudinal and lateral association of molecules are indicated

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been identified. Lamins are involved in nuclear envelope breakdown and reassembly during mitosis, interphase chromatin organization, transcription, replication, cell cycle progression, and apoptosis, and they also seem to have a role in nuclear membrane growth (for review, see Gruenbaum et al. (2005), Schirmer and Foisner (2007), Stewart et al. (2007), Dechat et al. (2008)). Therefore, lamins can be seen as multifunctional proteins that form a stable nuclear envelope scaffold and are critically involved in many fundamental cellular and developmental processes. This view is supported by the fact that a variety of severe human diseases (i.e., laminopathies) as certain muscular dystrophies, lipodystrophies, cardiomypathy, neuropathy, and premature aging syndromes are linked to mutations in A-type lamins (Dauer and Worman 2009).

12.2

Mammalian Germ Line-Specific Lamin Isoforms

When compared with somatic cells, the nuclear envelope of mammalian spermatogenic cells shows a series of peculiarities. Particularly interesting are the findings concerning the nuclear lamina (Fig. 12.2). Among somatic lamins, lamin B1 is the only one that is expressed during spermatogenesis. Lamins A, C, and B2 are not present in any phase of this differentiation process (Vester et al. 1993). Instead, two short gametogenesis-specific lamins appear at different stages to become part of a therefore specifically modified lamina. These are the A-type lamin C2 that is selectively expressed during meiotic prophase I (Smith and Benavente 1992; Furukawa et al. 1994; Alsheimer and Benavente 1996) and lamin B3, a short lamin B2 isoform (Furukawa and Hotta 1993), whose expression is restricted to postmeiotic stages (Sch€ utz et al. 2005a).

A-type lamins lamin A lamin C lamin C 2 B-type lamins lamin B1 lamin B2 lamin B3

Fig. 12.2 Schematic representation of lamin expression pattern in mammalian spermatogenic cells

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Compared to their somatic counterparts, both gametogenesis-specific lamins are N-terminally truncated versions, that is, they lack the complete N-terminal head including a significant part of the rod domain (Fig. 12.1). These domains are replaced by short unique sequences. In case of lamin C2, it is only six amino acids long (Furukawa et al. 1994; Alsheimer et al. 2000). In lamin B3, the specific domain consists of 84 unique amino acids (Furukawa and Hotta 1993; Fig. 12.1). The N-terminal truncation in both lamins is remarkable, as it concerns the front part of the rod domain, which was shown to be crucial for the assembly into higher order structures. Therefore, mammalian germ line-specific lamins can be seen as “natural deletion mutants” of their somatic counterparts that possess unique properties due to the lack of the head of the rod domain. Their stage-specific appearance and behavior that is linked to the most crucial events during spermatogenesis makes those good candidates for playing a major role during these processes. In the following text, we summarize available information about meiotic and postmeiotic mammalian lamins, largely rodent lamins, and discuss their possible functions.

12.3

Nuclear Lamins in Mammalian Meiosis

As mentioned above, the composition of the nuclear lamina during meiotic prophase I of mammals differs significantly from that of somatic cells as it is formed by lamin B1 together with the meiosis-specific A-type isoform C2. Additional differences are the relative low amount of lamins in meiotic cells (Vester et al. 1993; Alsheimer et al. 1999) as well as the relative lower structural stability of the meiotic nuclear lamina in comparison to that of somatic cells (Stick and Schwarz 1982). The relative low stability of the meiotic lamina structure is most likely due to the general low amount of lamins and the unique properties of lamin C2 (see also below). There is no indication that lower stability might be related to the fact that we are dealing with dividing (prophase) cells. In somatic cells, lamins become selectively hyperphosphorylated during mitotic prophase, a modification that results in nuclear lamina depolymerization and nuclear envelope breakdown. We have shown that during meiotic prophase, the isoelectric points of lamin B1 and LAP2 of the inner nuclear envelope are indistinguishable from those of interphase somatic cells, an indication that these proteins are not hyperphosphorylated during meiotic prophase (Alsheimer et al. 1999). Changes in migration properties of lamin B1 and LAP2 which would be consistent with hyperphosphorylation are detectable later during transition from prophase to metaphase (von Glasenapp and Benavente 2000). Lamin C2 presents a series of peculiarities that are consistent to the notion that it fulfils specific functions, different from that of other A-type isoforms. As mentioned above, it is selectively expressed during the first meiotic prophase. Later in

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meiosis, it is no longer detectable with immunocytochemical and biochemical methods (Alsheimer and Benavente 1996; von Glasenapp and Benavente 2000). In contrast to somatic lamins, lamin C2 lacks a C-terminal CaaX box. Instead, it contains a myristoylation site corresponding to the first six N-terminal amino acids of the molecule (Fig. 12.1: GNAEGR; Alsheimer et al. 2000). Direct mutation of this site combined with immunolocalization and mass spectrometry revealed that lamin C2 is in fact myristoylated and that this hydrophobic moiety is essential for nuclear envelope association. Lamin C2 molecules lacking a myristoyl moiety remain diffusely distributed in the nucleoplasm (Alsheimer et al. 2000). Among known vertebrate and invertebrate lamins lamin C2 is the only one requiring myristoylation for nuclear envelope association. Remarkably, lamin C2 also lacks the first 120 amino acids that, in somatic A-type lamins, correspond to nonhelical N-terminus and the head of the rod domain. As shown in Fig. 12.1, these domains are essential for proper lateral and head-to-tail interactions between molecules. These differences suggest that polymerization properties of lamin C2 strongly differ from those of other A-type isoforms. Two sets of experiments support this assumption. When overexpressed in somatic cells, lamin A polymerizes to highly organized paracrystalline structures showing a characteristic repeat organization (Stuurman et al. 1998; Krohne 1998). Under the same experimental conditions, lamin C2 forms fibrillar aggregates that, however, are less organized and lack a paracrystalline organization (Pr€ ufert et al. 2005). More recently, we have compared the properties of lamin C2 and somatic lamins in FRAP and FLIP experiments. As previously reported (Broers et al. 1999), somatic lamins were virtually immobile. In contrast, lamin C2 showed a significantly higher mobility that, as we could determine, is a consequence of the lack of the globular N-terminus and the head of the rod domain (Jahn et al. 2010). What is the functional significance of the conspicuous nuclear lamina of meiotic cells? During the extended first meiotic prophase, homologous chromosomes pair, synapse, and recombine. These processes are essential to ensure a correct segregation during the reductional first meiotic division (for review, see Zickler and Kleckner (1998, 1999), Benavente and Volff (2009)). Interestingly, in early meiotic prophase, chromosomes become attached to the nuclear envelope via their telomeres. This attachment is highly dynamic as it allows telomeres to move within the nuclear envelope and to congregate at a pole of the nuclear envelope to form a so-called bouquet and to move apart later during pachytene. Telomere movements at the plane of the nuclear envelope are suggested to allow chromosome recognition as well as the stable pairing of the homologues (Zickler and Kleckner 1998, 1999; Alsheimer 2009). Nuclear lamins is somatic cells are rather homogeneously distributed at the nuclear periphery. Remarkably, this is clearly not the case for lamin C2, as it is highly enriched at the attachment sites of the telomeres (Alsheimer et al. 1999; Schmitt et al. 2007). Because of this specific localization pattern and on basis of the proposed influence of the N-terminal truncation on the polymerization properties of lamin C2, lamin C2 might lead to a local flexibilization of the nuclear envelope at

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the attachment sites that in turn enables the movement of telomeres and therefore supports pairing, recombination, and synapsis. Support for this hypothesis came from the study of a mouse model that was generated by Sullivan et al. (1999). Because of a targeted disruption of the LMNA gene, these animals lack A-type lamins, including lamin C2. Examination of the testes of Lmna-/- mice revealed that spermatogenesis is dramatically affected, as almost no postmeiotic stages appeared within the seminiferous tubules (Alsheimer et al. 2004). Correspondingly, virtually no sperm were found in the epididymis. Lmna-/- mice showed massive failures during the process of pairing and synapsis of the homologous chromosomes. This is reflected by the appearance of univalents, long unpaired axial elements, and nonhomologous pairing in pachytene spermatocytes. These defects are most likely the reason for the dramatic increase in apoptosis observed during pachytene stage which in turn leads to spermatogenesis disruption. These results allowed defining A-type lamins as important determinants of male fertility (Alsheimer et al. 2004). Experiments are in progress in our laboratory to determine the contribution of lamin C2 deficiency to this phenotype.

12.4

Nuclear Lamins in Mammalian Spermiogenesis

During spermiogenesis, postmeiotic cells undergo a profound remodeling, that is, among other changes, characterized by a species-specific nuclear reshapening from round to oblong (Benavente 2004; Hermo et al. 2010a, b). This also includes a successive replacement of histones with protamines, transcriptional inactivation, and chromatin condensation (for review, see Hermo et al. (2010a, b)). Remarkably, these processes are accompanied by dramatic changes within the nuclear envelope, including the nuclear lamina, which have been investigated to a much lesser extent. The functional significance of nuclear envelope changes, however, is largely unknown. The composition of the nuclear lamina in postmeiotic cells differs significantly from that of somatic cells and of meiotic cells. The nuclear lamina of spermatids is composed of lamin B1 and the spermiogenesis-specific lamin B3. In contrast to meiotic stages, no A-type lamins are detectable at any time (Fig. 12.2; Vester et al. 1993; Sch€ utz et al. 2005a). A further major difference deals with lamin distribution. In round spermatids, lamin B3 was detected at the nuclear periphery as well as diffusely distributed in the nucleoplasm. Later, lamin B3 of nuclear envelope and nucleoplasma becomes progressively polarized at the posterior pole of elongating spermatid nuclei where they accumulate before becoming undetectable at the end of spermiogenesis (Sch€ utz et al. 2005a, and below). As already mentioned, lamin B3 is a short isoform encoded by LMNB2, the gene that also codes for somatic lamin B2. A schematic representation of the lamin B3 molecule is shown in Fig. 12.1. In comparison to lamin B2, lamin B3 lacks the first

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240 amino acids corresponding to the nonhelical N-terminus as well as a significant part of the rod domain. Instead, lamin B3 contains a unique N-terminal end that in the mouse is 84 amino acids long (Furukawa and Hotta 1993; Sch€utz et al. 2005b). As in the case of lamin C2, lamin B3 lacks part of the domains that in somatic lamins have been shown to be important for the assembly of stable polymeric structures. To investigate possible peculiarities in the polymerization properties, lamin B3 was ectopically expressed in somatic cells. Under these experimental conditions, lamin B3 causes severe nuclear deformations. Nuclei are transformed from round to oblong (Furukawa and Hotta 1993; Sch€ utz et al. 2005b). In addition, we observed a dual localization, that is, lamin B3 localized both to the nuclear periphery and the nuclear interior as well (Sch€ utz et al. 2005b), which resembles exactly the situation for the endogenous lamin B3 in early round spermatids (Sch€ utz et al. 2005a). Nucleoplasmic localization and nuclear deformation leading to hook-shaped elongated nuclei are specific for lamin B3 as they were not observed in the case of lamin C2, that is, the other mammalian short lamin (Alsheimer et al. 2000). In cell fractionation experiments, it was demonstrated that, compared with lamin B2, lamin B3 shows a clearly reduced binding strength. This could be corroborated and extended by FRAP and FLIP approaches which revealed that, in contrast to the situation for ectopically expressed somatic lamin B2, lamin B3 molecules within the nuclear periphery are mobile and interchange with those of the nucleoplasmic pool. Furthermore, expression of different mutant constructs evidenced that these effects are solely due to the shortened rod domain and not a consequence of the unique N-terminal head domain (Sch€ utz et al. 2005b). Consistent with these results, recent polymerization studies showed that human lamin B3 is unable to form long filaments but only short structures (von Moeller et al. 2010). The functional role of the unique 84 amino acids at the N-terminus of lamin B3 is not known. Particularly intriguing is the general low evolutionary conservation of the corresponding mammalian sequences, even between rat and mouse. What is the functional significance of the conspicuous nuclear lamina composition and reorganization in spermiogenesis? The transfection and cell fractionation experiments as well as the immunolocalizations summarized here indicate that lamin B3 has peculiar polymerization properties that would provide the nuclear periphery of spermatids with a flexible condition, which in turn would be required for nuclear reshaping. As mentioned above, lamin B3 becomes polarized as spermiogenesis progresses. Interestingly, a similar polarization could be observed for lamin B1 and the inner nuclear membrane proteins LAP2 and LBR (Alsheimer et al. 1998; Mylonis et al. 2004; Sch€ utz et al. 2005a). At present, the molecular basis and functional significance of this redistribution is unclear. However, because of the already described chromatin-binding capability of most of the nuclear envelope proteins involved in redistribution, it can be speculated that nuclear envelope remodeling would contribute to the process of sperm-specific chromatin reorganization (Alsheimer et al. 1998; Mylonis et al. 2004).

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Nuclear Lamins in Spermatogenesis of Nonmammalian Vertebrates

In comparison to mammals, very little is known about lamin expression during spermatogenesis of nonmammalian vertebrates. Nevertheless, the emerging picture reveals that lamin expression in spermatogenic cells of nonmammalian vertebrates also differs from that of somatic cells (Benavente and Krohne 1985; von Moeller et al. 2010). In the amphibian Xenopus laevis, we detected the expression of spermatogenesis-specific lamin LIV that shows a patchy distribution at the nuclear periphery of sperms (Benavente and Krohne 1985). Very recently, von Moeller et al. (2010) have characterized lamin LIV in detail. They could demonstrate that lamin LIV is a splice variant of the lamin LIII gene. This gene also codes for lamin LIII, the major lamin isoform expressed in Xenopus oocytes (Stick 1988). Lamin LIV is longer than lamin LIII (as well as other Xenopus lamins) due to an extension within the rod domain of the molecule. Interestingly, this extension results in altered polymerization properties of LIV similar to those of mammalian germ line-specific lamins. The findings in amphibians are consistent with the notion that sperm head formation in vertebrates requires a flexible nuclear periphery, but that germ line-specific lamins followed different evolutionary paths (von Moeller et al. 2010).

12.6

Outlook

Mammalian germ line-specific lamins can be seen as “natural deletion mutants” of their somatic counterparts that possess unique properties due to the lack of the N-terminus and the head of the rod domain. Their stage specific appearance and behavior that is linked to the most crucial events during spermatogenesis makes them good candidates for playing a major role during these processes. Hopefully, the generation of knockout mice lacking lamins C2 and B3, respectively, as well as the investigation of their protein-binding partners in the future will provide more insight into the functional relevance of these unique isoforms. Acknowledgments Supported by the Priority Programme 1384 of the DFG, the Graduate School “Organogenesis,” and the Elitenetzwerk Bayern.

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€g C, Stewart CL, Alsheimer M (2007) Transmembrane Schmitt J, Benavente R, Hodzic D, H€ oo protein Sun2 is involved in tethering mammalian meiotic telomeres to the nuclear envelope. Proc Natl Acad Sci USA 104:7426 7431 ¨ llinger R, Benavente R (2005a) Nuclear envelope remodelling during Sch€utz W, Alsheimer M, O mouse spermiogenesis: postmeiotic expression and redistribution of germ line lamin B3. Exp Cell Res 307:285 291 Sch€utz W, Benavente R, Alsheimer M (2005b) Dynamic properties of germ line specific lamin B3: the role of the shortened rod domain. Eur J Cell Biol 84:649 662 Smith A, Benavente R (1992) Identification of a short nuclear lamin protein selectively expressed during meiotic stages of rat spermatogenesis. Differentiation 52:557 563 Stewart CL, Roux KJ, Burke B (2007) Blurring the boundary: the nuclear envelope extends its reach. Science 318:1408 1412 Stick R (1988) cDNA cloning of the developmentally regulated lamin LIII of Xenopus laevis. EMBO J 7:189 197 Stick R, Schwarz H (1982) The disappearance of the nuclear lamina during spermatogenesis: an electron microscopic and immunofluorescence study. Cell Differ 11:235 243 Stuurman N, Heinz S, Aebi U (1998) Nuclear lamins: their structure, assembly, and interactions. J Struct Biol 122:42 66 Sullivan T, Escalante Alcalde D, Bhatt H, Anver M, Bhat N, Nagashima K, Stewart CL, Burke B (1999) Loss of A type lamin expression compromises nuclear envelope integrity leading to muscular dystrophy. J Cell Biol 147:913 919 Vester B, Smith A, Krohne G, Benavente R (1993) Presence of a nuclear lamina in pachytene spermatocytes of the rat. J Cell Sci 104:557 567 von Glasenapp E, Benavente R (2000) Fate of meiotic lamin C2 in rat spermatocytes cultured in the presence of okadaic acid. Chromosoma 109:117 122 von Moeller F, Barendziak T, Apte K, Goldberg MW, Stick R (2010) Molecular characterization of Xenopus lamin LIV reveals differences in the lamin composition of sperms in amphibians and mammals. Nucleus 1:1 11 Zickler D, Kleckner N (1998) The leptotene zygotene transition of meiosis. Annu Rev Genet 32:619 697 Zickler D, Kleckner N (1999) Meiotic chromosomes: integrating structure and function. Annu Rev Genet 33:603 754

Part IV

Fundamental Aspects of Gene Expression Regulation During Gametogenesis

Chapter 13

Specific Transcription Regulatory Mechanisms of Male Germ Cells Irwin Davidson

Abstract Spermatogenesis is a remarkable process in which spermatogonial stem cells differentiate first into primary spermatocytes that after two successive meiotic divisions undergo a major biochemical and structural reorganization of the haploid cells to generate mature elongate spermatids. Each of these steps is controlled by precise transcriptional regulatory programs that orchestrate this complex series of events. Genetic and biochemical approaches in model organisms such as Drosophila melanogaster and mouse have identified many of the factors involved and revealed mechanisms of action that are unique to male germ cells. Defects in these transcriptional programs have dramatic and often very specific effects on male fertility. As many of these factors and pathways are conserved in human spermatogenesis, the lessons learned from the study of these model organisms are likely to provide important insights into understanding the molecular mechanisms of human infertility. Keywords Chromatin Drosophila Meiosis Spermatogenesis TATA-binding protein TFIID

13.1

Introduction

Spermatogenesis is a remarkable differentiation process that takes place continuously or seasonally during the adult life of many organisms. In mammals, spermatogenesis occurs in the seminiferous epithelium of the testis under a complex endocrine control. Spermatogenesis initiates when cells from the spermatogonia stem cell population commit to differentiation rather than remaining in the selfrenewing population. These cells give rise to early primary spermatocytes that

I. Davidson Institut de Ge´ne´tique et de Biologie Mole´culaire et Cellulaire. CNRS/INSERM/UDS, 1 Rue Laurent Fries, 67404 Illkirch, France e mail: [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 13, # Springer Verlag Berlin Heidelberg 2011

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replicate their DNA, giving rise to leptotene spermatocytes. During this premeiotic phase in primary spermatocytes, aligned homologous chromosomes pair, the synaptonemal complex is formed and crossing over takes place. This is followed by two successive meiotic divisions to generate the haploid round spermatids (Clermont 1972; de Rooij 2001). During the postmeiotic spermiogenesis phase, the round spermatids undergo a major morphological and biochemical transformation involving the loss of most somatic histones, the condensation of the genome into a tightly packed protamine DNA complex, flagellar formation, and finally cytoplasmic exclusion (Kimmins and Sassone-Corsi 2005). Transcription is repressed during the elongation phase as the histone octamer-based chromatin organization of the somatic cells is lost and replaced by the incorporation of transition proteins and protamines. In mouse and in humans, however, a small fraction of the genome retains a nucleosomal organization (Gatewood et al. 1990). It has been speculated that inheritance of these histones and their covalent modifications may confer epigenetic information that influences gene expression following fertilization (Albert and Peters 2009; van der Heijden et al. 2008). Each stage of the spermatogenic process is controlled by complex regulatory programs directing the expression of specific sets of genes at different developmental stages. In mouse, a first set of germ cell-specific genes are expressed in pachytene spermatocytes followed by a second wave of postmeiotic transcription in round spermatids where most genes required for morphological and biochemical reprogramming are expressed (Sassone-Corsi 2002). Study of transcription regulation in spermatogenesis is hampered by the lack of an in vitro differentiation model. The most important discoveries have been possible due to the power of genetics in model organisms such as Drosophila and mouse where many of the factors and mechanisms involved have been elucidated (Fuller 1998; Sassone-Corsi 2005). These studies have revealed that transcription regulation in male germ cells often involves unique and specialized mechanisms mediated by paralogs of components of the general transcription machinery or mechanisms that bypass the normal regulatory pathways used in somatic cells. In this chapter, I will discuss some of the specialized aspects of the male germ cell transcription program and how defects in these regulatory pathways may provide insights into the physiopathology of human fertility.

