The Mitochondrion in the Germline and Early Development (Current Topics in Developmental Biology Volume 77)

Current Topics in Developmental Biology Volume 77 Series Editor Gerald P. Schatten Director, PITTSBURGH DEVELOPMENTA...

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Current Topics in Developmental Biology

Volume 77

Series Editor Gerald P. Schatten Director, PITTSBURGH DEVELOPMENTAL CENTER Deputy Director, Magee-Women’s Research Institute Professor and Vice-Chair of Ob-Gyn Reproductive Sci. & Cell Biol.-Physiology University of Pittsburgh School of Medicine Pittsburgh, Pennsylvania 15213

Editorial Board Peter Gru¨ss Max-Planck-Institute of Biophysical Chemistry Go¨ttingen, Germany

Philip Ingham University of Sheffield, United Kingdom

Mary Lou King University of Miami, Florida

Story C. Landis National Institutes of Health National Institute of Neurological Disorders and Stroke Bethesda, Maryland

David R. McClay Duke University, Durham, North Carolina

Yoshitaka Nagahama National Institute for Basic Biology, Okazaki, Japan

Susan Strome Indiana University, Bloomington, Indiana

Virginia Walbot Stanford University, Palo Alto, California

Founding Editors A. A. Moscona Alberto Monroy

The Mitochondrion in the Germline and Early Development Current Topics in Developmental Biology Volume 77 Edited by

Justin C. St. John The Medical School The University of Birmingham Birmingham, United Kingdom

AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier

Cover Photo Credit: Compliments of Justin St. John and Emma Bowles, The University of Birmingham and Keith Campbell and Ramiro Alberio, The University of Nottingham. Cover is a 10-cell embryo generated using a goat donor cell and a sheep enucleated oocyte. It has been stained with antibody against COXI and a FITCconjugated secondary antibody (both molecular probes). The embryo was imaged with a confocal microscope (Leica Microsystems Ltd., Buckinghamshire, UK) using an X10 objective and X63 digital zoom. FITC excitation was at 488 nm and detection was between 500 and 535 nm. Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald’s Road, London WC1X 8RR, UK

This book is printed on acid-free paper. Copyright ß 2007, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (www.copyright.com), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2007 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0070-2153/2007 $35.00 Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (þ44) 1865 843830, fax: (þ44) 1865 853333, E-mail: [email protected]. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting ‘‘Support & Contact’’ then ‘‘Copyright and Permission’’ and then ‘‘Obtaining Permissions.’’ For information on all Elsevier Academic Press publications visit our Web site at www.books.elsevier.com ISBN-13: 978-0-12-373662-8 ISBN-10: 0-12-373662-5 PRINTED IN THE UNITED STATES OF AMERICA 07 08 09 10 9 8 7 6 5 4 3 2 1

Contents

Contributors Preface xiii

ix

Section I: The Mitochondrion and Gametes

1

1 The Role of the Mitochondrion in Sperm Function: Is There a Place for Oxidative Phosphorylation or Is This a Purely Glycolytic Process? ´ ´ ´ Eduardo Ruiz-Pesini, Carmen Dı´ez-Sanchez, Manuel Jose´ Lopez-P erez, and Jose´ Antonio Enrı´quez I. II. III. IV. V. VI.

Introduction 3 Spermiogenesis: Removing Everything But the Essential 4 Sperm Mitochondria Are Required for Functional Purposes Is Glycolysis Required for Sperm Motility? 9 Substrates Available for the Supply of Energy 11 Concluding Remarks 13 Acknowledgments 14 References 14

6

2 The Role of Mitochondrial Function in the Oocyte and Embryo ´ Dumollard, Michael Duchen, and John Carroll Remi I. II. III. IV. V. VI. VII. VIII. IX. X.

Introduction 22 Mitochondrial Generation and Distribution in Eggs and Embryos 25 Studying Mitochondria in Eggs and Embryos 27 Mitochondria and Energy Production in the Embryo 31 Mitochondria and Ca2þ Homeostasis in Eggs and Embryos 33 Ca2þ-Induced Mitochondrial Activity: Matching ATP Supply and Demand 34 Mitochondria and Redox Metabolism in the Embryo 34 Impact of Mitochondrial Activity on Embryonic Development 37 Mitochondria and Apoptosis in the Embryo 40 Conclusions and Perspectives 41 Acknowledgments 42 References 42 v

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3 Mitochondrial DNA in the Oocyte and the Developing Embryo Pascale May-Panloup, Marie-Franc¸oise Chretien, Yves Malthiery, and Pascal Reynier I. II. III. IV.

Introduction 52 Mitochondrial DNA Structure and Function 54 mtDNA Content in Oocytes 57 Variation of mtDNA Content and Transcripts in Early Embryonic Development 65 V. mtDNA Mutations in Oocytes and Embryos 71 VI. Conclusions and Perspectives 74 References 74

Section II: Transmission

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4 Mitochondrial DNA and the Mammalian Oocyte Eric A. Shoubridge and Timothy Wai I. II. III. IV. V. VI. VII. VIII. IX.

Introduction 88 mtDNA: Structure, Organization, Replication, and Expression Origin and Development of the Germ Line 92 Transmission of mtDNA in the Female Germ Line 95 OXPHOS, Atresia, and Oocyte Quality 98 mtDNA Copy Number in the Oocyte 99 Male Transmission of mtDNA 101 Clinical Perspectives 103 Conclusions and Future Prospects 105 Acknowledgments 106 References 106

5 Mitochondrial Disease—Its Impact, Etiology, and Pathology R. McFarland, R. W. Taylor, and D. M. Turnbull I. II. III. IV. V. VI.

Introduction 114 Mitochondrial Genetics 116 Impact 123 Etiology 129 Pathology 137 Conclusions 143 Acknowledgments 143 References 143

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6 Cybrid Models of mtDNA Disease and Transmission, from Cells to Mice Ian A. Trounce and Carl A. Pinkert I. What Can Be Learned from Cybrids About mtDNA Disease? 158 II. Animal Modeling of mtDNA Transmission and Human Disease: From Transgenesis to Transmitochondrial Cybrid Models 169 III. Summary 176 References 176

Section III: The Use of Assisted Reproductive Technologies to Regulate mtDNA Disease 185

7 The Use of Micromanipulation Methods as a Tool to Prevention of Transmission of Mutated Mitochondrial DNA Helena Fulka and Josef Fulka, Jr. I. II. III. IV.

Biological Material: Oocytes and Zygotes 188 Technical Equipment 189 Germinal Vesicle Transfer 190 Short- and Long-Term Storage of Nuclear Material and Asynchronous GV to Cytoplast Transfer 194 V. Chromosome Group Transfer 198 VI. Transfer of Pronuclei 203 VII. Conclusions 207 Acknowledgments 207 References 208

8 Difficulties and Possible Solutions in the Genetic Management of mtDNA Disease in the Preimplantation Embryo J. Poulton, P. Oakeshott, and S. Kennedy I. Heteroplasmy of mtDNA Presents Problems for Chorionic Villus Sampling 214 II. The Mitochondrial Bottleneck 217 III. Management Options Beyond CVS 220 References 223

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Section IV: The Effects of Invasive Assisted Reproductive Technologies on mtDNA Transmission 227

9 Impact of Assisted Reproductive Technologies: A Mitochondrial Perspective of Cytoplasmic Transplantation A. J. Harvey, T. C. Gibson, T. M. Quebedeaux, and C. A. Brenner I. Why Are Invasive Assisted Reproductive Technology Procedures Important? 230 II. Cytoplasmic Transfer: Animal Models That May Assist Human Infertility Therapeutics 232 III. The Mitochondrial Genome and Mitochondrial Biogenesis 233 IV. Molecular Control of Mitochondrial Function in Preimplantation Embryos 237 V. Mitochondrial Copy Number, Deletions, and Mutations 239 VI. Oocyte and Embryo Mitochondrial Localization Patterns 242 VII. Implications of Mitochondrial Heteroplasmy in Infertility, Mitochondrial Disease, and Stem Cell Therapeutics 243 References 244

10 Nuclear Transfer: Preservation of a Nuclear Genome at the Expense of Its Associated mtDNA Genome(s) Emma J. Bowles, Keith H. S. Campbell, and Justin C. St. John I. II. III. IV. V. VI.

Introduction 252 Adaptations to NT 252 Applications of NT 255 Health Problems Associated with NT 257 Possible Causes of NT Failure 258 Are All the Potential Reasons for Low Success Rates in NT Interlinked? 274 VII. Conclusions 277 References 277 Index 291 Contents of Previous Volumes

307

Contributors Numbers in parentheses indicate the pages on which the authors’ contributions begin.

Jose´ Antonio Enrı´quez (3), Departamento de Bioquı´mica, Biologı´a Molecular y Celular, Universidad de Zaragoza, 50013 Zaragoza, Spain Emma J. Bowles (251), The Mitochondrial and Reproductive Genetics Group, The Division of Medical Sciences, The Medical School, The University of Birmingham, Birmingham B15 2TT, United Kingdom C. A. Brenner (229), Department of Biology, University of New Orleans, New Orleans, Louisiana 70148; Pennington Biomedical Research Center, Baton Rouge, Louisiana 70808 Keith H. S. Campbell (251), Animal Development and Biotechnology Group, School of Biosciences, University of Nottingham, Sutton Bonington, Loughborough Leics LE12 5RD, United Kingdom John Carroll (21), Department of Physiology, University College London, London WC1E 6BT, United Kingdom Marie-Franc¸oise Chretien (51), Poˆle de Biologie, De´partement de Pathologie Cellulaire et Tissulaire, Centre Hospitalier Universitaire, F-49033 Angers, France Carmen Dı´ez-Sa´nchez (3), Departamento de Bioquı´mica, Biologı´a Molecular y Celular, Universidad de Zaragoza, 50013 Zaragoza, Spain Michael Duchen (21), Department of Physiology, University College London, London WC1E 6BT, United Kingdom Re´mi Dumollard (21), Department of Physiology, University College London, London WC1E 6BT, United Kingdom; Laboratoire de Biologie du De´veloppement, UMR 7009 CNRS/UPMC, Station Zoologique, Observatoire, 06230 Villefranche-sur-Mer, France Helena Fulka (187), Institute of Experimental Medicine, Videnska 1083, CS-142 20 Prague 10, Czech Republic Josef Fulka, Jr. (187), Center for Cell Therapy and Tissue Repair, VUZV Laboratories, Pratelstvi 815 CS-104 01 Prague 10, Czech Republic T. C. Gibson (229), Department of Biology, University of New Orleans, New Orleans, Louisiana 70148

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Contributors

A. J. Harvey (229), Department of Biology, University of New Orleans, New Orleans, Louisiana 70148; Pennington Biomedical Research Center, Baton Rouge, Louisiana 70808 Justin C. St. John (251), The Mitochondrial and Reproductive Genetics Group, The Division of Medical Sciences, The Medical School, The University of Birmingham, Birmingham B15 2TT, United Kingdom Manuel Jose´ Lo´pez-Pe´rez (3), Departamento de Bioquı´mica, Biologı´a Molecular y Celular, Universidad de Zaragoza, 50013 Zaragoza, Spain S. Kennedy (213), NuYeld Department of Obstetrics and Gynaecology, The Women’s Center, University of Oxford, Oxford OX3 9DU, United Kingdom Yves Malthiery (51), INSERM U694, F-49033 Angers, France; Poˆle de Biologie, De´partement de Biochimie et Ge´ne´tique, Centre Hospitalier Universitaire, F-49033 Angers, France Pascale May-Panloup (51), INSERM U694, F-49033 Angers, France; Poˆle de Biologie, De´partement de Pathologie Cellulaire et Tissulaire, Centre Hospitalier Universitaire, F-49033 Angers, France R. McFarland (113), Mitochondrial Research Group, School of Neurology, Neurobiology, and Psychiatry, The Medical School, University of Newcastle upon Tyne, Newcastle upon Tyne NE2 4HH, United Kingdom P. Oakeshott (213), Community Health Sciences, St. George’s University of London, London SW17 0RE, United Kingdom Carl A. Pinkert (157), Department of Pathology and Laboratory Medicine, Center for Aging and Developmental Biology, University of Rochester Medical Center, Rochester, New York 14642 J. Poulton (213), NuYeld Department of Obstetrics and Gynaecology, The Women’s Center, University of Oxford, Oxford OX3 9DU, United Kingdom T. M. Quebedeaux (229), Department of Biology, University of New Orleans, New Orleans, Louisiana 70148 Pascal Reynier (51), INSERM U694, F-49033 Angers, France; Poˆle de Biologie, De´partement de Biochimie et Ge´ne´tique, Centre Hospitalier Universitaire, F-49033 Angers, France Eduardo Ruiz-Pesini (3), Departamento de Bioquı´mica, Biologı´a Molecular y Celular, Universidad de Zaragoza, 50013 Zaragoza, Spain Eric A. Shoubridge (87), Department of Human Genetics, Montreal Neurological Institute, McGill University, Montreal, Quebec H3A 2B4, Canada

Contributors

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R. W. Taylor (113), Mitochondrial Research Group, School of Neurology, Neurobiology, and Psychiatry, The Medical School, University of Newcastle upon Tyne, Newcastle upon Tyne NE2 4HH, United Kingdom Ian A. Trounce (157), Center for Neuroscience, University of Melbourne, Victoria 3010, Australia D. M. Turnbull (113), Mitochondrial Research Group, School of Neurology, Neurobiology, and Psychiatry, The Medical School, University of Newcastle upon Tyne, Newcastle upon Tyne NE2 4HH, United Kingdom Timothy Wai (87), Department of Human Genetics, Montreal Neurological Institute, McGill University, Montreal, Quebec H3A 2B4, Canada

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Preface The transmission, segregation, and inheritance of mitochondrial DNA (mtDNA) have been increasingly investigated over the last 20 years. mtDNA is transmitted through the oocyte, embryo, and fetus to the live oVspring. However, before this mtDNA can be transmitted, the whole process is initially dependent on the quality of the oocyte prior to fertilization and the fertilizing sperm. The first section of this volume involves extensive discussion of the importance of sperm and oocyte mitochondria in the processes prior to fertilization and elicits their contributions to the fertilization process. It then discusses the role of mitochondria during preimplantation development and how mtDNA contributes to this process. It also defines how transcription and replication of this genome are tightly regulated during preimplantation embryogenesis. The second section concentrates on mtDNA transmission through the oocyte. It draws on findings from animal and cellular experimental model systems to explain how mutated mtDNA are segregated and can consequently result in severely debilitating disease or can even be fatal. The rationale for such scientific investigation is supported by a detailed account documenting and discussing the impact, etiology, and pathology of human mitochondrial disease. Section three describes in detail the several assisted reproductive techniques that have been proposed to prevent the transmission of mitochondrial disease. It also demonstrates how these techniques can be successfully tested and developed prior to potential consideration in the clinical environment. It is followed by a discussion of various screening approaches that might be utilized to determine whether a woman is at risk of transmitting a mitochondrial disease to her oVspring prior to and shortly after implantation. The final section concentrates on the problems associated with some of these sophisticated and controversial reproductive technologies. It discusses the potential that these techniques have for violating the strict unimaternal transmission of mtDNA and its highly regulated transcription and replication mechanisms. It also raises concerns about the use of such techniques for the generation of tailor‐made embryonic stem cells for patient‐specific cell therapies and suggests mechanism at the mitochondrial level for overcoming such anomalies.

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The investigators contributing to each of these sections are experienced and accomplished. They have provided a wealth of knowledge to provide an all encompassing account of mtDNA from the oocyte to the live oVspring. Justin C. St. John University of Birmingham Birmingham, United Kingdom

Section I The Mitochondrion and Gametes

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The Role of the Mitochondrion in Sperm Function: Is There a Place for Oxidative Phosphorylation or Is This a Purely Glycolytic Process? Eduardo Ruiz‐Pesini, Carmen Dı´ez‐Sa´nchez, Manuel Jose´ Lo´pez‐Pe´rez,1 and Jose´ Antonio Enrı´quez1 Departamento de Bioquı´mica, Biologı´a Molecular y Celular Universidad de Zaragoza, 50013 Zaragoza, Spain

I. Introduction II. Spermiogenesis: Removing Everything But the Essential III. Sperm Mitochondria Are Required for Functional Purposes A. OXPHOS Is a Major Pathway for Sperm Motility IV. Is Glycolysis Required for Sperm Motility? V. Substrates Available for the Supply of Energy VI. Concluding Remarks Acknowledgments References

We review here the current knowledge related to the metabolic pathways used by spermatozoa to meet their high demands for ATP. This is discussed with special emphasis on one of their key roles, motility. We believe that the controversy among glycolytic and oxidative phosphorylation supporters is artificial and, as it happens in many other cell types, the source of ATP is multiple and depends on external inputs. ß 2007, Elsevier Inc.

I. Introduction Which is the major biochemical pathway for the supply of energy that mediates sperm motility? Is this glycolysis or oxidative phosphorylation (OXPHOS)? This is a long‐lasting debate in the field of gamete research (Ford, 2006). We believe that the underlying cause of this never‐ending dispute is not the lack of suYcient body of experimental data, but that the question itself does not make sense. To understand this statement, we can slightly modify the question by substituting ‘‘sperm motility’’ by ‘‘muscle contraction.’’ 1

These two authors act together as senior authors.

Current Topics in Developmental Biology, Vol. 77 Copyright 2007, Elsevier Inc. All rights reserved.

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0070-2153/07 $35.00 DOI: 10.1016/S0070-2153(06)77001-6

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Firstly, we should be aware that the requirements for sperm motility and muscle contraction are a basal ATP supply and an adequate rate of replacement for the hydrolyzed ATP when it is needed. Following our comparison with muscle, it is well established that several overlapping metabolic pathways contribute to the supply of ATP and that the preponderance of a given pathway at a particular time point depends on a variety of parameters, amongst others: a carbon source and oxygen supply, rate of lactate clearance, and so on. In fact, most cell types are capable of both glycolysis and OXPHOS, and they rely on one or other pathway as the major ATP supplier according to external inputs. Secondly, glycolysis and OXPHOS are interdependent metabolic processes. Thus, glycolyzable substrates reach OXPHOS through glycolysis; however, for ATP production, anaerobic glycolysis is not very eYcient and because of that, it is mainly used under hypoxic conditions or when glycolytic substrates are very abundant. Whenever oxygen is not limiting and the availability of glucose is restricted, cells rely on the highly eYcient OXPHOS to cover their demands of ATP. In this chapter, we focus our attention on the following aspects. (1) Genetic and molecular programs aimed to keep and maintain OXPHOS‐competent mitochondria during diVerentiation of the gamete. (2) The available data supporting the argument that the spermatozoon is capable of degrading glucose to CO2 and H2O through glycolysis, tricarboxylic cycle, and OXPHOS. (3) The current knowledge on substrate availability (carbon source and oxygen) to spermatozoa throughout their race to reach the oocyte. (4) The evidence demonstrating that sperm motility is impaired whenever the glucose catabolic route (either glycolysis or OXPHOS) is impaired.

II. Spermiogenesis: Removing Everything But the Essential To fulfill its role, the spermatozoon requires a tightly packed haploid genome to reach the target and deliver its load. Thus, a refined diVerentiation program is implemented. Sperm maturation is therefore an amazing and exquisite exercise of economy relinquishing every structure unnecessary for the unique aim of the male gamete, namely, bringing together the paternal contribution to the maternal one. Sperm structure is simple and well defined, a head and a tail. In the head, the acrosome contains hydrolytic enzymes needed to fuse and deliver the condensed nucleus inside the ovum. The tail or flagellum includes the midpiece tightly packed with mitochondria and the principal piece, both required for the sperm motility. In support of this, three diVerent types of defects in flagellum structure have been described in decapitated spermatozoa. In two of them the mitochondrial sheath is absent or incompletely

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developed. These flagella contain a certain amount of cytoplasm in the proximal end but motility is low or nonexistent. In the third type, however, the mitochondrial sheath is perfectly formed and, the motility of these flagella is very high. Therefore, complete diVerentiation of the midpiece is required for motility (Baccetti et al., 1984; Holstein et al., 1986; Perotti et al., 1981; Toyama et al., 1995). Moreover, the tail length correlate with the fraction occupied by the midpiece as does mitochondrial volume with flagellum length or beating frequency (Cardullo and Baltz, 1991). Not surprisingly then, asthenozoospermic humans have smaller midpieces with lower number of mitochondrial gyres and anomalous organization of the organelles (Mundy et al., 1995). Notably, mitochondria are not only maintained in the fully diVerentiated spermatozoon but they also undergo substantial programed modifications throughout spermatogenesis. They modify their cellular localization, morphology, and metabolism. Thus, spermatogonial organelles are oval in shape with lamellar crista and are located around the nucleus (De Martino et al., 1979). However, mitochondria are totally condensed at the late pachytene state and a few cristae occupy most of the mitochondrial volume (De Martino et al., 1979). During this process, mitochondria elongate and divide to produce small clusters of round organelles randomly distributed in the cytoplasm (De Martino et al., 1979). Individual mitochondria are close to the plasma membrane in spermatids. Throughout spermatid maturation, mitochondria develop convolved crista, the intracrista space is reduced and some of them move to the flagellum. The remaining undiVerentiated mitochondria and the rest of cellular organelles are lost to the residual body. In testicular spermatozoa, mitochondria are wrapped around the flagellum and the cristae and matrix form a concentric system (Baradi and Rao, 1979; Cieciura and Klimek, 1988; De Martino et al., 1979). All these structural changes during spermatogenesis mirror metabolic diVerentiation. Thus, some OXPHOS subunits such as cytochrome c and subunit VIb‐2 of the cytochrome c oxidase (COX) are exclusively expressed in germinal cells (Hess et al., 1993; Huttemann et al., 2003). Moreover, COX activity is higher in spermatozoa than in pachytene spermatocyte (De Martino et al., 1979) and, as expected, mitochondrial membrane potential increases by up to sixfold throughout spermatogenesis because this potential is dependent on OXPHOS activity (Petit et al., 1995; Saunders et al., 1993). These observations contrast with those from erythrocyte diVerentiation. These cells do not require mitochondria for their functional role. Therefore, they simply eliminate them. Then, why does a cell that dramatically reduces its volume by removing any superfluous structure retain very specialized mitochondria? Mitochondria contain their own genome, the mitochondrial DNA (mtDNA), which encodes essential genes for OXPHOS function

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(Enriquez et al., 1999). Therefore, it is possible that spermatozoa retain their mitochondria to deliver a full set of male genetic information, including mtDNA, to the oocyte. However, it has been demonstrated that mitochondria and their mtDNAs are committed to degradation within the zygote or embryos (Sutovsky et al., 1999). In normal circumstances, mtDNA is only maternally inherited (Giles et al., 1980). Therefore, delivering mtDNA to the oocyte is not the purpose for retaining mitochondria in sperm cells.

III. Sperm Mitochondria Are Required for Functional Purposes Mitochondria host critical metabolic pathways. They regulate the levels of important intracellular second messengers, such as calcium and reactive oxygen species, and they are also involved in the control of apoptosis (Kroemer et al., 1997). Other important mitochondrial biochemical pathways are the Krebs cycle and the OXPHOS system. These biochemical pathways are critical for the integration of cell metabolism. Thus, they connect sugar, fatty acid, and amino acid catabolism with anabolic pathways like pyrimidine synthesis, urea cycle, and so on. However, spermatozoa are terminal cells and mitochondrial anabolic pathways are likely not very relevant. OXPHOS is not only 15 times more eYcient than anaerobic glycolysis for ATP production but it also increases the number and variety of suitable substrates. Therefore, considering the eYciency of the energy supply, OXPHOS as a support for sperm motility would be the best explanation for understanding the strong investment in mitochondria during spermatogenesis.

A. OXPHOS Is a Major Pathway for Sperm Motility Analysis of the implications for OXPHOS in the supply of energy for sperm motility is straightforward. First, there is a great variety of OXPHOS‐ specific inhibitors and their eVect on flagellum activity and ultimately on sperm motility is easy to evaluate. Second, OXPHOS biogenesis depends on the maintenance and expression of mtDNA as well as well‐defined nuclear‐ encoded genes (Fernandez‐Silva et al., 2003). Then, genetic approaches are also attainable. Mitochondrial activity correlates with sperm motility (Gopalkrishnan et al., 1995). Thus, normozoospermic individuals show higher sperm motility and vitality and also a higher mitochondrial activity than oligozoospermic individuals. Motility and sperm concentration are improved when mitochondrial

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membrane potential (D m) is high (Donnelly et al., 2000; Marchetti et al., 2002; Troiano et al., 1998). Moreover, a high D m increases the in vitro fertilization (IVF) rate. These results suggest that in humans, the analysis of mitochondrial D m is the most sensitive test to determine sperm quality and that sperm motility is associated with the functional status of mitochondria (Marchetti et al., 2002). Similar results have also been obtained for other species such as rat or ram (Auger et al., 1989, 1993; Evenson et al., 1982; Gravance et al., 2001; Kramer et al., 1993; Ronot and Auger, 1990; Windsor, 1997). Indirect biochemical analysis also provides strong support for the dependence of sperm motility on OXPHOS. Oxygen consumption and sperm motility correlate (Ford and Harrison, 1981; Halangk et al., 1990). This correlation was also found in cells with modified metabolism by using metabolic inhibitors (Halangk and Bohnensack, 1986). Cyclic nucleotides and phosphodiesterase inhibitors stimulate respiration and increase sperm motility (Garbers et al., 1971). On the other hand, the higher transmission rate of mouse spermatozoa with the mutant allele tn, an allele of the T locus transmitted in a non‐Mendelian frequency, is accompanied with a lower NADH/NADþ ratio, an increase in oxygen consumption and higher motility (Ginsberg and Hillman, 1974). Mitochondrial D m and oxygen consumption depend on the activity of the mitochondrial electron transport chain (ETC) and the ATP synthase (both constituting the OXPHOS system). The four respiratory complexes (I–IV) form the ETC. The activity of all four respiratory complexes is higher when sperm motility increases (Ruiz‐Pesini et al., 1998). Moreover, the use of drugs that specifically inhibit individual respiratory complexes dramatically decreases sperm motility. In particular, rotenone, an inhibitor of respiratory complex I depleted ATP and decreased sperm motility in humans (de Lamirande and Gagnon, 1992; Halangk et al., 1985a; Rikmenspoel, 1965; Ruiz‐Pesini et al., 2000). Rotenone, but not glycolytic inhibitors, reduces the fertility of cervically inseminated sheep, suggesting that mitochondrial respiration plays an important role in the penetration of the cervix by the spermatozoa (Windsor, 1997). Antimycin A, an inhibitor of the respiratory complex III, promotes similar eVects on sperm motility (Ford and Harrison, 1981; Krzyzosiak et al., 1999; Ruiz‐Pesini et al., 2000). Nitric oxide (NO) by acting on ferroproteins of the ETC is able to decrease ATP levels and aVect sperm motility (McKinney et al., 1995; Weinberg et al., 1995). Inhibition of complex IV activity either by NO (Brown, 1995) or by KCN strongly reduces sperm motility (Halangk and Bohnensack, 1986; Pascual et al., 1996; Ruiz‐Pesini et al., 2000). Gossypol that inhibits respiration also decreases sperm motility (Breitbart et al., 1989; Kim et al., 1984). To complete the picture, oligomycin, a drug that directly blocks ATP synthesis by mitochondria, negatively aVects sperm motility (Dreanno et al., 1999; Halangk et al., 1985a).

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Four of the five enzymatic complexes that constitute the OXPHOS system are partially encoded by mtDNA. Thus, mutations in mtDNA genes that impair the expression of one or more proteins encoded in the mtDNA can promote diseases in humans. Since these diseases are very often severe, not very much attention has been paid to the sperm quality of patients with mitochondrial diseases. Systematic analysis on patient fertility has allowed reporting that a single deletion (Kao et al., 1995) or multiple deletions within the mtDNA are associated with diminished fertility and motility of human sperm. Moreover, dysfunction in the proper adjustment of the sperm mtDNA level either in terms of depletion (Kao et al., 2004) or excess amounts (Diez‐Sanchez et al., 2003a; May‐Panloup et al., 2003) seems to be detrimental to motility. Folgero et al. (1993) investigated the quality of sperm in a patient suVering from mitochondrial encephalopathy with lactic acidosis and stroke (MELAS) syndrome who harbored the mtDNA A3243G mutation. They were able to demonstrate reduced sperm motility in this patient (Folgero et al., 1993). This observation was corroborated in a second MELAS patient (Spiropoulos et al., 2002). More interestingly, it has been shown that some human mtDNA population variants can influence sperm motility (Montiel‐Sosa et al., 2006; Ruiz‐Pesini et al., 2000). We have previously proposed that some of these variants could decrease OXPHOS coupling eYciency, and subsequently aVect sperm motility (Ruiz‐Pesini et al., 2004). Genetically modified mice also illustrate the relevance of OXPHOS to sperm motility. Thus, a testis‐specific cytochrome c null mouse has been generated (Narisawa et al., 2002). These mice produce properly diVerentiated spermatozoa. Unfortunately the lack of OXPHOS activity in these spermatozoa was not demonstrated. Despite that, and when compared to wild‐type sperm, the proportion of immotile spermatozoa is greater, more than 50%. This would categorize such mice as asthenozoospermic when assessed according to human semen analysis criteria (WHO, 1999). Moreover, sperm from mutant mice contain reduced level of ATP and showed a fourfold decrease eYciency for IVF assays (Narisawa et al., 2002). Interestingly, despite their evident impairment in motility, these males are still fertile. To properly analysis this, we have to keep in mind that fertilization in laboratory animals is assessed without competition between males (one single male with several females) and that mouse females are naturally highly fertile, with multiple ovulations. In this way, conditions that result in a subfertile or infertile human couple would still allow fertile mice. Unfortunately, no estimation of the in vivo fertility eYciency (as average size of the litters, rate of pregnancy failures after copulation, and so on) for these mice is available. A second mouse model provides additional insights into the role of mitochondria in sperm function. Trifunovic et al. (2004) developed a homozygous knock‐in in mouse that expresses a proofreading‐deficient version of polymerase gamma (POLG), the nuclear‐encoded catalytic subunit of mtDNA‐specific

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polymerase. These animals manifest an mtDNA mutator phenotype with a substantial increase in the level of mtDNA mutations as well as deletions. They develop and grow normally up to the age of 25 weeks. After that an impressive phenotype of premature aging is expressed. Very interestingly, a reduction in fertility of mtDNA‐mutator mice of both sexes was found. However, all young females were fertile and became prematurely infertile probably as part of the general premature aging phenotype. To the contrary, young and still healthy males were virtually infertile, only 1 litter from 8 males bred with 16 wild‐type females (Trifunovic et al., 2004). In humans, POLG has been proposed to have a role in male infertility (Jensen et al., 2004; Rovio et al., 2001), although this association has been questioned by others (Aknin‐Seifer et al., 2005; Brusco et al., 2006). All theses observations indicate that pathological genetic alterations of OXPHOS‐related genes impair sperm quality and particularly motility.

IV. Is Glycolysis Required for Sperm Motility? Recent reports present evidence that glycolysis is needed for sperm motility. However, since ATP production from glucose involves both glycolysis and OXPHOS, it is necessary to establish whether the ATP needed for sperm motility can have a pure glycolytic origin (fermentation). This has been proposed because: (1) the absence of mitochondria at the end of the sperm tail and the potential diYculty in meeting satisfactorily flagellum ATP demands at the tail and (2) some published observations interpreting their findings as evidence that OXPHOS is not required for sperm motility. The delivery of ATP at the appropriate rate to the dynein ATPase along the flagellum is a logistic problem that it is not fully understood. It has been proposed that physical forces that originate cytoplasmic waves may facilitate the ATP supply to the end of tail. These forces would be derived from the flagellum’s movement itself. In addition, specific metabolic shuttles such as adenylate kinase and phosphoglycerate kinase would contribute to the ATP supply (Ford, 2006). Both mechanical and biochemical contributions will promote rapid diVusion of mitochondrial ATP from the midpiece to the tail. Miki et al. (2004) provide the strongest evidence for the importance of glycolysis for sperm motility. They generated a knockout mouse for the sperm‐ specific isoform of glyceraldehyde 3‐phosphate dehydrogenase (GAPDHs), a glycolytic enzyme. Homozygote knockout males were infertile and had profound defects in sperm motility, exhibiting sluggish movement without forward progression (Miki et al., 2004). Motility was not abolished by the absence of glycolysis, only about 3% of sperm showed progressive motility immediately after removal from the epididymis, and this was not maintained after 2 hours incubation. However, up to 60% of spermatozoon from GAPDHs/ mice

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kept some nonprogressive movement even after 4 hours incubation. Very interestingly, the authors highlighted a characteristic flagella bending in the middle piece that was not propagated eVectively along the principal piece of the sperm tail. In other words, flagella beating seemed to be initiated at the proximal tail but was not propagated along the flagellum (Miki et al., 2004). The GAPDHs/ mice showed another striking characteristic, a dramatically low level of ATP (10% of control). This was attributed solely to the lack of ATP synthesis by glycolysis and as evidence suggesting that ATP is not produced by sperm mitochondria. However, GAPDHs/ spermatozoa do maintain the glycolytic reactions that consume ATP. In fact, sperm from mutant mice use ATP to phosphorylate glucose and fructose 6‐phosphate in the first steps of glycolysis and they finally accumulate glyceraldehyde 3‐phosphate (G3P) by up to fourfold more than controls (Miki et al., 2004). If ATP in spermatozoa is solely produced by glycolysis, what is the source of ATP that generates such abnormal accumulation of G3P? Consequently, an additional source of ATP other than glycolysis has to be used and this source should be OXPHOS. Since GAPDHs and the remaining glycolytic enzymes are located at the fibrous sheath of the flagellum (Bunch et al., 1998) away from mitochondria, the accumulation of G3P also indicates that mitochondrial ATP should diVuse toward the distal part of the flagellum. Interestingly, vanadate, a dynein ATPase inhibitor aVects sperm motility and decreases mitochondrial respiration (Halangk et al., 1985b). ADP is the major stimulator of OXPHOS activity. Therefore, inhibition of the dynein ATPase would decrease ADP production and subsequently mitochondrial stimulation, meaning this ADP would return to the mitochondria and would not be consumed in the flagellum by glycolysis. Therefore, these mutant mice transform glycolysis from an ATP‐generating pathway to an ATP‐consuming one (Fig. 1). As a result, glycolysis competes against dynein ATPase for the ATP, contributing severely to the motility defect. In fact, the incubation of spermatozoa from GAPDHs/ in the presence of glucose further reduces ATP concentrations from 10% to 1.9% and eliminates any trace of progressive spermatozoa (Miki et al., 2004). Spermatazoa from many species including human can remain motile in glucose‐free media. Interestingly, by using an inhibitor of the GAPDHs, it was found that mammalian spermatozoa were immotile in the presence of glucose but they were motile when glucose was replaced by respiratory substrates (Ford, 2006). A number of recent papers suggest spermatozoa might be capable of gluconeogenesis and synthesis of glycogen to allow cell motility. Motile spermatozoa consume energy and it would be contradictory that opposite pathways, one catabolic (glycolysis) and the other anabolic (gluconeogenesis and glycogen synthesis), worked simultaneously in the same cell compartment

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Figure 1 Glycolysis. The glycolytic pathway contains two phases: one consumes ATP and the other produces ATP. GAPDHs knockout mouse only maintain the ATP‐consuming phase.

and using the same substrate. Moreover, gluconeogenesis consumes three times more ATP than that produced by glycolysis (Pilkis and Granner, 1992). It would be absurd to activate gluconeogenesis to produce glucose for use in glycolysis. However, it would probably be beneficial to activate gluconeogenesis in those scenarios where enough energy obtained from respiration is used in sperm motility and residual levels would be used for energy storage.

V. Substrates Available for the Supply of Energy A major issue in defining the metabolic process responsible for the ATP production for sperm motility is the availability of the proper substrate. This is very often overlooked in the models proposed to explain the variety of observations available. Thus, glycolytic substrates, respiratory substrates, or both are required to be available for spermatozoa in suYcient amounts and at the right time. Respiratory substrates and oxygen levels (Max, 1992) are

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A

1.6–21.0 H 9.7 S 7.1–10.0 M Oviduct

B 60 mmHg Oviduct

Uterus 40 mmHg

Ovarium

Uterus 1.9 H 15.4 M

Endocervix

Endocervix Vagina 150 mmHg Blood 40–100 mmHg

Ovarium

Vagina Plasma 0.2 H 0.5 S 0.4 M

Figure 2 Oxygen, lactate, and glucose levels in the female reproductive tract. (A) Oxygen levels. (B) Lactate/glucose ratio for diVerent anatomical portions of the female reproductive tract: human (H) (Dickens et al., 1995; Gardner et al., 1996; Tay et al., 1997), swine (S) (Nichol et al., 1992), and mouse (M) (Harris et al., 2005). Lactate is also richer in endocervix and vagina, where glycolyzable sugar levels are poor (Jones, 1998; Windsor, 1997). Lactate concentration is much higher than that in blood plasma but glucose levels are lower.

abundant along male and female reproductive tracts to support OXPHOS activity (Fig. 2). The stimulation of Sertoli cells by follicle stimulating hormone (FSH) produces an increase in pyruvate and lactate secretion to the seminiferous epithelium lumen (Jutte et al., 1983; Sylvester and Griswold, 1994) and lactate is the central energy metabolite used by germ cells (Grootegoed et al., 1984; Mita et al., 1982). RNA and protein syntheses, in addition to oxygen consumption of isolated spermatocytes and spermatids, are stimulated by exogenous lactate but not by glucose (Boussouar and Benahmed, 2004). Spermatids possess all enzyme activities for the glycolytic pathway. However, glucose metabolism cannot maintain cellular ATP content, and exposure of isolated spermatids to glucose without other energy substrates soon results in ATP depletion (Grootegoed et al., 1984; Mita et al., 1982). The use of lactate as the main energetic substrate during spermatogenesis might be a mechanism to check a potential deficiency of OXPHOS activity when germ cells are still able to respond because mature spermatozoa are transcriptionally inert (Diez‐Sanchez et al., 2003b). Moreover, the mammalian epididymis is rich in lactate but reducing sugars are not present in the epididymal lumen (Jones and Murdoch, 1996). Seminal plasma rich in fructose stays in the vagina and only spermatozoa reach the cervical mucus (Haas and Beer, 1986). Lactobacillus has long been

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considered the protective flora in the vagina. Lactic acid produced by this microorganism decreases vaginal pH and acts against vaginal pathogens. Therefore, the vagina is enriched in lactate. Interestingly, in diabetic women, an association has been found between glycemia and prevalence of fungi (Nowakowska et al., 2004). Thus, the levels of glycolyzable sugars are probably low in vagina. Considering the rest of the female reproductive tract, it has been shown that the ewe cervix and uterine washings of swine are poor in glucose and glycolyzable substrates (Jones, 1998; Windsor, 1997). Lactate levels are 15‐, 10‐, and 38‐fold higher than glucose in mouse uterine, oviduct, and follicular fluids, respectively (Harris et al., 2005) and bovine oviductal fluid contains less than 100 mM glucose (Galantino‐Homer et al., 2004). All this evidence supports an important role for lactate, a respiratory substrate, as an important energy source for spermatozoa. In agreement with this, there is a testis‐specific lactate dehydrogenase isozyme [lactate dehydrogenase C4 (LDH‐C4)]. This is first expressed at the beginning of spermatogenic activity in gametogenic cells from preleptotene spermatocytes up to spermatids. It represents the predominant form of LDH in mature spermatozoa and has a dual localization, in the cytosol of spermatocytes, spermatids, and spermatozoa, as well as in the matrix of sperm‐type mitochondria (Burgos et al., 1995). Moreover, there is a correlation between LDH‐C4, the number of motile spermatozoa, and mitochondrial activity (Sawane et al., 2002). It is lactate, rather than pyruvate, which enters the mitochondria thereby constituting a lactate–pyruvate transport system in these cells for regenerating cytoplasmic nicotinamide adenine dinucleotide (Jones, 1997). Another sperm‐specific isoenzyme downstream of the glycolytic pathway is pyruvate dehydrogenase (PDH). Malate increases markedly sperm PDH activity. The diVerential sensitivity to L‐malate appears to be a particular regulatory property of the PDH complex in gametes (Gerez de Burgos et al., 1994). Very interestingly, acetoacetate and ‐hydroxybutyrate stimulate sperm motility. Glycolysis inhibitors stop the motility of sperm mediated by glucose but not by ketone bodies (Tanaka et al., 2004). Moreover, a novel mitochondrial succinyl CoA transferase (SCOT‐t), required for ketone body metabolism, has been described as being specifically expressed in testicular germ cells and spermatozoa and substitutes for the somatic cell isoform (Tanaka et al., 2002). It should be noted that ketone bodies can only be consumed by the OXPHOS system.

VI. Concluding Remarks Ejaculated spermatozoa are not under the same homeostatic regulation as the common cell in the body. When ejaculated, they have to survive alone with their own baggage and they have to maximize their chances to fulfill their

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function, to reach the ovum. The available evidence indicates that spermatozoa are set to take advantage of a wide‐ranging and eYcient cohort of metabolic options to survive under diVerent energy sources with OXPHOS being a prominent one. This versatility is critical to ensure fertilization success.

Acknowledgments We would like to thank Santiago Morales for his technical assistance. Our work was supported by the Spanish Ministry of Education (SAF2003‐00103), the Instituto de Salud Carlos III (REDMitoEspan˜a‐G03/011, REDEMETH‐G03/054, REDCIEN C03/06‐Grupo RC‐N34‐3, and Research Project FIS‐PI‐050647), the EU (EUMITOCOMBAT‐LSHM‐CT‐2004‐503116), and by the Diputacio´n General de Arago´n (Grupo de Excelencia DGA‐B55 and Grupo consolidado DGA‐B33 and PM‐078/2006).

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Perotti, M. E., Giarola, A., and Gioria, M. (1981). Ultrastructural study of the decapitated sperm defect in an infertile man. J. Reprod. Fertil. 63, 543–549. Petit, J. M., Ratinaud, M. H., Cordelli, E., Spano, M., and Julien, R. (1995). Mouse testis cell sorting according to DNA and mitochondrial changes during spermatogenesis. Cytometry 19, 304–312. Pilkis, S. J., and Granner, D. K. (1992). Molecular physiology of the regulation of hepatic gluconeogenesis and glycolysis. Annu. Rev. Physiol. 54, 885–909. Rikmenspoel, R. (1965). The inhibition by amytal of respiration and motility of bull spermatozoa. Exp. Cell Res. 37, 312–326. Ronot, X., and Auger, J. (1990). Flow and image cytometry for quality assessment of fresh and frozen human sperm samples. Anal. Cell. Pathol. 2, 313–322. Rovio, A. T., Marchington, D. R., Donat, S., Schuppe, H. C., Abel, J., Fritsche, E., Elliott, D. J., Laippala, P., Ahola, A. L., McNay, D., Harrison, R. F., Hughes, B., et al. (2001). Mutations at the mitochondrial DNA polymerase (POLG) locus associated with male infertility. Nat. Genet. 29, 261–262. Ruiz‐Pesini, E., Diez, C., Lapena, A. C., Perez‐Martos, A., Montoya, J., Alvarez, E., Arenas, J., and Lopez‐Perez, M. J. (1998). Correlation of sperm motility with mitochondrial enzymatic activities. Clin. Chem. 44, 1616–1620. Ruiz‐Pesini, E., Lapena, A. C., Diez‐Sanchez, C., Perez‐Martos, A., Montoya, J., Alvarez, E., Diaz, M., Urries, A., Montoro, L., Lopez‐Perez, M. J., and Enriquez, J. A. (2000). Human mtDNA haplogroups associated with high or reduced spermatozoa motility. Am. J. Hum. Genet. 67, 682–696. Ruiz‐Pesini, E., Mishmar, D., Brandon, M., Procaccio, V., and Wallace, D. C. (2004). EVects of purifying and adaptive selection on regional variation in human mtDNA. Science 303, 223–226. Saunders, P. T., Millar, M. R., West, A. P., and Sharpe, R. M. (1993). Mitochondrial cytochrome C oxidase II messenger ribonucleic acid is expressed in pachytene spermatocytes at high levels and in a stage‐dependent manner during spermatogenesis in the rat. Biol. Reprod. 48, 57–67. Sawane, M. V., Kaore, S. B., Gaikwad, R. D., Patil, P. M., Patankar, S. S., and Deshkar, A. M. (2002). Seminal LDH‐C4 isoenzyme and sperm mitochondrial activity: A study in male partners of infertile couples. Indian J. Med. Sci. 56, 560–566. Spiropoulos, J., Turnbull, D. M., and Chinnery, P. F. (2002). Can mitochondrial DNA mutations cause sperm dysfunction? Mol. Hum. Reprod. 8, 719–721. Sutovsky, P., Moreno, R. D., Ramalho‐Santos, J., Dominko, T., Simerly, C., and Schatten, G. (1999). Ubiquitin tag for sperm mitochondria. Nature 402, 371–372. Sylvester, S. R., and Griswold, M. D. (1994). The testicular iron shuttle: A ‘‘nurse’’ function of the Sertoli cells J. Androl. 15, 381–385. Tanaka, H., Kohroki, J., Iguchi, N., Onishi, M., and Nishimune, Y. (2002). Cloning and characterization of a human orthologue of testis‐specific succinyl CoA: 3‐Oxo acid CoA transferase (Scot‐t) cDNA. Mol. Hum. Reprod. 8, 16–23. Tanaka, H., Takahashi, T., Iguchi, N., Kitamura, K., Miyagawa, Y., Tsujimura, A., Matsumiya, K., Okuyama, A., and Nishimune, Y. (2004). Ketone bodies could support the motility but not the acrosome reaction of mouse sperm. Int. J. Androl. 27, 172–177. Tay, J. I., Rutherford, A. J., Killick, S. R., Maguiness, S. D., Partridge, R. J., and Leese, H. J. (1997). Human tubal fluid: Production, nutrient composition and response to adrenergic agents. Hum. Reprod. 12, 2451–2456. Toyama, Y., Kazama, T., Fuse, H., and Katayama, T. (1995). A case of decapitated spermatozoa in an infertile man. Andrologia 27, 165–170. Trifunovic, A., Wredenberg, A., Falkenberg, M., Spelbrink, J. N., Rovio, A. T., Bruder, C. E., Bohlooly, Y. M., Gidlof, S., Oldfors, A., Wibom, R., Tornell, J., Jacobs, H. T., et al. (2004).