13.1.1

Transcriptional Regulation in Male Germ Cells of Drosophila Melanogaster

As described above, many critical insights into the transcription regulatory mechanisms of male germ cells have come from genetic studies of model organisms such as Drosophila and mouse. While the murine models are obviously more relevant to humans, some interesting paradigms have been established in Drosophila that are

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relevant for other organisms. The governing principal of the use of germ cellspecific paralogs of general transcription factors and unique regulatory mechanisms was first described in this organism. Unlike mammals, where there is significant postmeiotic transcription in round spermatids (Potrzebowski et al. 2008; Schultz et al. 2003), in Drosophila, the vast majority of genes required for spermatogenesis are expressed in spermatocytes. For example, protamines are only detected late in elongation; however, protamine transcripts accumulate in primary spermatocytes (Jayaramaiah Raja and Renkawitz-Pohl 2005). This large set of testis-expressed genes are regulated in primary spermatocytes by the concerted actions of two protein complexes, tMAC [testis meiotic arrest complex, (Beall et al. 2007; White-Cooper 2009)] and tTAFs (testis-specific TBP-associated factors), each of which are defined both biochemically and genetically. While the existence and function of a mammalian tMAC complex has not been studied, the tTAF complex illustrates the important paradigm. The general RNA polymerase II transcription factor TFIID is formed by the TATA-binding protein (TBP) and 13 14 TBP-associated factors (TAFs) (Cler et al. 2009; Matangkasombut et al. 2004; Tora 2002). The composition and organization of TFIID is highly conserved through evolution. TFIID has a modular structure with a core domain formed by TAF5 and a set of histone fold-containing TAFs, upon which a second module comprising TAF1, TAF7, and TBP associates to form TFIID [(Gangloff et al. 2001; Papai et al. 2009) and see Fig. 13.1 and Table 13.1]. TBP and TAFs are expressed in almost all embryonic and adult tissues. However, male germ cells express paralogs of TBP and several TAFs that play critical but distinct roles in spermatogenesis in Drosophila and mouse. In Drosophila, the can-class meiotic arrest genes can, mia, nht, rye, and sa encode testis-specific paralogs of the somatically expressed TAFs that form the TFIID core (Hiller et al. 2001, 2004). These tTAFs share the same structural domains (histone-folds, WD40 repeats) and form heterodimers similar to those described for the corresponding somatic core TAFs [(Kolthur-Seetharam et al. 2008) and see Table 13.1]. In addition, a testis-specific splice isoform of TAF1 has been described (Metcalf and Wassarman 2007). The tTAFs do not appear to act by forming a testis-specific TFIID complex, but their function is rather to sequester the Polycomb Repression Complex 1 [PRC1, (Saurin et al. 2001)] in the nucleolus. Polycomb and other PRC1 components localize exclusively to chromatin in spermatogonia, but relocalize to the nucleolus in primary spermatocytes (Chen et al. 2005) along with the tTAFs. Hence, the primary function of tTAFs is to remove the PRC1 repressor complex from the promoters of testisspecific genes and sequester it in the nucleolus, allowing the expression of these genes in spermatocytes. These results show that Drosophila male germ cells specifically express a set of TAF paralogs that do not function primarily through formation of a germ cell TFIID complex but by an alternative derepression mechanism.

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9

6

10

11 13

3 1311 6 9 4 5 12

8 5 3 10

8

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4

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+

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10

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Elongate spermatid

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Round spermatid

12 C

ALF 1/1L TBP IIA

4 TRF2

ALF IIA

TRF2 TIPT HP1γ

Fig. 13.1 Schematic description of TFIID. The assembly of the TFIID from its constituent TBP and TAF subunits is depicted. The subunits assemble into the core and TBP TAF1 modules, which

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Table 13.1 Composition and organization of TFIID. The TFIID subunits that form the TFIID core are listed and their mammalian and Drosophila tTAFs paralogues shown. The names of the corresponding Drosophila genes are shown in brackets. The square brackets indicate the known heterodimerizations between the core domain TAFs containing histone fold motifs. Below, the components of the TBP TAF1 module are indicated along with the mammalian TBP paralogs TFIID subunits TAF3

Mammalian Paralogues

Drosophila tTAFs

TAF10 dTAF8L (sa)

TAF8 TFIID core domain

TAF4

TAF4b

TAF12

dTAF4L (nht) dTAF12L (rye)

TAF6

TAF6L

TAF9

TAF9b

dTAF6L (mia)

TAF11 TAF13

TAF1-TBP module

TBP Familly

TAF5

TAF5L

TAF1

TAF1L

dTAF5L (can)

TAF2 TAF7

TAF7L

TBP

TRF2 TRF3 (TBP2)

ä Fig. 13.1 (continued) associate to form the full TFIID complex. The schematic view is superposed on the structure of TFIID as determined from immunoelectron microscopy (EM) experiments. Murine spermatocytes predominantly contain fully assembled TFIID complexes containing TAF4 and/or TAF4b. In haploid cells, TAF4 downregulation leads to loss of full TFIID and accumulation of the TBP TAF1 TAF7L submodule where TBP interacts with either TFIIA or ALF. TRF2 also interacts with either TFIIA or ALF to regulate transcription or interact with chromatin via TIPT

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13.1.2

Genes Encoding Paralogs of Subunits of the TFIID Complex Are Essential for Normal Spermatogenesis in Mammals

13.1.2.1

TBP-Related Factors

The Drosophila tTAFs described above and the derepression mechanism are not conserved in mammals. Nevertheless, the mammalian genome encodes paralogs of several other TFIID components that are specifically expressed in testis and are critical for normal fertility. The TBP-related factors TRF2 and TRF3 provide an interesting illustration of how evolution has selected a specific function for TFIID paralogs in the germ line. TBP has a bipartite structure with a poorly conserved N-terminal region and a highly conserved C-terminal domain that folds as a molecular saddle-shaped structure and binds DNA via the concave surface (Burley 1996). Vertebrate genomes encode two TBP-related proteins, TRF2 and TRF3 (also designated as TLF and TBP2, respectively), which share the conserved C-terminal domain (Bartfai et al. 2004; Dantonel et al. 1999; Jallow et al. 2004; Persengiev et al. 2003; Rabenstein et al. 1999; Teichmann et al. 1999). These factors play important and general roles in embryonic development in the zebrafish D. rerio or Xenopus (Bartfai et al. 2004; Hart et al. 2007; Jacobi et al. 2007; Muller et al. 2001; Veenstra et al. 2000). However, in mammals, TRF2 is expressed almost uniquely in male germ cells, while TRF3 expression is restricted to the ovary (Martianov et al. 2001; Xiao et al. 2006). Genetic knockout experiments in mice show that TRF2-null mice are viable and females are fertile (Martianov et al. 2001; Zhang et al. 2001). In contrast, male animals are sterile with an almost complete arrest of spermiogenesis due to apoptosis of step 7 round spermatids. TRF2 mutant round spermatids are characterized by the fragmentation of the chromocenter, a nuclear structure comprising the centromeric heterochromatin from each chromosome (Martianov et al. 2002). In normal spermatids, a single dense nuclear structure is observed, while in TRF2 mutant cells, 3 4 large foci are found (Fig. 13.2). The chromocenter is proposed to act as a chromatin organizer prior to the elongation phase (Zalensky et al. 1995). The TRF2 mutant spermatids undergo apoptosis at the beginning of the elongation phase, suggesting that chromatin disorganization activates a checkpoint at this step. Loss of TRF2 also perturbs spermatid nuclear polarity as evidenced by mislocalization of H1FNT, also known as H1T2 or HANP1, a germ cell-specific variant of histone H1 expressed in round spermatids (Martianov et al. 2005; Tanaka et al. 2005). In wild-type spermatids, H1FNT specifically localizes to the region of the nucleus below the acrosome at the apical pole (Fig. 13.2). Inactivation of H1FNT leads to impaired nuclear condensation and cytoplasmic exclusion during the elongation phase, resulting in reduced sperm count, motility, and fertility. In TRF2 mutant spermatids, H1FNT adopts a bipolar localization showing a loss of spermatid nuclear polarity (Catena et al. 2006). Thus, loss of TRF2 results in

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-/Trf 2

297 Trf 2-/-

Fig. 13.2 TRF2 knockout disrupts chromatin organization. Merged Hoechst and H1FNT immu nostaining of purified stage 7 round spermatids from wild type or Trf2/ animals. The white arrows indicate the chromocenter that is fragmented in the mutant spermatids. The yellow arrows indicate H1FNT that localizes to the apical pole in the wild type but show bipolar localization in many of the mutant spermatids. This can be also seen by superposing and H1FNT immunostaining on the differential interference contrast image shown on the right panel

chromatin disorganization evidenced both by the fragmentation of the chromocenter and the mislocalization of H1T2. Interestingly, cloning of the human H1fnt gene revealed the existence of five alleles with single nucleotide polymorphisms in the Japanese population (Tanaka et al. 2006). Three of these SNPs lead to amino acid substitutions, but so far no significant association with male infertility has been found. What is the molecular function of TRF2? Biochemical experiments show that TRF2, like TBP, interacts with TFIIA (Teichmann et al. 1999). Moreover, TRF2 interacts with ALF (Ozer et al. 2000; Upadhyaya et al. 1999), a paralog of the TFIIA a/b subunit, which is specifically expressed in late pachytene spermatocytes and in haploid round spermatids (Catena et al. 2005). Like TFIIA, ALF is cleaved into a and b subunits that associate with TFIIAg. Immunoprecipitation experiments show that male germ cells contain not only TFIIAa/b/g and ALFa/b-TFIIAg complexes but also the full combination of hybrid complexes between the TFIIA/ ALF a and/or b subunits (Catena et al. 2005). The TRF2 TFIIA/ALF interaction suggests that TRF2 may function as a transcription factor. However, at present, no TRF2 binding sequence has been identified, and there has been no demonstration that TRF2 is bound in vivo in testis to the target promoters. Thus, the effects of TRF2 on gene expression could be indirect. TRF2 may also have a more direct role on chromatin as two hybrid experiments have shown an interaction with TIPT (TRF2 interacting protein in testis), a novel uncharacterized testis-specific protein that is specifically expressed in late pachytene and haploid round spermatids (Brancorsini et al. 2008). TIPT also interacts with the heterochromatin proteins HP1a and HP1g and colocalizes with HP1g in the euchromatin compartment. Although the function of TIPT in testis remains to be determined, these observations provide a potential direct link between TRF2 and chromatin organization. Alternatively, a second isoform TIPT2 has been suggested

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to be directly involved in transcription through interactions with Geminin and TBP or TRF2 (Pitulescu et al. 2009). It is noteworthy that in Drosophila, TRF2 is also required for spermatogenesis. Mutation of TRF2 leads to defects in meiotic chromatin condensation and chromosome segregation (Kopytova et al. 2006). TRF2 forms a high molecular weight protein complex with the DNA binding factor DREF and proteins such as ISWI involved in chromatin remodeling providing further evidence that TRF2 may directly modulate chromatin organization (Hochheimer et al. 2002). The specific role of TRF2 in male germ cells is mirrored by that of TRF3 in female germ cells. During reproductive life, oocytes are continuously selected from a pool of primordial follicles to develop via a growth phase during which they are arrested at diplotene of prophase I. TBP is expressed in the primordial follicles, but it is rapidly downregulated prior to arrest in metaphase I (Gazdag et al. 2007). In contrast, TRF3 is strongly expressed throughout folliculogenesis, declines at the oocyte stage, and disappears following fertilization. These results reveal a developmental switch where TRF3 replaces TBP during oocyte maturation, suggesting that TRF3 may be required to direct the specific gene expression program required in oocytes. Genetic knockout of TRF3 in mice shows results in female infertility, deregulation of oocyte specific genes, and altered chromatin organization (Gazdag et al. 2009). In contrast, inactivation of TBP in the female germ line does not affect oocyte formation or female fertility, demonstrating that it is TRF3 and not TBP that is essential for the oocyte transcription program. The above results highlight the fact that both TRF2 and TRF3 have evolved to play essential roles in the male and female germ lines where they are required to direct specific gene expression programs and/or play a role in chromatin organization.

13.1.2.2

Genes Encoding TAF Paralogs

The role of TFIID paralogs in germ cell transcription is not limited to TRF2 and TRF3. The mouse and human genomes encode TAF4b and TAF7L, which are paralogs of the somatic TAF4 and TAF7, respectively, and the genomes of old world monkeys, apes, and humans additionally contain a retrotransposed copy of the TAF1 gene that encodes the TAF1L protein with functional properties similar to those of the somatic TAF1 (Wang and Page 2002). Genetic studies in mice show that the TAF4b and TAF7L proteins play distinct but essential functions in spermatogenesis. TAF4 and TAF4b share the conserved CR-I (TAFH) domain (Wang et al. 2007; Wei et al. 2007) and the CR-II domain containing a histone fold motif mediates heterodimerization with TAF12 and incorporation into TFIID (Dikstein et al. 1996; Gangloff et al. 2000; Mengus et al. 1997; Thuault et al. 2002). Electron microscopy studies suggest that TAF4b-containing TFIID may have a different conformation from that containing TAF4 and hence a specialized function (Liu et al. 2008). TAF4b was first identified as a lymphoid B-cell enriched factor (Dikstein et al. 1996). Studies of expression in mouse, however, have shown that it is widely

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expressed in many tissues, and its mRNA is strongly enriched in ovary and testis (Freiman et al. 2001). In females, the TAF4b protein is present in the granulosa cells, and in males, in spermatogonia and premeiotic spermatocytes (Falender et al. 2005; Freiman et al. 2001). The TAF4b protein is not, however, specific to the germ line or gonads as it is found in a wide variety of tissues and cell lines. Genetic studies in mice show that TAF4b is required for both male and female fertility (Falender et al. 2005; Freiman et al. 2001). In females, TAF4b controls the granulosa-cell-specific expression of the proto-oncogene c-jun, and these proteins regulate transcription of ovary-selective promoters (Geles et al. 2006). Loss of TAF4b leads to the deregulation of genes of the inhibin-activin signaling pathway and altered cellular morphologies and interactions during follicle growth that likely contribute to the female infertility observed in TAF4b-null female mice. Young TAF4b-mutant male animals are fertile, but with age, the animals become progressively sterile due to a cell autonomous effect in the germ cell population. TAF4b is not required for embryonic gonad differentiation, but shortly after birth, spermatogonial proliferation is impaired, leading to a loss of the germ cell population (Falender et al. 2005). This phenotype may be explained by the diminished expression of genes involved in spermatogonia stem cell proliferation, such as the tyrosine kinase c-Ret, involved in GDNF signaling and the zinc finger transcription factor PLZF. Hence, while the expression of TAF4b is not germ cellspecific, it is critical for the normal expression of genes involved in maintaining spermatogonia proliferation and thus adult spermatogenesis. It is important to note that unlike TRF2, loss of TAF4b does not lead to arrested spermatogenesis, but to more subtle reductions in target gene expression and a progressive infertility. TAF7L is a protein with high sequence similarity to somatically expressed TAF7 that is encoded on the X chromosome and specifically expressed in male germ cells. (Wang et al. 2001). TAF7L is the ancestral intron-containing gene, while the somatic TAF7 is encoded by a retrotransposed autosomal copy (Zhou and Chiang 2001). The TAF7L protein is expressed in spermatogonia and in early primary spermatocytes where it is localized in the cytoplasm (Pointud et al. 2003). From mid-pachytene stage onwards, TAF7L is imported into the nucleus and accumulates strongly in postmeiotic round spermatids. The import of TAF7L into the nucleus is coordinated both with a loss of TAF7 expression, so that in round spermatids only TAF7L is expressed, and a potent upregulation of TBP. In addition, TAF4, which is essential for TFIID assembly and stability, is also strongly downregulated in haploid round spermatids. These observations are remarkable in two respects. Firstly, in pachytene cells, the sex chromosomes are heterochromatinized and physically separated from the autosomes in the XY body (Wang 2004). This state persists for up to 6 days in mouse and 15 days in human. A widely accepted idea is that to bypass the silencing of X-linked genes that are essential at this stage, retrotransposed autosomal expressed copies have been selected encoding functionally equivalent proteins. In contrast, TAFL7 is testis-specific, while the retrotransposed TAF7 is widely expressed. Also, despite the sex chromosome-linked repression of TAF7L in

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pachytene cells, the protein level is in fact upregulated as it is imported into the nucleus, suggesting additional control at the translation step (Pointud et al. 2003). Secondly, although TAF7L and TBP are upregulated, TAF4 that is essential for assembly and stability of TFIID (Wright et al. 2006) is strongly downregulated, suggesting that little fully assembled TFIID exists in these cells. The TBP and TAF7L in haploid cells may rather be present in a subcomplex along with TAF1. Further evidence for this idea comes from the observation that a retrotransposed copy of TAF1 is present in the genomes of old world monkeys, apes, and humans (Wang and Page 2002). The retrotransposed TAF1L is specifically expressed in human germ cells and appears fully functional as it can complement a temperaturesensitive mutation of TAF1 in a hamster cell line and interacts with TBP. It is therefore likely that primates evolved a retrotransposed copy of TAF1 to compensate the loss of its expression due to prolonged X chromosome inactivation during meiosis. TBP and TAF7L both interact with TAF1 and presumably with TAF1L. These observations lend weight to the idea that in contrast to premeiotic cells containing fully assembled TFIID, haploid cells contain a TFIID subcomplex of TBP, TAF7L, and TAF1, and in the case of human germ cells TAF1L, which is required for transcription in these cells (Fig. 13.1). To test this idea, TAF7L has been inactivated in mice by homologous recombination (Cheng et al. 2007). Loss of TAF7L does not lead to arrested spermatogenesis but to a strongly reduced sperm count and reduced fertility. Sperm from mutant animals are characterized by abnormal morphology, reduced motility, and folded tails. Transcriptome analysis of wild-type and TAF7L-mutant testis, however, identified only six transcripts with significantly reduced expression, including FSCN1 involved in actin bundling that may be required for normal sperm morphology and motility. These relatively minor changes in gene expression are nevertheless unlikely to fully account for the observed phenotype. Although further transcriptome studies on purified germ cell populations may perhaps reveal more profound effects on gene expression, the phenotype of the TAF7L knockout animals can also be interpreted in the context of the “promiscuous” transcription state of haploid cells that express elevated levels of many mRNAs and generally higher overall mRNA levels (Schmidt and Schibler 1995). TBP mRNA levels are almost 100-fold higher in haploid cells compared to somatic tissues, principally due to use of alternate promoters (Schmidt et al. 1997; Schmidt and Schibler 1997). TBP protein levels are eight- to ten-fold higher in testis than in somatic cells. Similarly, TFIIB, RNA polymerase II, TFIIA, and ALF are also strongly expressed in haploid cells (Schmidt and Schibler 1995). The overexpression of these critical components of the general transcription machinery may be required to produce the large amounts of some mRNAs (protamines, transition proteins, etc) that have to be synthesized and stored for translation during the elongation and remodeling phase and for lower transcription of a wide but not ubiquitous set of genes (Schmidt and Schibler 1995). It has been suggested that this promiscuous expression may facilitate DNA repair of the transcribed genes before transmission to the next generation, since transcription and DNA repair are coupled (Citterio et al. 2000; Laine and Egly 2006).

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Haploid cells may therefore express germ cell-specific paralogs of the general factors for two distinct purposes, to orchestrate the expression of a specific gene expression program and at the same time to ensure sufficient levels of basal factors to promote generally elevated levels of transcription. Hence, rather than regulating expression of specific genes in haploid cells, TAF7L may be required to ensure generally elevated transcription. This interpretation would explain the paucity of specific changes on the Affymetrix array and account for a phentotype with a generalized reduction in spermatogenesis rather than arrest at a specific stage. Together, the above studies describe the importance of paralogs of the TFIID complex subunits in male fertility. Further mouse knockouts will be required to determine the role of ALF, the TFIIA paralog in male germ cells and additional paralogs such as TCEA2, a paralog of the initiation and elongation factor TFIIS (Guglielmi et al. 2007; Kim et al. 2007; Sikorski and Buratowski 2009) that is selectively expressed at the late pachytene and round spermatid stages (Ito et al. 1996; Umehara et al. 1997), have been identified and their functions remain to be addressed.