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Premature ageing in mice expressing defective mitochondrial DNA polymerase. Nature 429, 417–423. Troiano, L., Granata, A. R., Cossarizza, A., Kalashnikova, G., Bianchi, R., Pini, G., Tropea, F., Carani, C., and Franceschi, C. (1998). Mitochondrial membrane potential and DNA stainability in human sperm cells: A flow cytometry analysis with implications for male infertility. Exp. Cell Res. 241, 384–393. Weinberg, J. B., Doty, E., Bonaventura, J., and Haney, A. F. (1995). Nitric oxide inhibition of human sperm motility. Fertil. Steril. 64, 408–413. Windsor, D. P. (1997). Mitochondrial function and ram sperm fertility. Reprod. Fertil. Dev. 9, 279–284. WHO (1999). ‘‘World Health Organization Laboratory Manual for the Examination of Human Semen and Semen‐Cervical Mucus Interaction,’’ p. 138. The Press Syndicate Cambridge, Cambridge.

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The Role of Mitochondrial Function in the Oocyte and Embryo Re´mi Dumollard,*,{ Michael Duchen,* and John Carroll* *Department of Physiology, University College London London WC1E 6BT, United Kingdom { Laboratoire de Biologie du De´veloppement, UMR 7009 CNRS/UPMC Station Zoologique, Observatoire, 06230 Villefranche‐sur‐Mer, France

I. II. III. IV. V. VI. VII. VIII. IX. X.

Introduction Mitochondrial Generation and Distribution in Eggs and Embryos Studying Mitochondria in Eggs and Embryos Mitochondria and Energy Production in the Embryo Mitochondria and Ca2þ Homeostasis in Eggs and Embryos Ca2þ‐Induced Mitochondrial Activity: Matching ATP Supply and Demand Mitochondria and Redox Metabolism in the Embryo Impact of Mitochondrial Activity on Embryonic Development Mitochondria and Apoptosis in the Embryo Conclusions and Perspectives Acknowledgments References

Mitochondria have long been known to be the powerhouses of the cell but they also contribute to redox and Ca2þ homeostasis, provide intermediary metabolites and store proapoptotic factors. Mitochondria have a unique behavior during development. They are maternally transmitted with little (if any) paternal contribution, and they originate from a restricted founder population, which is amplified during oogenesis. Then, having established the full complement of mitochondria in the fully grown oocyte, there is no further increase of the mitochondrial population during early development. The localization of mitochondria in the egg during maturation and their segregation to blastomeres in the cleaving embryo are strictly regulated. Gradients in the distribution of mitochondria present in the egg have the potential to give rise to blastomeres receiving diVerent numbers of mitochondria. Such maternally inherited diVerences in mitochondrial distribution are thought to play roles in defining the long‐term viability of the blastomere in some cases and embryonic axes and patterning in others. Mitochondria may also regulate development by a number of other means, including modulating Ca2þ signaling, and the production of ATP, reactive oxygen species, and intermediary metabolites. If the participation of mitochondria in the Current Topics in Developmental Biology, Vol. 77 Copyright 2007, Elsevier Inc. All rights reserved.

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0070-2153/07 $35.00 DOI: 10.1016/S0070-2153(06)77002-8

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regulation of sperm‐triggered Ca oscillations is now well established, the role of other properties of mitochondrial function during development remain largely unexplored probably due to the diYculty of accessing the mitochondrial compartment in an embryo. Maintaining a functional complement of maternally derived mitochondria is vital for the early embryo. Mitochondrial dysfunction may not only compromise developmental processes but also trigger apoptosis in the embryo. This dual role for mitochondria (to maintain life or to commit to cell death) may well represent a quality control system in the early embryo that will determine whether the embryo proceeds further into development or is quickly eliminated. ß 2007, Elsevier Inc.

Abbreviations ATP: adenosine trisphosphate; ½Ca2þ c : cytosolic Ca2þ concentration; ½Ca2þ m : mitochondrial Ca2þ concentration; Cn: cyanide; ER: endoplasmic reticulum; GFP: green fluorescent protein; GSH: reduced glutathione; GV: germinal vesicle; H2O2: hydrogen peroxide; IICR: IP3‐induced Ca2þ release; IP3: inositol 3,4,5 trisphosphate; IRP: intracellular redox potential; mtDNA: mitochondrial DNA; NADPH: reduced nicotinamide adenine dinucleotide phosphate; O2: oxygen; O2: superoxide; ROS: reactive oxygen species.

I. Introduction Long known to be the ‘‘powerhouses’’ of the cell, mitochondria are now understood to be central to diverse cellular functions. The complexity and pervasiveness of mitochondrial activity is reflected in their contribution to diverse signaling pathways and intracellular processes. In fact, in addition to producing most of the cell’s ATP (Ernster and Schatz, 1981), mitochondria can sequester and release Ca2þ, cytochrome c, and proteins (Duchen, 2000; Kroemer, 2003). They also produce reactive oxygen species (ROS), reducing equivalents [such as NAD(P)H/NAD(P)þ] and intracellular metabolites (such as Krebs cycle intermediates) (Brookes et al., 2004; Duchen, 2000; MacDonald et al., 2005; Pagliarini and Dixon, 2006; Turrens, 2003). These mitochondrial activities ensure that mitochondria play a central role in cellular processes such as Ca2þ handling and Ca2þ signaling, the regulation of intracellular redox potential (IRP), and control of apoptosis, and they may well be the mediators of cellular and organismal aging (Balaban et al., 2005). One peculiarity of mitochondria in development is that they are inherited maternally and independently of the nuclear genome (Cummins, 2000; Dawid and Blackler, 1972; Jansen, 2000; Shoubridge, 2000). Strikingly, the growth of the mitochondrial population in the embryo is discontinuous with

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a growing burst during oogenesis followed by an arrest of replication during cleavage stages until mitochondrial replication resumes after gastrulation (Dumollard et al., 2006a and references therein). Thus, embryonic mitochondria all originate from a restricted founder population present in the primordial germ cell, which is amplified during oogenesis. This pattern of mitochondrial replication provides a genetic ‘‘bottleneck’’ ensuring a homoplasmic population of mitochondria in the embryo even though mitochondrial DNA is prone to mutations (reviewed by Shoubridge and Wai in Chapter 4, this volume). Such homoplasmy is vital for the survival of each blastomere of the cleaving embryo, which depends on inheriting a functional complement of mitochondria from the egg. Another consequence of such segregation of mitochondria during cleavage is that any heterogeneity in the distribution of mitochondria in the mature egg will be maintained during early development to give rise to blastomeres with diVerent mitochondrial load, thereby potentially conferring a diVerent developmental fate. An extreme example of this phenomenon is found in the determination of germ cells. Oocytes of many organisms (amphibians, fishes, insects, planarians, chaetognaths, nematodes, and some mammals) possess a cytoplasmic structure rich in mitochondria (called the germ plasm or nuage or Balbiani body or pole plasm) that segregates with the germ line and is necessary for its specification (Kloc et al., 2004). The mitochondria of the germ plasm are selected during oogenesis and aggregate with germ line determinants before they are relocated in the vegetal cortex of the egg (reviewed in Dumollard et al., 2006a; Kloc et al., 2004). Other eggs and embryos display gradients in the distribution of somatic mitochondria that may impact embryo patterning and such gradients may well be maternal factors influencing embryonic cell fate. The activity of mitochondria in early mammalian embryos has been under investigation for many years. Oocytes and embryos have a relatively low oxygen consumption and electron microscopy shows that embryonic mitochondria have fewer and shorter cristae than mitochondria from metabolically active cells found in adult tissues (Trimarchi et al., 2000 and references therein). These observations led to the hypothesis that embryonic mitochondria are immature and with little capacity for respiratory activity (Houghton and Leese, 2004; Trimarchi et al., 2000). However, the pattern of consumption of energetic substrates seen in mammalian embryos (Biggers et al., 1967; Brinster, 1965; Johnson et al., 2003; Quinn and Wales, 1973) as well as pharmacological approaches undertaken on mammalian and ascidian embryos (Dumollard et al., 2003, 2004, 2006a) suggest that mitochondrial activity is crucial for the activation of development and for embryonic survival. Indeed, even though each mitochondrion may have a low level of metabolism, the concerted action of the large complement of embryonic

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mitochondria (from tens of thousands to tens of millions per egg or early embryo) is absolutely critical for the supply of energy in the embryo. After being unanimously recognized in the 1990s as crucial players in intracellular Ca2þ signaling of somatic cells, mitochondria have been shown to be involved in the regulation of sperm‐triggered Ca2þ oscillations during ascidian and mouse fertilization (reviewed in Dumollard et al., 2006a). These studies revealed intricate relationships between mitochondrial activity and Ca2þ dynamics. In particular, endoplasmic reticulum (ER) Ca2þ release stimulates mitochondrial respiration and, in return, mitochondrial activity supports long‐lasting sperm‐triggered Ca2þ oscillations (Dumollard et al., 2006a). Such interactions are advantageous to mitochondria by allowing a fine matching of energetic supply to energetic demand, which may be necessary for embryonic mitochondria to survive until mitochondrial renewal starts. The oxidative metabolism associated with mitochondrial activity also confers a major role for mitochondria in the regulation of the IRP and oxidative load in the embryo. Mitochondria can have an impact on the IRP by regulating the NAD(P)H/NAD(P)þ ratio and by producing ROS (Brookes et al., 2004; Duchen, 2000; Hansford, 1994; Turrens, 2003) and intermediary metabolites used in the cytosol for the regeneration of NADH and NADPH [and hence of reduced glutathione (GSH)] (MacDonald et al., 2005). Several transcription factors involved in diverse developmental processes are now known to be regulated by oxidative stress or by the IRP (Dickinson and Forman, 2002; Funato et al., 2006; Imai et al., 2000; Liu et al., 2005; Rahman et al., 2004; Zhang et al., 2002). The recent discovery that the activity of these factors can be sensitive to oxidation by ROS, or S‐glutathionylation, or requires NAD(P)H (the reduced form) or NAD(P)þ (the oxidized form) is opening new roles for mitochondria in the regulation of embryonic development. The diverse and primary roles for mitochondria in the regulation of apoptosis (Duchen, 2000; Kroemer, 2003) emphasize another aspect of the impact of mitochondrial activity on embryonic development. The implication of mitochondrial‐dependent apoptosis has been established during oogenesis and remodeling of the embryo (digitation, cardiac remodeling, metamorphosis) (Cecconi and Gruss, 2001; Nakajima et al., 2005; Zuzarte‐Luis and Hurle, 2002). Noteworthy, actors in the mitochondrial‐dependent apoptotic pathway are expressed in oocytes and continue to be present during early development. However, despite such early expression of the components of the apoptotic cascade, apoptosis is suppressed after fertilization in zebrafish and Xenopus embryos (reviewed in Greenwood and Gautier, 2005). Altered mitochondrial activity has been associated with embryo demise at diVerent stages of development. It has been hypothesized that dysfunctional mitochondria mediate maternal and postovulatory oocyte aging,

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but evidence supporting these hypotheses remains scarce and with its widespread acceptance has not been achieved (Bartmann et al., 2004; Fissore et al., 2002). Many of these ideas are borne out of the knowledge that embryonic mitochondria are very sensitive to oxidative stress (Liu et al., 2000) and it is tempting to hypothesize that environmental insults leading to oxidative stress in the embryo impair development by aVecting mitochondrial activity. One experimentally tested system has indicated that miscarriages observed in diabetic women may be due to mitochondrial dysfunctions in the blastocyst (Chi et al., 2002). After briefly describing how mitochondria are generated and distributed in eggs and embryos, this chapter will discuss some of the experimental approaches and technical issues of studying diVerent mitochondrial functions in eggs and embryos. We will then describe the roles played by mitochondria in energy production, Ca2þ homeostasis, and redox metabolism in the embryo. Finally, we will discuss the influence of mitochondrial activity on patterning and survival of the embryo.

II. Mitochondrial Generation and Distribution in Eggs and Embryos The production of mitochondria is discontinuous during embryonic development. Mitochondria are produced during oogenesis before replication ceases in the fully grown oocyte. The number of mitochondria remains static during preimplantation development before mitochondrial replication resumes around the time of implantation (Dumollard et al., 2006a; Jansen, 2000; Poulton and Marchington, 2002). The dramatic amplification of a founding population of approximately 10–100 mitochondria in a human primordial germ cell to several hundred thousands in a mature oocyte (Poulton and Marchington, 2002) raises a number of important considerations. This amplification of germ cell mitochondria provides that all embryonic mitochondria are derived from a limited number of precursors and ensures that they all have the same genetic origin and, therefore, much the same metabolic potential. Moreover, as mitochondria are not replicated during cleavage, the presence of a homogeneous population of mitochondria in the oocyte is very important to maintain an equivalent viability of each blastomere, which will receive only a subset of the oocyte’s mitochondria. It has been observed that the total amount of mtDNA per embryo does not change until gastrulation in fishes (Wang and Yan, 1992) before the pluteus stage in sea urchin (Matsumoto et al., 1974), before the swimming tadpole stage in frogs (Chase and Dawid, 1972), or before larval stage in nematodes (Tsang and Lemire, 2002). The mtDNA copy number is also constant until

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implantation in the mouse (Piko and Taylor, 1987), but variations in mtDNA content have been observed during preimplantation development of bovine embryos (May‐Panloup et al., 2005). Further studies in other mammalian species are required to reevaluate this question of mtDNA turnover during mammalian early development and to confirm whether mtDNA content is constant or changes during mammalian early development. Since mitochondria in the embryo possess only one mtDNA molecule (Piko and Taylor, 1987), this observation suggests that the number of mitochondria is constant during early development. Furthermore, inhibiting mitochondrial DNA replication in mouse embryos with ethidium bromide (Piko and Chase, 1973) or knocking out transcription factors required for mitochondrial biogenesis, such as the mitochondrial transcription factor A (mTFA), the peroxisome proliferator‐activated receptor gamma coactivator 1 (PGC‐1), and nuclear respiratory factor 1 (NRF1), does not prevent development until around the time of implantation (Huo and Scarpulla, 2001; Johnson et al., 2003; Larsson et al., 1998). These studies further demonstrate that mitochondrial biogenesis appears to be dispensable for early development. Mitochondrial proteins appear to be remarkably stable during early development. Murine embryos lacking pyruvate dehydrogenase (PDH) activity (pdh1a KO; Johnson et al., 2001) or cytochrome c (cytochrome c KO; Li et al., 2000) or dihydrolipoamide dehydrogenase (Dld KO, Johnson et al., 1997), all ultimately required for oxidative phosphorylation, survive until implantation. Presumably these mutations do not aVect viability due to the maternal component of these proteins being suYcient to support early development. Further evidence for a slow rate of turnover of mitochondrial proteins may be inferred from a study in which morpholinos against the diVerent cytochrome oxidase subunits (COX5a, 5b, or 6b1) were injected in mouse oocytes before the oocytes were fertilized and development assessed (Cui et al., 2006). This study revealed that inhibiting translation of these COX proteins during early development decreased their levels only moderately and did not inhibit development to the blastocyst stage (Cui et al., 2006). Therefore, because of such slow turnover of mitochondrial proteins, the morpholino approach is inadequate to study the role of mitochondrial function in mammalian early development. Together, these observations indicate that the mitochondria and protein components present in the egg have a low rate of turnover and, as such, a long life in the preimplantation embryo. During oogenesis, the growing population of mitochondria is relocated in diVerent regions of the oocyte. The early phases of mitochondrial location seem conserved between species, whereas the final phases of mitochondrial relocation during maturation are more divergent and generate the variable patterns of mitochondrial distribution observed in mature oocytes of diVerent species (reviewed in Dumollard et al., 2006a).

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In the mature egg, there exist marked diVerences in mitochondrial density at diVerent poles of the egg. Provided that these asymmetric polar distributions are retained through the cleavage divisions, the result will be blastomeres hosting a varying mitochondrial load. As mentioned earlier, the most striking example is found in species displaying a mitochondria‐rich germ plasm located in the vegetal cortex of the mature egg and subsequently in the most vegetal blastomeres of the embryo (reviewed in Dumollard et al., 2006a; Kloc et al., 2004). In eggs not harboring mitochondria‐rich germ plasm, gradients of mitochondria also exist. In ascidians, a mitochondria‐rich domain spanning the vegetal subcortex of the egg has been found to be maintained during development and to segregate exclusively with the muscle lineage (Dumollard et al., 2002; Roegiers et al., 1999). Sea urchin eggs have been shown to have a higher density of mitochondria in a region that is segregated to the oral pole of the embryo and artificial manipulations of this gradient of mitochondria can reorientate the oral‐aboral axis of the embryo (CoVman et al., 2004). In the mouse, diVerent patterns of mitochondrial distribution have been described (Acton et al., 2004; Calarco, 1995; Dumollard et al., 2004, 2006a; Nagai et al., 2004; Van Blerkom and Runner, 1984; Van Blerkom et al., 2003). All these studies have revealed heterogeneity in the distribution of mitochondria in eggs and early embryos, some of which would be consistent with a spatial patterning of the egg by these mitochondria (Section VIII). The disparity of the pattern of mitochondrial distribution observed in mouse oocytes and embryos in the diVerent studies makes it very diYcult to establish the physiological pattern of mitochondrial distribution and emphasizes the diYculty of accessing the mitochondrial compartment in the mouse embryo with techniques devised (and validated) on somatic cells.

III. Studying Mitochondria in Eggs and Embryos During the past 40 years, advances in microscope technology and the development of numerous intracellular indicators have opened new avenues into studying mitochondrial function in living cells (Bjornberg et al., 2006; Chiesa et al., 2001; Duchen et al., 2003; Kuznetsov et al., 1998; Manfredi et al., 2002; Miyawaki, 2003; Passamaneck et al., 2006). These advances in imaging in living cells have uncovered novel roles for mitochondria in cell signaling that could not be observed using traditional biochemical approaches. Mitochondria have a very hyperpolarized electrical potential (around 160 mV) and this unique property of intracellular organelles is used to access the mitochondrial compartment. Lipophilic cationic fluorescent compounds such as rhodamine 123 (Rhod 123) (Chen, 1988), TMRE (tetramethylrhodamine ethyl ester; Ehrenberg et al., 1988), or JC1 (5,50 ,6,60 ‐tetrachloro‐1,10 ,3,30 ‐ tetraethyl‐benzimidazolocarbocyanine iodide; Smiley et al., 1991) were first

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used to monitor the mitochondrial electrical potential in somatic cells and then in eggs and embryos (Rhod 123: Dumollard et al., 2004; Van Blerkom et al., 2003; TMRE: Dumollard et al., 2004; Jouaville et al., 1995; Saelim et al., 2004; JC1: Acton et al., 2004; Van Blerkom et al., 2002; Wilding et al., 2001). These dyes have been used as markers of active mitochondria as their selectivity is dependent on the ability of mitochondria to generate a larger electrical potential than any other organelle. The common vital mitochondrial markers called MitoTrackers (manufactured by Molecular Probes), all use the same principle even though some MitoTrackers can be retained in the mitochondria independently of the electrical potential. In contrast, the cationic JC1 accumulates in mitochondria until it forms a J‐aggregate that is irreversible and cannot exit mitochondria anymore (Smiley et al., 1991). Numerous studies have used these potentiometric dyes to image mitochondrial distribution in eggs and embryos (fruit flies: Akiyama and Okada, 1992; Cox and Spradling, 2003; sea urchin: CoVman et al., 2004; Dumollard et al., 2006a; ascidians: Dumollard and Sardet, 2001; Prodon et al., 2006; Roegiers et al., 1999; frogs: Dumollard et al., 2006a; Jouaville et al., 1995; Machado et al., 2005; Marchant et al., 2002; hamster: Ludwig et al., 2001; Squirrell et al., 2001; mouse: Acton et al., 2004; Calarco, 1995; Dumollard et al., 2004, 2006a; Nishi et al., 2003; Van Blerkom and Runner, 1984; Van Blerkom et al., 2003; pig: Sun et al., 2001; primates: Squirrell et al., 2003; human: Van Blerkom et al., 2002; Wilding et al., 2001). A number of studies have been performed on mammalian oocytes and embryos and the pattern of mitochondrial distribution appears to diVer between studies. Studies using JC1 show perinuclear mitochondria and highly polarized cortical mitochondria (Acton et al., 2004; Van Blerkom et al., 2002, 2003; Wilding et al., 2001), other studies using mitotracker or TMRE have not documented the presence of highly active cortical mitochondria (Dumollard et al., 2004, 2006a; Newhall et al., 2006; Nishi et al., 2003; Fig. 1). These discrepancies cannot be readily explained since both indicators accumulate according to the mitochondrial potential and further work is necessary to determine whether the diVerences are due to mitochondrial membrane potential or the behavior of the indicators. The specificity of mitochondrial staining may be confirmed by comparing the vital mitochondrial staining with the distribution of known mitochondrial markers or of mitochondria‐targeted GFPs (Cox and Spradling, 2003; Dumollard et al., 2004; Marchant et al., 2002; Figs. 1 and 2). Confirmation that mitochondria are the source of the fluorescence may be further confirmed by high resolution imaging that will allow the resolution of single mitochondria, which are rod‐shaped or oblong in eggs and early embryos (2‐ to 4‐mm long) (Dumollard et al., 2002, 2003, 2004; Marchant et al., 2002; Prodon et al., 2006). GFP constructs can be expressed in eggs and embryos by injection into the oocyte or early embryo of an mRNA (Aida et al., 2001; Halet et al., 2004; Levasseur and McDougall, 2000), or of a DNA plasmid (Marchant et al., 2002;

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Figure 1 Distribution of mitochondria in oocytes. (A–C) Confocal images of fully grown mouse GV oocytes showing distribution of NAD(P)H (A) (excitation: 365 nm, emission: 435–485 nm), of mitochondrial FADþþ (B) (excitation: 458 nm, emission: 505–550 nm), and of mitochondria expressing a mitochondrially targeted GFP (C) (excitation: 488 nm, emission: 505–550 nm). At this stage, the mitochondria accumulate around the centrally placed germinal vesicle. Scale bars: 10 mm. (D–F) Confocal images of mature metapahse II‐arrested oocytes showing distribution NAD(P)H (D), mitochondrial FADþþ (E), and mitochondria expressing a mitochondrially targeted GFP (Mito‐GFP) (F). At this stage, the mitochondria are organized in clusters dispersed in the inner cytoplasm and no accumulations of mitochondria in the cortex of the oocyte can be observed. Scale bars: 10 mm. (G, H) Images of a mature sea urchin egg showing the distribution of mitochondria stained with TMRE (H) (excitation: 543 nm, emission: 580 nm). The mitochondria are rich in NAD(P)H as seen in G and are organized in clusters dispersed throughout the egg. Scale bar: 15 mm.

Vintersten et al., 2004) or by generating transgenic animals (Hadjantonakis and Papaioannou, 2004; Nagai et al., 2004; Sobkow et al., 2006; Zeller et al., 2006). A transgenic mouse expressing a mitochondrial GFP has been generated and the pattern of mitochondrial distribution observed revealed that most

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Figure 2 Distribution of mitochondria in embryos. (A) Confocal image showing the distribution of mitochondria in a mouse one cell embryo expressing mito‐GFP. Most of the mitochondria are accumulated around the nucleus. Scale bar: 10 mm. (B) Confocal FADþþ image showing the distribution of mitochondria in a two‐cell embryo. Most of the mitochondria are accumulated around the nucleus of each blastomere. Scale bar: 10 mm. (C) Confocal image showing the distribution of mitochondria in a sea urchin embryo (gastrula) expressing mito‐ GFP. Scale bar: 30 mm. (D) Close‐up of the image in C showing the punctate pattern of the GFP fluorescence that is characteristic of mitochondria. Scale bar: 15 mm. (E) Confocal image showing the distribution of mitochondria in the ascidian embryo (early tailbud). Most of the embryonic mitochondria revealed by immuno‐cytochemistry against the F1 subunit of the ATP synthase (see Patalano et al., 2006 for protocol) are in the muscle cells of the tail. Scale bar: 30 mm. (F) Confocal FADþþ image of living ascidian embryo (just before hatching) where the mitochondria‐rich muscle cells have a strong FADþþ signal emanating from the mitochondria. Scale bar: 30 mm. (G) Confocal NAD(P)H image of the same embryo as in F. The tail muscle cells are seen to be rich in NAD(P)H but other areas of the embryo which contain less mitochondria are also fluorescent. The NAD(P)H image in this example cannot be used to follow mitochondrial distribution in the embryo. Scale bar: 30 mm.

mitochondria have a perinuclear localization in the early embryo (Nagai et al., 2004). In addition, the use of this GFP‐based system together with a potentiometric indicator may provide opportunities for developing a ratiometric technique for monitoring mitochondrial electrical potential and making comparisons between cells. A convenient endogenous mitochondrial marker that can be imaged in living eggs and embryos is mitochondrial autofluorescence (Duchen et al, 2003; Dumollard et al., 2003, 2004; Kuznetsov et al., 1998; Figs. 1 and 2). Under UV excitation, the pyridine nucleotides NADH and NADPH are highly fluorescent in mitochondria, while the oxidized flavoproteins (FADþþ) contained in

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dehydrogenases of the Krebs cycle and in the mitochondrial electron transport chain fluoresce green on blue light excitation (Duchen et al., 2003; Kuznetsov et al., 1998). Images of NAD(P)H or FADþþ fluorescence in mouse and sea urchin eggs show a similar pattern to TMRE staining or to mitochondrial GFP, confirming that autofluorescence signal is a good mitochondrial marker (Dumollard et al., 2004; Fig. 1). This illustrates the benefits of using multiple markers to study mitochondrial distribution in eggs and embryos. The development of GFP‐based Ca2þ, pH, or redox indicators or luciferase‐based ATP indicators (Bjornberg et al., 2006; Chiesa et al., 2001; Manfredi et al., 2002; Miyawaki, 2003) allows the specific targeting of these indicators to the mitochondrial matrix. Applying these strategies to eggs and early embryos will permit the characterization of the diverse mitochondrial functions during early development as it is now known in somatic cells.

IV. Mitochondria and Energy Production in the Embryo The vast majority of studies on energy production in the embryo concern mammalian embryos, which utilize nutrients found in their environment within the oviduct. In contrast, in species presenting an external development, their embryos consume almost exclusively internal stores present in the lipid and protein‐rich yolk. Whilst in reptiles, birds, or fishes, oxidation of yolk fatty acids accounts for the vast majority of energy production (Groscolas et al., 2003; Thompson and Speake, 2002), in frog oocytes and early embryos, amino acids derived from yolk proteins are the main source of energy (Dworkin and Dworkin‐Rastl, 1989). In these embryos, mitochondria have a major role in energy production as fatty acids and amino acids are oxidized in the mitochondria (Stryer, 1970; Fig. 3). Numerous biochemical studies have investigated the substrate requirements for mammalian embryo development. These studies show that the embryo possesses a very specific energetic metabolism which is tightly regulated during early development (Biggers et al., 1967; Brinster, 1965; Gardner et al., 2002; Johnson et al., 2003; Summers and Biggers, 2003). In vivo, mammalian preimplantation embryos develop in an undefined, complex milieu containing nutrients, including pyruvate, glucose, lactate, and amino acids; ions and macromolecules, many of which may be used to satisfy metabolic requirements (Houghton and Leese, 2004). Like most mammalian cells, preimplantation embryos derive their ATP predominantly from oxidative phosphorylation, initially from pyruvate, lactate, and amino acids. This long‐established role for pyruvate as the major energy substrate in early development was confirmed using imaging techniques in living mouse oocytes and zygotes (Dumollard et al., 2006b). By imaging mitochondrial autofluorescence, it was found that pyruvate is rapidly metabolized by

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NADH

ATP H+ NAD+ Krebs cycle

NADH Pyruvate Pyruvate

= −1 5

0m

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NADH

O2

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H2O2 O2− H2O

NADPH ATP

PDH

MCT

Ac-CoA

CiC

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GPx

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Isocitrate NADP ICDH

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Citrate

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Figure 3 Redox and energetic metabolism in the mammalian embryo before compaction. Schematic representation of the metabolic pathways producing NADH, NADPH, and GSH functioning in the cytosol and mitochondria. See main text for details. Am. Ac., amino acid; GSH, reduced glutathione; GSSG, oxidized glutathione; GPx, glutathione peroxidase; GR, glutathione reductase; H2O2, hydrogen peroxide; O2, superoxide; ‐KG, alpha‐ketoglutarate; OAc, oxaloacetate; Ac‐CoA, acetyl coenzyme A; CiC, citrate carrier; LDH, lactate dehydrogenase; ALT, alanine amino transferase; NAD(P)þ, oxidized nicotinamide adenine dinucleotide (phosphate); NADH, reduced nicotinamide adenine dinucleotide; NADP‐ICDH, NADP‐ dependent isocitrate dehydrogenase; PDH, pyruvate dehydrogenase; MCT, monocarboxylate transporter.

mitochondria whereas glucose is not. Surprisingly, lactate‐derived pyruvate appears to be poorly metabolized by mitochondria (Dumollard et al., 2006b). As amino acids are metabolized to pyruvate, oxaloacetate, or alpha‐ ketoglutarate (‐KG) before entering Krebs cycle (Fig. 3), it seems that ATP production in the early embryo is provided by the mitochondria at least until compaction. After compaction, glucose becomes an important substrate, but in quantitative terms, makes only a modest contribution to ATP generation. At the blastocyst stage, glycolysis is sharply increased concomitantly with oxygen consumption (Houghton and Leese, 2004; Trimarchi et al., 2000), this suggests that glucose is metabolized to pyruvate in the cytosol by glycolysis followed by oxidation of glycolytic pyruvate by

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mitochondria. The importance of glucose metabolism at the blastocyst stage is illustrated by the fact that downregulation in early embryos of glut transporters (by RNAi or by high glucose exposure) triggers apoptosis in the blastocyst (Riley and Moley, 2006). Together, these observations unequivocally demonstrate the central role of mitochondria in energy production in the mammalian preimplantation embryo. The low oxygen consumption measured in early embryos compared to blastocysts has been associated with small and sparse cristae in the mitochondria of early embryos (Trimarchi et al., 2000 and references therein). Therefore, mammalian embryos have been proposed to have adopted a quiet energetic metabolism (Leese, 2002). Such quiet metabolism of mitochondria may serve to lower their production of oxidants, thereby limiting the exposure of mammalian embryos to oxidative stress (Dumollard et al., 2004; Gardner et al., 2002; Houghton and Leese, 2004). Fertilization triggers the transition from a relatively quiescent oocyte into an actively dividing embryo. The activation of development is thus associated with an increase in energy demand that needs to be provided by mitochondria. Indeed, an increase in mitochondrial respiration has been observed during fertilization of a few species and such stimulation of mitochondrial respiration is mediated by the sperm‐triggered Ca2þ signals.

V. Mitochondria and Ca2þ Homeostasis in Eggs and Embryos One of the most common features of egg activation is the triggering of a single or repetitive Ca2þ waves on sperm entry (reviewed in Dumollard et al., 2002; Stricker, 1999). These sperm‐triggered Ca2þ signals are mediated by IP3‐dependent Ca2þ release from the ER. Similar to somatic cells, mitochondria are consistently found in clusters in close proximity to ER membranes in eggs and embryos (Dumollard et al., 2006a). Such close proximity between ER and mitochondria allows for an eYcient transmission of cytosolic Ca2þ signals into the mitochondria and an eYcient supply of ATP from the mitochondria to the Ca2þ pumps of the ER (SERCAs) (Duchen, 2000; Dumollard et al., 2002, 2006a; Hansford, 1994). In somatic cells, mitochondrial Ca2þ uptake and release as well as mitochondrial ATP production are now known to participate in the regulation of intracellular Ca2þ signaling (reviewed in Duchen, 2000). Mitochondrial ROS production was also found to be necessary for Ca2þ oscillations (Camello‐Almaraz et al., 2006). The roles of mitochondria in Ca2þ homeostasis and in the regulation of sperm‐triggered Ca2þ oscillations have been investigated in ascidian and mouse eggs (reviewed in Dumollard et al., 2006a). In ascidian eggs, both mitochondrial Ca2þ cycling and mitochondrial ATP production participate in the regulation of sperm‐triggered Ca2þ oscillations (Dumollard

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et al., 2003). In contrast, in mouse eggs, mitochondrial ATP production is necessary to maintain the resting ½Ca2þ c and support sperm‐triggered Ca2þ oscillations, whereas mitochondrial Ca2þ cycling is dispensable for sperm‐triggered Ca2þ oscillations (Dumollard et al., 2004). In sea urchin eggs, an increase in mitochondrial Ca2þ has been measured after fertilization, but whether this impacts on the dynamics of the single sperm‐triggered Ca2þ wave is not known (Eisen and Reynolds, 1985). Further studies in other species are needed to determine whether mitochondria universally regulate sperm‐triggered Ca2þ oscillations and to determine the respective contribution of mitochondrial Ca2þ cycling, mitochondrial ATP, and ROS production in the regulation of sperm‐triggered Ca2þ signals.

VI. Ca2þ‐Induced Mitochondrial Activity: Matching ATP Supply and Demand Mitochondria are a major target of cytosolic Ca2þ transients which are readily transmitted to the mitochondrial matrix at fertilization (Duchen, 2000; Dumollard et al., 2006a). Indeed, Ca2þ in the mitochondria is a pivotal ‘‘multisite’’ activator of oxidative phosphorylation as it stimulates several dehydrogenases of the Krebs cycle (Hansford, 1994), the electron transport chain (Gunter et al., 1994), and has a direct action on the F0–F1 ATP synthase (Territo et al., 2000). Ca2þ in the mitochondria thus stimulates the production of NADH used to generate ATP and ROS. During fertilization of starfish, ascidian, and mouse egg, stimulation of mitochondrial respiration by sperm‐ triggered Ca2þ transients has been observed (Campbell and Swann, 2006; Dumollard et al., 2003, 2004; Schomer and Epel, 1998). Such signaling by mitochondrial Ca2þ allows tight coupling of ATP supply and demand, which provides a major advantage for early mammalian development. The long life span of mitochondria in oocytes and early embryos requires that mitochondria are protected from potentially damaging ROS. The maintenance of a low level of oxidative phosphorylation that can be stimulated by an increase in ATP demand provides one means of lowering mitochondrial oxidative stress. Further studies in other species are required to confirm whether such a mechanism is ubiquitously involved at fertilization.

VII. Mitochondria and Redox Metabolism in the Embryo Mitochondrial activity has a major impact on the IRP and oxidant load. This impact is complex and multifaceted as mitochondrial function influences the levels of NAD(P)H, intermediary metabolites as well as ROS. The contribution

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of these factors will depend on the physiological environment of the cell and as such is subject to change as oocytes and embryos develop. The IRP is set by the ratio of the numerous redox couples present in cells. The three most abundant redox couples in the cell are reduced and oxidized glutathione (GSH/GSSG) and the pyridine nucleotides NADH/NADþ and NADPH/NADPþ. The IRP can thus be estimated by measuring intracellular GSH/GSSG or NAD(P)H/NAD(P)þ ratios. Oxidative stress is mediated by ROS and results in an imbalance of the IRP toward an oxidized potential (Balaban et al., 2005). The resting cytoplasmic redox potential is reduced and is maintained by the glutathione/glutaredoxin and the thioredoxin systems (Holmgren et al., 2005). These systems are subject to regulation by a number of factors that can have a major impact on the IRP, some of which are mitochondrial. Glutaredoxin and thioredoxin systems both use electrons from NADPH (but not NADH) to regenerate GSH. This makes NADPH an important factor essential for antioxidant defense (Dickinson and Forman, 2002). However, in a twist to this role, NADPH can promote the formation of ROS in cells containing NADPH oxidases (Geiszt and Leto, 2004). Therefore, NADPH may have a dual impact on the oxidant load in the cell. In contrast, NADH cannot be used by the glutaredoxin and thioredoxin systems but can be oxidized by intracellular oxidases to generate ROS, NADH will thus increase the oxidant load in the cell. The intracellular oxidases producing most of the ROS in the cell are NAD(P)H oxidases (Geiszt and Leto, 2004), xanthine/xanthine oxidases, and the oxidases of the mitochondrial respiratory chain (Balaban et al., 2005; Brookes et al., 2004). Active cytoplasmic NAD(P)H oxidases are present in sea urchin and echiuroid eggs and early embryos (Schomer and Epel, 1998; Wong et al., 2004) as well as in sperm of numerous species (Baker and Aitken, 2004) but have not been found in ascidians or vertebrates eggs or early embryos. Thus, in embryos, NADH will be mostly oxidized by the mitochondrial respiratory complexes to produce ROS (mainly from complexes I and III, Balaban et al., 2005; Brookes et al., 2004; Turrens, 2003; Fig. 3) and ATP. As such, it seems that in mammalian embryos at least, NADPH will supply antioxidant defense, while NADH will promote oxidative stress via mitochondrial oxidative phosphorylation. Therefore, the balance between mitochondrial NADH and NADPH production may ensure eYcient ATP production, while limiting oxidative stress that is quickly detrimental to mitochondrial function (Balaban et al., 2005; Brookes et al., 2004). The shuttling of NADH between the mitochondrial and cytosolic compartments is essential for normal cell function. As intact mitochondria are impermeable to NADH, the transfer of NADH across the mitochondrial membrane is performed by an indirect pathway, involving the movement of electrons from NADH across the mitochondrial membranes. One active

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component of this pathway that shuttles mitochondrial NADþ to the cytosol and cytosolic NADH into the mitochondria is the malate‐aspartate shuttle (MAS) (Lane and Gardner, 2005; MacDonald et al., 2005). All of the enzymes involved in this activity, mitochondrial aspartate aminotransferase, cytoplasmic aspartate aminotransferase, mitochondrial malate dehydrogenase, and cytoplasmic malate dehydrogenase have been shown to be expressed in mouse embryos (Lane and Gardner, 2005). The shuttle appears to be active from the two‐cell stage where it is thought to be necessary to supply the NADþ required for the cytosolic metabolism of lactate (Lane and Gardner, 2005). Mitochondria also synthesize Krebs cycle intermediates (through a process called anaplerosis) that will be exported from the mitochondria and used in the cytosol (through a process called cataplerosis; MacDonald et al., 2005). These intermediates may be used to regenerate NADPH and thereby increase cytosolic antioxidant defense. Strikingly, mitochondria are the only producers of citrate inside the cell (MacDonald et al., 2005) and the regeneration of cytosolic NADPH via citrate metabolism is vital to counteract the oxidant load building up during ischemia/reperfusion (Mallet and Sun, 2003) or cell aging (Kil et al., 2006). Mouse preimplantation embryos contain high levels of citrate which are maintained by mitochondrial metabolism of pyruvate (Barbehenn et al., 1974). It was found in mouse eggs that inhibiting the cytosolic NADPH‐producing enzyme NADP‐isocitrate dehydrogenase (NADP‐ICDH) decreased cytosolic NADPH levels, suggesting that isocitrate (arising from citrate exported from the mitochondria) is metabolized in the cytosol to maintain NADPH levels (Dumollard et al., 2006b; Fig. 3). Cytosolic NADPH is used to regenerate GSH in these eggs and such cytosolic production of NADPH via NADP‐ICDH might be vital to preserve the antioxidant defense of the embryo after fertilization (Dumollard et al., 2006b). Indeed, during oocyte maturation, GSH levels are maintained both by ATP‐dependent de novo synthesis and by NADPH‐dependent regeneration whereas, during early development, ATP‐dependent synthesis of GSH is switched oV (Gardiner and Reed, 1995; Luberda, 2005) and GSH can only be maintained by the NADPH‐dependent pathway. Moreover, the pentose phosphate pathway (the major supplier of cytosolic NADPH in somatic cells) does not participate in the maintenance of cytosolic NADPH levels (Dumollard et al., 2006b), further emphasizing the critical role of citrate metabolism by cytosolic NADP‐ICDH for the maintenance of NADPH (and hence GSH) levels (Fig. 3). Mitochondrial metabolism of pyruvate may also regulate the cytosolic NADH/NADþ ratio indirectly. The cytosolic NADH/NADþ ratio is set by the ratio of lactate to pyruvate in the cytosol and the steady‐state level of cytosolic pyruvate is balanced both by the flux of pyruvate from the medium and by the import of pyruvate into the mitochondria (Dumollard

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et al., 2006b). Such import of pyruvate into the mitochondria is necessary to maintain a reducing cytosolic redox potential as blocking pyruvate entry into mitochondria induces a sharp LDH‐dependent oxidation of cytosolic NADH (Dumollard et al., 2006b). The complex oxidative metabolism in mitochondria may thus have a dual impact on intracellular redox metabolism in the embryo, and the net eVect of mitochondrial metabolism on the IRP and oxidant load is very diYcult to predict. In somatic cells, mitochondrial metabolism of pyruvate can have a net oxidant action (via generation of ROS in the electron transport chain; Xu and Finkel, 2002) or a net antioxidant action (via anaplerosis and cytosolic NADP‐ICDH; Mallet and Sun, 2003). In early mouse embryos, intracellular metabolism of pyruvate seems to have an overall oxidant action that must be limited to preserve developmental potential (Dumollard et al., 2006b). Indeed, culturing embryos in a medium containing a high pyruvate concentration impairs early development (Dumollard et al., 2006b; Wales and Whittingham, 1970). Such inhibition of development is most probably due to an excessive oxidant load in the embryo since it is reversed by adding a cell‐permeant antioxidant to the culture medium (Dumollard et al., 2006b). These observations support the notion that quiet energetic metabolism limiting the oxidant action of mitochondria favors early development (Leese, 2002). In addition to regulating ½Ca2þ c and [ATP]c, mitochondria regulate the IRP and oxidant load in the embryo. These three functions of mitochondria are essential for the transition from oocyte to embryo and for early embryonic development.