13.1.3 CREM, ACT, and KIF17b Cooperate in an Integrated Regulatory Mechanism Required for Spermatogenesis in Mouse and Humans Cyclic AMP response element (CRE) binding protein (CREB) and cyclic AMP response element modulator (CREM) are highly related proteins that regulate transcription in response to various stress, metabolic, and developmental signals (Hummler et al. 1994; Sassone-Corsi 1995). These two proteins bind to the consensus palindromic CRE 50 -TGACGTCA-30 or half-CRE 50 -TGACG-30 and 50 -CGTCA-30 elements present in the promoters of target genes. The CREM locus encodes multiple repressor or activator protein isoforms (Foulkes and Sassone-Corsi 1992). In male germ cells of prepubertal animals, low expression of the repressor isoforms is observed. At puberty, follicle stimulating hormone (FSH) acts via the Sertoli cells to modulate the usage of alternative polyadenylation sites such that several destabilizer signals in the 30 untranslated region of the CREMt activator isoform mRNA are eliminated, leading to increased stability and the accumulation of the CREMt protein to high levels in postmeiotic round spermatids (Foulkes et al. 1992; Foulkes et al. 1993). The precise molecular mechanism by which Sertoli cells signal to the spermatids and regulate the alternative polyadenylation site usage remains to be determined. CREM knockout leads to early apoptosis of haploid cells, around step 4 of their development, and male sterility showing that CREM plays an essential role in spermiogenesis (Blendy et al. 1996; Nantel et al. 1996; Nantel and Sassone-Corsi 1996). Among the direct CREM targets are the genes encoding protamines, transition proteins, and outer dense fiber protein, whose expression initiates in round

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spermatids but whose gene products are required only later during the elongation stage. Diminished expression of these genes may account for the reduced sperm count and abnormalities seen in heterozygous CREM mutant mice but do not readily explain the apoptosis. ChIP from adult testis coupled to high throughput sequencing shows that CREM occupies the promoters of more than 6,000 target genes in haploid cells (Martianov et al. 2010 in press). Comparison with transcriptome data shows that only a small subset of these genes are deregulated following CREM inactivation. The deregulated genes include the antiapoptotic factor Bcl6b and genes involved in signal transduction and metabolic processes (Beissbarth et al. 2003; Martianov et al. 2009). It is rather the deregulation of these genes that accounts for the apoptotic phenotype seen upon CREM inactivation. The role of CREM in human fertility has also been addressed. Like in mouse and rats, human (h) CREMt is expressed in round spermatids and undergoes the isoform switch from the repressor to activator (Peri et al. 1998; Weinbauer et al. 1998). Expression of the CREMt activator isoform is strongly reduced or undetectable in patients with round spermatid maturation arrest, but it was detectable in patients with other syndromes where spermiogenesis proceeded through the elongation phase. These results suggest, but do not formally demonstrate that the loss of CREM results in maturation arrest in humans. The loss of CREM could also be a consequence of impaired spermatogenesis. FSH secretion and bioactivity in these patients is normal; hence, the cause of loss of CREM expression in these patients remains to be determined. There have also been attempts to correlate mutations in the human CREM locus with round spermatid arrest (Vouk et al. 2005). Several SNPs and insertions have been found in both the exons and promoter regions. However, clear evidence linking these changes to infertility has not yet been definitive obtained. In somatic tissues, a variety of stimuli and several different kinases converge to phosphorylate a serine residue (S133 or S117) in the activation domains of CREB and CREM, respectively (Johannessen et al. 2004). Serine phosphorylation results in recruitment of the p300 and CREB binding protein (CBP) coactivators (Chrivia et al. 1993; Nakajima et al. 1997). Alternatively, CREB can also activate transcription of a subset of target genes through a phosphorylation-independent interaction with the TORC (transducers of regulated CREB activity) family of coactivators via the CREB bZIP domain (Conkright et al. 2003; Xu et al. 2007). In male germ cells, CREM bypasses the requirement for phosphorylation through interaction with the LIM-only domain protein ACT (activator of CREM in testis) encoded by the Fhl5 gene (Fimia et al. 1999) specifically expressed in haploid cells. ACT functions as a coactivator in yeast and in mammalian cells where it stimulates CREM transcriptional activity. Inactivation of ACT in mice does not phenocopy loss of CREM as there is no block in spermatogenesis and males are fertile (Kotaja et al. 2004). ACT knockout does, however, lead to a severe reduction in the number of mature sperms and the appearance of frequent malformations of the sperm head and flagellum. These observations suggest only a subset of CREM target genes are regulated using ACT as a coactivator or that ACT is required for a general fine-tuning of CREM target transcript levels.

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To evaluate the role of ACT in human infertility, its expression and variant alleles have been investigated in infertile patients. In one study of 96 azoo- or oligospermic patients, 12 SNPs in the Flh5 gene were described, all of which were present in both the infertile patients and the fertile controls, suggesting that they do not individually contribute to the pathogenesis of male infertility (Christensen et al. 2006). However, the frequency of the nucleotide changes, and the distribution of some of the associated haplotypes did show significant differences between the patients and controls. In a second, more limited study, ACT expression was found to be reduced in three of four patients with round spermatid maturation arrest (Steger et al. 2004). The activity of the CREM-ACT pathway is controlled by KIF17b, a testisspecific form of the kinesin protein KIF17, which interacts with ACT (Macho et al. 2002). KIF17b colocalizes with ACT in the nucleus of haploid cells, but at the onset of the elongation phase, KIF17b and ACT cotranslocate to the cytoplasm. KIF17b also transports a set of CREM-regulated mRNAs (Chennathukuzhi et al. 2003) and interacts with the MIWI component of the chromatoid body implicated in RNA metabolism (Kotaja et al. 2006; Kotaja and Sassone-Corsi 2007). This interaction may play a role in the transportation of mRNAs to the chromatoid body. Thus, KIF17b links the processes of transcription and transport of specific mRNAs in male germ cells. Intracellular translocation of KIF17b is mediated by a microtubule-independent mechanism that requires neither the motor domain nor microtubules (Kotaja et al. 2005) and is modulated by the cyclic AMP-dependent protein kinase A (PKA) that phosphorylates KIF17b to regulate its subcellular localization. Male germ cells therefore utilize a unique and integrated mechanism by which CREM, ACT, and KIF17b act together to regulate transcription in developing male germ cells (summarized in Fig. 13.2).

13.2

What Have Mouse Knockout Studies Told Us About the Physiopathology of Human Infertility?

Approximately 15% of couples are infertile, and amongst these couples, male infertility accounts for approximately 50% of causes. The cause of a majority of the cases can be traced to hypothalamic or pituitary failure and defective hormonal signaling. The current estimate is that about 30% of infertile men show oligozoospermia or azoospermia despite unimpaired reproductive hormone secretion and lack of other known determinants of infertility. These patients are categorized as suffering from idiopathic infertility, whose causes often still remain elusive. The mouse genetic models have defined a large number of proteins including the transcription factors described here that could potentially be involved in human male infertility (Yan 2009). Among the factors discussed here, the best evidence for a role in human infertility is described for CREM, where changes in its expression and SNPs at the CREM locus can be correlated with round spermatid arrest.

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In contrast, while the spermatogenic functions of TAF7L, TAF4b, and HFNT1 have been defined in mouse, little information is available concerning their expression or potential roles in human spermatogenesis. Assuming a conservation of their functions, the phenotypes observed upon inactivation of these proteins, characterized by quantitative and/or qualitative reductions in sperm count and quality, may be more relevant to some human subfertility or infertility conditions than a complete arrest of spermatogenesis. Although no studies have so far implicated these genes in human infertility, the above model emphasizes the idea that proper expression levels of these transcription factors and their target genes are essential for normal fertility. Any condition that impacts their expression or function, for example, SNPs in the coding or regulatory regions or hormonal influences, could lead to diminished fertility. Future studies will be required to establish whether these genes are involved in the physiopathology of human fertility. Acknowledgments Work in the Davidson laboratory is supported by grants from the CNRS, the INSERM, the Association pour la Recherche contre le Cancer, the Ligue Nationale et De´partementale Re´gion Alsace contre le Cancer, the INCA, the ANR Regulome project grant, and the EuTRACC program of the European union. ID is an e´quipe labe´llise´e of the Ligue Nationale contre le Cancer.

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Chapter 14

The Chromatoid Body: A Specialized RNA Granule of Male Germ Cells Ippei Nagamori, Adam Cruickshank, and Paolo Sassone-Corsi

Abstract RNA processing and miRNA pathways have been shown to exert a central control on a wide variety of cellular functions. The epigenetic program of male germ cells is highly specialized, including RNA processing pathways, which play an important role in the male germ cell lineage. Male germ cells differentiate through a remarkable process of cellular restructuring. One highly specialized structure is the chromatoid body (CB), a male reproductive cell-specific organelle that appears in spermatocytes and spermatids. It is a finely filamentous, lobulated perinuclear granule located in the cytoplasm of male germ cells. The molecular composition and function of the chromatoid body have remained elusive for a longtime. Accumulating evidence indicates that the CB is involved in RNA storing and metabolism, being related to the RNA processing body (P-body) of somatic cells. We propose that the CB operates as an intracellular nerve-center of the microRNA pathway. The role of the chromatoid body underscores the importance of posttranscriptional gene regulation and of the microRNA pathway in the control of postmeiotic male germ cell differentiation. Keywords Chromatoid body Epigenetics micro RNA Post-transcriptional regulation Spermatogenesis VASA

14.1

Introduction

Spermatogenesis displays unique features that distinguish it from all other cellular differentiation programs (Ewing et al. 1980; Kimmins and Sassone-Corsi 2005; Oakberg 1956; Seydoux and Braun 2006). These include: (a) Meiosis and the generation of haploid cells; (b) Direct hormonal control, via the hypothalamicpituitary axis; (c) Persistence in the adult organism, with a cyclic, ongoing

I. Nagamori, A. Cruickshank, and P. Sassone Corsi (*) Department of Pharmacology, School of Medicine, University of California, Irvine, CA 92697, USA e mail: [email protected]

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production of differentiating germ cells. These remarkable features delineate the necessity for specialized strategies to control gene expression (Eddy and O’Brien 1998; Hecht 1998; Sassone-Corsi 2002). Most importantly, since germ cells are able to give rise to totipotent diploid zygotes, it is crucial that their genome is errorfree. Thus, it is not surprising that these cells have evolved a specialized transcription machinery to ensure the accurate transmission of the genome and for the activation of the enormous number of genes that are required in embryogenesis. Germ cells have developed highly specialized transcriptional mechanisms that ensure stringent stage-specific gene expression (Kimmins and Sassone-Corsi 2005; Braun 1998; Kleene 1993; Sassone-Corsi 1997). These transcriptional onsets are intimately coupled to prominent posttranscriptional events, including alternative splicing, alternative polyadenylation, and translational regulation specific for the male germ cells. Posttranscriptional control of mRNA translation appears especially significant toward the end of spermatogenesis, at a time when it is generally believed that global transcription ceases (Sassone-Corsi 2002).

14.1.1 RNA Processing in Male Germ Cells There are several levels of control of gene expression that are implemented posttranscriptionally. These include a variety of RNA processing pathways devoted in regulating the abundance levels of specific transcripts. Increasing evidence underscores the importance played by these posttranscriptional events in germ cells that are transcriptionally silent. In the highly transcriptionally active spermatocytes, transcripts accumulate at extraordinary levels. Consequently, mRNA storage and translational activation are key steps in controlling the synthesis of many spermatid and spermatozoa proteins, which are produced in late stages of germ cell maturation (Kimmins and Sassone-Corsi 2005; Kleene 1993). The length of the poly(A) tail of mRNA expressed in the testis changes during spermiogenesis, indicating that the tail length may affect translation of mRNAs (Foulkes et al. 1993). In accordance with this notion, the deletion of the testis-specific, cytoplasmic poly(A) polymerase TPAP in TPAP-deficient mice impairs the expression of haploid-specific genes and causes the incomplete elongation of the poly(A) tails of particular transcription factor mRNAs (Kashiwabara et al. 2002). Many mRNAs, such as protamines and transition protein transcripts, are translationally repressed in early spermatids with long poly(A) tracts and are sequestered in cytoplasmic ribonucleoprotein particles for up to a week. Translation subsequently takes place in late spermatids after mRNAs undergo a poly(A) shortening by deadenylation (Lee et al. 1995). There are several RNA-binding proteins, such as testis-brain RNA-binding protein (TB-RBP/transilin), Protamine-1 RNA-binding protein (Prbp), MSY2 and 4, a spermatid perinuclear RNA-binding protein (Spnr), poly(A)-binding protein 2 (PABP2), and Sam 68, which appear to be critical in the regulation of translation in male germ cells (Braun 1998; Paronetto et al. 2009). MSY2 has been suggested to mark specific mRNAs in the nucleus for cytoplasmic storage by binding to the Y-box motifs, thereby linking transcription and mRNA storage in meiotic and

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postmeiotic male germ cells (Yang et al. 2005a). In mice lacking msy2, spermatogenesis is blocked in postmeiotic germ cells, and oogenesis is also disrupted (Yang et al. 2005b). Deletion of prbp gene in the mouse leads to male sterility due to disturbed translational activation of the mRNAs encoding the protamines (Zhong et al. 1999). Interestingly, a human Prbp homolog, TRBP, interacts with the endonuclease Dicer that is a crucial component of RNA interference (RNAi) and microRNA (miRNA) pathways and is required for optimal RNA silencing mediated by small interfering RNAs (siRNAs) and endogenous miRNAs (Haase et al. 2005). These results indicated that translational regulation in male germ cells involves siRNA and miRNA pathways. An important aspect of these studies is how specificity may be achieved. An interesting example is provided by TB-RBP/translin and translin-associated factor X (TRAX), which bind to specific mRNAs, such as the mRNAs of protamines. It has been proposed that it can be subsequently transported from the nucleus and across intercellular bridges between haploid spermatids (Chennathukuzhi et al. 2003b; Morales et al. 2002). TB-RBP/translin complexed with specific mRNAs, including CREM target RNAs, is transported by the kinesin KIF17b through nuclear pores to the cytoplasm for storage (Chennathukuzhi et al. 2003a). Intriguingly, the KIF17b kinesin was originally described as implicated in the regulation of CREM-ACT-mediated transcription (Macho et al. 2002) and subsequently suggested that it could link transcriptional regulation and mRNA transport in testis (Chennathukuzhi et al. 2003a; Kotaja et al. 2005). While the molecular and physiological implications of these findings are not fully understood, they seem to provide an interesting lead to further investigations.

14.1.2 A Highly Specialized RNA Granule The central role of cytoplasmic RNA granules in germ cells (polar and germinal granules), somatic cells (stress granules and processing bodies), and neurons (neuronal granules) has emerged in the posttranscriptional regulation of gene expression. RNA granules contain various ribosomal subunits, translation factors, decay enzymes, helicases, scaffold proteins, and RNA-binding proteins, and they have been implicated in the control of localization, stability, and translation of their RNA cargo (Anderson and Kedersha 2006). In postmeiotic germ cells, a cytoplasmic organelle has been the focus of various exciting studies. The chromatoid body (CB) appears to occupy a strategic position within the regulatory pathways that control the spermatogenic process via posttranscriptional mechanisms (Kotaja and Sassone-Corsi 2007). Germ cells of various organisms are characterized by the accumulation of dense fibrous material into a cytoplasmic structure, called germplasm or nuage. In Drosophila, Caenorhabditis elegans, and Xenopus, the germplasm has been shown to be responsible for the early specification of the germline lineage (Liang et al. 1994; Parvinen 2005; Raz 2000; Styhler et al. 1998). In mammals, cytoplasmic factors do not seem to have a

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Fig. 14.1 The chromatoid body appears as a cloud like, dense structure in the cytoplasm of male germ cells. Its perinuclear localization is evident in this differential interference contrast image

DIC

MVH/DNA

direct role in germ cell specification, but germ cells are induced through cellular interactions during gastrulation. No germplasm-like structure is detectable in unfertilized and fertilized eggs in mice. However, on the basis of its structural features and protein composition, the so-called CB in the cytoplasm of haploid male germ cells is believed to be the mammalian counterpart of nuage (Parvinen 2005). The CB appears to be composed of thin filaments that are consolidated into a compact mass or into dense strands of varying thickness that branch to form an irregular network (Fig. 14.1). The dynamic evolution of the CB during spermatogenesis and its intriguing intracytoplasmic movements point to a potential function in RNA-processing pathways (Kotaja and Sassone-Corsi 2007). This hypothesis is supported by recent findings showing that the CB in postmeiotic spermatids is composed of elements of the microRNA (miRNA) and RNA decay pathways (Kotaja et al. 2006a). Thus, despite that, the existence of the CB has been acknowledged for several decades; only now, its function in male germ cell differentiation begins to be revealed.

14.1.3

Unique Features of the CB

A number of studies have shown that the CB forms dynamically along the germ cell differentiation program, changing its size, density, and location. The CB appears for the first time in the cytoplasm of mouse meiotic pachytene spermatocytes as a fibrous-granular structure in the interstices of mitochondria clusters (Fujiwara et al. 1994). Postmeiotically, the CB condenses to one single finely filamentous, lobulated perinuclear granule in round spermatids, which stays as a distinctive feature in the cytoplasm of postmeiotic spermatids until the nucleus starts to elongate (Parvinen 2005). The varying localization of the CB during successive stages of spermatogenesis is one of its primary features. It appears at the nuclear envelope in association with nuclear pore complexes during early spermiogenesis and associates with the annulus later in spermiogenesis (Parvinen 2005; Fawcett et al. 1970). Intriguingly, the CB moves around the Golgi complex and has frequent contacts with it. The CB also moves perpendicularly to the nuclear envelope, and some

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reports even indicate that it may transfer through cytoplasmic bridges to the neighboring spermatids (Parvinen 2005). Recent advances have provided important clues on the molecular composition of the CB. One component, however, has been well-characterized and has become a hallmark of germ cells: this is the ATP-dependent DEAD-box RNA helicase, VASA (in the mouse MVH, for Mouse VASA Homolog), present in both Drosophila nuage and mouse CB (Styhler et al. 1998; Fujiwara et al. 1994; Toyooka et al. 2000). Importantly, VASA and MVH are present exclusively in the nuage and CB, respectively. In Drosophila, vasa is required for the formation of germ cells and oogenesis, and VASA protein was found to play a central role in translational regulation of many important mRNAs (Fujiwara et al. 1994). Notably, targeted mutation of the mvh gene in the mouse yields a drastic interruption of spermatogenesis by zygotene stage (Tanaka et al. 2000). This finding demonstrates that MVH and thereby the CB are essential for a normal progression of the differentiation program of male germ cells. The ATP-dependent RNA helicase function of MVH strongly suggests a direct involvement in RNA processing, although its specific mode of action is still elusive. Its critical presence in the CB is also indicating that, through a number of potential interactions with various RNAbinding proteins and RNA as well, its function may be controlled by a variety of regulatory steps, including posttranslational modifications (Saunders et al. 1992; Soderstrom et al. 1976).