VIII. Impact of Mitochondrial Activity on Embryonic Development Mitochondria regulate developmental potential through many of the pathways described above, including generation of ATP, regulation of Ca2þ, and the maintenance of the IRP. In mammals, the production of ATP by mitochondria is absolutely necessary for the oocyte to mature (Takeuchi et al., 2005) and for the fertilized egg to proceed through development. Inhibiting mitochondrial ATP production in a fertilized egg quickly provokes a deregulation of Ca2þ homeostasis, culminating in a sustained high cytosolic [Ca2þ] that precedes apoptotic cell death (Dumollard et al., 2004; Liu et al., 2000, 2001). Graded mitochondrial injury can be induced in mouse oocytes by fluorophore photosensitization of mitochondria during which a fluorescent dye targeted to the mitochondria is illuminated to generate free radicals, thereby creating damaging mitochondrial oxidative stress (Takeuchi et al., 2005; Thouas et al., 2004, 2006). These studies show that the extent of

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mitochondrial destruction determines whether the oocyte will die by oncotic cell death (after extensive destruction) or apoptotic cell death (after milder destruction) or whether the embryo will proceed through the early stages of development before embryonic cell death occurs (Thouas et al., 2004, 2006). These diVerences in type or timing of cell death are probably due to the diVerential ATP depletion induced by such graded mitochondrial insults. Oncosis, a process that does not need ATP, would occur in oocytes strongly depleted in ATP, whereas apoptosis in the egg or in the early embryo (a process requiring ATP; Skulachev, 2006) would occur in damaged eggs and embryos that could still maintain a minimal ATP level. These observations are most readily explained by impaired mitochondrial ATP production; however, the techniques used to inhibit mitochondrial function can also impair mitochondrial Ca2þ cycling and mitochondrial redox metabolism. As such, it is diYcult to isolate the precise mechanisms underlying cell death in these experiments. The mitochondrial regulation of Ca2þ uptake and release may also be an important factor in the regulation of oocyte and embryo development. As discussed previously, mitochondrial Ca2þ cycling regulates the sperm‐ triggered Ca2þ signals in ascidian zygotes, thereby allowing the activation of development in this species (Dumollard et al., 2003, 2006a). Further studies are required to establish whether mitochondrial Ca2þ cycling has a role during fertilization in other species. In the mouse, while mitochondrial ATP production is essential to maintain baseline Ca2þ levels, mitochondrial Ca2þ uptake and release may not be necessary during fertilization (Dumollard et al., 2004). Ca2þ‐induced ATP increases in mouse oocytes at fertilization appear to persist for a number of hours (Campbell and Swann, 2006; Dumollard et al., 2004). This increase may have long‐term eVects on embryo development but direct evidence for this idea is lacking. Increases in Ca2þ are also known to take place at diVerent stages of development, including during cleavage to the two‐cell stage (FitzHarris et al., 2005) and during pattern formation and organogenesis (Whitaker, 2006), but to date there have been no studies performed to determine whether any eVects may be mediated by mitochondrial Ca2þ handling. Mitochondria may also regulate development by their ability to modulate the IRP and oxidant load in the embryo. Interestingly, it was shown in sea urchin embryos that the oxidant action of mitochondria, which are asymmetrically distributed in the early embryo, may determine the position of the oral (ventral) pole of the embryo (CoVman et al., 2004). Redox gradients were measured a long time ago in numerous invertebrate embryos (reviewed in Blackstone, 2006) and asymmetries in the distribution of mitochondria in the embryo or in the egg may create such redox gradients. In mammals, while the impact of oxidative stress on embryo survival has been extensively studied, the regulation of early developmental processes by

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the IRP or mitochondrial oxidant load remains largely unexplored even though candidate factors exist. Oxidative stress and IRP are known to regulate a number of transcription factors, some of which are critical in early development (Funato et al., 2006; Imai et al., 2000; Liu et al., 2005; Rahman et al., 2004). The transcription factor NF‐B and the glycogen synthase kinase‐3 (GSK3 ) both can be activated by mitochondrial ROS production (Hughes et al., 2005; Nemoto et al., 2000). NF‐B, which is known to play a pivotal role in formation of Drosophila ventral structures and during limb morphogenesis (Bushdid et al., 1998; Gordon et al., 2005) and in ascidian notochord determination (Kawai et al., 2005), is expressed throughout preimplantation development in the mouse (Nishikimi et al., 1999). GSK3 is required to establish the dorsal‐ventral axes in vertebrates, anterior‐posterior segment polarity in Drosophila, animal–vegetal patterns in sea urchin, and endoderm/mesoderm cell fates in nematodes (Kim and Kimmel, 2000), but no study has yet addressed the possibility that mitochondrial ROS production stimulates NF‐B or GSK3 during these developmental processes. C‐terminal‐binding proteins (CtBPs) are transcriptional corepressors of mediators of Notch, Wnt, and other signaling pathways (Chinnadurai, 2002). Thus, they are potential players in the control of several developmentally important processes, including segmentation, somitogenesis, and neural tube and limb patterning. Agents capable of increasing NADH levels stimulate CtBP binding to its partners in vivo and potentiate CtBP‐mediated repression. CtBP can thus detect changes in nuclear NADH/NADþ ratio and behaves as a sensor of the IRP for transcription (Zhang et al., 2002). The latest class of histone deacetylases (HDAC III) called sirtuins require NADþ for their enzymatic activity and are known to modulate histones but also the myogenic determinant MyoD and p53 activities (Imai et al., 2000). It was shown in myogenic cell lines that the IRP is reduced as muscle cells diVerentiate and that oxidation of the IRP inhibits muscle diVerentiation by stimulating sirtuin activity (Fulco et al., 2003). Some of these sirtuins are expressed from the two‐cell stage during mouse early development (McBurney et al., 2003), suggesting that the impact of the IRP on these HDAC may regulate early development. S‐glutathionylation of many proteins occurs as a consequence of an oxidation of the IRP. A microarray study in a cell line revealed that 95 genes are upregulated and 25 are downregulated by S‐glutathionylation, 34 of which are transcription factors (Fratelli et al., 2005). Here again, the ability of mitochondria to modulate the IRP can be reflected in S‐glutathionylation of proteins that will change their activity and change gene expression. Mammalian oocytes have a high concentration of GSH (10 mM) that is generated during oocyte maturation. Ninety percent of this GSH will be consumed during early development (see Luberda, 2005 for review) but

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the high levels of GSH suggest that in addition to protecting against oxidative stress during preimplantation development, GSH may be used for S‐glutathionylation for the purposes of regulating gene expression. Although proteomic approaches remain technically challenging when faced with the relatively small quantities of protein present, a proteomic analysis of S‐glutathionylated proteins in early mammalian embryos may reveal new means of mitochondrial regulation of early development that remain completely unexplored. Altered mitochondrial Ca2þ cycling and mitochondrial ROS production are also determining factors for the decision to commit to cell suicide and to execute the apoptotic program (Brookes et al., 2004; Duchen, 2000). Furthermore, mitochondrial‐dependent apoptotic pathways are used during oogenesis and tissue remodeling (Cecconi and Gruss, 2001; Kim and Tilly, 2004; Nakajima et al., 2005; Zuzarte‐Luis and Hurle, 2002), while mitochondrial dysfunctions can induce apoptosis in the egg and early embryo (Liu et al., 2000, 2001), thus emphasizing another major role for mitochondria during development.

IX. Mitochondria and Apoptosis in the Embryo The involvement of the mitochondrial‐dependent pathway of apoptosis has been characterized during oogenesis, embryonic remodeling, and tail resorption in mammals as well as in other vertebrates and invertebrates (Cecconi and Gruss, 2001; Greenwood and Gautier, 2005; Kim and Tilly, 2004; Nakajima et al., 2005; Zuzarte‐Luis and Hurle, 2002). During the mitochondrial‐dependent pathway, an apoptotic signal stimulates the translocation of Bcl2 family proteins to the mitochondria which triggers mitochondrial depolarization. Such mitochondrial depolarization is followed by oxidation of mitochondrial NADH and subsequent release of cytochrome c that stimulates caspases, or release of apoptosis‐inducing factor (AIF) and endonuclease G that cleave DNA or release of smac/diablo that will inhibit inhibitors of apoptosis proteins (Cecconi and Gruss, 2001; Kroemer, 2003). Mitochondria are thus a store of proapoptotic factors in the cell. Mitochondrial‐dependent apoptosis is necessary for oocyte apoptosis during follicular atresia (Kim and Tilly, 2004) and may be involved in postovulatory aging in mammals (Fissore et al., 2002). Injection of extra numerous follicular granulosa cell mitochondria into mouse oocytes decreases the incidence of apoptosis, thus suggesting that these mitochondria have an antiapoptotic action during postovulatory aging (Perez et al., 2000). Furthermore, injection of mitochondria isolated from adult cells into a mouse early embryo inhibits development to the blastocyst, while injection of oocyte cytoplasm (containing mitochondria) does not aVect early development (Takeda et al., 2005). This observation suggests that mitochondria from adult cells promote apoptosis in

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the early embryo, while ooplasmic mitochondria do not. Further studies are required to establish whether mitochondria acquire a proapoptotic activity during development. Involvement of mitochondrial‐dependent apoptosis has been characterized during tail regression in ascidian, Xenopus (Chambon et al., 2002; Nakajima et al., 2005; Sachs et al., 2004), and during neuronal development in mammals (Akhtar et al., 2004), illustrating the widespread use of mitochondrial‐dependent apoptosis during normal development. The prominent regulatory role for mitochondria in apoptosis induction puts mitochondria in a crucial position to determine whether early development will proceed or not. Such a role for mitochondria might be critical in pathological conditions or during in vitro/environmental insults. In the mouse egg, oxidative stress can provoke mitochondrial dysfunctions that trigger apoptosis (Liu et al., 2000). Hyperglycemia is known to be responsible for miscarriages and has been shown to induce apoptosis in the blastocyst (Chi et al., 2002; Riley and Moley, 2006). Hyperglycemia acts via downregulation of glucose transporters and a decrease in intracellular glucose levels. By measuring single embryo concentrations of Krebs cycle and glycolytic metabolites, changes that occur as a result of exposure to high‐ glucose conditions were observed. Embryos exposed to elevated glucose conditions experienced significantly lower fructose bisphosphate, suggesting decreased glycolysis, significantly higher pyruvate, suggesting increased pyruvate uptake by the embryos in response to decreased glycolysis, and increased Krebs metabolites, suggesting an inability to oxidize the pyruvate and a slowing of the Krebs cycle. It was thus speculated that the glycolytic changes lead to mitochondrial dysfunction that results in an abnormal Krebs metabolite pattern and triggers the apoptotic event (Chi et al., 2002). A higher incidence of ROS generation, principally in the inner cell mass cells, has been observed in embryos cultured in presence of high levels of glucose, suggesting that oxidative stress (maybe mitochondrial) also mediates apoptosis induced by hyperglycemia (Leunda‐Casi et al., 2002). Therefore, preserving mitochondrial functions during early development will be critical to maintaining a low incidence of apoptosis and support development.

X. Conclusions and Perspectives The importance of mitochondrial function in embryonic development and embryo survival has now been known for a long time, but the characterization of the multifaceted influence of mitochondrial metabolism on diVerent developmental processes is only beginning to emerge. The development of protein‐based intracellular indicators that can be targeted to the mitochondrial compartment should further our understanding of the

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role of mitochondrial function in early development. Finally, the recent discovery of a redox regulation for several proteins expressed during early development illustrates a new aspect of the mitochondrial regulation of development. A challenge for the future will be to study the physiology of mitochondria in early live embryos together with the molecular characterization of the relevant genes in order to unravel at the molecular level how mitochondrial function regulates embryonic development.

Acknowledgments Work in the laboratories of J.C. and M.D. is supported by the MRC and the Wellcome Trust, respectively.

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Mitochondrial DNA in the Oocyte and the Developing Embryo Pascale May‐Panloup,*,{ Marie‐Franc¸oise Chretien,{ Yves Malthiery,*,{ and Pascal Reynier*,{ *INSERM U694, F‐49033 Angers, France { Poˆle de Biologie, De´partement de Pathologie Cellulaire et Tissulaire Centre Hospitalier Universitaire, F‐49033 Angers, France { Poˆle de Biologie, De´partement de Biochimie et Ge´ne´tique Centre Hospitalier Universitaire, F‐49033 Angers, France

I. Introduction II. Mitochondrial DNA Structure and Function A. The Mitochondrial Genome Is a Vestige of Its Endosymbiotic Prokaryotic Ancestor B. mtDNA Structure C. mtDNA Maintenance and Expression D. Maternal Transmission of mtDNA III. mtDNA Content in Oocytes A. Oogenesis Is Associated with Drastic Restriction/Amplification of mtDNA Content B. The Oocyte Has the Largest Cellular mtDNA Content of the Organism C. The mtDNA Content of Oocytes Is Highly Variable D. The Mitochondrial Content of Oocytes Could AVect Their Fertilizability E. Ovarian InsuYciency Is Associated with Major mtDNA Depletion F. How Does mtDNA Content Influence Oocyte Quality? IV. Variation of mtDNA Content and Transcripts in Early Embryonic Development V. mtDNA Mutations in Oocytes and Embryos VI. Conclusions and Perspectives References

Mitochondria play a primary role in cellular energetic metabolism, homeostasis, and death. They possess their own multicopy genome, which is maternally transmitted. Mitochondria are directly involved at several levels in the reproductive process since their functional status influences the quality of oocytes and contributes to the process of fertilization and embryonic development. This chapter discusses recent findings concerning mitochondrial DNA content and its expression during oogenesis, fertilization, and early embryonic development. ß 2007, Elsevier Inc.

Current Topics in Developmental Biology, Vol. 77 Copyright 2007, Elsevier Inc. All rights reserved.

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0070-2153/07 $35.00 DOI: 10.1016/S0070-2153(06)77003-X

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I. Introduction Mitochondria are present in the cytoplasm of almost all eukaryotic cells. These organelles may vary considerably in size, shape, mass, and distribution but they usually form a functional tubular network adapted to the energetic needs of the cell (Okamoto and Shaw, 2005). The dynamics of the network are driven by the balance between mitochondrial fission and fusion (Rube and van der Bliek, 2004), whereas the mitochondrial mass reflects the balance between mitochondrial biogenesis and autophagy (Kroemer and Jaattela, 2005). Mitochondria are directly involved in cellular energetic metabolism, homeostasis, growth, and death. The best known mitochondrial function is the production of adenosine triphosphate (ATP) by oxidative phosphorylation (OXPHOS), a mechanism coupling the oxidation of reduced nutrients and organic molecules with the phosphorylation of adenosine diphosphate (ADP) (Nelson and Cox, 2000; Wallace, 2005). OXPHOS is supported by the mitochondrial respiratory chain that consists of four complexes located on the inner mitochondrial membrane (Fig. 1). Complex I oxidizes nicotinamide

Cytoplasm Outer membrane

H+

H+

H+

H+

C Q

Inner membrane

I

II

III

IV

V O2

Matrix

NADH Electron flow Proton flow

Succinate ADP

ADP + Pi

Figure 1 Mitochondrial respiratory chain. The mitochondrial respiratory chain is organized in five complexes situated in the inner mitochondrial membrane. Complex I: NADH‐dehydrogenase, complex II: succinate‐dehydrogenase, complex III: ubiquinone‐cytochrome c‐reductase, complex IV: cytochrome c‐oxidase, and complex V: ATP synthase. Q, quinine; C, cytochrome c.

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dinucleotide (NADH) produced by several catabolic pathways such as cytosolic glycolysis, the pyruvate dehydrogenase complex, the tricarboxylic acid cycle (TCA), and fatty acid ‐oxidation. Complex II oxidizes flavine adenine dinucleotide (FADH2) produced by TCA. Complexes I and II transfer electrons from NADH and FADH2 to ubiquinone, a lipophilic benzoquinone which is soluble in the inner mitochondrial membrane. Reduced ubiquinone transfers the electrons to complex III, which reduces cytochrome c, a mobile hydrophobic hemoprotein. Cytochrome c then transfers the electrons to complex IV which reduces O2 to produce H2O. The exergonic (releasing energy) transfer of electrons along the respiratory chain is exploited by complexes I, III, and IV to expel protons to the intermembrane space generating an energetic protonic gradient. Complex V, ATP synthase, which is not involved in electron transfer, provides a channel for the entrance of protons into the mitochondrial matrix while the exergonic protonic flux drives the phosphorylation of ADP to ATP. Exergonic hydrolysis of ATP to ADP þ Pi (inorganic phosphate) provides energy for most of the energy‐dependent cell processes and is essential for cell survival, growth, division, and several other functions. In addition to the catabolism of nutrients and energy supply, mitochondria play an important role in cell survival. They are involved, at least partially, in several biosynthetic pathways such as those leading to the syntheses of heme, iron–sulfur proteins, nucleotides, steroid hormones, amino acids, and, more generally, organic acids (Nelson and Cox, 2000). Mitochondria also regulate calcium homeostasis by buVering acute intracellular calcium concentration variations (Gunter et al., 2004). Although mitochondria produce reactive oxygen species (ROS), they also participate in their detoxification through a specific ROS‐detoxifying enzymatic pathway (Fleury et al., 2002). The catabolism of nutrients leads to the production of ATP but it also generates a reducing power essential to cell survival, mainly supported by reduced NADPH (Nelson and Cox, 2000). This cofactor provides the electrons necessary for cellular biosyntheses (reducing processes) and for detoxification of ROS. In addition, mitochondria play an essential role in heat production (Mozo et al., 2005), oxygen sensing (Lahiri et al., 2006), and programed cell death (Green and Kroemer, 2004). The pleiotropic properties of mitochondria explain the central role of these organelles in the phenomena of aging as well as in several genetic and neurodegenerative diseases, cancer, obesity, and diabetes (Wallace, 2005). There is increasing evidence that mitochondria also contribute to fertility and that mitochondrial defects could be at least partially responsible for male or female infertility. Indeed, mitochondrial defects appear to aVect not only the gametes but also the process of fertilization and early embryonic development (Cummins, 2004; Smith et al., 2005; Van Blerkom, 2004).

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II. Mitochondrial DNA Structure and Function A. The Mitochondrial Genome Is a Vestige of Its Endosymbiotic Prokaryotic Ancestor The mitochondrion probably originates from a prokaryotic ancestor related to the ‐proteobacteria phyla by a mechanism of endosymbiosis (Gray, 1989). One of the most surprising and unique features of this organelle is persistence of a specific, multicopy genome. Most of the genes of the prokaryotic ancestor were probably lost or transferred to the nuclear genome of the eukaryotic host during the course of the evolution of the endosymbiont (Gray et al., 1999). To date, the only known function of the remaining mitochondrial DNA (mtDNA) is to produce 13 of the 1500 estimated mitochondrial proteins. The maintenance (replication and repair) and the expression (transcription and translation) of mitochondrial genomes in the organelles necessitate the biosynthesis of hundreds of diVerent proteins encoded by the nucleus. The persistence of the mitochondrial genome in these organelles is thought to be due to the highly hydrophobic nature of the 13 proteins that it encodes (Von Heijne, 1986). Indeed, it would seem diYcult to transport such proteins from the cytoplasm to the mitochondrial matrix across the inner and outer membranes.

B. mtDNA Structure The mitochondrial genome is a double‐stranded, circular, DNA molecule containing 16,569 base pairs (Anderson et al., 1981). This molecule is organized in nucleoproteic complexes called nucleoids that are devoid of histones (Legros et al., 2004; Satoh and Kuroiwa, 1991). The mitochondrial genome is very compact due to the lack of intronic or intergenic sequences and the absence of significant 50 and 30 untranslated regions (UTRs). Most of the sequences are coding sequences, involving both the DNA strands. The two main noncoding regions encompass the two origins of replication. The control region (also called the D‐Loop) contains 1200 base pairs organized as a triple‐stranded structure and includes the main regulatory sequences of replication and transcription. mtDNA carries 37 genes: 13 protein‐encoding genes, 22 tRNA genes, and 2 rRNA genes, the products of the RNA genes being necessary for the translation of the 13 proteins. Seven of these proteins are complex I subunits, one is a complex III subunit, three are complex IV subunits, and two are complex V subunits. Although mtDNA is highly polymorphic, its gene content and organization appear to be well conserved since all the animals studied so far have been found to carry identical sets of mitochondrial genes (Macino et al., 1980; Saccone et al., 2000) (Fig. 2).

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ATPase ATPase8

Figure 2 Mitochondrial genome. CO, cytochrome oxidase; Cyt, ubiquinone‐cytochrome c reductase; ND, NADH dehydrogenase; ATPase, ATP synthase. tRNAs are represented as points.

C. mtDNA Maintenance and Expression The replication of the mitochondrial genome is independent of the cell cycle and occurs continuously in cells including postmitotic cells. The DNA duplication process is driven either by bidirectional and asynchronous replication of both strands (Shadel and Clayton, 1997) or by the unidirectional progression of the replication loop (Holt et al., 2000). According to the former model, the replication of the first strand starts at the origin of replication of the heavy strand (OH) located in the control region. The progression of the replication of the heavy strand then triggers the replication of the second strand starting at the origin of replication of the light strand (OL) (Clayton, 1982). The expression of the mitochondrial genome is closely linked to its replication since transcription provides the primer sequence necessary to initiate replication. mtRNA polymerase promotes the synthesis of an RNA

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starting from the light strand promoter (LSP) present in the control region (Montoya et al., 1982). This transcript is then cleaved by a specific RNase (MRP) at specific sequences (conserved sequences blocks, CBS) before acting as a replication primer and being elongated by the mitochondrial‐ specific DNA polymerase (POLG). The replication of the heavy strand is frequently interrupted in the control region at the termination‐associated sequences (TAS), thus generating the characteristic triplex structure of the control region (Shadel and Clayton, 1997). The transcription of mtDNA is bidirectional starting at each of the promoters, that is, the heavy strand promoter (HSP) and the LSP, situated in the control region. The two main polycistronic RNA strands are entirely transcribed and then cleaved into 13 mRNAs, 22 tRNAs, and 2 rRNAs that are finally matured (Fernandez‐Silva et al., 2003; Ojala et al., 1981). The translation is performed by mitochondrial‐specific tRNA and mitoribosomes, leading to the in situ synthesis of 13 subunits that are finally assembled with nuclear‐encoded subunits in the respiratory chain complexes. A surprising feature of the mitochondrial translational apparatus is that four of the codons have a significance that diVers from the universal genetic code in humans (Barrell et al., 1980). Mitochondrial transcription factor A (TFAM) is a nuclear‐encoded protein of the High Mobility Group with a dual function since it acts as an mtDNA‐packaging factor (Fisher et al., 1992) and as a transcription factor, promoting and regulating both replication and transcription (Chang and Clayton, 1984). Two additional mitochondrial transcription factors are necessary to regulate and activate mtDNA transcription: TFB1M and TFB2M (transcription factors B1 and B2) (Scarpulla, 2006). TFAM interacts with both LSP and HSP promoters and facilitates the interaction of mtRNA polymerase with promoters (Gaspari et al., 2004). The expression of mitochondrial and nuclear‐encoded subunits of the respiratory chain complexes must be closely coordinated. The nuclear respiratory factors 1 and 2 (NRF1 and NRF2) are the main factors responsible for this coordination. NRF1 is recognized by the promoters and activates transcription of several mitochondrial nuclear‐encoded proteins such as certain subunits of all respiratory complexes, the RNase MRP, and TFAM (Scarpulla, 1997). The NRF1‐induced transcription activation of TFAM coordinates the transcription of both the mitochondrial and nuclear genomes. TFAM and NRF1 are expressed ubiquitously. At a higher level of regulation, several factors coordinate the metabolic status of the cell (glucose uptake, glycolysis, ‐oxidation, thermogenesis, and so on) and mitochondrial biogenesis (Lin et al., 2005; Naar et al., 2001). One of the best known of these coordinating factors, PGC‐1 (PPAR gamma coactivator 1, existing as and isoforms), is mostly expressed in brown adipose tissue, skeletal muscle, heart, kidneys, and brain. PGC‐1 enhances NRF1 and NRF2 expression and directly interacts

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with NRF1, increasing its ability to activate target genes (Puigserver et al., 1998). A third coactivator, Pgc‐1‐related coactivator (PRC) is ubiquitously expressed and also interacts with NRF1, increasing its ability to activate its targets (Andersson and Scarpulla, 2001). The maintenance and repair of the mitochondrial genome are carried out by the same type of enzymes as in the nucleus. However, mitochondria lack some of these enzymes and the mitochondrial repair system is therefore thought to be less eYcient than that of the nucleus (Larsen et al., 2005). Taken together with the proximity to ROS production and the absence of protective histone proteins, this less eYcient repair system may be responsible for the high mtDNA mutation rate (Parsons et al., 1997; Wallace et al., 1987). Indeed, numerous studies on aging cells and tumors have described the instability of mitochondrial genomes associated with the accumulation of somatic mutations.

D. Maternal Transmission of mtDNA The mitochondrial genome is maternally inherited in animals since it is exclusively transmitted by oocytes (Giles et al., 1980). In mammals, although the mitochondria of spermatozoa are found in the oocyte just after fertilization (Ankel‐Simons and Cummins, 1996; Cummins, 1998), it has been shown in mouse that they are specifically destroyed before the four‐cell stage of embryogenesis (Kaneda et al., 1995). It has been reported in cow that the mechanism of paternal mitochondrial elimination involves the proteasome of oocytes, which recognizes mitochondria that have been ubiquitinated during spermatogenesis (Sutovsky et al., 1999, 2000). However, evidence for occasional paternal inheritance has been reported in human (Schwartz and Vissing, 2002), in Drosophila (Kondo et al., 1992), in mouse (Gyllensten et al., 1991), and in sheep (Zhao et al., 2004), whereas in the marine mussel Mytilus edulis, the paternal transmission of mtDNA is more common (Sutherland et al., 1998).

III. mtDNA Content in Oocytes A. Oogenesis Is Associated with Drastic Restriction/Amplification of mtDNA Content Oogenesis involves both growth and diVerentiation of oocytes and takes place in several steps occurring over several years. Oogenesis starts during fetal life, the primordial germinal cells becoming oogonia that proliferate and produce primary oocytes. The growth and diVerentiation of oocytes

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take place at puberty and are intimately linked to folliculogenesis. The follicle is generated from satellite cells of the ovarian stroma. The follicle cells play a trophic and endocrinous role in oocyte maturation and ovulation. At the time of ovulation, oocytes are contained in preovulatory follicles. They are composed of liquid cavity called antrum and of about a million follicular cells surrounding the oocyte (cumulus oophorus), the cell layer in contact with oocytes being the corona radiata. Cytoplasmic communications between follicular cells and oocytes allow exchanges of substances necessary to oocyte growth and maturation. Studies on mtDNA segregation in pedigrees that are aVected by mtDNA mutations show that when a point mutation occurs, there may be a complete switching of mutants to high levels within a single generation (Hauswirth and Laipis, 1982; Marchington et al., 1997; Poulton et al., 1998). Taking into account the large number of mitochondrial genomes in oocytes, it has been postulated that a very small number of mtDNAs could populate the oocyte and thus the organism (Hauswirth and Laipis, 1985). This idea led to the bottleneck theory (Hauswirth and Laipis, 1985). According to this theory, the mitochondrial contingent could be drastically reduced before being strongly amplified during oogenesis. The premigratory primordial germ cells are supposed to contain only a few copies (less than 10) of the founder mitochondrial genomes selected to populate the organism. This very small number of mitochondria per cell could lead to the rigorous selection of cells with the best mitochondrial profile accompanied by the elimination of cells with defective mitochondria. In most cases, this mechanism would tend to eliminate mutated mitochondrial genomes and homogenize mtDNA populations, preserving mitochondrial integrity over the generations. Deleterious mutations such as deletions would be eYciently eliminated and not transmitted maternally to the oVspring. In addition, this restriction/amplification mechanism would drastically reduce or accidentally amplify the rate of point mutations that might be transmitted by the mother (Marchington et al., 1998). Thus, the bottleneck theory tends to explain how mtDNA, which is involved in aging and severe genetic diseases, is ‘‘refreshed’’ and ‘‘purified’’ from one generation to another when it passes through the ‘‘narrow neck’’ of the mitochondrial population during early oogenesis (Jansen and de Boer, 1998). Following the sharp reduction of mtDNA content associated with the bottleneck, germ cells could drastically amplify the selected set of mitochondria by clonal expansion, allowing individuals to carry a homogenous set of mtDNAs. Primordial germ cells into the gonadal ridge could contain up to 100 copies of mtDNA and diVerentiated oogonia as many as 200 copies (Jansen and de Boer, 1998). From the 15th week to the 7th month of fetal life, oogonia are amplified by somatic mitosis. During this step, the mitochondrial mass increases exponentially. After division, the oogonia are transformed into oocytes I that are surrounded by follicle cells. Oocytes I have started their first meiotic

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division and are blocked at the prophase I stage. At this stage, the mtDNA content could be about 6000 copies per cell (Jansen and de Boer, 1998). Besides, oocytic atresia occurs at the same time, leading to the decrease of the follicular stock from 2 million oocytes at birth to about 400,000 at puberty, the process continuing until the stock is finally exhausted at menopause. Interestingly, it has been shown that in mice with a high rate of oocytic atresia, the injection of about 5000 mitochondria (from follicular granulosa cells) into the oocytes strongly decreases their apoptotic rate (Perez et al., 2000). The spontaneous rate of apoptosis significantly falls from 70% in control oocytes (not injected with mitochondria or injected only with a buVer) to only 36% in oocytes that are injected with mitochondria. This suggests that eYcient mitochondrial biogenesis could be an important factor of oocyte selection and survival at this stage. After several years of quiescence (up to 50 years until menopause), during the follicular recruitment for ovulation, the oocyte undergoes a final spectacular growth spurt, its diameter increasing from 30 to 120 mm. At this stage, the oocyte accumulates all the proteins, RNAs, energetic substrates, and organelles that are essential for early embryonic development. The cytoplasmic growth of the oocyte is accompanied by a drastic increase in the mitochondrial mass. For example, mature human oocytes contain an average of 2 105 copies of mtDNA. B. The Oocyte Has the Largest Cellular mtDNA Content of the Organism The mtDNA content varies greatly from cell to cell. Although precise quantifications have not been reported according to all the diVerent types of cell in the human organism, it is estimated that leukocytes and monocytes contain hundreds of mtDNA copies whereas cells with high energy requirements, such as neurons and muscle cells, contain thousands of these molecules, with cells such as fibroblasts and epithelial cells containing an intermediate number (Casula et al., 2005; Miller et al., 2003; unpublished personal data). Mitochondrial quantification in mouse oocytes by means of electron microscope morphometry led to an estimation of 92,500 mitochondria per fertilized oocyte (Piko and Matsumoto, 1976). Later, using the dot blot technique, Piko and Taylor (1987) found an average of 119,000 mtDNA copies in pooled mouse oocytes. Using hybridization techniques on pooled cow oocytes, Michaels et al. (1982) found that oocytes contain an average of 260,000 copies. The emergence of more sensitive techniques such as the polymerase chain reaction (PCR) has allowed quantification in single oocytes. Chen et al. (1995) using competitive PCR and Steuerwald et al. (2000) using real‐time quantitative PCR have quantified the mtDNA content in 9 and 18 single human oocytes, respectively collected after in vitro fertilization (IVF) failure. These studies found 138,000 and 315,000 copies per oocyte, respectively, and

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underscored the large inter‐oocyte variation of the mtDNA content. In order to investigate the potential role of mitochondrial biogenesis in fertility, we collected 113 oocytes that were not fertilized after 72 hours of incubation in an IVF procedure in 43 patients (Reynier et al., 2001). These oocytes were blocked in metaphase II since they presented only one polar body and absence of pronuclei. We quantified mtDNA content in these oocytes by using real‐ time quantitative PCR and found an average of 193,000 (122,000) molecules per cell. In a later study, we found that the average number of mtDNA per cell (256,000 213,000) was not significantly diVerent in 116 immature oocytes (without the first polar body) obtained after the intracytoplasmic sperm injection (ICSI) procedure (May‐Panloup et al., 2005a). Besides, there were no significant diVerences in mitochondrial content between oocytes blocked in prophase I (276,000 238,000) and those blocked between prophase I and metaphase II (227,000 166,000). Thus, the mtDNA content is fixed at the time of ovulation, and is not influenced by nuclear maturation at this stage, that is, at the resumption of meiosis. Overall, in our studies on 229 oocytes, the average number of mtDNA copies is about 225,000. Other reports on the average mtDNA content in human oocytes vary from 138,000 to 795,000 copies (Almeida Santos et al., 2006; Barritt et al., 2002; Chen et al., 1995; May‐Panloup et al., 2005a; Reynier et al., 2001). The discrepancies between these diVerent studies on human oocytes could be due to several factors, in particular, the absence of standardized protocols of DNA extraction and quantification. In addition, the small number of oocytes reported in some of the studies may have biased the results. Moreover, the oocytes analyzed may have constituted heterogeneous samples in terms of the age of the patients, the type of infertility, the protocols for ovarian stimulation, and the criteria used for the selection of oocytes. When added together, the mitochondrial genomes of the oocyte, estimated at about 200,000, contain a total number of nucleotides similar to that of the nuclear genome (16,569 200,000 ¼ 3.3 109). This underlines the importance of mtDNA in these haploid cells since, if we exclude the presence of the first polar body, it represents about half the total DNA content, although the mitochondrial genome qualitatively represents only 0.1% of the human genome sequence. This is not surprising considering the vast cytoplasm of these cells. Although the oocyte cytoplasm contains many other constituents, that is, mRNA, proteins, other organelles, and energetic reserve molecules, the mitochondria is one of the most important, at least in terms of quantity.

C. The mtDNA Content of Oocytes Is Highly Variable One of the most surprising features concerning the mtDNA content of oocytes is the high variability. The 229 oocytes we have studied contained

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from 11,000 to 903,000 mtDNA molecules (May‐Panloup et al., 2005a; Reynier et al., 2001). This 80‐fold variation of mitochondrial content between the diVerent oocytes also concerned those collected from the same women (cohorts). Other studies have also reported considerable inter‐oocyte variation (14,000–700,000) in mtDNA content (Almeida Santos et al., 2006). Although the compensatory increase of mitochondrial mass is frequently associated with mtDNA diseases, such a high variation of mitochondrial content is not usual in other tissues. Taking into account the cell growth that occurs in the later stages of oogenesis, we may postulate that this variation in mtDNA content is directly related to the exceptional rate of cytoplasmic growth. Indeed, it is generally admitted that diVerent levels of maturation coexist in the cohorts of oocytes collected for IVF (Flood et al., 1990), these diVerences in maturation levels could therefore explain the high variability in the mtDNA content of oocytes. Despite the development of techniques used in medically assisted reproduction, the average pregnancy rate obtained after transfer of embryos generated by IVF remains low (about 25%), the implantation rate per embryo being only about 10%. One of the limits of IVF is the diYculty of appreciating oocyte quality. This is of particular importance since embryogenesis stems directly from oogenesis and the ability to produce a developing embryo depends on oocyte quality (Gosden, 2002). Although the maturity of the oocyte nucleus is easily attested by the presence of the first polar body, the essential maturation of the oocyte cytoplasm is difficult to appreciate. The mtDNA content reflecting the mitochondrial mass could be a useful indicator of oocyte maturity (and quality) in experimental models. Unfortunately, although the mtDNA content can be readily determined, the method used leads to the destruction of the oocytes examined. Nevertheless, mtDNA measurements carried out on the unfertilized oocytes of cohorts might prove useful to pinpoint the cytoplasmic immaturity of oocytes in women with repeated IVF failures.

D. The Mitochondrial Content of Oocytes Could Affect Their Fertilizability Oocyte quality ensures its fertilizability and its capacity to produce embryos by IVF. To investigate the relationship between the oocyte mtDNA copy number and its fertilizability, we compared several groups of unfertilized oocytes (Reynier et al., 2001) (Fig. 3). The average mtDNA content measured in unfertilized oocytes was significantly lower (152,000 90,000) in cohorts suVering from fertilization failure (fertilization <20%) compared to cohorts with a normal rate of fertilization (209,000 122,000). Moreover, cohorts with fertilization failure that seemed to be due to defective oocytes

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A

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500

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400 300 200 100 0

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* mtDNA content × 1000

mtDNA content × 1000

600

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Group 1

Group 2

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Group 1

Figure 3 mtDNA content and oocyte fertilizability. (A) Comparison of mtDNA content in oocytes from Group 1 (cohorts with fertilization failure due to presumed oocyte defects) and Group 2 (cohorts with a normal rate of fertilization) (p < 0.02). Results are presented in box‐ and‐whisker form: the box comprises 50% of the values, the line in the box gives the median value, and the whiskers represent the distribution of values (mean 2 SD) (* are outliers). (B) Comparison of mtDNA content in oocytes from Groups 1 and 2 (cohorts with fertilization failure due to sperm defects) (p < 0.0001).

showed significantly lower levels (p < 0.0001) of mtDNA content than cohorts with fertilization failure suspected to be mainly due to a severe sperm defect (255,000 110,000) (Reynier et al., 2001). Significantly, six of the seven oocytes with fewer than 50,000 copies of mtDNA belonged to the group that underwent fertilization failure for unknown reasons. These results suggested that inadequate mitochondrial biogenesis could be linked to abnormal cytoplasmic maturation aVecting oocyte quality and leading to IVF failure. Almeida Santos et al. (2006) also found a significant diVerence concerning mtDNA copy numbers in unfertilized oocytes according to the male or female cause of infertility. This team compared the average mtDNA copy numbers in 35 fertilized human oocytes (from 21 patients) and 65 unfertilized oocytes (from 36 patients); the mean mtDNA copy numbers in these two groups were 250,454 and 163,698, respectively. These results confirm the link between mtDNA content and oocyte fertilizability. Moreover, the influence of mitochondria on fertilization has been experimentally demonstrated in the pig (El Shourbagy et al., 2006). These authors found a significant diVerence in mtDNA content between fertilized and unfertilized pig oocytes. In particular, they showed that the supplementation of oocytes with mitochondria from maternal relatives significantly improved fertilization outcome following both IVF and ICSI. This latter result has to be correlated to several studies that showed that intracytoplasmic injection of donor ooplasm or mitochondria could restore or improve the embryonic

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development or fertility. In animal experiments, normal developmental potential has been restored to eggs with ooplasmic deficiencies by the transfer of ooplasm from normal eggs (Flood et al., 1990; Levron et al., 1996). In humans, inter‐oocyte cytoplasmic transfer has been used in an attempt to overcome pregnancy failure due to poor oocyte quality (Cohen et al., 1997). Although the precise cellular structures and macromolecules thus introduced into the oocyte are unknown, mitochondria could represent an important component. Indeed, the transfer of isolated mitochondria between oocytes is known to increase ATP production in the recipients (Van Blerkom et al., 1998). It is interesting to observe here that mitochondrial biogenesis during spermatogenesis is strictly inverse compared to that occurring during oogenesis. Indeed, spermatogenesis is associated with a drastic reduction in mtDNA content. Mature spermatozoa have been reported to contain mtDNA copy numbers ranging from a few thousand down to ten (Alcivar et al., 1989; Hecht et al., 1984; Kao et al., 2004; Manfredi et al., 1997; Shitara et al., 2000). Spermatozoa with a low mtDNA content appear to have the best fertilization ability (Diez‐Sanchez et al., 2003; May‐Panloup et al., 2003). In our experiment, the spermatozoa with the best fertilization ability were almost totally devoid of mtDNA, whereas those with a higher mtDNA content were of poor quality. Thus, as in the case of oocytes, mtDNA content could be related to the maturation of spermatozoa and thus constitute a marker of eYcient gametogenesis and spermatozoal quality.

E. Ovarian Insufficiency Is Associated with Major mtDNA Depletion In order to further investigate the relationship between defective mitochondrial biogenesis and oocyte immaturity or impairment of oocyte quality, we quantified the mtDNA content of 116 unfertilized oocytes collected from 47 women undergoing the ICSI procedure according to their ovarian profile (May‐ Panloup et al., 2005a). In order to determine the best protocol for ovarian stimulation and oocyte collection, candidates for IVF are classified according to the results of a set of clinical, biological, and ultrasound investigations (Hazout, 1999). The three main categories correspond to women with a normal ovarian profile, an ovarian dystrophy profile, and an ovarian insuYciency profile. The precise pathophysiological mechanisms responsible for ovarian dystrophy and ovarian insuYciency are not well understood. However, ovarian dystrophy is related to the defective quality, fertilization, and cleavage rates of oocytes (Aboulghar et al., 1997; Dor et al., 1990; Engmann et al., 1999; Plachot et al., 2003). Similarly, ovarian insuYciency is associated with cycle cancellation, poor oocyte quality, and low IVF success rates (Jenkins et al., 1991; Pellicer et al., 1987), resulting from a qualitative and quantitative decrease of

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mtDNA content ⫻ 1000

400

300

317

305

n = 39 n = 47

200

100

100 n = 30 Group 1 (normal)

Group 2 Group 3 (ovarian (ovarian dystrophy) insufficiency)

Figure 4 mtDNA content and ovarian profile. Comparison of mtDNA content in oocytes from three groups of patients: Group 1 (normal ovarian profile), Group 2 (ovarian dystrophy), and Group 3 (ovarian insuYciency). There were significantly fewer mtDNA copy numbers in Group 3 than in the other two groups (p < 0.0001).

ovarian reserve (Bancsi et al., 2002; Nasseri et al., 1999). We compared the mtDNA content of oocytes from the three groups of women presenting these diVerent ovarian profiles. The mean mtDNA copy number was not significantly diVerent in ovarian dystrophy compared with controls, but it was significantly lower in oocytes from women with ovarian insuYciency (100,000 99,000, p < 0.0001) (Fig. 4). The two ranges of mtDNA levels were distinct, with no overlapping between patients with ovarian insuYciency and those with a normal profile. These results showed that the depletion of mtDNA content is associated with the impaired oocyte quality observed in ovarian insuYciency. This indicates abnormal mitochondrial biogenesis during oocyte growth and probably reflects cytoplasmic immaturity. In the case of ovarian dystrophy, impaired oocyte maturity could be unrelated to mitochondrial biogenesis. This diVerence in oocyte maturity may explain the clinical results observed in these two syndromes. Indeed, although a general immaturity exists in the cohort of oocytes displaying ovarian dystrophy, a subset of mature oocytes can still develop into good quality embryos, once fertilized (Dor et al., 1990; Engmann et al., 1999; Harman, 1972). This is very diVerent from the case of ovarian insuYciency, which is characterized by poor oocyte quality as well as poor embryo quality. According to recent reports, mtDNA depletion in immature oocytes associated with ovarian insuYciency could be related to mutations in the specific mtDNA polymerase (POLG1) that may be responsible for premature

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menopause (Luoma et al., 2004; Pagnamenta et al., 2006). Taking into account that POLG1 plays a fundamental role in mtDNA replication and that strong links exist between ovarian insuYciency and premature menopause, it is very tempting to speculate that the poor constitutive replication ability of mtDNA due to POLG1 mutations could lead to the same type of mtDNA depletion that we observed.

F. How Does mtDNA Content Influence Oocyte Quality? We have seen that low mtDNA content in oocytes is closely associated with fertilization failure, ovarian insuYciency, and poor oocyte quality. The decreased mtDNA content could be just an indicator of perturbed oogenesis or the main abnormality responsible for poor oocyte quality. It is tempting to think that insuYcient mitochondrial biogenesis and oocyte maturation could be linked to poor energetic production. Indeed, during the precompaction stage, ATP synthesis is largely dependent on OXPHOS (Bavister and Squirrell, 2000; Thompson et al., 2000; Van Blerkom et al., 1998, 2000). The variation of ATP content in human oocytes has been associated with the developmental competence of embryos (Van Blerkom et al., 1995; Wilding et al., 2001). We found no correlation between mtDNA copy numbers and ATP content (unpublished data) that was constant at a concentration of about 2 pmol per oocyte, as described in literature by Van Blerkom et al. (1995). However, since this result does not preclude energy production abnormalities, we believe it would be better to measure the ATP production after ADP stimulation of the oocyte. Moreover, mitochondria are involved in many essential cellular processes other than ATP production so that several functions could be impaired by a low mitochondrial mass.

IV. Variation of mtDNA Content and Transcripts in Early Embryonic Development Fertilization of the oocyte marks the beginning of embryonic development characterized by a succession of cell divisions before implantation and the appearance of the first cell diVerentiation. The first embryonic cell division takes place 11–20 hours after fertilization (Menezo and Renard, 1991). At the 8‐cell stage in mice and at the 16/32‐cell stage in humans and cattle, cell contacts are established and blastomeres are compacted. The embryo is then called the morula. At this stage, each blastomere is polarized, this process being under the control of cell contacts. At the 32‐cell stage in mice and at the 80/100‐cell stage in humans and cattle, some liquid accumulates

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in the central cavity of the embryo which is then called a blastocyst (Menezo and Renard, 1991). At this stage, the peripheral cells of the embryo diVerentiate into epithelial cell types to become the trophectoderm, the internal mass forming the embryo itself. After hatching, that is, the exit from the zona pellucida, the trophectoderm establishes contacts with the maternal endometrium allowing implantation of the embryo. In all mammals, the mitochondria of metaphase II oocytes are small spherical organelles with a few cristae surrounding the electron‐dense matrix (Van Blerkom, 2004). This aspect suggests a relatively weak energetic activity and low ATP production (Dvorak and Tesarik, 1985). Mitochondria appear to be disseminated throughout the ooplasm at this stage (Au et al., 2005). After fertilization, the internal structure of the mitochondria does not change but these organelles undergo a peripronuclear relocalization in response to increased energetic demand. The perinuclear localization appears to be maintained in blastomeres during the first embryonic cell divisions (Barnett and Bavister, 1996; Stojkovic et al., 2001). Inadequate redistribution of mitochondria may result in poor oocyte fertilization and compromise embryo development since it has been shown that there is a decreased number of mitochondria around pronuclei in arrested embryos (Au et al., 2005). Before compaction, most of the ATP requirement is met by mitochondrial OXPHOS but this depends on the species and culture conditions (Biggers and Stern, 1973; Brison and Leese, 1991; Gott et al., 1990; Leese et al., 1993). ATP production, oxygen consumption, and aerobic use of glucose by the embryo start to increase as soon as first divisions occur (Biggers and Stern, 1973; Sturmey and Leese, 2003; Trimarchi et al., 2000; Van Blerkom et al., 1998). At the same time, morphological changes are observed in mitochondria. The mitochondrial matrix is decondensed and the mitochondria acquire tubular structures and the number of cristae increases. Simultaneously, the number of mitochondria with high membrane potential increases (Acton et al., 2004). Mitochondrial activation occurs at diVerent stages according to the species but is most frequently associated with the maternal‐embryo transition (MET) when the embryo itself takes control, as it were, of its own development (Gandolfi and Gandolfi, 2001). In humans, these metabolic and morphologic modifications of mitochondria occur progressively at the morula stage (Dvorak and Tesarik, 1985). During compaction, a metabolic shift occurs leading to the preferential consumption of glucose by anaerobic glycolysis to provide ATP. This metabolic shift is concomitant to the transit of the embryo in the uterine Fallopian tube where the partial oxygen pressure is drastically decreased (Fischer and Bavister, 1993; Thompson et al., 2000). The variation of the mtDNA content during embryogenesis has been evaluated in mouse embryos by the Southern blot technique (Piko and Taylor, 1987). This study was performed on pooled embryos and showed that the mtDNA content was constant from the oocyte stage to the implantation

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stage of the embryo. These results, confirmed in mice by quantitative PCR, suggest that there is no replication of mtDNA before implantation (Thundathil et al., 2005). However, a recent report showed that mtDNA replications occur in pronuclear and two‐cell stage mouse embryos, so the constant level of mtDNA may rather reflect a balance between the degradation and neosynthesis of mitochondrial genomes (McConnell and Petrie, 2004). In addition, knock out of the POLG in mice leads to the arrest of embryonic development at 8 days post‐coitum, suggesting the importance of mtDNA replication at this stage (Hance et al., 2005). In the bovine model, we found a significant drop in mtDNA content between the two‐ and four/eight‐cell stages (135,000 28,000) (Fig. 5), although we did not find any diVerence in mtDNA content between metaphase II oocytes and two‐cell embryos (mean 373,000 63,000 and 371,000 52,000, respectively) (May‐Panloup et al., 2005b). The mtDNA copy number remains constant through the 8/16‐cell stage (163,000 36,000) and the morula stage (180,000 26,000), increasing dramatically at the blastocyst stage (688,000 50,000). These results are in contradiction with

mtDNA copy number (×103)

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Figure 5 Variation of mtDNA content in bovine embryogenesis. Comparison of mtDNA content in bovine oocytes and embryos at various stages of development. The mtDNA content decreases between the 2‐ and 4/8‐cell stages (p ¼ 0.0008); in contrast, it increases sharply between the morula and the blastocyst stages (p < 0.0001). Bars with diVerent superscripts diVer significantly.