14.1.4 Components of the Chromatoid Body In an attempt to elucidate the function and dynamic regulation of the CB, a number of researchers have embarked in analyses aimed at identifying CB components. While a systematic proteomic analysis has not been performed yet, several components, as well as their interactions, have been identified. Specifically, recent studies provide an attractive experimental gateway for the understanding of the physiological role played by this organelle. Indeed, it has been recently shown that miRNA pathway components, such a miRNA-processing endonuclease Dicer, Argonaute (Ago) proteins, and miRNAs are concentrated in CB (Kotaja and Sassone-Corsi 2007; Kotaja et al. 2006a). Additionally, the components of RNA processing bodies (P-bodies) accumulate in CBs, indicating that the germ cell-specific CBs and somatic P-bodies are functionally related (Kotaja et al. 2006a). As P-bodies in somatic cells are thought to integrate RNA decay and RNA regulation by miRNAs, we have proposed that CBs function as platforms for miRNA pathway (Kotaja and Sassone-Corsi 2007; Parvinen et al. 2007). These notions strengthen the hypothesis of RNA interference and miRNA pathways being mechanisms essential for gene regulation in spermatogenesis. These findings questioned whether components of the RISC complex accumulate in the CB to exert a specialized function in the mRNA maturation pathway during spermatogenesis. Importantly, a hallmark of spermatogenesis is the massive

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wave of transcription that occurs postmeiotically (Eddy and O’Brien 1998; SassoneCorsi 2002). This involves a large number of genes, which encode products with specialized functions during the developmental process that leads to the shaping of mature sperm cells (spermiogenesis). Intriguingly, many postmeiotic transcripts accumulate in significant amounts and are stored in the cytoplasm of spermatids for 4 5 days (Sassone-Corsi 2002; Braun 1998; Kleene 1993). These regulatory events lead to translational repression, a phenomenon whose regulatory mechanisms and physiological control remain elusive. Interestingly, polyA-containing mRNAs accumulate in the CB of spermatids and have suggested that the process of translational repression is organized within this cytoplasmic structure (Kotaja et al. 2006a) This scenario is further supported by the coordinate presence in the CB of proteins that are essential for the effector phase of mRNA silencing (Kotaja et al. 2006a). Moreover, it is notable that mutation of the Drosophila melanogaster nuage component Maelstrom (MAEL) causes mislocalization of the miRNA pathway proteins Dicer and Ago2 (Costa et al. 2006). Finally, the TUDOR-related protein Tdrd1/Mtr-1 also localizes in the CB and is essential for male germ cell differentiation (Chuma et al. 2006). Similarly, Tdrd6 and Tdrd7 are also localized in the CB (Hosokawa et al. 2007; Table 14.1). Proteins with TUDOR domains are often able to bind RNA and are implicated in RNA metabolism. MTR-1 co-localizes with Sm proteins, which are components of snRNPs in the CBs located in the perinuclear region (Biggiogera et al. 1990; Chuma et al. 2003; Moussa et al. 1994), suggesting that MTR-1 functions in assembling snRNP into cytoplasmic granules including the CBs in germ cells. Importantly, Tudor is genetically downstream of vasa and encodes a component of the Drosophila “nuage” (Bardsley et al. 1993; Golumbeski et al. 1991). In this respect, the recent finding that Tdrd6, another protein with multiple TUDOR domains, is essential for CB architecture and regulation of miRNA expression is highly interesting. Importantly, mice lacking Tdrd6 display an interrupted spermatogenesis and a misplaced localization of MIWI, MAEL, and MVH in the CB (Vasileva et al. 2009).

14.1.5

The Critical Roles of MIWI and PIWI

The mouse PIWI family proteins (MILI, MIWI, and MIWI2) play pivotal roles in spermatogenesis through transcriptional and posttranscriptional gene regulation. MIWI, an RNA-binding protein, belongs to the Argonaute/PIWI family proteins that share PAZ and PIWI domains and are classified into two subfamilies depending on sequence similarity to either Arabidopsis thaliana Ago1 or D. melanogaster PIWI (Carmell et al. 2002; Sasaki et al. 2003). In mammals, the Argonaute1 subfamily members, Ago1 to Ago4 are widely expressed in many tissues and are essential components of RISC complexes in the RNAi and miRNA pathways (Carmell et al. 2002; Sasaki et al. 2003). By contrast, all four members of the PIWI subfamily are expressed mainly in the testis, and three of them, MIWI, MIWI2, and MILI, are required for the progress of spermatogenesis in mouse

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Table 14.1 Components of the mammalian chromatoid body Component Function MVH Dead box RNA helicase that is required for spermatogenesis MIWI

RanBPM

Kif17b

Tdrd1

Tdrd6

Tdrd7 p48 and p52 Ago2 and Ago3 Dicer

Dcp1a

GW182

RNA miRNA

mRNA snRNP

GRTH

Argonaute/PIWI family RNA binding protein, which is required for spermatogenesis. MIWI and MILI repress transposon element mediated by piRNA RanGTP binding protein that is involved in microtubule nucleation. It associates with MVH Kinesin motor protein that is involved in the transport of the co activator protein ACT and mRNAs Tudor domain containing protein that is required for spermatogenesis. It interacts with MILI Tudor domain containing protein that is required for spermatogenesis and CB formation Tudor domain containing protein Germ cell specific RNA binding proteins Main catalytic engine of RISC Cytoplasmic RNase that produces siRNA and miRNA from their precursors P body localized decapping enzyme that remove 50 cap of mRNA P body localized RNA binding protein that is required for RNAi Small RNA that represses complement target mRNA expression Translation template Essential component of the spliceosomal complex that function in pre mRNA processing Dead box RNA helicase that is required for spermatogenesis. Deficient of GRTH shows impaired CB formation

317

Ref Chuma et al. (2003), Kotaja et al. (2006a, b), Kotaja and Sassone Corsi (2007) Kotaja et al. (2006a, b), Deng and Lin (2002), Grivna et al. (2006b)

Shibata et al. (2004)

Kotaja et al. (2006b)

Kojima et al. (2009), Wang et al. (2009), Hosokawa et al. (2007) Vasileva et al. (2009)

Hosokawa et al. (2007) Olsen et al. (1997) Kotaja et al. (2006a) Kotaja et al. (2006a)

Kotaja et al. (2006a)

Kotaja et al. (2006a)

Kotaja et al. (2006a) Kotaja et al. (2006a)

Kotaja et al. (2006a), Saunders et al. (1992) Biggiogera et al. (1990), Moussa et al. (1994), Paniagua et al. (1985) Tsai Morris et al. (2004)

(continued)

318 Table 14.1 (continued) Component H2B H4 Acetylated H3 Acetylated H4 MTR1

Actin Vimentin Ubiquitin E2 p52 pA700 Cytochrome C COX1 HSP70 F1alpha F1beta LDH Enolase

PHGPx Ca2þ, Mg2þ

I. Nagamori et al.

Function Chromatin formation Chromatin formation Chromatin regulation Chromatin regulation Tudor domain containing protein that is involved in the assembly of snRNPs Cytoskeletal protein Intermediate filament Targeting signal for several pathways Ubiquitin conjugating enzyme 26S proteasome subunit Proteasome activator Electron transfer component Cytochrome C oxidase subunit 1 Protein chaperon ATP synthase subunit alpha ATP synthase subunit beta Dehydrogenation of 2 hydroxybutylate Conversion from 2 phosphogylcerate to phosphoenoylpyruvate Reduces lipid hydroperoxides in membranes

Histocompatibility antigen LAMP1 and LAM2 Acid phosphatase Cathepsins B,D, H, and L LAP DNase RNase NADPase

Immune system Lysosome localized glycoproteins Lysosome localized phosphatase Protease Leucine aminopeptidase Hydrolysis of DNA Hydrolysis of RNA Hydrolysis of NADP

CMPase

Hydrolysis of monophosphate

Polysaccharides

Ref Haraguchi et al. (2005) Werner and Werner (1995) Haraguchi et al. (2005) Haraguchi et al. (2005) Chuma et al. (2003)

Haraguchi et al. (2005), Walt and Armbruster (1984) Haraguchi et al. (2005) Haraguchi et al. (2005) Haraguchi et al. (2005) Haraguchi et al. (2005) Haraguchi et al. (2005) Hess et al. (1993) Haraguchi et al. (2005) Haraguchi et al. (2005) Haraguchi et al. (2005) Haraguchi et al. (2005) Haraguchi et al. (2005) Haraguchi et al. (2005)

Haraguchi et al. (2005) Andonov and Chaldakov (1989), Rouelle Rossier et al. (1993) Head and Kresge (1985) Haraguchi et al. (2005) Anton (1983) Haraguchi et al. (2005) Haraguchi et al. (2005) Haraguchi et al. (2005) Haraguchi et al. (2005) Tang et al. (1982), Thorne Tjomsland et al. (1988) Tang et al. (1982), Thorne Tjomsland et al. (1988) Krimer and Esponda (1980)

(Sasaki et al. 2003; Carmell et al. 2007; Deng and Lin 2002; Kuramochi-Miyagawa et al. 2004). Although all the Argonaute/PIWI family members bind to small RNAs through their conserved PIWI domains, there are differences in their activities, for example, in their abilities to possess a siRNA-dependent mRNA cleavage, or slicer, activity. These findings indicate that the Argonaute/PIWI proteins could have

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different functions in diverse, dynamically forming complexes (Liu et al. 2004; Meister et al. 2004; O’Donnell and Boeke 2007). A role for MIWI in the structure and function of the CB has been proposed (Kotaja et al. 2006b). MIWI is found in ribonucleoprotein complexes (RNP) in male germ cells (Deng and Lin 2002) and colocalizes with the other Argonaute/ PIWI family members, Ago2 and Ago3, in the CB (Kotaja et al. 2006a). We have therefore speculated that MIWI might operate as a male germ cell-specific component of the machinery that is involved in mRNA regulation in the testis (Kotaja et al. 2006b). Whether MIWI and Ago proteins act on the same or functionally distinct pathways is not fully understood yet. An important discovery is related to the finding of a novel class of small RNAs. These are PIWI-interacting RNAs (piRNAs) and are germline-specific. Compared to miRNAs, piRNAs are slightly longer and interact with the mammalian PIWI family members MIWI and MILI but not with Ago proteins (Aravin et al. 2006; Girard et al. 2006; Grivna et al. 2006a, b; Lau et al. 2006; Watanabe et al. 2006). The function of piRNAs is still unclear, and it remains to be explored whether they act together with PIWI family members in the germ cell-specific regulation of translation and whether they colocalize with MIWI in the CB. Interestingly, MIWI was demonstrated to interact with piRNAs and mRNAs both in polysomes and in RNPs, suggesting a possible involvement of MIWI and piRNAs in the control of translation and/or mRNA stability (Grivna et al. 2006b). However, the unchanged global translation profile in testes from miwi-null mice indicates that MIWI might play a very specific role in translational regulation (Grivna et al. 2006b). The mammalian Piwi protein MILI is also localized in the cytoplasm of spermatogenic cells and is thought to be associated with the CB as well as with polysomes (Unhavaithaya et al. 2009). In a search for MILI interacting proteins, the association with the Tudor domain-containing protein 1 (Tdrd1) was shown (Chen et al. 2009; Kojima et al. 2009; Reuter et al. 2009; Vagin et al. 2009; Wang et al. 2009). Tdrd1 is a germline protein with multiple Tudor domains that when mutated in the mouse shows a defective spermatogenic phenotype similar to the one described for MILI (Wang et al. 2009). This control may implicate regulatory feedback loops since MILI positively regulates the expression of the Tdrd1 mRNA. Differently from MILI, Tdrd1 is not required for the biogenesis of piRNAs, indicating that its critical role in spermatogenesis may be related to a yet undefined RNA processing pathway. Genetic studies using mice carrying targeted mutations of specific genes have provided critical insights. MIWI is a critical CB component, and the structure of the CB is disrupted in the round spermatids of miwi-null mice (Kotaja et al. 2006b). Lack of the MIWI protein results either in the absence of CBs or in a drastic reduction in the compaction of the CB. It has been suggested that the CB originates from thin fibrous cytoplasmic structures that consolidate into a dense particle in late step 1 spermatids (Parvinen 2005; Fawcett et al. 1970). Electron microscopy analysis has shown that very thin threads of CB material are still present in round spermatids from miwi-null mice, but these filaments do not successfully condense into mature CBs. Despite the problems in condensation, some CB components,

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such as MVH, are present in spermatids of miwi-null mice in the cytoplasmic area, which is normally occupied by the CB, indicating that they can be recruited to the CB independently of MIWI (Kotaja et al. 2006b). The functional role and molecular mechanisms of MIWI and MIWI-related proteins in the formation of the CB are currently unclear. Importantly, MIWI2 is directly involved in spermatogenesis (Carmell et al. 2007) and MILI in transposon control via the piRNAs (Aravin et al. 2007; Kuramochi-Miyagawa et al. 2008). Thus, an attractive possibility is that the CB may be critical in the spatial and temporal organization of the dynamic events that control RNA metabolism in germ cells.

14.1.6

Dynamic Movements of the CB

During the differentiation of male germ cells, the CB changes its size, density, and location. In addition, at the time of full maturity, the CB displays active and seemingly nonrandom movements in the cytoplasm of round spermatids. The CB moves rapidly both in parallel and perpendicular fashion related to the nuclear envelope (Kotaja and Sassone-Corsi 2007; Parvinen and Parvinen 1979; Ventela et al. 2003). Electron microscopy has indeed revealed material continuities between the nucleus and the CB as if the CB had the capacity to selectively collect nuclear material (Soderstrom and Parvinen 1976; Parvinen and Parvinen 1979). This hypothesis is supported by the observation that nuclear pores tend to be more concentrated in the area adjacent to the CB (Fawcett et al. 1970). The CB has been also observed to move through cytoplasmic bridges to neighboring cells as if this organelle would operate mechanistically in the sharing of haploid products between adjacent spermatids (Ventela et al. 2003). Microtubular network is suggested to be involved in the mobility of the CB as microtubule depolymerizing drugs disturb its movement (Ventela et al. 2003). The molecular mechanisms and cellular pathways involved in CB movements within the cell remain undeciphered. A series of intriguing findings point to a possible molecular mechanism that could account for the intracellular CB movements. KIF17b, a testis-specific isoform of the brain kinesin-2 motor KIF17 (Macho et al. 2002), localizes in the CB (Kotaja et al. 2006b). KIF17b was first described as a kinesin able to shuttle between cytoplasmic and nuclear compartments and transports the activator of CREM (cAMP-response element modulator) in the testis (Fimia et al. 1999; Kotaja et al. 2004) from the nucleus to the cytoplasm, therefore regulating CREM-dependent transcription (Macho et al. 2002; Kotaja et al. 2005). In addition to the transport of ACT, KIF17b has been implicated in the transport of RNA. KIF17b binds to RNA protein complexes containing specific CREM-regulated mRNAs through an interaction with the testis brain RNA-binding protein (TB-RBP) and transports these particles between the nucleus and the cytoplasm (Chennathukuzhi et al. 2003a). In this respect, it is tempting to speculate that via the association with different RNA-binding proteins, KIF17b could function as a transporter that carries RNA from the nucleus to the CBs in round spermatids. Indeed, the

The Chromatoid Body: A Specialized RNA Granule of Male Germ Cells

MVH

Merge

Kif17b

Fig. 14.2 Immunostaining of chromatoid bodies showing the presence of MVH, MIWI, and KIF17b (green) in the CB. DAPI stains the nuclei (blue) as reference. The localization of the CB’s components is highly specific within the cytoplasmic perinuclear organelle. Bar, 5 mm

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MIWI

14

microtubule-binding kinesin protein, KIF17b, accumulates in CBs (Figs. 14.2 and 14.3). Interestingly, although KIF17b function is microtubule-independent (Kotaja et al. 2005), its presence in the CB could also offer a possible mechanistic explanation for the active movements of the CB. It is also reasonable to speculate that in addition to RNA transport, KIF17b could also be involved in the transport of specific CB protein components.

14.1.7 Regulation by Critical Interactions The list of protein components of the CB has been increasing, begging the question of how these proteins may be organized within such compact structure (Table 14.1). The reciprocal interactions among various CB components are likely to dictate the physiological functions of these proteins and of the CB itself. In this respect, MVH, the hallmark component of the CB, may be critical. Importantly, it has been shown that MVH physically interacts with Dicer (Kotaja et al. 2006a) and with the Argonaute/PIWI family members (Kuramochi-Miyagawa et al. 2004). Could the DEAD-box RNA helicase MVH be a germ cell-specific element of the RISC complex? Interestingly, turnover of the RISC cleavage activity is dependent on ATP, which indicates that release of the cleaved mRNA halves may involve ATPase activities (Haley and Zamore 2004; Sontheimer 2005). Our hypothesis is that MVH acts as a germ cell-specific regulator for providing

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I. Nagamori et al. Cytoplasm Chromatoid body m7 G

(A)n KIF17b

Dcp1a MIWI/MILI MVH Dicer

Ago2 GW182

Pre-miRNA

Nucleus

miRNA

KIF17b

m 7G

m7G

Ago2

(A)n

KIF17b

(A)n

Fig. 14.3 Model of chromatoid body (CB) function in haploid male germ cells. In addition to the male germ specific CB components MVH and MIWI, CB contains the components of both microRNA pathway, such as Ago proteins and the endonuclease Dicer, and RNA decay pathway, such as the decapping enzyme Dcp1a and GW182 (Kotaja et al. 2006a). CB moves around the nucleus and makes frequent contacts with the nuclear pores, collecting material from the nucleus. MicroRNAs and mRNAs that are synthesized in the nucleus are transported to the cytoplasm and loaded directly to the CB through nuclear pores. We favor a view where the shuttling kinesin protein KIF17b plays an important role in transporting RNA between the nucleus and the CB (Kotaja et al. 2006b). In the CB, mRNAs are directed either to decay or translational repression by an unknown mechanism

helicase/ATPase activity to the RISC complex. On the other hand, DEAD-box RNA helicases have been reported to be involved in remodeling of RNA-protein interactions, suggesting that MVH could possibly function as a regulator of RNP complexes in the CB (Jankowsky and Bowers 2006). As many interactions are also likely to be dynamic, the interaction of KIF17b with MIWI (Kotaja et al. 2006b) is of particular interest. KIF17b and MIWI are both found in RNP complexes with CREM-target mRNAs (Chennathukuzhi et al. 2003a; Deng and Lin 2002). Interestingly, mice deficient for either MIWI or CREM display a similar block in spermatogenesis at the very early stages of spermiogenesis, suggesting that these proteins could function on the same pathway (Deng and Lin 2002; Nantel et al. 1996). Localizations of KIF17b and MIWI overlap both temporally and spatially in postmeiotic round spermatids: MIWI is a cytoplasmic protein that is concentrated in CBs, whereas KIF17b is shuttling between the two compartments, whose presence in the CB is transient. MIWI is not required for KIF17b localization in the CB (Kotaja et al. 2006b); therefore, the interaction

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between these proteins might be required for some other functions. We have speculated that the interaction may be one step of the process of exchange of material between KIF17b and MIWI (Kotaja and Sassone-Corsi 2007). Further studies aimed at revealing the interactions among the various elements composing the CB will be essential to unravel the function of the CB, the physiology of RNA processing in testis, and the molecular pathways that lead to the differentiation of male germ cells.

14.1.8 RNA Granules in Somatic Versus Germ Cells It is now established that a variety of RNA granules exist in all cells (Anderson and Kedersha 2009), a notion that begs the question of what would be the differences and relationships between all these granules, both from a functional and molecular point of view. The presence of the CB and nuage begs the question of whether these organelles are unique to germ cells. Interestingly, somatic cells also contain morphologically and biochemically distinct cytoplasmic bodies with unique functional characteristics. Processing bodies, or P-bodies, have been described in yeast and mammalian cells, capturing significant attention (Eulalio et al. 2007). These bodies concentrate the enzymes that are involved in the RNA decay pathway and accumulate mRNAs targeted to translational repression and degradation (Eulalio et al. 2007; Brengues et al. 2005; Cougot et al. 2004; Sheth and Parker 2003). In P-bodies, which function as concentration sites for Ago proteins, miRNAs, and miRNA-repressed mRNAs, functional miRNA pathways operate (Liu et al. 2005a, b; Pillai et al. 2005; Sen and Blau 2005). It was proposed that mammalian miRNAs inhibit the initiation of translation of mRNAs that subsequently accumulate in P-bodies for storage (Pillai et al. 2005). A functional link between P-bodies and the ability of miRNAs to repress expression of a target mRNA was provided by a study, which showed that silencing of the P-body component GW182 delocalizes P-body proteins and impairs the function of miRNA pathways (Liu et al. 2005a). In a parallel manner, several P-body markers, such as the decapping enzyme Dcp1a and GW182, were shown to be highly concentrated in CBs, therefore providing evidence of a functional analogy with the somatic P-bodies (Kotaja et al. 2006a). One revealing experiment further strengthened this parallel: we have ectopically expressed MVH, which is male germ cell-specific, in fibroblasts and found that its localization is targeted to P-bodies (Kotaja et al. 2006a). Importantly, the absence of ribosomal RNA in the CB (Kotaja et al. 2006a) correlates well with the current hypothesis about the function of P-bodies (Brengues et al. 2005; Coller and Parker 2005; Kedersha et al. 2005; Teixeira et al. 2005). P-bodies are thought to temporarily store a pool of translationally repressed mRNARNP particles. The favored scenario proposes that there is equilibrium with the translating pool, and that translationally repressed RNPs are timely released to be translated into the protein for which there is a physiological need (Eulalio et al. 2007; Rossi 2005). This equilibrium could be altered by miRNAs and associated

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proteins. In accordance, mRNAs have been shown to physically move out of P-bodies and transported to the translation apparatus for protein synthesis (Brengues et al. 2005). The analogies outlined above suggest that the CB could function in a similar manner in male germ cells. Interestingly, the CB moves very actively and appears to release small satellite granules that can be associated with other cellular organelles. This raises the possibility for a transport mechanism that could be utilized to move mRNAs between the pools of repressed and active translation. Future studies will be aimed at addressing these questions and to unravel the signaling pathways that control the functions of CB components in the regulation of RNA processing in germ cells.