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the recent report showing that mtDNA remains stable throughout the preimplantation stage in mouse model; however, the high variability of mtDNA content between individual blastocysts could suggest that some embryos initiate their mtDNA replication before implantation (Thundathil et al., 2005). Other authors have found that the mtDNA content in the eight‐cell stage embryo was much higher than that in unfertilized oocytes in humans, suggesting early active replication (Lin et al., 2004). In our bovine model, the activation of the mtDNA replication is disconnected from the implantation stage (which occurs at 21 days post‐coitum in the cow) and occurs early in embryogenesis. The sudden, sharp reduction of about 60% of the mtDNA content that we observed between the two‐ and four/eight‐cell stages is in favor of an active destruction of mtDNA rather than a reduced turnover of the molecules. It is interesting to observe that the few mitochondria transmitted by the spermatozoa are actively eliminated at the same stage by a mechanism mediated by the ubiquitin/proteasome system. We may speculate that this active destruction concerns a proportion of altered maternal mitochondria. If this was the case, the decrease of mtDNA content could constitute another ‘‘bottleneck,’’ that is, a reduction of mtDNA content followed by the drastic amplification of selected mitochondria, allowing the homogenization of mitochondrial genomes. This hypothesis is supported by the suggestion that the restriction/amplification of mtDNA could occur at multiple steps during oogenesis and embryogenesis (Smith et al., 2002). Preexisting RNAs, proteins, and organelles present in oocytes before fertilization are critical components for early embryonic development before the genome of the embryo begins to be expressed. A minor activation of the embryonic genome expression starts as early as the 2‐cell stage (in cows), whereas a major activation of the embryonic transcription (MET) only occurs at the 8/16‐cell stage (Memili et al., 1998). Indeed, mRNA levels markedly increase at the blastocyst stage. Thus, during the first embryo cell divisions, there is a balanced coexistence and expression of oocyte and embryo transcripts. According to several reports, the onset of mitochondrial transcriptional activity appears to occur at the same time as the MET, whereas the mitochondrial mRNAs remain constant before MET. Interestingly, the inhibition of mtDNA transcription does not impair embryonic development in the mouse until the blastocyst stage (Piko and Chase, 1973). The active transcription of the mitochondrial genome could start at diVerent stages of embryo development, depending on the species. It has been shown to start at the 2‐cell stage in mice, at the 4/8‐cell stage in humans, and at the 8/16‐cell stage in cattle (Telford et al., 1990; Thompson et al., 2000). The expression of factors involved in this activation of mitochondrial expression has recently been investigated. It has been shown that the increase of mRNAs corresponding to NRF1 (the main nuclear factor involved in the coordinated expression of genes coding the mitochondrial proteins) and

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Abundance of transcripts (arbitrary unit)

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Figure 6 Variation of mRNAs content in bovine embryogenesis. Comparison of COX1, TFAM, and NRF1 transcript levels in oocytes and at various stages of bovine embryonic development. Levels of COX1 mRNA remain constant from the oocyte to the 8/16‐cell stage, and then increase sharply from the morula stage onward (p ¼ 0.002). TFAM mRNA was not detected before the morula stage. The abundance of this transcript increased dramatically in blastocysts (p < 0.0001). The abundance of NRF1 transcripts remained practically constant from the oocyte to the morula stage, after which it increased significantly up to the blastocyst stage (p < 0.0001).

TFAM (the main mtDNA transcription/replication factor) occurred at the 8‐cell stage in the mouse embryo, just before the activation of mtDNA transcription (Thundathil et al., 2005). In bovine embryos, we observed that the rate of COX1 mRNAs (mtDNA encoded) was constant in oocytes and fertilized embryos up to the morula stage after which it dramatically increases (May‐Panloup et al., 2005b) (Fig. 6). A similar expression pattern has been shown for cytochrome b (Cyt b) (encoded by mtDNA) mRNAs in cattle (Bilodeau‐Goeseels and Schultz, 1997) and for ND2 and ND6 (encoded by mtDNA) mRNAs in mice (Thundathil et al., 2005). In our study, TFAM mRNA levels were undetectable in oocytes and up to the 8/16‐cell stage embryos. TFAM mRNA started to appear at the morula stage and increased sharply at the blastocyst stage, suggesting that de novo synthesis of this factor was essential for the activation of mtDNA transcription. Indeed, TFAM mRNAs appear at the same time as mtDNA transcription is activated and

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just before the increase of mtDNA content (transcription and replication are coupled in mitochondria). In mice, TFAM transcripts were also almost undetectable and began to increase significantly at the eight‐cell and morula stages (Thundathil et al., 2005). Contrary to TFAM, high levels of NRF1 transcripts were found to preexist in oocytes and remain practically constant after fertilization and up to the morula stage, after which NRF1 transcripts increased significantly up to the blastocyst stage. In the mouse model, NRF1 transcripts also preexist in metaphase II oocytes and were found to remain at a steady state from the eight‐cell to the blastocyst stages (Thundathil et al., 2005). In mice, it was also shown that high levels of mitochondrial RNA polymerase transcripts preexisted in oocytes, whereas transcripts corresponding to the two units of POLG as well as those coding for the MRP RNase and for the single‐strand‐binding protein (SSB), which were very few in metaphase II oocytes, increased significantly at the morula stage (Thundathil et al., 2005). NRF1 and TFAM are essential ubiquitous factors for mtDNA replication and expression. Their critical role in early embryo development has been demonstrated by transgenic experiments showing depletion of mtDNA in TFAM (Larsson et al., 1998) and NRF1 (Huo and Scarpulla, 2001) homozygous knockout mice. TFAM knockout mice showed impaired mitochondrial biogenesis and embryonic development, whereas NRF1 knockout mouse embryos died around the time of implantation. Homozygous NRF1 knockout embryos died significantly earlier (3–6.5 days post‐coitum) than homozygous TFAM knockout mouse embryos (10.5 days post‐coitum). The preexistence of maternal NRF1 mRNAs before MET activation could therefore be essential to the mtDNA activation pathway. We hypothesized that these preexisting maternal NRF1 mRNAs play an essential role in the triggering of mitochondrial biogenesis. These maternally encoded mRNAs could be progressively unmasked to become fully functional and promote the activation of TFAM gene transcription and thus the activation of mtDNA replication and transcription. Finally, ubiquitous factors driving mitochondrial biogenesis also play an essential role in early embryonic development. Contrary to the report on pigs (El Shourbagy et al., 2006), we found no diVerences in mtDNA content between post‐insemination, uncleaved oocytes, and two‐cell stage embryos. However, we observed that bovine oocytes that had failed to cleave contained significantly less COX1 and NRF1 transcripts than two‐cell stage embryos. This finding is in accordance with a study that showed a generalized decreased expression of mitochondrial genes (ND2, COI, COII, ATPase 6, COIII, ND3, ND6, and Cyt b) in unfertilized human oocytes and in arrested embryos (Hsieh et al., 2004). These results support the idea that mitochondrial quality is closely related to the fertilizability of oocytes and to the developmental capacity of the embryo. They are also in agreement with a study that showed that the developmental potential of the embryo depends strongly on the ATP content (Van Blerkom et al., 1995).

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V. mtDNA Mutations in Oocytes and Embryos Cell aging is associated with mtDNA instability particularly in postmitotic tissues (Wallace, 2005) that would be unable to eliminate cells containing damaged mitochondria by cell division and selection. Several reports have described the accumulation of multiple deleted mitochondrial genomes, as well as point mutations and duplications, in various postmitotic tissues and in tissues with a low rate of regeneration (Arnheim and Cortopassi, 1992; Cortopassi and Arnheim, 1990; Lee et al., 1994; Simonetti et al., 1992). The accumulation of mtDNA mutations could be an important mechanism involved in the physiology of aging cells and in pathophysiology of age‐ related diseases. According to the free radical theory of aging, ROS generated through normal metabolism, and especially by the mitochondria respiratory chain, could be responsible for irreversible cell damage (Harman, 1956). mtDNA could be particularly vulnerable to such damage due to its close position to the respiratory chain, the lack of protective histone proteins, and the limited ability of accurate repair mechanisms. A vicious cycle may occur during aging in which somatic mtDNA mutations could lead to respiratory chain dysfunction and enhanced production of ROS that in turn could induce accumulation of mtDNA mutations. The heteroplasmy rate of each age‐related mtDNA mutation is usually very low (less than a few percent) in postmitotic tissues. However, mitochondrial function could be aVected at the individual cellular level by clonal expansion of such mutations (Khrapko et al., 1999; Kopsidas et al., 1998). Recent transgenic mouse models with mtDNA‐mutator phenotype due to a homozygous knockin, expressing a proofreading‐deficient mitochondrial POLG, have demonstrated that accumulation of mtDNA leads to premature aging and death (Kujoth et al., 2005; Trifunovic et al., 2004). These models display a progressive and random somatic accumulation of mtDNA deletions and point mutations that, in turn, cause a progressive respiratory chain deficiency and lead to a premature aging phenotype. Furthermore, it was suggested that the aging phenotype was associated with increased apoptosis (Kujoth et al., 2005). Taken into account its role in mtDNA maintenance, POLG is probably a key protein of the reproductive system. Indeed, in addition to reduced life span and multiple age‐related organ dysfunctions, it is interesting to observe that these mutator mice have reduced fertility (Trifunovic et al., 2004). Moreover, germ line mutations of POLG have been linked to a hereditary syndrome associating premature menopause and parkinsonism. Several authors have suggested that polymorphisms of POLG are associated with male infertility but this result remains controversial (Aknin‐Seifer et al., 2005; Brusco et al., 2006; Jensen et al., 2004; Rovio et al., 2001). Finally, it has been shown that POLG was absolutely essential for mammalian embryonic development

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(Hance et al., 2005). Indeed, disruption of POLG in mice was found to cause an early developmental arrest between 7.5 and 8.5 days post‐coitum, associated with severe reduction of the mtDNA content. Since oocytes are quiescent for several years, they are likely to accumulate such mtDNA mutations. These mutations could lead to mitochondrial dysfunction, to energetic failure, and finally to apoptosis. One of the more frequent mtDNA deletions, the so‐called common deletion of 4977 base pairs, was investigated in oocytes. Chen et al. (1995) found 49% of 104 analyzed unfertilized oocytes (31 donors) containing the common deletion without correlation to maternal age. In a similar experiment, the presence of this common deletion was found in 32.8–47% of human oocytes without any link with maternal age (Brenner et al., 1998). In contrast, unfertilized oocytes from older women were found to be more likely to contain deleted mtDNA (93% of the oocytes from 38‐year‐old patients harbored the common deletion) than oocytes from younger women (28% of the oocytes from <38‐year‐old patients harbored the common deletion) (Keefe et al., 1995). More recently, the increased presence of the common deletion has been reported in metaphase II oocytes from women older than 35 (Chan et al., 2005). These authors also found that the mtDNA copy number was negatively correlated to the donor’s age, although we and others did not find this result (Barritt et al., 2002). The consequence of this mutation on oocyte quality remains hypothetical since the number of the common deletions was low, representing only 0.01–0.1% of the total mtDNA content (Chen et al., 1995). Total amplification of mtDNA by long PCR allows the exhaustive determination of the presence of mtDNA deletions, including the common deletion. The analysis of 35 oocytes by this technique showed that 60% of them did not carry any deletion (Reynier et al., 1998). Taking into account the long life span of female gametes, this result attests to a relatively good stability and maintenance of oocyte mtDNA. However, 40% of the oocytes studied carried between one and three types of large deletion. Incidentally, it was observed that the diVerent oocytes from the same donors carried diVerent types of deletions. This suggests that deletions might occur before division of the precursor cells and may be randomly segregated during early oogenesis, in accordance with the bottleneck theory. Interestingly, the proportion of human embryos harboring the common deletion was significantly lower than that of oocytes (8–20.2%) (Brenner et al., 1998). Similarly, Hsieh et al. (2002) found the common deletion in 66.1% of the unfertilized human oocytes and in only 34.8% of the arrested embryos. Decreased mutated mtDNA was also observed in large mitochondrial rearrangements other than the common deletion (Barritt et al., 1999). Taken together, these results suggest the existence of a strong filtering mechanism, similar to the bottleneck previously evoked, by limiting the transmission of mtDNA rearrangements.

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Assisted medical procreation techniques induce artificial superovulation, using exogenous gonadotropin to produce mature oocytes. The consequences of such ovarian stimulation on mitochondrial function are just beginning to be studied. It was shown that repeated ovarian stimulations could induce oxidative damage and mtDNA mutations (Chao et al., 2005). Indeed, it was found that high ovarian stimulation in mice produced poor quality oocytes associated with a high proportion of multiple rearrangements of mtDNA in the ovaries. Similarly, an increased proportion of mtDNA deletions was found in stimulated oocytes and embryos from Rhesus macaques compared with immature unstimulated oocytes (Gibson et al., 2005). mtDNA is highly polymorphic. Single nucleotide polymorphisms (SNP) are associated with each other in haplogroups and subhaplogroups that reflect the phylogenic evolution of species and migrations (Ruiz‐Pesini et al., 2004). There are an increasing number of reports showing that mtDNA haplogroups, which are probably associated with various mitochondrial respiratory chain activities, can influence human diseases. For instance, haplogroup K is likely to play a protective role in Parkinson’s disease (Ghezzi et al., 2005), whereas haplogroup J could increase the penetrance of mitochondrial optic neuropathies associated with pathogenic mtDNA mutations (Torroni et al., 1997). The influence of mtDNA haplogroups on human fertility has been demonstrated and a correlation has been found between diVerent haplogroups and sperm quality. For example, the haplogroup T is overrepresented in patients with reduced sperm motility and impaired activity of complexes I and IV of the respiratory chain (Ruiz‐Pesini et al., 2000). Similarly, the haplogroup U has been associated with reduced sperm motility and vitality (Montiel‐Sosa et al., 2005). It seems that the molecular consequences of mtDNA polymorphisms could lead to biochemical abnormalities in specific cell types or when associated with environmental factors. The eventuality of the influence of mtDNA haplogroups on oocytes and embryonic competence remains an open question. Polymorphisms in the mtDNA control region have been associated with the proportion of oocytes with high developmental ability recovered by ovum pick up (OPU) in cattle as well as with the rate of transferable nuclear‐transfer embryos (BruggerhoV et al., 2002). In addition, haplogroups have been associated with calving rate in beef cattle (Sutarno et al., 2002). In order to tackle this question, we collected oocytes from donor cows whose oocytes are known to diVer in their developmental capacity, measured by the blastocyst formation rate (Tamassia et al., 2003). To investigate the potential role of mtDNA on oocyte quality, the control region of the six donor cows was sequenced and, after pairwise comparison of polymorphisms, the cows were grouped into two major haplogroups that diVered significantly in ATP content (Tamassia et al., 2004). Moreover, the rate of blastocyst production

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also diVered significantly between the two haplogroups. These results suggest that the developmental competence of the embryo might be influenced by the mitochondrial haplogroups, but this hypothesis remains to be tested at a larger scale.

VI. Conclusions and Perspectives The study of the interaction between the nuclear and the cytoplasmic genetic and epigenetic factors of the oocyte is of special interest and will present a great challenge in reproductive biology research over the coming years. Mitochondria unquestionably play a crucial role in the processes of gamete maturation, fertilization, and embryonic development. Mitochondrial dysfunctions intervene at multiple levels of the reproductive process and are likely to be involved in human fertility defects. The analysis of the molecular mechanisms involved in oocyte mitochondrial biogenesis and metabolism can be expected to lead to new insights in reproductive physiology. Among the numerous oocyte mitochondrial functions and components, mtDNA is of central importance considering its maternal transmission, the strong variation of its content in gametogenesis, and its propensity to accumulate mutations. Future therapeutic strategies aimed against infertility will surely have to take into account the importance of oocyte mitochondrial functions and biogenesis in reproduction.

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Section II Transmission

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Mitochondrial DNA and the Mammalian Oocyte Eric A. Shoubridge and Timothy Wai Department of Human Genetics, Montreal Neurological Institute McGill University, Montreal, Quebec H3A 2B4, Canada

I. II. III. IV. V. VI. VII. VIII. IX.

Introduction mtDNA: Structure, Organization, Replication, and Expression Origin and Development of the Germ Line Transmission of mtDNA in the Female Germ Line OXPHOS, Atresia, and Oocyte Quality mtDNA Copy Number in the Oocyte Male Transmission of mtDNA Clinical Perspectives Conclusions and Future Prospects Acknowledgments References

In mammals, mitochondria and mitochondrial DNA (mtDNA) are transmitted through the female germ line. Mature oocytes contain at least 100,000 copies of mtDNA, organized at 1–2 copies per organelle. Despite the high genome copy number, mtDNA sequence variants are observed to segregate rapidly between generations, and this has led to the concept of a developmental bottleneck for the transmission of mtDNA. Ultrastructural investigations of primordial germ cells show that they contain approximately 10 mitochondria, suggesting that mitochondrial biogenesis is arrested during early embryogenesis, and that the mitochondria contributing to the germ cell precursors are simply apportioned from those present in the zygote. Thus, as few as 0.01% of the mitochondria in the oocyte actually contribute to the oVspring of the next generation. Mitochondrial replication restarts in the migrating primordial germ cells, and mitochondrial numbers steadily increase to a few thousand in primordial oocytes. Genetic evidence from both heteroplasmic mice and human pedigrees suggests that segregation of mtDNA sequence variants is largely a stochastic process that occurs during the mitotic divisions of the germ cell precursors. This process is essentially complete by the time the primary oocyte population is diVerentiated in fetal life. Analysis of the distribution of pathogenic mtDNA mutations in the oVspring of carrier mothers shows that risk of inheriting a pathogenic mutation increases with the

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proportion in the mother, but there is no bias toward transmitting more or less of the mutant mtDNAs. This implies that there is no strong selection against oocytes carrying pathogenic mutations and that atresia is not a filter for oocyte quality based on oxidative phosphorylation capacity. The large number of mitochondria and mtDNAs present in the oocyte may simply represent a genetic mechanism to ensure their distribution to the gametes and somatic cells of the next generation. If true, mtDNA copy number, and by inference mitochondrial number, may be the most important determinant of oocyte quality, not because of the eVects on oocyte metabolism, but because too few would result in a maldistribution in the early embryo. ß 2007, Elsevier Inc.

I. Introduction ATP produced by oxidative phosphorylation (OXPHOS) is essential for the normal function of most eukaryotic cells. This process requires the activity of five multimeric enzyme complexes located in the inner mitochondrial membrane. Electron transport along complexes I–IV of the respiratory chain creates an electrochemical gradient for protons, which is used to drive the synthesis of ATP from ADP and inorganic phosphate by complex V, ATP synthase. The subunits of the OXPHOS complexes are encoded by both the nuclear and mitochondrial (mtDNA) genomes. Of the approximately 80 structural subunits of OXPHOS, 13 are mtDNA encoded, and all are essential for function. Mitochondria and mtDNA are maternally inherited in mammals, and as mitochondria cannot be made de novo, but rather only elaborated from other mitochondria, all of our mitochondria ultimately derive from those in one of our mother’s oocytes. Paternal mitochondria containing mtDNA are present in the zygote; however, they are rapidly degraded in the preimplantation embryo by a process that remains poorly understood. Mutations in mtDNA in humans are an important cause of a group of multisystem disorders whose birth prevalence has been estimated at 1:5000 (Chinnery et al., 2000a; Thorburn, 2004). An understanding of the mechanisms of transmission and segregation of mtDNA in the female germ line is important for genetic counseling and clinical management of patients aVected by these disorders. Further, the introduction of new reproductive technologies, such as cytoplasmic transfer and direct sperm injection, have forced a reevaluation of the potential contribution of exogenous or paternally derived mitochondria to the next generation. Here we review the current knowledge in these areas.