References Anderson P, Kedersha N (2006) RNA granules. J Cell Biol 172(6):803 808 Anderson P, Kedersha N (2009) RNA granules: post transcriptional and epigenetic modulators of gene expression. Nat Rev Mol Cell Biol 10(6):430 436 Andonov MD, Chaldakov GN (1989) Morphological evidence for calcium storage in the chroma toid body of rat spermatids. Experientia 45(4):377 378 Anton E (1983) Association of Golgi vesicles containing acid phosphatase with the chromatoid body of rat spermatids. Experientia 39(4):393 394 Aravin A, Gaidatzis D et al (2006) A novel class of small RNAs bind to MILI protein in mouse testes. Nature 442(7099):203 207 Aravin AA, Sachidanandam R et al (2007) Developmentally regulated piRNA clusters implicate MILI in transposon control. Science 316(5825):744 747 Bardsley A, McDonald K et al (1993) Distribution of tudor protein in the Drosophila embryo suggests separation of functions based on site of localization. Development 119(1):207 219 Biggiogera M, Fakan S et al (1990) Immunoelectron microscopical visualization of ribonucleo proteins in the chromatoid body of mouse spermatids. Mol Reprod Dev 26(2):150 158 Braun RE (1998) Post transcriptional control of gene expression during spermatogenesis. Semin Cell Dev Biol 9(4):483 489 Brengues M, Teixeira D et al (2005) Movement of eukaryotic mRNAs between polysomes and cytoplasmic processing bodies. Science 310(5747):486 489 Carmell MA, Xuan Z et al (2002) The Argonaute family: tentacles that reach into RNAi, developmental control, stem cell maintenance, and tumorigenesis. Genes Dev 16(21): 2733 2742 Carmell MA, Girard A et al (2007) MIWI2 is essential for spermatogenesis and repression of transposons in the mouse male germline. Dev Cell 12(4):503 514 Chen C, Jin J et al (2009) Mouse Piwi interactome identifies binding mechanism of Tdrkh Tudor domain to arginine methylated Miwi. Proc Natl Acad Sci USA 106(48):20336 20341 Chennathukuzhi V, Morales CR et al (2003a) The kinesin KIF17b and RNA binding protein TB RBP transport specific cAMP responsive element modulator regulated mRNAs in male germ cells. Proc Natl Acad Sci USA 100(26):15566 15571 Chennathukuzhi V, Stein JM et al (2003b) Mice deficient for testis brain RNA binding protein exhibit a coordinate loss of TRAX, reduced fertility, altered gene expression in the brain, and behavioral changes. Mol Cell Biol 23(18):6419 6434 Chuma S, Hiyoshi M et al (2003) Mouse Tudor Repeat 1 (MTR 1) is a novel component of chromatoid bodies/nuages in male germ cells and forms a complex with snRNPs. Mech Dev 120(9):979 990

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Styhler S, Nakamura A et al (1998) vasa is required for GURKEN accumulation in the oocyte, and is involved in oocyte differentiation and germline cyst development. Development 125(9):1569 1578 Tanaka SS, Toyooka Y et al (2000) The mouse homolog of Drosophila Vasa is required for the development of male germ cells. Genes Dev 14(7):841 853 Tang XM, Lalli MF et al (1982) A cytochemical study of the Golgi apparatus of the spermatid during spermiogenesis in the rat. Am J Anat 163(4):283 294 Teixeira D, Sheth U et al (2005) Processing bodies require RNA for assembly and contain nontranslating mRNAs. RNA 11(4):371 382 Thorne Tjomsland G, Clermont Y et al (1988) Contribution of the Golgi apparatus components to the formation of the acrosomic system and chromatoid body in rat spermatids. Anat Rec 221(2):591 598 Toyooka Y, Tsunekawa N et al (2000) Expression and intracellular localization of mouse Vasa homologue protein during germ cell development. Mech Dev 93(1 2):139 149 Tsai Morris CH, Sheng Y et al (2004) Gonadotropin regulated testicular RNA helicase (GRTH/ Ddx25) is essential for spermatid development and completion of spermatogenesis. Proc Natl Acad Sci USA 101(17):6373 6378 Unhavaithaya Y, Hao Y et al (2009) MILI, a PIWI interacting RNA binding protein, is required for germ line stem cell self renewal and appears to positively regulate translation. J Biol Chem 284(10):6507 6519 Vagin VV, Wohlschlegel J et al (2009) Proteomic analysis of murine Piwi proteins reveals a role for arginine methylation in specifying interaction with Tudor family members. Genes Dev 23(15):1749 1762 Vasileva A, Tiedau D et al (2009) Tdrd6 is required for spermiogenesis, chromatoid body architecture, and regulation of miRNA expression. Curr Biol 19(8):630 639 Ventela S, Toppari J et al (2003) Intercellular organelle traffic through cytoplasmic bridges in early spermatids of the rat: mechanisms of haploid gene product sharing. Mol Biol Cell 14(7):2768 2780 Walt H, Armbruster BL (1984) Actin and RNA are components of the chromatoid bodies in spermatids of the rat. Cell Tissue Res 236(2):487 490 Wang J, Saxe JP et al (2009) Mili interacts with tudor domain containing protein 1 in regulating spermatogenesis. Curr Biol 19(8):640 644 Watanabe T, Takeda A et al (2006) Identification and characterization of two novel classes of small RNAs in the mouse germline: retrotransposon derived siRNAs in oocytes and germline small RNAs in testes. Genes Dev 20(13):1732 1743 Werner G, Werner K (1995) Immunocytochemical localization of histone H4 in the chromatoid body of rat spermatids. J Submicrosc Cytol Pathol 27(3):325 330 Yang J, Medvedev S et al (2005a) The DNA/RNA binding protein MSY2 marks specific transcripts for cytoplasmic storage in mouse male germ cells. Proc Natl Acad Sci USA 102(5):1513 1518 Yang J, Medvedev S et al (2005b) Absence of the DNA /RNA binding protein MSY2 results in male and female infertility. Proc Natl Acad Sci USA 102(16):5755 5760 Zhong J, Peters AH et al (1999) A double stranded RNA binding protein required for activation of repressed messages in mammalian germ cells. Nat Genet 22(2):171 174

Chapter 15

Sperm RNA: Reading the Hidden Message David Miller

Abstract In the early 1990s, an obstetrician colleague and I came to the conclusion that looking for sperm-specific RNA by RT-PCR in the ejaculate of recently vasectomized men might be a more sensitive (and convenient) way of determining whether the operation had been a success. A pilot study was conducted (never published), which showed clearly that molecular methods could indeed be a more efficient and accurate alternative for detecting residual spermatozoa in a sterilized ejaculate than manually scanning through many semen samples under the microscope. Shortly thereafter, the first reports appeared in the literature documenting the presence of mRNA in human spermatozoa by RT-PCR. Sperm RNA was an unexpected discovery for many andrological biologists because it had been generally assumed long before that RNA was lost or degraded during the extensive cellular remodeling that spermatids undergo during spermiogenesis, when nuclear condensation is accompanied by the loss of most of the cytoplasm (and cytoplasmic RNA) and the termination of protein synthesis. Hence, the detection of fully processed intact mRNA in mature ejaculate spermatozoa was treated with some scepticism. Some critics considered that sperm RNA was an artifact caused by immature round or somatic cell contamination of semen samples. Others felt that sperm RNA was (at best) a minor residual component of the mature gamete that served no useful purpose. Both criticisms were subsequently challenged by compelling experiments showing that sperm RNA is unequivocally spermatozoal in origin and that regardless of its passive diagnostic potential in a fertility context, sperm RNA probably performs active functions both within the cell itself and perhaps more intriguingly, once it gains access to the ooplasm. The focus of this chapter is on the discovery of sperm RNA and the historical and contemporary research that has provided some important clues to its function. It will include discussion of its utility in a diagnostic context (for male infertility) and of recent interesting work suggesting that sperm RNA is part of and evidence for a wider

D. Miller Reproduction and Early Development Group, Leeds Institute of Genetics and Health Therapeutics (LIGHT), University of Leeds, Clarendon Way, Leeds, West Yorkshire LS2 9JT, UK e mail: [email protected]

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9 15, # Springer Verlag Berlin Heidelberg 2011

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mechanism for reshaping the genome that augments its more obvious epigenetic potential as detailed elsewhere in this volume. Keywords Embryonic genome activation Epigenetics Gene expression Repetitive elements Reverse transcriptase Sperm RNA

15.1

Introduction

Female gametes frequently retain many of the characteristics of a generalized somatic cell, in that they have abundant cytoplasm with an intact and functional gene expression machinery. In contrast, male gametes show a very wide variety of morphological forms ranging from the amoeboid sperm of nematodes to the highly specialized flagellates of most vertebrate species (LaMunyon and Ward 2002; Swallow and Wilkinson 2002) and Fig. 15.1. Many plants have flagellate sperm (the ferns for example), although in all ‘higher’ plants, free living motile sperm have given way to immotile sperm encased in pollen that rely on transporting factors such as the wind and insects to bring male and female gametes into contact. In all complex sexually reproducing metazoans studied to date, the male gamete is responsible for transmitting the paternal genome to the female gamete and in so doing, supports syngamy, zygote formation, and assuming all is well, subsequent embryonic development. In terms of understanding their formation, the most studied sperm of vertebrates are undoubtedly those of rodents, particularly the rat and mouse, although because of the massive expansion in assisted reproduction procedures in the past 15 years, the humans have also enjoyed considerable attention in this regard (Elder and Dale 2001). Sperm of numerous other vertebrate species have also enjoyed more attention, often for reasons of conservation (Garcia-Rosello et al. 2009; Kouba et al. 2009; Morrow et al. 2009). With the exception of certain insects, notably, Drosophila, (Dorus et al. 2006; Rathke et al. 2007; White-Cooper 2009), we know very little about the sperm of invertebrate species including the round worm, Caenorhabditis elegans, despite the fact that this animal’s developmental body plan where one can follow the origin and fate of every cell, including the gametes, is better known than in any other species (Hope 1994). In mammals, spermiogenesis (the postmeiotic phase of spermatogenesis) accommodates the shutting down of nuclear gene expression to facilitate the generalized substitution of somatic and germ-line nucleohistones with the male gamete-specific nucleoprotamines (Hecht 1988, 1998; Braun 2001; Wykes and Krawetz 2003; D’Occhio et al. 2007). During this substitution and thereafter, gene expression is exclusively under translational control, utilizing mRNAs transcribed and stored earlier in spermatogenesis (Gu et al. 1996; Hecht 1998). A similar mechanism operates in the o€ ocyte, where the maternal genome is shut down by the germinal vesicle breakdown stage and the cell relies on mRNAs

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Fig. 15.1 Line drawings and photomicrographs of some plant spermatozoa. (a) Line drawings of flagellate spermatozoa from various nonflowering plants (not to scale). Following from top left to bottom right, these show gametes from typical phytoplankton (Diatom), brown algae (Laminaria, Fucus); Bryophytes (Liverwort, Moss); ferns and fern like plants (Lycopodium, Selaginella, Psilotum, Fern, Equisetum). Source: Wikimedia Commons, from Raven et al. (2005). (b) DAPI stained pollen grain from Maize plant (Zea mays) showing both sperm nuclei (brightly staining; one nucleus contributes to the zygote, the other to the endosperm) and the fainter vegetative nucleus. From Engel et al.; plant gamete gene expression (http://www.pgec.usda.gov/.../Gametes/ Gametesindex.htm). (c) streamlined spermatozoid from Equisetum arvense showing multiple flagella. Courtesy of Renzaglia at http://www.phytoimages.siu.edu/

produced and stored at earlier growth phases to support protein synthesis (Briggs et al. 1999). Although the maternal genome is not repackaged by protamines, its shutdown is governed by similar principles arising from chromatin modification and condensation that accompanies the formation of the MII metaphase spindle after GV breakdown (Briggs et al. 1999). The RNA store is also particularly important in the early zygote, prior to embryonic genome activation, when the embryo relies solely on it to support development (McLay and Clarke 1997; Adjaye et al. 2007). Indeed, it is the embryonic reliance on maternal RNAs coupled with the comprehensive remodeling of the spermatid during spermiation and the consequent shedding of its cytoplasm that have been the main obstacles to the recognition of sperm RNA by the scientific community. As a terminally differentiated cell with a shut down, densely packed haploid genome, the mammalian sperm is exquisitely

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specialized (and optimized) to deliver that genome to the egg. With no fresh RNA synthesis likely and no translational machinery available to support its translation, any RNA remaining in the mature spermatozoon is residual and of no useful function. Not surprisingly, anything that has no function is either ignored, forgotten, or both. This chapter aims to introduce the reader to the original controversy over sperm RNA as it arose in the 1960s and 1970s and to examine more recent reports on its composition, utility, and possible functions. In the light of the postgenomic, epigenetic, and bioinformatic era that we are now entering, I shall also attempt to show how the older and more recent research are linked by some common threads that could indicate a male gamete with a more active nucleus than originally thought. If correct, this more active spermatozoan dynamic might be responsible for wider epigenetic phenomena that could have a profound influence on how we view the male gamete’s role as a passive delivery vehicle for the paternal genome and on the use of immature sperm in assisted conception procedures.

15.2

Gene Expression Studies in Mature Spermatozoa: Historical Aspects

The writer first became interested in sperm RNA in the early 1990s as a marker for successful vasectomy. Currently, the standard test (in the UK) is for a pathologist to examine postoperative semen samples for signs of sperm (Hancock and McLaughlin 2002). The screening procedure is normally carried out manually, requires at least two sequential samples from a vasectomized patient, and is rather time consuming. We reasoned that a suitable molecular marker could do the job more quickly and accurately. Protamine 2 mRNA was chosen as the target transcript because it is haploid-expressed and testis-specific (Domenjoud et al. 1991). The assay was based on standard RT-PCR protocols of the time and was successful at detecting a 110 bp cDNA product derived from the spliced 30 and 50 ends of PRM2’s exons 2 and 5, respectively, in normozoospermic control samples. Only b-actin as a positive control marker was detected in postvasectomized men and PRM2 in a few patients with incomplete clearance of spermatozoa. The results of this small trial were never published, but an associated study reporting on the efficacy of sperm RNA as a proxy for the testis (Miller et al. 1994) led to the revisiting of earlier work on the gene expression capacities of mature spermatozoa and some interesting attributes of the sperm that at the time of their appearance were somewhat obscure and mysterious but with new evidence may shed light on functional significance. Most early experiments were aimed at understanding the flow of genetic information in mature spermatozoa compared with typical somatic cells. It was generally assumed that in the former, this information flow was unnecessary and had essentially ceased. Reports appearing in the late 1950s and early 1960s, showing that spermatozoa in semen samples were capable of taking up radioactive tracers into acid

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precipitable material (Bhargava 1957; Abraham and Bhargava 1963), were dismissed as artifacts of bacterial contamination (Markewitz et al. 1967). In a later series of landmark experiments, several independent studies concluded that bovine spermatozoa were transcriptionally active, but that this activity was confined to their mitochondria (Premkumar and Bhargava 1972; MacLaughlin and Terner 1973; Hecht and Williams 1978). A more recent report confirmed these observations in human spermatozoa (Grunewald et al. 2005). These early studies essentially concluded much of the original research on sperm nucleic acid metabolism until the appearance of a report on the presence of RNA in the nucleus of the Scolopendrium (fern) spermatozoid detected by an electron microscopy based in situ hybridization technique (Rejon et al. 1988). Pessot later demonstrated nuclear RNA in rat and human sperm (Pessot et al. 1989) and resolved small sperm nuclear RNAs by gel electrophoresis. Kumar et al. (1993) provided the first evidence for the presence of c-MYC mRNA in human spermatozoa localized to the mid-piece and tail regions. Numerous mRNAs including HSP 70, 90, and b-actin were detected in ejaculate swum-up spermatozoa using both standard RT-PCR and RNA display techniques [Miller 1997, (Fig. 15.2)], and in a complementary study, Wykes et al. (1997) localized PRM1, PRM2, and TNP2 RNA in or around the cell nucleus using in situ hybridization (Fig. 15.3). Additional reports of mRNAs in human ejaculate spermatozoa include HLA (Chiang et al.

ST

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sperm testis

liver

ovary endo’m RBC DNA

ER2 actin HSP90 PRM2 (+I) HSP70 PRM2 (–I)

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Fig. 15.2 Identification of specific mRNAs in human ejaculate spermatozoa. Agarose gel electro phoresis of RT PCR generated amplicons using primers directed to sequences on ER2a, HSP 70, HSP 90, protamine 2 (PRM2), and b actin on RNA isolated from sperm, testis, liver, ovary, endometrium, and red blood cells (RBCs). PRM2 primers span a 162 bp intron, giving rise to a 312 bp product from genomic DNA [PRM2 (þI)] and a 150 bp product from testis and sperm RNAs [PRM2 ( I)]. From Miller et al. (1994)

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Fig. 15.3 Localization of RNA in Rat (a d) and human (e i) sperm nuclei. Electron micrographs of nuclei labeled with colloidal gold tagged RNAse in the absence (a, b and d) and presence (c) of tRNA or with colloidal gold tagged BSA (d and f). Light micrographs of silver grains in nuclei labeled with 35S anti sense RNAs to protamine 1 (g), protamine 2 (h) or transition protein 2 (i). Panels a f from Pessot et al. (1989). Panels g i from Wykes et al. (1997)

1993, 1994), L-type calcium channel and N-cadherin (Goodwin et al. 2000a, b), estrogen and progestin-like receptors (Durkee et al. 1998; Sachdeva et al. 2000; Aquila et al. 2004; Lambard et al. 2004b), cyclic nucleotide phosphodiesterases (PDE) (Richter et al. 1999), integrins (Rohwedder et al. 1996), aromatase (Aquila et al. 2003; Lambard et al. 2003), nitric oxide synthase (NOS) (Lambard et al. 2004a), and surprisingly, insulin (Aquila et al. 2005), among others (Dadoune et al. 2005). Complex RNA populations have also been reported in the sperm of bovines (Gilbert et al. 2007; Lalancette et al. 2008; Bissonnette et al. 2009) and porcines (Curry et al. 2009; Yang et al. 2009) and the sperm of plant pollen (Engel et al. 2003; Okada et al. 2007; Borges et al. 2008; Bayer et al. 2009; Gou et al. 2009; Haerizadeh et al. 2009). Indeed, RNA carriage in the sperm of mature pollen is a rapidly expanding area of research at the time of writing, and epigenetic effects of its transmission to ovules will be considered briefly in a later section. Most interesting are the recent reports on the detection of sperm mRNAs in the testis and/or spermatozoa of the water buffalo, Bubalus bubalis, containing small microsatellite repeat sequences including GACA, GATA, and CACCTCTCCACCTGCC (minisatellite 33.15), some of which are sperm-specific

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(Srivastava et al. 2006, 2009). These minisatellites are mainly confined to noncoding regions of the genome, and their presence in some mRNAs suggests a role in the regulation of their expression. The majority of GACA and all GATA-tagged transcripts are either highly or exclusively detectable in the testis/sperm of this species compared with somatic tissues. This group obtained similar results with mRNAs containing the longer 33.15 minisatellite sequences, which were highly detectable in buffalo testis or sperm. Moreover, the presence of orthologous DNA sequences from several other mammalian species suggests that their role is likely to be conserved. Unfortunately, the authors did not go so far as to look for the presence of orthologous mRNA sequences in the mature spermatozoa from these species. Nevertheless, these data accord strikingly with results from our own laboratory showing that transcripts for other repetitive sequences including non long terminal repeat (non-LTR) and short (SINE) and long (LINE) interspersed nuclear repeat elements are abundant in mature human spermatozoa (Miller 2000). The presence of antisense and miRNAs in the sperm of many species including human (Ostermeier et al. 2005), mouse (Amanai et al. 2006), pig (Curry et al. 2009), and plants (Grant-Downton et al. 2009) suggests that sperm RNA has the potential to influence the phenotype of progeny, a feature that is better understood in plant than in mammalian reproduction. These observations are interesting in the light of work showing that sperm chromatin released from epididymal mouse spermatozoa following activation of an endogenous endonuclease is enriched in DNA sequences for both SINE and LINE elements (Pittoggi et al. 1999). Both the transcriptional and endonuclease susceptibility data from these studies suggest that repetitive elements are preferentially located in the minor nucleohistone packaged chromatin of the sperm nucleus, which is also enriched in gene regulatory sequences carrying a strong embryonic developmental ontology (Arpanahi et al. 2009).

15.3

Gene Expression Studies in Mature Spermatozoa: In Relation to Sperm RNA Carriage

In comparison with other developmental systems, gametogenesis is unique in accommodating meiosis, where gene expression supporting germ cell formation must continue while DNA recombination and haploid reduction is taking place. More than 50 years can elapse in the human between the fetal block at diakinesis (MI) and the adult initiation of MII, during which time, gene expression is under both transcriptional and translational levels of control (exclusively translational following germinal vesicle breakdown; Briggs et al. 1999; Evans and Hunter 2005; Piccioni et al. 2005). There is no gap between MI and MII in spermatogenesis, and unlike oogenesis, there is an extended posthaploid period (spermiogenesis) when active gene expression switches from a mixed transcriptional/translation control mechanism to one based solely on translation of stored mRNAs (Hecht 1998). This extended period is required to accommodate the repackaging of DNA from the usual histone-based nucleosomal configuration to a protamine-based

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nucleotorroidal conformation that is almost 20 times more efficient at DNA condensation and enables the complete shutdown of the spermatid nucleus (Bench et al. 1996; Brewer et al. 2002; Balhorn 2007). The premeiotic and meiotic stages, usually referred to as spermatocytogenesis and spermatidogenesis, take approximately 40 days to complete in the human (14 days in the mouse). The gradual shutdown of RNA transcription begins during meiosis when the paired sex chromosomes are accommodated in the male germ cell-specific sex body, leading to the repression of gene expression from the X and Y chromosomes (FernandezCapetillo et al. 2003; de la Fuente et al. 2007). Autosomal retrogenic paralogs of X-linked genes are switched on and replace the sex chromosome-derived gene products as required (Marques et al. 2005; Vinckenbosch et al. 2006). The switch from X-linked PGK1 to the autosomal retrogene, PGK2, is a studied example (Danshina et al. 2010; McCarrey and Thomas 1987; Zhang et al. 1999). The gross posthaploid RNA-dependent cellular remodeling that accompanies DNA supercondensation in spermiogenesis occurs in the absence of transcription. The control of gene expression during these late-spermatid developmental stages involve both 30 and 50 UTR RNA binding proteins and by changes in polyA tail length (Fujimoto et al. 1988; O’Brien et al. 1994; Kleene 1996; Gu et al. 1998; Dai et al. 2001; Luetjens et al. 2004; Miller and Ostermeier 2006). We now also recognize that the Argonaute family of testis-specific MILI binding PIWI pRNAs may play a role in controlling translation via these routes by interacting with mRNAs and their binding proteins in the nucleus, the cytoplasm, and in the chromatoid body (Aravin et al. 2006; Grivna et al. 2006; Kotaja et al. 2006). This unusual structure or ‘nuage’ is a prominent feature in the cytoplasm of meiotic and postmeiotic spermatocytes, although there is evidence for an equivalent structure in oocytes (Chemes et al. 1978; Morales et al. 1991; Parvinen 2005; Reunov 2006). Taken together, the complexity of spermatogenesis has led to the evolution of a robust system of gene expression control, ensuring that uninterrupted sperm production can take place for the duration of the male’s life following sexual maturity. As a terminally differentiated cell, the ejaculate (mammalian) spermatozoon is exquisitely specialized for delivering the paternal genome to the egg. Most gene expression events in the germ line and supporting Sertoli cells are geared towards achieving this goal. Hence, the presence of RNA in the sperm nucleus as reported by various laboratories is paradoxical if one assumes that it serves no function. However, even this assumption can be accommodated if the RNA is solely residual. Considering the high-energy expenditure of spermatogenesis compared with oogenesis, it makes energetic ‘sense’ for the sperm to rely on the egg to discard any surplus constituents that are carried into the ooplasm alongside the paternal genome. Evidence of extensive ubiquitination of structural proteins in the sperm suggests that they are tagged for recycling by the oocyte’s proteasomal machinery, and sperm RNA could be destroyed by equivalent targeted RNAse activities (Sutovsky et al. 1999). Judging by the difficulties encountered in extracting the small quantities of available RNA from human spermatozoa (estimated at between 10 and 14 fg) and its frequent contamination by DNA

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(Miller et al. 1999), it is likely that sperm RNA is an integral constituent of the nucleus. In support of this conclusion, in situ hybridization assays have located the RNA to the sperm head (Rejon et al. 1988; Pessot et al. 1989; Wykes et al. 1997; Fig. 15.3). Such a location is fully compatible with a residual presence of sperm RNA, which, formerly active in the condensing and transcriptionally silent spermatid, becomes trapped within the nucleus as nucleocytoplasmic transport activity ceases (Fig. 15.3). Indeed, it is also possible that the mRNA is involved in the transcriptional shut down of the sperm nucleus via an RNAi effect, feeding back on promoter sequences that may escape histone replacement (Miller and Ostermeier 2006; Fig. 15.4). Several lines of evidence, however, suggest that the RNA of the mature spermatozoon is not residual. The absence of introns indicates that mRNA is fully processed, which would not be expected of hnRNA (Miller et al. 1999; Ostermeier et al. 2005). Moreover, the selective retention of mRNAs and siRNAs when most cytoplasmic RNA is lost to the residual body (normally destroyed during sperm preparation) during remodeling argues against passive trapping as does the evidence that sperm RNA can support protein synthesis de novo during capacitation

Fig. 15.4 Potential roles for spermatozoal RNA. Sperm RNA may be associated with the small proportion of sperm chromatin remaining packaged by nucleosomes [indicated by yellow patches (a)]. Recently discovered antisense spermatozoal RNAs could provide an epigenetic mark that is necessary for establishing and/or maintaining paternal imprints (b). Some cytoplasmic mRNAs are apparently translated de novo, suggesting the active maintenance of the gamete during its journey to the egg (c). Paternal RNAs may provide epigenetic marks to the developing embryo that influence the phenotype of the offspring (d). Panel e illustrates possible RNA carriage by the paternally derived centrosome. From Miller and Ostermeier (2006)

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(Naz 1998; Gur and Breitbart 2006). This Ca2+-dependent process makes ready the spermatozoon for the acrosome reaction (Luconi et al. 1998; Breitbart 2003; Suarez 2008). The concept of active RNA-dependent translation in capacitating sperm is supported by reports showing that specific sperm RNAs are ‘consumed’ during manual processing that supports capacitation of normal viable sperm in vitro (Naz 1998; Lambard et al. 2004a; Aoki et al. 2006). An effect of sperm RNA on nonMendelian inheritance of coat color in the mouse has also been reported (Rassoulzadegan et al. 2006). The mechanism here appeared to be via an RNA-mediated epigenetic and heritable paramutation effect.

15.4

Sperm RNA as a Diagnostic Resource

One should not underestimate the damaging effects of male factor infertility, which is an important and increasingly serious human problem (Balen 2008). Unfortunately, the rapid expansion of assisted reproduction technologies (ART) aimed at relieving the condition has not been paralleled by a concomitant expansion in studies into its causes. There are several reasons for this impasse. While positional gene mapping studies based on family cohorts have provided some evidence of a heritable component in male factor infertility (Lilford et al. 1994), the condition often remains hidden within consanguineous populations on which the best gene studies rely. However, it is difficult to conclude that traditional linkage disequilibrium studies would be of much help because most cases of male infertility are unlikely to arise from single gene defects. Indeed, few, if any, reports identifying a mutation or deletion in any autosomal or X-linked gene involved in murine spermatogenesis have appeared in a corresponding human study [see the review of Yan (2009)] for a more general overview and (Roest et al. 1996; Steger et al. 2003; Grootegoed et al. 1998; Hagaman et al. 1998; Adham et al. 2001; Cho et al. 2001; Miki et al. 2002 for some specific examples). There is also an understandable lack of tissue available for molecular investigation. Testicular biopsy, for example, is not justified or ethically appropriate unless a clearly abnormal semen profile is reported, which is not the case in the majority of cases. Paradoxically, by effectively circumventing natural barriers to conception that would ordinarily exclude genetic anomalies underlying the infertile male phenotype, the advent of intracytoplasmic sperm injection (ICSI) is likely to increase the prevalence of an infertile or subfertile genotype in the general population. The reasons for this likelihood will become apparent from the following sections in this chapter. To gain a better understanding of the complexity of the RNA population in human ejaculate spermatozoa, Ostermeier et al. (2002) used arrays to detect over 3,000 mRNA species in ejaculate human spermatozoa. The sperm transcripts were a subset of over 7,000 identified in a testis cDNA library. Alongside variations in the expression of two sperm RNAs coding for LDHC transcript variant 1 and TPX1 reported in men with poor sperm motility (asthenozoospermia; Wang et al. 2004), these studies provided the first array-based experimental evidence that sperm

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RNA as a useful proxy for the testis can be used diagnostically. In relation to the latter study, semi-quantitative measurements of the RNA profiles from ejaculate human spermatozoa by target-directed (PRM1, PRM2, and eNOS) RT-PCR provided evidence for mRNA consumption or turnover in response to motility and/or capacitative function of sperm taken from different layers in a differential density gradient (Lambard et al. 2004a). Interestingly, a latent translational capacity in spermatozoa based on its abolition by cycloheximide had been reported previously (Naz 1998). Serial Analysis of Gene Expression (SAGE) revealed 389 clustered genes with a highly selective grouping among the most abundant SAGE tags (Zhao et al. 2006). Almost 25% of these (96) related to DNA-dependent transcription or regulation of transcription. More sophisticated arrays based on Affymetrix and Illumina platforms successfully distinguished between normozoospermic and teratozoospermic (abnormal forms) using clustering algorithms and showed a major disturbance of ubiquitin-mediated protein recycling pathways in the abnormal semen samples (Platts et al. 2007; Fig. 15.5). A related report based on the data from these samples revealed the presence of a core set of 30 transcript pairs in the sperm from 24 normozoospermic men that are highly and invariably co-expressed in all samples (Lalancette et al. 2009). This core set of transcripts has roles in the regulation of transcription, development, metabolism, phosphorylation, protein synthesis, ubiquitination, and nuclear mRNA splicing. The implication is that the core set could be used as the baseline for studies aimed at uncovering deregulated gene expression pathways in the spermatozoa of infertile men. Upwards of 3,000 spermatozoal transcripts (Ostermeier et al. 2002) should contain a considerable body of information regarding testicular events underpinning spermatogenesis and by extension, male fertility. Although much emphasis has been placed on eliminating any somatic or immature germ cell-derived transcripts in the target preparations for analysis, there is no logic in doing so for diagnostic purposes. Careful preparation of spermatozoa was an absolute priority for establishing beyond any doubt the presence of mRNA in the ejaculate cell (Miller 2000). By excluding other cell types including immature round cells and leucocytes, however, one loses potentially important information on the condition of the testis and its associated organs including the prostate and seminal vesicles as well as information on inflammatory conditions. Using complementation assays, one report so far has identified mutations in the KLHL10 gene in men with oligozoospermia using total RNA isolated from whole ejaculates (Yatsenko et al. 2006). KLHL10 is expressed exclusively in the cytoplasm of elongating and elongated spermatids, and Klhl10 null mice are infertile due to a failure of sperm maturation (Yan et al. 2004). The use of microarrays or new sequencing technologies are obvious ways forward in the search for gene pathways and networks associated with male factor infertility. (Ostermeier et al. 2002; Wang et al. 2004). Moreover, by characterizing the set of sperm-specific RNAs delivered to the oocyte, we can begin to ask questions about what these RNAs may be doing in the fertilized egg (Ostermeier et al. 2004). An additional and currently overlooked potential of whole semen RNA profiling is in the early detection of carcinoma in situ (CIS) related to testicular

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Fig. 15.5 Diagnostic potential of sperm RNA. It is possible to cluster semen profiles into normozoospermic (N) and teretozoospermic (T) categories based on analysis of sperm transcripts from fertile (13 samples) and infertile (8 samples) men. The two statistical tools illustrated here are unsupervised hierarchical clustering (a) and principal component analysis (b). The subcategories in a refer to different models or modes of analysis. From Platts et al. (2007)

cancer (TC). Current methods relying on immunocytochemical detection of germ cell markers such as OCT3/4 and NANOG show that semen cell profiling can be informative in the detection of CIS (Hoei-Hansen et al. 2007; van Casteren et al. 2008). However, semen RNA profiling on microarrays may be more informative for both the detection of CIS/TC and also for indicating the stage of the disease. Moreover, RNA-based assays should be more sensitive than assays based on immunocytochemistry. TC is an increasing problem among younger men worldwide, particularly in developed countries (Virtanen et al. 2005). The condition is likely to be caused by embryonic germ cells that are not cleared from the fetal testis and which are symptomatic of a testicular dysgenesis that also leads to lowered sperm counts (reviewed in (Sonne et al. 2008)). Finally, the roles that selectively retained mRNA plays in spermatozoon development, chromatin repackaging, and genomic imprinting are all important goals deserving the attention of andrologists and other specialists in the years to come.

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Functions for Sperm RNA

There are no available studies on the presence of RNA in the sperm of birds, reptiles fish, or invertebrates, although based on its existence in the sperm of mammals and plants, we can predict with some confidence that future studies will confirm its presence in these classes too. Providing clues to the RNAs’ function is challenging as the spermatozoal nucleus is considered inert (Grunewald et al. 2005 and Fig. 15.4). The recent updating of Bhargava’s (1957) and Naz’s (1998) original work by Gur and Breitbart (2006) demonstrated the incorporation of labeled amino acids into proteins by capacitating spermatozoa and indicated that this renewed synthesizing activity was dependent on translation of endogenous mRNAs de novo. Some specific aspects of this study with wider implications for our understanding of protein synthesis in general were most intriguing. De novo protein synthesis was abolished by the mitochondrial uncoupler carbonylcyanide p-trifluoromethoxyphenyl hydrazone (FCCP), which is as predicted of an ATPdependent mechanism. However, the antibiotic, chloramphenicol, which targets prokaryotic (and mitochondrial) 55S ribosomes, also abolished protein synthesis, while contrary to the Naz (1998) study, cycloheximide, which targets cytoplasmic (80S) ribosomes, had no effect. These findings suggest that spermatozoal mRNA is translated on mitochondrial polysomes. It remains unclear how cytoplasmic mRNAs, which carry distinctive 50 recognition sequences designed for 80S ribosomes, are translated on mitochondrial machinery, and this warrants further investigation, including confirmation in other species and by other laboratories (Zhao et al. 2009). A requirement for spermatozoal RNA in supporting protein synthesis de novo is understandable. However, several reports, mainly from studies on plant pollen suggest that sperm RNA may also have postfertilization functions. Although small RNAs are dealt with elsewhere in this volume, it is worth considering briefly here. Based on a heterologous model system, we know that the spermatozoon delivers its RNA cargo to the oocyte (Ostermeier et al. 2004). Rassoulzadegan’s study (Rassoulzadegan et al. 2006) showed that a c-Kit-derived heritable effect on hair color in the mouse was strongly influenced by the presence of aberrant levels of ‘scambled,’ noncoding c-Kit RNA transferred by the spermatozoa of an affected individual. Under normal circumstances, animals homozygous for the functional wild type alleles arising from heterozygous crosses should be unaffected by this trait. However, even these animals were affected, indicating a strong, nonMendelian, transgenerational paramutational epigenetic effect of sperm RNA. This work was followed by a study showing that pathological overexpression of Cdk9, a key regulator of cardiac growth in the mouse, could be induced and heritably transmitted following the intraooplasmic injection of RNAs targeting the gene, including the sequence-related miR-1 miRNA. miR-1 is among a number of inhibitory small RNAs that are present in sperm (Wagner et al. 2008). Paramutation is commonly observed in some plant species but rarely described in animals (Ashe and Whitelaw 2007). These new data coupled with the

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increasing interest in the role of paternal mRNAs and siRNAs in controlling zygotic development in plants including rockcress, maize, and leadwort (Engel et al. 2003; Borges et al. 2008; Chambers and Shuai 2009; Gou et al. 2009; Haerizadeh et al. 2009) suggest that paternal transmission of epigenetic effects including paramutation may be a widespread feature of sexual reproduction. As gynegenones (and other naturally occurring parthenotes) suggest that a male contribution is not strictly necessary for successful embryonic development (Kono et al. 2004), the phenomenon of sperm RNA transmission may seem superfluous. However, this author considers it possible that in mammals, the RNA preserves the imprinting profile of the paternal genome and prevents undesirable parthenogenetic activation. The preferred RNA-dependent mechanism may also be used by testisactive transposable ‘selfish’ elements that can only be transmitted to and expand in subsequent generations following introduction by sperm. Indeed, much of the Argonaute-directed repression of retrotransposon expression in the testis is likely to have evolved in response to that challenge (Ma et al. 2009). In the light of these findings, the original suggestion that sperm RNA is a relic of the gradual shut down of the nuclear transcriptional apparatus that marks the replacement of histones by protamines in spermatozoal chromatin is no longer tenable (Miller et al. 1999, 2005). This notion arose because RNAs (and proteins) are free to move between spermatids that remain connected to their neighbors during this period (Caldwell and Handel 1991), and on chromatin condensation, the RNA becomes trapped in the nucleus. The above reports and a later study (Lalancette et al. 2008) showing that sperm RNA is closely associated with the nuclear matrix (itself involved in regulating DNA replication and RNA synthesis; Jackson 1997; Shaman et al. 2007; Schofer and Weipoltshammer 2008) makes a residual presence seem rather unlikely.

15.6

Is the Spermatozoon Nucleus More Active Than We Think?

The recent work demonstrating de novo translation (and turnover) of spermatozoal RNA begs the question of whether the sperm nucleus is as shut down as original reports suggest. One unlikely source of support for a more active sperm nucleus (or at least a nucleus that can be activated under certain circumstances) comes from studies examining the mechanism of sperm-mediated gene transfer (SMGT) as reviewed recently by Smith and Spadafora (2005). These studies suggested that RNA and DNA are interconvertible de novo in mature spermatozoa. One study showed that RNA can be absorbed and reverse-transcribed into DNA and that absorbed DNA can be transcribed into fully processed RNA (Sciamanna et al. 2003). DNA- and RNA-dependent (presumably reverse transcriptase) polymerase activities were described originally in mature spermatozoa (and seminal plasma) back in the 1970s (Witkin and Bendich 1977; Witkin et al. 1977) although the role of these synthesizing complexes was unknown. The more recent work used SMGT

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with a hybrid murine leukemia virus/virus-like 30S (MLV/VL30) b-galactosidase (b-gal) gene-containing vector that was detected as DNA in murine spermatozoal extracts, indicating the activity of an operational reverse transcriptase in mature spermatozoa (Sciamanna et al. 2003, 2009). It is perhaps more intriguing that a DNA construct containing an intronic sequence can be processed by spermatozoal machinery such that the intron is correctly spliced out. Topoisomerase may be implicated in these processes (Shaman et al. 2006). These reports support the notion of a much more active sperm nucleus than is traditionally assumed. They suggest that under certain conditions, sperm RNA can be translated or reverse transcribed to cDNA. Certainly, compared with other tissues, the high number of testis-expressed processed pseudogenes suggests that the paternal germ line is a favorable environment for the generation of extra genetic diversity (Miller et al. 2007). Evidence to date, however, shows that cDNA copies of genes in testicular extracts are not present at a sufficiently high level to support the (high) activity of an endogenous reverse transcriptase in the testis (Zhong and Kleene 1999). But what other purposes could endogenous polymerases and nucleases serve the spermatozoon? One possibility is that the nucleases (including topoisomerases) help disable any spermatozoon that comes into contact with and absorbs exogenous nucleic acids. This is most likely to occur in the female genital tract where DNA released from other cell types (including bacteria) are encountered (Sotolongo et al. 2003; Ward and Ward 2004). Such an apoptosis-like mechanism would ensure that under normal circumstances, foreign DNA is unlikely to reach the egg (however, see Moreira et al. (2005) for one report describing inadvertent SMGT of bacterial DNA following murine ICSI). Certainly, activation of these endogenous nucleases releases DNA from sperm, making the cells unviable. The relationship between reverse transcriptase (RT) in spermatozoa on the one hand and sperm RNA on the other may be difficult to decipher, particularly in view of the observations that oocytes and early embryos may also have RT activities of their own (Evsikov et al. 2004). Interestingly, proviral particles, which are derived from endogenous LTR-containing retrotransposons are a rich source of active RT enzyme and have been observed in murine epididimides and seminal fluid (Kiessling 1989), in oocytes (Nilsson et al. 1981) and in fetal tissues (Mondal and Hofschneider 1982). Hence, endogenous LTR and nonLTR retrotransposon activity may be the general features of reproductive cells and tissue. If retrotransposons are indeed the source of active RT enzyme in spermatozoa and oocytes (Beraldi et al. 2006), it must be a special property of germ cells, because in all somatic cells (with the notable exception of some malignant cell lines), their expression is almost completely repressed by DNA methylation. Keeping these elements under control is also one potential function of the testisspecific PIWI RNAs (Thayer et al. 1993; Ma et al. 2009). Compelling evidence for the existence of an RT-dependent mechanism involving gametes and embryos has come from experiments where the enzyme’s activity was blocked with antiretroviral agents. In (niverapine) treated embryos, for example, irreversible developmental arrest was observed at the 2 4 cell stage, coincident with the timing of embryonic genome activation (ZGA) (Pittoggi et al. 2003).

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Moreover, embryo implantation in mice can be blocked using antisense morpholinos specifically targeting the LINE 1 RT (Beraldi et al. 2006), suggesting a wider role for this enzyme in reproductive function (see below). Understanding what RT is doing under these circumstances and why its abrogation is capable of abolishing ZGA is partially helped by our knowledge of what the enzyme does during the normal replication of LINE 1 elements. We know that through the expression of the p40 ribonucleoprotein encoded by the element’s ORF 1, for example, these elements are expressed in the testis, despite being under strict repression control (Branciforte and Martin 1992). Their pathological insertion into and inactivation of essential genes is observed from time to time (Kazazian et al. 1988). The mechanism of replication is also reasonably well understood and we can infer from this mechanism and observations of insertional events generated in vitro that LINE 1-mediated exon shuffling is a potentially powerful force for reshaping the genome (Moran et al. 1999). As indicated above, LINE 1 transcripts are also present in spermatozoa and in the ooplasm, although there is apparently a much greater load of LTR retrotransposon-derived RNA in oocytes and early cleavage stage embryos. Indeed, one study estimated that over 10% of the RNA in these embryos is of retrotransposon origin. Such elements may play a role in the regulation of gene expression in oocytes and early embryos by providing alternative promoter and first exon sequences for a number of genes (Peaston et al. 2004, 2007). All mammals carry a heavy retrotransposon load although their expression appears to be under strictly controlled regulation in all tissues, including the testis. Expression, however, is observed under permissive conditions (Larsson et al. 1981; Mondal and Hofschneider 1982; Nilsson et al. 1992; Sichangi et al. 2002), and there is good evidence that somatic tissues have taken advantage of their presence in the distant past. For example, the syncytin protein that mediates the fusion of placental cytotrophoblast into syncytiotrophoblast has been co-opted to this task from an endogenous retrotransposon ancestor (Mi et al. 2000). One final comment in relation to the above is the suggestion that tumor suppressor genes (TSGs) in higher metazoans including APC, BRCA, p53, RE, and WT1 arose from mating factor genes in lower organisms by either vertical or horizontal transmission (the latter via mobile elements) (Gosden et al. 1998). The report highlighting this hypothesis provided intriguing evidence that horizontal transmission of TSGs most likely occurred in oocytes (although there was no reason to exclude fertilizing spermatozoa from this scenario). This original hypothesis has gained considerable credence given the high levels of RT later found in gametes and the suggested role of their retrotransposon source in regulating host genes in oocytes and early embryos (Peaston et al. 2004). Moreover, the conclusion that transposable elements (probably retrotransposons) are frequently involved in tumorigenesis is also supported by the more recent evidence showing that RT-inhibitors can reverse a malignant phenotype by promoting redifferentiation and reestablishment of normal growth patterns (Sciamanna et al. 2005; Serafino et al. 2009).

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Conclusions

We have come a long way since the first reports of RNA in mature spermatozoa appeared in the literature. In both waves (the 1960s 1970s and 1990s), the subject sparked some controversy because the presence/synthesis of RNA did not fit well with the accepted paradigm of the sperm as a vehicle for packaging and delivering the paternal genome and nothing else. As new research has shed light on older observations, it is now clear that sperm are capable of far more dynamic nuclear responses than was considered possible hitherto. One aspect linking all of the experimental observations so far is that the machinery driving these events works independently of gene expression. There is no evidence that, for example, endogenous sperm RNA or the widely reported polymerases, nucleases, and transcription factors associated with sperm nuclei are involved in the expression of the paternal genome per se (except, possibly in the male pronucleus; Aoki et al. 1997). Instead, they seem to be part of an extragenomic mechanism active in the maintenance of the sperm during its journey to the egg and perhaps in the zygote itself. Translation of stored spermatozoal RNA de novo as a means of ‘topping up’ cognate proteins that are consumed or degraded during capacitation is one possible function, although disappointingly, this phenomenon has not been reported more widely. The siRNAs and antisense RNAs of mature sperm could have both local and remote regulatory roles in the spermatozoon and zygote, respectively. Locally, they could be involved in silencing the paternal genome during DNA repackaging or in helping to organize the repackaged chromatin into regional domains based on a potential for expression (in the male pronucleus, for example; Fig. 15.4). Their delivery alongside the paternal genome could also affect the zygote, marking the embryo for some paternally mediated epigenetic alteration in response, perhaps to local perturbations in the testis or epididymis. Also intriguing are the endogenous polymerase and endonuclease activities of mature sperm. These are normally observed under conditions when the cell is either capacitating or coming into contact with foreign DNA (or RNA) molecules. While it is relatively easy to see these reactions as part of an apoptotic-like mechanism aimed at protecting the zygote from foreign DNA intrusion or as responses to prolonged capacitation, the active conversion of RNA to cDNA is harder to explain. While there is evidence that an enzyme encoded by the LINE 1 element mediates this activity, alternative, endogenous retrovirally originated enzymes are also available in the CVBD egg and early embryo. One possibility, supported by experimental evidence, is that RT is somehow involved in triggering ZGA. Conceivably, this activity could involve exon shuffling that may be the key to the retrotransposon-mediated control of gene expression detected in oocytes and early embryos (Miller 2007) and that may even extend to tumorigenesis in later life. Sperm RNA (and/or RT) may be playing various active roles in the process from epigenetically marking the embryonic genome to acting as a template for some RT-mediated event, including ZGA.

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Regardless of any active role played by sperm RNA, both sperm RNA and total semen RNA offer considerable potential for routine investigations into the causes of male infertility and subfertility. In the majority of idiopathic cases where testicular biopsy is simply not warranted, they offer unique windows into the testis and its accessory organs that continue to be underexploited clinically despite a raft of publications showing clear efficacy in this context. The undisputed complexity of the nucleic acid cargo delivered to the egg by the sperm should also be given greater consideration by IVF practitioners who often resort to using testicular sperm in ICSI procedures. Such sperm may be successful at supporting normal development and the birth of live offspring, but the longer term consequences of their routine use have not been tested.

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Glossary

Acetylation

Agouti gene

Alleles

Assisted reproduction technologies (ART) 5-Azacytidine

Bisulfite genomic sequencing

Bivalent chromatin

The introduction, via an enzymatic reaction, of an acetyl group to an organic compound, for instance, to histones or other proteins. The agouti gene (A) controls fur color through the deposition of yellow pigment in developing hairs. Several variants of the gene exist, and for one of these (Agouti Variable Yellow, Avy), the expression levels can be heritably modified by DNA methylation. Different variants or copies of a gene. For most genes on the chromosomes, there are two copies: one copy inherited from the mother and the other from the father. The DNA sequence of each of these copies may be different because of genetic polymorphisms. The combination of approaches that are being applied in the fertility clinic, including IVF and ICSI. A cytidine analog in which the 5 carbon of the cytosine ring has been replaced with nitrogen. 5-azacytidine is a potent inhibitor of mammalian DNA methyltransferases. A procedure in which bisulfite is used to deaminate cytosine to uracil in genomic DNA. Conditions are chosen so that 5-methylcytosine is not changed. PCR amplification and subsequent DNA sequencing reveals the exact position of cytosines that are methylated in genomic DNA. A chromatin region that is modified by a combination of histone modifications such that it represses gene transcription, but at the same

S. Rousseaux and S. Khochbin (eds.), Epigenetics and Human Reproduction, Epigenetics and Human Health, DOI 10.1007/978 3 642 14773 9, # Springer Verlag Berlin Heidelberg 2011

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356

BrdU

Bromo domain

Brno nomenclature

Cell fate

Cellular memory (epigenetic)

ChIP ChIP on chip

ChIP Seq

Chromatid

Glossary

time retains the potential of acquiring gene expression. Bromodeoxyuridine (5-bromo-2-deoxyuridine, BrdU) is a synthetic nucleoside that is an analog of thymidine. BrdU is commonly used in the detection of proliferating cells in living tissues. Protein motif found in a variety of nuclear proteins including transcription factors and HATs involved in transcriptional activation. Bromo domains bind to histone-tails carrying acetylated lysine residues. Regulation of the nomenclature of specific histone modifications formulated at the Brno meeting of the NoE in 2004. Rules are:

<modification type>. Example: H3K4me3 ¼ trimethylated lysine-4 on histone H3. The programed path of differentiation of a cell. Although all cells have the same DNA, their cell fate can be different. For instance, some cells develop into brain, whereas others are the precursors of blood. Cell fate is determined in part by the organization of chromatin DNA and the histone proteins in the nucleus. Specific active and repressive organizations of chromatin can be maintained from one cell to its daughter cells. This is called epigenetic inheritance and ensures that specific states of gene expression are inherited over many cell generations. see chromatin immuno-precipitation. After chromatin immunoprecipitation, DNA is purified from the immunoprecipitated chromatin fraction and used to hybridize arrays of short DNA fragments representing specific regions of the genome. Sequencing of the totality of DNA fragments obtained by ChIP to determine their position on the genome. Sequencing is usually preceded by PCR amplification of ChIP-derived DNA to increase its amount. In each somatic cell generation, the genomic DNA is replicated in order to make two copies

Glossary

Chromatin

Chromatin immunoprecipitation (ChIP)

Chromatin remodeling

Chromo domain (chromatin organization modifier domain)

Chromosomal domain

CpG dinucleotide

CpG island

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of each individual chromosome. During M phase of the cell cycle, these copies called chromatids are microscopically visible one next to the other, before they get distributed to the daughter cells. The nucleoprotein complex constituting the chromosomes in eukaryotic cells. Structural organization of chromatin is complex and involves different levels of compaction. The lowest level of compaction is represented by an extended array of nucleosomes. Incubation of chromatin fragments comprising one to several nucleosomes, with an antiserum directed against particular (histone) proteins or covalent modifications on proteins. After ChIP, the genomic DNA is purified from the chromatin fragments brought down by the antiserum and analyzed. Locally, the organization and compaction of chromatin can be altered by different enzymatic machineries. This is called chromatin remodeling. Several chromatin remodeling proteins move nucleosomes along the DNA and require ATP for their action. Protein protein interaction motif first identified in Drosophila melanogaster HP1 and polycomb group proteins. Also found in other nuclear proteins involved in transcriptional silencing and heterochromatin formation. Chromo domains consist of approx. 50 amino acids and bind to histone tails that are methylated at certain lysine residues. In higher eukaryotes, it is often observed that in a specific cell type, chromatin is organized (e.g., by histone methylation) the same way across hundreds to thousands of kilobases of DNA. These “chromosomal domains” can comprise multiple genes that are similarly expressed. Some chromosomal domains are controlled by genomic imprinting. A cytosine followed by a guanine in the sequence of bases of the DNA. Cytosine methylation in mammals occurs at CpG dinucleotides. A small stretch of DNA, of several hundreds up to several kilobases in size, which is particularly

358

Cytosine methylation

Deacetylation

DNA-demethylation

“de novo” DNA-methylation

DNA-methylation

DNA methyltransferase

Dosage compensation

Embryonic stem (ES) cells

Glossary

rich in CpG dinucleotides and is also relatively enriched in cytosines and guanines. Most CpG islands comprise promoter sequences that drive the expression of genes. In mammals, DNA methylation occurs at cytosines that are part of CpG dinucleotides. As a consequence of the palindromic nature of the CpG sequence, methylation is symmetrical, i.e., it affects both strands of DNA at a methylated target site. When present at promoters, it is usually associated with transcriptional repression. The removal of acetyl groups from proteins. Deacetylation of histones is often associated with gene repression and is mediated by histone deacetylases (HDACs). Removal of methyl groups from DNA. This can occur “actively”, i.e., by an enzymatically mediated process, or “passively,” when methylation is not maintained after DNA replication. The addition of methyl groups to a stretch of DNA that is not yet methylated (acquisition of “new” DNA methylation). A biochemical modification of DNA resulting from addition of a methyl group to either adenine or cytosine bases. In mammals, methylation is essentially confined to cytosines that are in CpG dinucleotides. Methyl groups can be removed from DNA by DNA-demethylation. Enzyme that puts new (de novo) methylation onto the DNA or that maintains existing patterns of DNA methylation. The X chromosome is present in two copies in the one sex and in one copy in the other. Dosage compensation ensures that in spite of the copy number difference, X-linked genes are expressed at the same level in males and females. In mammals, dosage compensation occurs by inactivation of one of the X chromosomes in females. Cultured cells obtained from the inner cell mass of the blastocyst, and for human ES cells, possibly also from the epiblast. These cells are pluripotent; they can be differentiated into all different somatic cell lineages. ES-like cells can be obtained by dedifferentiation in vitro of somatic cells (see iPS cells).

Glossary

Endocrine disruptor

Enhancer

Epialleles

Epigenesis

Epigenetics

Epigenetic code

Epigenetic inheritance

Epigenetic marks

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A chemical component that can have an antagonistic effect on the action of a hormone (such as on estrogen) to which it resembles structurally. Some pesticides act as endocrine disruptors and have been found in animal studies to have adverse effects on development, and for some, to induce altered DNA methylation at specific loci. A well-characterized endocrine disruptor is Bisphenol-A, a chemical used for the productions of certain plastics. A small, specialized sequence of DNA, which, when recognized by specific regulatory proteins, can enhance the activity of the promoter of a gene(s) located in close vicinity. Copies of a DNA sequence or a gene that differ in their epigenetic and/or expression states without the occurrence of a genetic mutation. The development of an organism from fertilization through a sequence of steps leading to a gradual increase in complexity through differentiation of cells and formation of organs. The study of heritable changes in gene function that arise without an apparent change in the genomic DNA sequence. Epigenetic mechanisms are involved in the formation and maintenance of cell lineages during development, and, in mammals, in X-inactivation and genomic imprinting, and are frequently perturbed in diseases. Patterns of DNA methylation and histone modifications can modify the way genes on the chromosomes are expressed. This has led to the idea that combinations of epigenetic modifications can constitute a code on top of the genetic code, which modulates gene expression. The somatic inheritance, or inheritance through the germ line, of epigenetic information (changes that affect gene function, without the occurrence of an alteration in the DNA sequence). Regional modifications of DNA and chromatin proteins, including DNA methylation and histone methylation, which can be maintained from one cell generation to the next and which may affect the way genes are expressed.

360

Epigenetic reprograming

Epigenome

Epigenotype

Epimutation

Euchromatin

FISH

FLIM

FRAP

Glossary

The resetting of epigenetic marks on the genome so that these become like those of another cell type or of another developmental stage. Epigenetic reprograming occurs, for instance, in primordial germ cells, to bring them back in a “ground state.” Epigenetic reprograming and dedifferentiation also occur after somatic cell nuclear transfer. The epigenome is the overall epigenetic state of a particular cell. In the developing embryo, each cell type has a different epigenome. Epigenome maps represent the presence of DNA methylation, histone modification, and other chromatin modifications along the chromosomes. The totality of epigenetic marks that are found along the DNA sequence of the genome in a particular cell lineage or at a particular developmental stage. A change in the normal epigenetic marking of a gene or a regulatory DNA sequence (e.g., a change in DNA methylation), which affects gene expression. A type of chromatin that is lightly staining when observed through the microscope at interphase. Euchromatic chromosomal domains are loosely compacted and relatively rich in genes. The opposite type of chromatin organization is heterochromatin. Fluorescence in situ hybridization is a cytogenetic technique used to detect and localize the presence or absence of specific DNA/RNA sequences in cells. Fluorescence lifetime imaging microscopy or FLIM is a powerful tool for producing an image based on the differences in the exponential decay rate of the fluorescence from a fluorescent sample. Fluorescence recovery after photobleaching exploits the photobleaching characteristics of fluorophores for measuring the mobility of fluorescently labeled proteins at a subcellular level.

Glossary

Genomic imprinting

Germ line-specific stem cells

GV

Heterochromatin

Histone acetylation

Histone acetyltransferase (HAT) Histone code

Histone deacetylase (HDAC)

Histone-Demethylase (HDM)

361

An epigenetic phenomenon that affects a small subset of genes in the genome and results in monoallelic gene expression in a parent-oforigin-dependent way (for a given pair of alleles, uniformly either the maternally or paternally derived copy is active). Cells derived from undifferentiated germ cells, which can be maintained without alterations in their characteristics through many cell divisions. The Germinal Vesicle is the nucleus of an oocyte toward the end of prophase of its first meiotic division. A type of chromatin that is darkly staining when observed through the microscope at interphase. Heterochromatic chromosomal domains, found in all cell types, are highly compacted, rich in repeat sequences, and show little or no gene expression. Extended regions of heterochromatin are found close to centromeres and at telomeres. Posttranslational modification of the e-amino group of lysine residues in histones catalyzed by a family of enzymes called histone acetyltransferases (HATs). Acetylation contributes to the formation of decondensed, transcriptionally permissive chromatin structures and facilitates interaction with proteins containing bromo domains. An enzyme that acetylates (specific) lysine amino acids on histone proteins. Theory that distinct chromatin states of condensation and function are marked by specific histone modifications or specific combinatorial codes (see also epigenetic code). An enzyme that removes acetyl groups from histone proteins. This increases the positive charge of histones and enhances their attraction to the negatively charged phosphate groups in DNA. Proteins catalyzing the active enzymatic removal of methyl groups from either lysine or arginine residues of histones. Prominent examples are LSD1 and Jumonji proteins.

362

Histone methylation

Histone methyltransferase (HMT) Hotspot

IAP

ImmunoFISH Imprinted genes

Imprinted X-inactivation

Imprinting Imprinting control region (ICR)

In vitro fertilization (IVF)

Induced pluripotent stem cells (iPS)

Glossary

Posttranslational methylation of amino acid residues in histones catalyzed by histone methyltransferases (HMTs). Histone methylation is found at arginine as mono- or dimethylation and lysine as mono-, di-, or tri-methylation. Modifications are described depending on the position and type of methylation (mono, di, trimethylation) according to the Brno nomenclature. Different types of methylation can be found in either open transcriptionally active or silent (repressive) chromatin (histone code). Methylated lysine residues are recognized by proteins containing chromo domains. Enzymes catalyzing the transfer of methyl groups from S-adenosyl-methionine (SAM) to lysine or arginine residues in histones. Recombination hotspots are small regions in the genome of sexually reproducing organisms that exhibit highly elevated rates of meiotic recombination. Intracisternal A-Particle elements constitute a family of retrovirus-like genetic elements, which code for virus-like particles. Combines FISH with multicolor immunofluorescence detection of proteins. Genes that show a parent-of-origin-specific gene expression pattern controlled by epigenetic marks that originate from the germ line. Preferential inactivation of the paternal X-chromosome in rodents (presumably also humans) during early embryogenesis and in the placenta of mammals. See genomic imprinting. Region that shows germ line derived parent of origin dependent epigenetic marking, which controls the imprinted expression of neighboring imprinted genes. Fertilization of a surgically retrieved oocyte in the laboratory, followed by a short period of in vitro cultivation before the embryo is transferred back into the uterus to allow development to term. Cells with an ES cell like pluripotent potential derived from differentiated somatic cells by in vitro reprograming. Reprograming is triggered

Glossary

Inner cell mass (ICM)

Intracytoplasmic sperm injection (ICSI) Isoschizomers

LINE

Locus control region (LCR)

LTR

Maintenance methylation

Maternal effects

363

by the activation of pluripotency factor genes and cultivation in ES-cell medium. iPS cells are capable to generate all cell types of an embryo. Cells of the inner part of the blastocyst forming the embryo proper. Inner cell mass cells are the source for ES cells. Capillary mediated injection of a single sperm into the cytoplasm of an oocyte followed by activation to promote directed fertilization. Restriction enzymes from different bacteria which recognize the same target sequence in DNA. Often, these enzymes respond differently to methylation of bases within their target sequence, which may make them important tools in DNA-methylation analysis. Thus, MspI cuts both CCGG and C5mCGG, whereas HpaII cuts only the unmethylated sequence. Long interspersed repetitive elements or Long interspersed nuclear elements are a group of genetic elements that are found in large numbers in eukaryotic genomes. Region marked by insulator functions and DNase hypersensitive sites. LCRs contain binding sites for insulator proteins and enhancer binding proteins. LCRs control the domain specific developmentally regulated expression of genes by long range interactions with gene promoters. Long terminal repeats are sequences of DNA that are repeated hundreds or thousands of times. They are found in retroviral DNA and in retrotransposons. Process that reproduces DNA-methylation patterns between cell generations. Depends in mammals critically (though not exclusively) on the activity of the “maintenance DNA-methyltransferase” Dnmt1. This enzyme preferentially methylates hemimethylated CpG-sites, generated by replication of symmetrically methylated CpG-sequences (see Cytosine methylation), while originally unmethylated sites remain unmethylated upon replication. Long-term effects on the development of the embryo triggered by factors in the cytoplasm of the oocyte.

364

Metastable epiallele

Methyl-binding domain (MBD)

Methyl-CpG-binding proteins (MBPs)

MSCI

Noncoding RNA (ncRNA)

Non-Mendelian inheritance

Nuclear periphery

Nuclear (chromosomal) territory

Nucleolus

Nucleosome

PAR

Glossary

Loci, whose epigenetic state is particularly labile, i.e., prone to be epigenetically modified in a variable and reversible manner. As a consequence of this lability, various phenotypes may derive from genetically identical cells, resulting in phenotypic mosaicism between cells (variegation) and also between individuals (variable expressivity). Protein domain in Methyl-CpG-binding proteins (MBPs) responsible for recognizing and binding to methylated cytosine residues in DNA. Proteins containing MBDs form a specific family of proteins with various molecular functions. Proteins containing domains (such as MBD) binding to 5-methyl-cytosine in the context of CpG dinucleotides. MBPs mostly act as mediators for molecular functions such as transcriptional control or DNA repair. Meiotic sex chromosome inactivation during spermatogenesis leads to the inactivation of the X and the Y chromosomes, microscopically visible as the sex body. RNA transcripts that do not code for a protein. ncRNA generation frequently involves RNA processing. Inheritance of genetic traits that do not follow Mendelian rules and/or cannot be explained in simple mathematically modeled traits. Region around the nuclear membrane characterized by contacts of the chromosomes with the nuclear lamina. Cell type-specific areas within the nucleus occupied by specific chromosomes during interphase (G1). Specific compartments within the nucleus formed by rDNA repeat domains. Nucleoli are marked by specific heterochromatic structures and active gene expression. Fundamental organizational unit of chromatin consisting of 147 base pairs of DNA wound around a histone octamer. The pseudoautosomal regions, PAR1 and PAR2, are regions that are homologous on the X and Y chromosomes.

Glossary

PGC

Pluripotency

Polycomb group proteins

Position effect variegation (PEV)

Primordial germ cell

Protamines

RNA interference (RNAi)

365

Primordial germ cells represent the early, premeiotic stages of gametogenesis, during which male and female germ cells are comparable as concerns chromatin remodeling. Capacity of stem cells to form all cell types of an embryo including germ cells but not extraembryonic lineages (see totipotency). Epigenetic regulator proteins forming multiprotein complexes (PRCs ¼ polycomb repressive complexes). Polycomb group proteins possess enzymatic properties to control the maintenance of a suppressed state of developmentally regulated genes, mainly through histone methylation and ubiquitination. A type of clonally heritable variability of gene expression, which relies on epigenetic lability (see also metastable epialleles) associated with the particular position of a gene within the genome. PEV has first been observed in the context of gene translocations from euchromatic to heterochromatic environments and is a consequence of variable formation of heterochromatin across the respective locus. PEV may give rise to patches of cells with altered expression profiles. A classical example is represented by certain mutations in Drosophila leading to variegated eye pigmentation (“mottled eyes”). Mammalian cells set aside during early embryogenesis, which migrate through the hind gut of the developing mammalian embryo into the “Gonadenanlagen” to form founder cells of the latter germ line. Small, arginine-rich proteins that replace histones late in the haploid phase of spermatogenesis (during spermiogenesis). They are thought to be essential for sperm head condensation and DNA stabilization. After fertilization, protamines are removed from paternal chromosomes in the mammalian zygote. Posttranscriptional regulatory effects on mRNAs (control of translation or stability) triggered by processed ds and ss small RNA (si-, mi-, pi RNAs) molecules. Effects are propagated by

366

SAHA

S-adenosylhomocysteine (SAH)

S-adenosyl methionine (SAM)

SCNT

SET domain

Sex body

Silencer

SINE

siRNAs

Somatic cell nuclear transfer (SCNT)

Glossary

enzymatic complexes such as RISC containing the small RNAs bound by Argonaute proteins. Suberoylanilide hydroxamic acid, an inhibitor of certain histone deactylases, leading to enhanced levels of histone acetylation. See also TSA. Hydrolyzed product formed after the methylation reaction catalyzed by DNA- and histone methyltransferases using SAM as methyl group donor. SAH is a competitive inhibitor of SAM for most methyltransferases. A cofactor for all DNA- (DNMTs) and histonemethyltransferases (HMTs) providing the methyl group added to either cytosines (DNA) or histones (arginine or lysine). Somatic cell nuclear transfer, or cloning, is achieved by introduction of a nucleus from a somatic cell into an enucleated oocyte, which can then be triggered to initiate development. A domain found in virtually all lysine-specific histone methyltransferases (HMTs). A protein protein interaction domain required for HMT activity and modulation of chromatin structure, frequently associated with cysteine-rich PreSET and Post-SET domains. During meiosis in male germ cells, the largely unpaired sex chromosomes become inactivated by MSCI. Their compaction is microscopically visible as sex bodies. Element in the DNA to which proteins bind that inhibit transcription of a nearby promoter. Silencer elements are recognized and bound by silencer proteins. Short interspersed repetitive elements or Short interspersed nuclear elements are short DNA sequences (<500 bases). small interfering RNAs, RNAs in the size range of 21 24 nucleotides derived from double stranded long RNAs cleaved by Dicer. siRNAs are incorporated into the RISC complex to be targeted to complementary RNAs to promote cleavage of these mRNAs. Transfer of the nucleus of a somatic cell into an enucleated oocyte using a glass capillary to form

Glossary

Spermatogenesis

Spermiogenesis

Stem Cell

Totipotency

Trithorax group proteins

Trophoblast TSA Uniparental Disomy

X chromosome inactivation

367

an SCNT-zygote. After the activation of the zygote, the genome of the nucleus derived from the somatic cells become reprogramed to start development. The process by which spermatogonia develop into mature spermatozoa. Spermatozoa (sperm) are the mature male gametes. Thus, spermatogenesis is the male version of gametogenesis. The final stage of spermatogenesis that sees the maturation of spermatids into mature, motile spermatozoa (sperm). During this stage, cells no longer divide and undergo a major morphological transformation. In addition, at most of the genome, histone proteins are replaced by the more basic protamines. Noncommitted cell, which has the capacity to self renew and divide many times, giving rise to daughter cells, which maintain the stem cell function. Stem cells have the property to differentiate into specialized cells. Capacity of stem cells to produce all cell types required to form a mammalian embryo, i.e., embryonic and extraembryonic cells (see Pluripotency). Totipotent cells are formed during the first cleavages of the embryo. Proteins containing a thritorax like bromodomain: They are usually involved in recognizing histone modifications marking transcriptionally active regions and contribute to maintenance of activity. Cells of the blastoderm forming the placental tissues in mammals. Trichostatin-A, an inhibitor of certain types of histone-deacetylases. The occurrence in the cell of two copies of a chromosome, or part of a chromosome, which are identical and of the same parental origin. Epigenetically controlled form of dosage compensation in female mammals resulting in transcriptional silencing of genes on surplus X-chromosomes. X-chromosome inactivation is triggered by the noncoding RNA Xist and manifested by various epigenetic modifications including histone methylation, histone deacetylation, and DNA-methylation.

Index

A ABI, 240, 246 Achondroplasia, 57 Acridine orange (AO), 58, 59 Acrosome, 197 Advanced age, 57 Aid, 105, 110 Alignment, 245 249 Aneuploidy, 137 Angelman syndrome (AS), 4, 6 8 Aniline blue, 63 Antioxidant therapeutic strategies, 66 68 Apert syndrome, 57 Arrays, 338, 339 ART. See Assisted reproductive technology Assembly, 243, 245 Assisted reproductive technology (ART), 3 14 outcomes, 65 B Beckwith Wiedemann syndrome (BWS), 4, 6 8, 11 Bioinformatics, 246 Blastocyst, 199, 203 Blimp1, 102 104, 107 110 BLIMP1/PRMT5, 21, 31 Bouquet, 161, 162, 164, 177 C Caenorhabditis elegans, 100, 107 Carcinoma in situ, DNA methylatioin, 29 Centromeres, 262 265, 267 269, 271, 273 Checkpoint, 161, 163 165, 167 169, 171, 172 ChIP Seq, 253, 254 Chromatin, 46 51, 54 56, 58, 59, 61, 63, 69, 236 fiber, 217, 218 spermatogenesis, 215, 216

spermiogenesis, 216, 217, 222 structure, 219, 223 Chromatin remodeling, 193, 196 200, 202, 204, 205 Chromatoid body, 311 324, 336 Chromocenter, 196, 198, 267, 268 Chromodomain, 48, 51 Chromomycin A3 (CMA3), 61 66 Chromosome hairpin, 268, 269, 271 positioning, 262, 263, 269 272 territory, 263 265, 267, 269, 271, 272 Chromosome number, 49, 56, 57 Cloning, 85 94 COMET, 52, 58, 60, 64, 65 Contigs, 243 CpG methylation, 203, 204 CREM, 301 303 Crossing over (CO), 121 124, 126 133, 143 146 Cryptorchidism, 25 27, 31 D Damaged DNA, 58, 64, 65, 68, 69 Density gradients, 67 Diplotene, 158, 163, 166, 168, 169, 176 Disulfide bonds, 46 48, 50, 57, 63, 64 DNA binding proteins, 48 DNA damage, 53, 54, 57, 58, 60, 64 66, 68 DNA demethylation, 100 103, 105, 106, 110 DNA double strand break (DSB), 159, 161 165, 167 172, 176 179 DNA fragmentation, 57, 58, 60, 63 65, 68 DNA helicase, 51 DNA integrity, 46, 49, 50, 52, 53, 57, 59, 63 64, 66 69 DNA methylation, 4 6, 8 14, 46, 47, 49, 53 55, 69, 235 236

369

370 DNA methyltransferase, 48, 51 DNA recombination, 195 DNA repair, 57, 188, 194, 199 DNA replication transcription repair, 47 DNA sequence, 49, 53, 56, 57 Dnmt1, 105, 112 Dnmt3a/b, 103, 105 DNMT3L, epigenetics, 29 Dosage compensation, 174 Double strand breaks (DSB), 122, 123, 129, 133, 136, 138, 140, 141, 144, 146, 147 Drosophila, 100, 292 296, 298 E ELAND, 241, 246 Electrophoresis, 48, 50, 58, 60, 63 65, 67, 68 Elongating Spermatid, 47 Embryo culture, 13 14 Embryonic development, 20, 24, 188, 199 202, 204 Embryonic germ cells, 99 113 Endocrine, 234, 235 Endocrine disrupters phthalates, 26, 27 vinclozolin, 26, 27 Epigenetic events, 4 6 Epigenetics, 46 49, 51, 54 56 asymmetry, 188, 202 effects, 202 inheritance, 188, 202 204 Epigenome, 46 56, 69 Epigenotype, 203, 204 Error, 240, 241, 243 246, 248 249 Error rate, 241, 243, 245 Evolution, 159 160, 170 F False discovery, 251 Female chromatin, 188, 205 Fertilization, 46 48, 51, 57, 63, 65, 66, 69 Flh5, 303 G G9a, 106, 109 111 Gametes development, 5 6 epigenetic, 332, 337 female, 330 male, 330, 332 manipulation, 12 13 paramutation, 341 paternal, 330, 332, 337 Genetic map, 124 129, 131

Index Genome, 46 57, 64, 66, 67 embryonic, 330, 331, 343, 345 maternal, 330, 331 paternal, 330, 332, 336, 342, 345 Genomic imprinting, 4, 5, 7, 12, 13 GLP, 103, 106, 111 H Halo size, 60 Heritability, 234 235 Heterochromatin, 192, 196 198, 201 HiC Seq, 242 Histone methylation H3K9me2, 102, 103, 106 112 H3K27me3, 103, 108 110 H4R3me2, 103, 109 Histones, 262, 265, 267, 268, 272, 273 acetylation, 216 219 acetyltransferase, 48, 51 canonical, 188 deacetylase, 51 histone fold domain, 217 hyperacetylation, 47 linker, 192, 194, 197, 198 methyltransferase, 48, 51 modifications, 56, 69 noncanonical, 188 post translational modification (PTM), 188, 201, 204 variants, 47, 56 H3K4Me3, 136, 138, 144 147 Homeodomain, 51 Hotspot, 123, 125, 128, 130 133, 136, 143 146, 148 Human inferlity, 56 64 Hypospadias, 23, 25, 27 I Illumina, 25, 239 241, 244, 246, 252 Impaired spermatogenesis, 25 Imprinted genes, 101, 103, 110 111 microRNAs, 56 Imprinting, 32 34, 202, 203, 235 236 Imprinting defects, 5, 8 12 Incidence, 26 Infertile, 50, 53, 54, 56 58, 60 64, 66, 68, 69 Infertility, 4, 7 11, 13, 14 Informatics, 231 254 Inner cell mass (ICM), 101, 102 Intracysternal A particle (IAP), 103, 111 In vitro fertilization (IVF), 197, 203, 204 In vitro growth (IVG), 13 In vitro maturation (IVM), 13

Index J Jumonji C (JmjC) domain containing protein, 106 L Lamins, 279 286 Leptotene, 161, 162, 168, 176, 178 Leucine zipper, 48, 51 LINE1, 103, 111 M Male chromatin, 188, 200, 202, 205 Mate pair, 240, 242 244 Meiosis, 120, 129 131, 133 143, 145, 146, 149, 262 266 Meiotic sex chromosome inactivation (MSCI), 159, 170 180, 194, 195 Meiotic silencing of unsynapsed chromatin (MSUC), 159, 171 173, 175, 179, 180, 196 5 Methyl cytosine, 53 Micrococcal nuclease, 55 Micro RNA, 313, 314, 322 Mimicking techniques, 12 14 miR 17, 34 miRNAs, 34 35, 237, 247, 248 Multigenerational effects, 204 Multiple testing, 251 Mutation 3, 193 N N50, 243 Nanos3, 104, 105 Next generation sequencing (NGS), 239 241, 454 GAII, 239 243 NGS. See Next generation sequencing Non seminoma, 24, 28 33 Nuclear, 45 69 Nuclear elongation, 193, 196 198 Nuclear matrix, 47, 49, 55 Nuclear proteome, 46, 48 54 Nucleohistone, 47, 55 Nucleoprotamine, 47 49, 55 Nucleosomal configuration, 335 Nucleosome, 188, 193 202, 204 Nucleosome associated regions, 55 Nucleotorroidal, 336 O OCT3/4, 30 Oligoasthenoteratozoospermia (OAT), 196, 204 Online resources, 235 238

371 Ontology, 335 Ovarian stimulation, 7, 11 Oxidate damage, 60 Oxidative stress, 57, 60, 64, 65 P Pachytene, 161, 164 166, 168 170, 173, 176, 179, 264 266 Packaging, 48, 55, 63 Paired end, 240 244, 252 Pairing, 157 180 Peak detection, 249 250 Pericentromeric regions, 248, 249 Pgc7/stella, 100, 101 piRNAs, 237, 238, 247 Placental abnormalities, 85 94 Polycomb, 293 Population exposure, 235 Post transcriptional regulation, 313, 316 Potential epigenetic effects, 11 PRDM9, 132 134, 136, 138, 145 148 Preimplantation embryo development, 6 Primary spermatocyte, 46 Primordial germ cells epigenetic reprogramming, 21 migration, 21 specification, 21 Prmt5, 108 109 Promoters, 56 Pronucleus/Pronuclei, 46, 47, 53, 56, 188, 198, 199, 202 Protamine, 197, 198, 201, 202 evolution, 220, 222 haploinsufficiency, 224 intrinsically disordered domain, 217 phosphorylation, 219 Protamine like intrinsically disordered domain, 217 phosphorylation, 219 Protamines, 46 50, 53 57, 60 66, 68, 69, 262, 267 269, 272, 273 Proteome, 46 56, 69 Proteome/epigenome, 46 56 Proviral, retrotransposons, 343 Pseudoautosomal region (PAR), 158 160, 165, 177 R Recombination, 119 149 Repair, 159, 161 165, 167 169, 171 173, 176 179 Repetitive sequences LINE, 335 SINE, 335

372 Reproductive technologies, 86 RNA, 48, 49, 54, 56, 69 mRNAs, 33, 333 nuclear, 333 sperm, 329 346 RNA polymerase II, 103, 107 Roche, 239 241, 244, 245 Round Spermatid, 46, 47 S Sanger, 238, 247 Scaffold, 242 245 SCD. See Sperm chromatin dispersion test SCSA. See Sperm chromatin structure assay Secondary spermatocyte, 47 Seminoma, 23, 24, 28 34 Sequencing depth, 243 245 Sex body ( XY body), 194 196 Sex chromatin, 196, 197 Sex chromosomes, 192, 194 196, 266 Sheep, 85 94 Silencing, 157 180 Single end, 240 242 siRNAs. See Small interacting RNAs Small interacting RNAs (siRNAs), 237, 247, 248 Small noncoding RNAs (sncRNA), 237, 248 Small RNAs, 237 238, 240, 242, 243, 247 249, 251 sncRNA. See Small noncoding RNAs Spermatids, 263 266 Spermatogenesis, 46 47, 51, 54, 56, 57, 64, 215, 216, 220, 261 274, 279 286, 311 316, 318 320, 322 infertility, 338 normozoospermic, 339, 340 spermiogenesis, 330, 335, 336 teratozoospermic, 339 Spermatogonia, 46, 47, 57 Spermatozoa, 46, 51, 54, 56, 61, 63 66, 68 Sperm chromatin dispersion test (SCD), 58 60, 65 Sperm chromatin structure assay (SCSA), 58 60, 63, 65 Sperm DNA quality, 56 60, 68 Spermiogenesis, 46, 47, 51, 216 219, 221, 222, 224 post meiotic gene expression, 222 Sperm mediated gene transfer (SMGT), ICSI, 343

Index Sperm nuclear basic proteins, infertility, 223 Sperm nuclear proteome, 46, 48, 49 Sperm nuclear quality, 45 69 Sperm nuclei/sperm nucleus, 47 49, 55, 56, 58, 61, 63, 68 Superovulation, 12 13 Swim up, 67 Synapsis, 159, 161, 162, 164 166, 168, 170, 172, 175 177 T Telomerase, 48, 51 Telomeres, 199, 200 proteins, 265 266 Testicular cancer, 23 28, 34, 35 Testicular dysgenesis syndrome, 19 36 TFIID, 293 301 Topoisomerase, 48, 51, 56, 199 Toroidal, 46, 47, 49, 55 Transcription factors, 48, 50, 53 Transition protein (TPs), 216 221, 223 225 knockout, 221, 224 TRF2, 295 299 TUNEL, 50, 52, 57, 58, 60, 65 U UCSC Genome Browser, 252 Uhrf1, 105 V VASA, 315, 316 Visualization, 251 252 X X chromosome, 22, 31, 32, 158 160, 165, 168 172, 174 176, 178, 179 X chromosome inactivation, 101, 103, 111 X inactive specific transcript (XIST), 31 32 XY body, 165 171, 173, 177, 179 Y Y chromosome, 158 160, 162, 165, 168 171, 173, 175, 177 179 Z Zinc finger, 48, 51 Zygosity, 243, 247 Zygote, 188, 195, 196, 200 203 Zygotene, 161, 165, 168 170, 176, 177