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II. mtDNA: Structure, Organization, Replication, and Expression mtDNA is a double‐stranded circular DNA molecule of approximately 16.5 kb in all mammals in which it has been sequenced (Fig. 1). The two stands are referred to as heavy (H) and light (L), reflecting their behavior in density gradients. Mammalian mtDNA codes for 13 polypeptides, all of which are subunits of the enzyme complexes of the OXPHOS system, and 22 tRNAs and 2 rRNAs, which constitute part of the dedicated mitochondrial translation machinery. The genome is exceedingly compact; there are no introns, and there is only one noncoding (control) region of approximately 1 kb that contains the replication origin for leading strand synthesis (OH), and the promoters for transcription of the H‐ and L‐strands. The mtDNA copy number in somatic cells is generally in the range of 103–104 copies per cell, packaged in a DNA‐protein structure called the nucleoid at approximately 2–10 copies per nucleoid (Legros et al., 2004; Satoh and Kuroiwa, 1991). Gametes are a notable exception: mature oocytes have approximately 105 mtDNAs (Piko and Taylor, 1987) and sperm about 102 (Hecht et al., 1984). Investigation of the protein constituents of the yeast mitochondrial nucleoid by mass spectrometry has revealed a large number of proteins, some of which have dual functions in nucleoid maintenance and tricarboxylic acid cycle activity (Chen et al., 2005; Kaufman et al., 2000). The mitochondrial nucleoid in higher eukaryotes is reported to contain TFAM, a mitochondrial transcription factor, single‐stranded binding protein, Twinkle (a helicase), and at least four additional inner membrane proteins (Bogenhagen et al., 2003; Garrido et al., 2003). TFAM is a basic protein of the HMG box family that is thought to package mtDNA (Alam et al., 2003). Decreasing TFAM levels results in loss of mtDNA; likewise, cells devoid of mtDNA contain no detectable TFAM, leading to the suggestion that mtDNA levels are controlled by TFAM (Ekstrand et al., 2004). The mechanism of mtDNA replication has been the subject of recent controversy. The conventional strand‐displacement model developed over the past two decades by Clayton and colleagues (Clayton, 1991; Shadel and Clayton, 1997) was suggested to be an artifact of biased incorporation of ribonucleotides into the L‐strand (Yang et al., 2002), and a coupled strand‐ synchronous mechanism was proposed in its stead (Bowmaker et al., 2003; Holt et al., 2000). In the strand‐displacement model, leading strand synthesis of the H‐strand starts from OH in the control region and proceeds about two‐ third the way around the molecule until a second origin (OL) is exposed, allowing synthesis of the L‐strand to proceed. The alternative model, based largely on evidence from 2‐D agarose gels, proposes that replication is strand‐ coupled, originating from a broad zone around OH (Bowmaker et al., 2003).

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Figure 1 Mitochondrial disease phenotypes associated with mutations in mtDNA‐encoded genes. Human mtDNA is a circular genome coding for 37 genes. The structural and rRNA genes are shown as shaded bars: COI‐III, subunits of cytochrome c oxidase (complex IV); ND1–5, subunits of NADH CoQ reductase (complex I); ATP6,8, subunits of ATP synthase (complex V); Cyt b, cytochrome b subunit of CoQ‐cytochrome c reductase (complex III). The tRNA genes are depicted as solid circles using the single‐letter amino acid code. The major clinical phenotypes associated with mutations in the individual genes are shown in boxes next to the position of the gene. The extent of the common deletion, which is associated with Kearns–Sayre syndrome and removes five tRNA genes, is indicated by lines inside the genome. CPEO, chronic progressive external ophthalmoplegia; LS, Leigh syndrome; PECM, progressive encephalomyopathy; MELAS, mitochondrial encephalomyopathy, lactic acidosis, and stroke‐like episodes; MERRF, myoclonus epilepsy and ragged red fibers; LHON, Leber’s hereditary optic neuropathy; NARP, neuropathy, ataxia, and retinitis pigmentosa; MILS, maternally inherited Leigh syndrome.

A study in which atomic force microscopy was used to examine mtDNA replicative intermediates in mouse liver was entirely consistent with the strand‐displacement model, and did not show any evidence of the theta structures that would be predicted from the strand‐coupled model (Brown

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et al., 2005). The same study also provided evidence for alternative L‐strand origins, and it was suggested that their presence, in addition to the propensity for branch migration in replicating DNA, might account for the patterns observed by 2‐D gel electrophoresis that were interpreted as evidence for strand‐coupled replication (Brown and Clayton, 2006). Replication is catalyzed by a distinct nuclear‐encoded polymerase, the

‐DNA polymerase, and leading strand synthesis at OH is primed by a short piece of RNA generated by transcription from the L‐strand promoter. Thus, replication of the mitochondrial genome is linked in some way to the expression of mtDNA genes. Both models require a primase to prime replication of the L‐strand, and although a primase activity has been recovered from human mitochondria (Wong and Clayton, 1985), its molecular identity remains unknown. Phylogenetic analysis of eukaryotic homologues of the Twinkle helicase suggests that it might retain a mitochondrial primase function, although the domains associated with this function are not well conserved in mammals (Shutt and Gray, 2006). mtDNA copy number varies widely from cell to cell and is tightly regulated in a cell‐specific fashion; however, replication of mtDNA is not tightly coupled to the cell cycle (Clayton, 1991). Thus, during mitosis some templates may replicate more than once, others not at all. This behavior, coupled with the random distribution of mtDNAs to daughter cells at cytokinesis, provides a mechanism for the segregation of mtDNA sequence variants. It is important to point out that replicative segregation of mtDNA also occurs in postmitotic cells, as mtDNA replication and turnover is an ongoing process throughout their lifetime (Gross et al., 1969). mtDNA is transcribed as three polycistronic units: the entire H and L strands and the two rRNAs, by a single subunit, phage‐like RNA polymerase (Shadel and Clayton, 1997). The rRNAs are transcribed 10–60 times more frequently than the entire H‐strand, a process controlled by a specific termination factor that binds to both the transcription initiation site and a termination site located 30 to the 16S rRNA in the gene coding for tRNALeu(UUR) (Fernandez‐Silva et al., 1997; Martin et al., 2005). Mitochondrial transcription has been reconstituted in vitro and requires only the presence of two transcription factors, TFAM and TFBM (either B1 or B2), and the single‐subunit RNA polymerase (Falkenberg et al., 2002; Gaspari et al., 2004). Maturation of the mitochondrial transcripts requires an RNase P activity, and it is thought that the tRNA genes, which are interspersed between many of the protein‐coding genes, act as signal sequences in this process (Ojala et al., 1981). Mitochondrial translation occurs on a dedicated apparatus located in the mitochondrial matrix that resembles that of prokaryotes (Spremulli et al., 2004). Mitoribosomes are sensitive to antibiotics that inhibit prokaryotic translation, but they are remarkable in that they have a much higher protein/ rRNA ratio (O’Brien, 2002). In fact, almost half of the mitochondrial

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ribosomal proteins have no obvious orthologues with either prokaryotic ribosomal proteins or those in cytoplasmic ribosomes. The initiation, elongation, and termination factors necessary for mammalian mitochondrial translation have all been cloned, but the regulation of translation remains poorly understood (Spremulli et al., 2004). All of the proteins involved in the replication and expression of mtDNA are encoded in the nuclear genome and must be targeted to the mitochondria.

III. Origin and Development of the Germ Line The emergence and development of the female germ line in mammals is a complex and dynamic process involving the interaction of the soma and the germ cells. Germ line development begins with the specification of the primordial germ cells (PGCs), which are induced in the primitive ectoderm of the pregastrulating embryo. In the mouse embryo, PGCs are first observed as a founding cluster of approximately 50 cells appearing posterior to the primitive streak between the endoderm and mesoderm of the ventral part of the amniotic fold. These large, round cells were first identified by the intense plasma membrane‐staining activity of tissue‐nonspecific alkaline phosphatase (TNAP), which still remains the de facto histochemical marker for PGCs (Chiquoine, 1954; Ginsburg et al., 1990). Beginning at E6.0, the earliest events of specification were investigated by single‐cell transcriptional profiling of nascent PGCs, leading to the identification of two genes, originally termed Fragilis and Stella (Saitou et al., 2002; Tanaka and Matsui, 2002), that are believed to define PGC competence prior to germ cell fate determination. Further investigation of Fragilis (also referred to as IFITM3), revealed a cluster located on human chromosome 11 containing two other family members, IFITM1 and ‐2 (Tanaka et al., 2005). IFITM1 expression mediates the repulsive activity that drives the migration of PGCs from the mesoderm to endoderm, and while the role of IFITM3 is less clear, it seems that its activity is important in the regionalization of PGCs to the posterior endoderm. After E7.0, signaling from the overlying ectoderm by BMP4/BMP8b (Lawson et al., 1999; Ying et al., 2001) to the precursor population of nascent PGCs is absolutely necessary for further specification of germ cell fate. Mice lacking BMP4 function are unable to produce PGCs, while BMP8b null mice have a severely reduced number of germ cells. Following PGC specification, these cells emigrate from the posterior primitive streak while expressing a subset of well‐characterized and spatiotemporally specific markers of the germ cell lineage. In addition to the IFITMs and Stella, Oct4, which belongs to the POU family of transcription factors, is required to maintain the pluripotency of the germ cell lineage. A transgenic mouse in which the PGC‐specific promoter elements of Oct4 are used to drive

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expression of a GFP reporter has served as a useful tool to track the movements of PGCs in vivo (Molyneaux et al., 2001; Szabo et al., 2002; Yoshimizu et al., 1999). At E8.5, the PGCs reside largely in the stalk of the allantois and the yolk sac splanchnopleure at the posterior end of the primitive streak. By E9.5, the PGCs have increased in number when they exit dorsally and become embedded in the wall of the hindgut endoderm, at that point there are approximately 200 TNAP‐positive cells (Ginsburg et al., 1990). Molyneaux et al. have used time‐lapse movies to show that the motility of PGCs is constant while migrating from the allantois, during which time cell division occurs every 16 hours (Molyneaux et al., 2001; Tam and Snow, 1981). From E9 until gonad colonization, the PGCs develop pseudopodia and other structures capable of ameboid movement (Buehr, 1997), suggesting an active migration through the dorsal mesentery to the genital ridge. However, the proliferation and motility of these cells are not exclusively a cell‐ autonomous process. On the contrary, the somatic cells adjacent to the PGCs also play a role in maintaining the survival of PGCs as they migrate to their final destination. It has been suggested that the invaginating hindgut of the embryo facilitates PGC migration by sweeping the PGCs passively during gastrulation (Buehr, 1997). PGCs apparently do not migrate independently, but rather form intercellular cytoplasmic processes, creating an extensive network (Fox et al., 1981). At E10.5, the gonads begin to form by proliferation of somatic cells at the genital ridge. PGCs begin to leave the hindgut endoderm and move along the dorsal mesentery toward the paired thickenings on the ventromedial surface of the mesonephros. At E11.5, the coalescence of germ and somatic cell leads to the formation of paired gonadal primordia. All the PGCs reach the gonad beginning around E12–13 in the mouse and E35 in humans (Wartenberg et al., 1998), at that time these cells are termed oogonia in females. In mice, the number of oogonia has been estimated at 4000–11,000 per gonad. Sexual dymorphism becomes apparent morphologically as these cells proceed along sex‐specific diVerentiation paths that are influenced by signaling events initiated by adjacent somatic cells of the mesonephros (Godin et al., 1990; Tam and Snow, 1981). Concomitant with these events is the proliferation of oogonia until they become oocytes. These germ cells enter meiosis at around E13.5 in the mouse when expression of markers such as TNAP and Oct4 are downregulated. Concomitant with meiotic arrest at the diplotene stage, the oocyte surrounds itself with a layer of pregranulosa cells of somatic origin, creating a primordial follicle. Primary follicles arise once the proliferating granulosa cells assume a cuboidal shape and the zona pellucida synthesis begins. Oocyte growth rates to this point are rapid, with follicles increasing from 40 (primary follicles) to 140 mm preovulatory follicles found in humans. The establishment of the oocyte pools is a function of three parameters: (1) the number of PGCs that reach the gonad, (2) the mitotic activity of dividing

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Figure 2 Diagrammatic representation of changes in the number of mitochondria during development of the female germ line. An estimate of the number of mitochondria is indicated at each stage of germ line development. A genetic bottleneck for the transmission of mtDNA occurs in the primordial germ cells. Development to the mature oocyte involves at least a 10,000‐fold increase in the number of mitochondria. It is not known how mtDNA copy number

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oogonia, and (3) the attrition of oogonia during oogenesis. The dogma that female germ line stem cells become extinct at birth has been challenged (Johnson et al., 2004), but the interpretation of the data in that study has been seriously questioned (Gosden, 2004). Remarkably, little is known about the metabolic activity or molecular biology of mitochondria during PGC specification and migration, and oogenesis. Ultrastructural studies of human PGCs show that the number of mitochondria increases from about 10 in premigratory PGCs to about 200 in oogonia, and to several thousand by the diplotene stage of meiosis where they arrest (Jansen and de Boer, 1998). There are no data on the fate of mtDNA copy number during this period of development. There are approximately 4500 cells in an E7 mouse embryo and 14,000 cells in an E7.5 embryo (Snow, 1977). If the approximately 100,000 mitochondria in the zygote were simply apportioned to all cells in the early embryo, and there were no mitochondrial replication, PGCs (and somatic cells) would contain between 7 and 22 mitochondria, remarkably similar to what has been reported from transmission EM studies.

IV. Transmission of mtDNA in the Female Germ Line Most mammals have a single mtDNA sequence variant in all of their cells, a condition referred to as mtDNA homoplasmy. The relative rarity of mtDNA heteroplasmy (the occurrence of more than one sequence variant in an individual) and the high degree of population polymorphism suggest that new mtDNA sequence variants are rapidly segregated in maternal lineages. This seemed paradoxical given the high mtDNA copy number (105) in mature oocytes, and the relatively small number of cell divisions in the development of the female germ line, and suggested the existence of a genetic bottleneck for the transmission of mitochondria and mtDNA (Fig. 2). The concept of a bottleneck was first proposed by Hauswirth and Laipis (1982) to explain the rapid segregation of an mtDNA D‐loop sequence variant in several maternal lineages of Holstein cows. Complete switching of the same allelic variant was observed in a single generation in 40% of the mother–daughter pairs examined in another study (Koehler et al., 1991). Segregation of many diVerent pathogenic mtDNA sequence variants has also been observed in a large number of human pedigrees; however, it is

changes during germ line development, but it is thought that there are 1–2 copies per mitochondrion in the mature oocyte. The segregation of mtDNA sequence variants is essentially complete by the time the primary oocytes are diVerentiated in fetal life. The numbers at the left indicate the approximate numbers of cells in the mouse.

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rarely as rapid as that observed in the D‐loop of cows (Chinnery et al., 2000b; Jenuth et al., 1996). The mtDNA copy number is not known in either PGCs or oogonia; however, it has been speculated that mtDNA copy number may be 1 copy per organelle at all stages of gametogenesis (Jansen and de Boer, 1998). The ultrastructural evidence clearly demonstrates a physical bottleneck in the number of mitochondria in the PGC population; however, it is possible that mtDNA replication is not coupled to mitochondrial biogenesis prior to specification of the PGC population such that each mitochondrion in the PGCs contains multiple mtDNAs. Replication of mtDNA does not appear to restart until after the blastocyst stage (Piko and Taylor, 1987; Thundathil et al., 2005), although one study suggests that mtDNA replication occurs during a very narrow window (one‐ to two‐cell stage) of development in the preimplantation embryo, albeit without a change in mtDNA copy number (McConnell and Petrie, 2004). Thus, it is likely that the number of copies of mtDNA in PGCs lies somewhere between 10 and 100. The 10,000‐fold reduction in organelle number from the zygote to the PGCs may simply be the result of a failure to restart zygotic replication of mitochondria, but whether that is the result of the absence of some key factors or mitochondrial biogenesis is actively repressed remains unknown. Studies of transmission of polymorphic mtDNA sequence variants in heteroplasmic mice using single‐cell PCR on PGCs and oocytes (Jenuth et al., 1996) have shown that segregation is essentially complete by the time the primary oocyte population is diVerentiated. Surprisingly, little intercellular variation in the degree of mtDNA heteroplasmy was found among individual PGCs sampled from individual embryos (although it must be said that the number of PGCs analyzed was quite small). These data imply that the PGCs contain a more or less representative sample of the mitochondria in the zygote and that the segregation of mtDNA sequence variants occurs by replicative segregation during the expansion of the oogonial population. The observation that the mean level of heteroplasmy in a large sample of oVspring from single mothers was not significantly diVerent than the level of heteroplasmy in the mother suggests that mtDNA segregation between generations occurs principally by random genetic drift (Jenuth et al., 1996). Studies of homopolymeric tract heteroplasmy in humans are consistent with the mouse studies, showing that segregation has occurred by the time oocytes are mature (Marchington et al., 1997). An attempt was made to estimate the eVective number of segregating units of mtDNA in the germ line using a population genetic model that takes into account the number of mitotic divisions in the development of the germ line, but assumes a constant mtDNA copy number during gametogenesis (Jenuth et al., 1996). While the latter is clearly not the case (mtDNA copy number steadily increases during germ line development), the analysis at least provides a

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basis for comparison, although it will tend to overestimate the number of segregating units, as the rate of mtDNA segregation is inversely related to copy number. Using this model, the number of segregating units of mtDNA was estimated at 200 in the mouse (Jenuth et al., 1996), in broad agreement with estimates obtained from human pedigrees segregating pathogenic point mutations, and nearly identical to that obtained by direct evaluation of heteroplasmy in the oocytes from a patient carrying a pathogenic tRNA mutation (Brown et al., 2001). However, it does not seem that all mutations are treated equally in the germ line. Two studies have investigated oocytes or early embryos from mothers carrying the T8993C neuropathy, ataxia, and retinitis pigmentosa (NARP) mutation in the ATP6 gene, and have shown almost complete segregation of this mutation, suggesting that the bottleneck for transmission of this mutation might be extraordinarily narrow (Blok et al., 1997; SteVann et al., 2006). Consistent with extreme skewing of heteroplasmy in the germ line in these cases, a disproportionate number of reports of apparently de novo mutations in the ATP6 gene have appeared (White et al., 1999). The pattern of transmission of mtDNAs has also been analyzed in a large number of pedigrees segregating six of the most common pathogenic mtDNA point mutations (Fig. 1): A8344G (MERRF, mitochondrial encephalomyopathy, and ragged red fibers), A3243G (MELAS, mitochondrial encephalomyopathy, lactic acidosis, and stroke‐like episodes), T8993C (NARP), T8993G (NARP), G11778A (LHON, Leber’s hereditary optic neuropathy), and G3460A (LHON) (Chinnery et al., 2000b). If the transmission of pathogenic mtDNAs was stochastic, then the proportions of mutant mtDNAs in the oVspring of carrier mothers should be symmetrically distributed about the value in the transmitting female; in other words, the proportion of mutant mtDNAs would be just as likely to increase as decrease. Transmission of the pathogenic mutation was not diVerent from that predicted by a stochastic model for two of the mutations, and for three of the mutations there was a significant increase in mutant mtDNAs in the oVspring. Only one showed a very small shift toward the wild‐type sequence. It has not been possible to construct animal models segregating specific pathogenic mtDNA, as a method to transform mammalian mitochondria has remained elusive. This problem has been circumvented in one instance by transferring naturally occurring large‐scale mtDNA deletions to one‐cell embryos using enucleated cytoplasts as the transfer vehicle (Inoue et al., 2000). In humans, these large deletions are usually associated with progressive external ophthalmoplegia (PEO) or Kearns–Sayre syndrome (KSS), both of which are nearly always sporadic diseases (DiMauro and Schon, 2003). The mice were able to transmit the deleted species of mtDNA at high levels (greater than 80% mutant mtDNAs in some animals) through three generations, clearly showing that there is no barrier to the transmission of pathogenic mtDNA mutations in this model, and no loss of oocytes with

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high proportions of mtDNA deletions. By contrast, the risk of transmitting a large‐scale mtDNA deletion in humans is only about 4% (Chinnery et al., 2004; SteVann et al., 2006). It is not known why the human and mouse diVer in their ability to transmit these particular mtDNA mutations, but it likely reflects the fact that humans with high proportions of large‐scale mtDNA deletions have a severe clinical phenotype and rarely reproduce. The above data, while still limited, suggest that the transmission of polymorphic and pathogenic mtDNA sequence variants in mammals is primarily a stochastic process and that the presence of a genetic bottleneck in the PGCs results in more rapid sorting out of new mutations than expected is based on the high genome copy number in the zygote.

V. OXPHOS, Atresia, and Oocyte Quality Most of the oocytes that are formed during fetal life die by atresia. About half die before birth and the process of wastage is continuous throughout life. Thus, in humans, of the roughly 106 primary oocytes present at birth, about 3 105 remain at sexual maturity, and 103 at menopause (Perez et al., 2000). An evolutionary argument has been advanced, suggesting that mitochondrial function might be the basis for atresia (Krakauer and Mira, 1999). The argument is based on the strong correlation between the size of the genetic bottleneck for mtDNA and the fraction of atretic cells determined in a number of diVerent animal species. Mitochondrial function has also been suggested to be an important contributor to oopause and oocyte aging (Jansen, 2000; Jansen and de Boer, 1998), and the introduction of mitochondria into rodent oocytes can reduce the number undergoing apoptosis in vitro (Perez et al., 2000). However, several lines of evidence argue against the idea that any of these eVects are based on the capacity for energy production by OXPHOS in the oocyte. First, mtDNA diseases are a significant medical problem. Full‐term, apparently normal babies, with very high proportions of pathogenic mtDNA mutants, are born to heteroplasmic carrier mothers. These babies only become symptomatic after birth when there is increased demand for OXPHOS and an upregulation of mitochondrial biogenesis. The simplest interpretation of these facts is that stringent selection for OXPHOS function does not exist, either during oogenesis or at any stage of fetal life. If atresia were a filter for OXPHOS capacity, it would eVectively prevent the transmission of oocytes with mtDNA mutations because the mutations would trigger a suicide response at some stage of oogenesis. Second, it is very diYcult to imagine how new germ line mutations could be selected against at any stage of oogenesis because they would initially be rare. The expression of a pathogenic phenotype due to an mtDNA mutation usually depends on reaching a critical

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threshold of mutant mtDNAs before a biochemical or clinical abnormality is observable. These thresholds can range from 30 to 85% of mutant mtDNAs (Boulet et al., 1992; Macmillan et al., 1993), so it is unlikely that a small proportion of mutant mtDNAs could be selected against. Even if there were a mechanism to select against such mutants, the number of oocytes harboring such mutations would be orders of magnitude less than the numbers of oocytes that die by atresia. Finally, atresia that occurs late in development is initiated by the somatic granulosa cells (Perez et al., 2000), and it is diYcult to see how the capacity for OXPHOS in the oocyte would initiate an apoptotic cascade in the surrounding follicular cells. What about mitochondrial function and oocyte aging? A number of studies have investigated the occurrence of mutations in aging oocytes, usually using as a marker the presence of the so‐called common deletion associated with KSS (Schon et al., 1989). Although these studies have generally been able to detect this mutation in some fraction of oocytes, the proportion of mtDNAs aVected has always been very small, and there is not a reproducible correlation with maternal age (Barritt et al., 1999; Brenner et al., 1998; Chan et al., 2005; Hsieh et al., 2002; Keefe et al., 1995). It is probably premature to completely discard the notion that some aspect of mitochondrial function is important in oocyte quality, but there is currently no compelling evidence to suggest that this correlates with the capacity for OXPHOS.

VI. mtDNA Copy Number in the Oocyte The idea that mitochondrial function is an important determinant of oocyte quality has led to a recent barrage of studies trying to demonstrate that oocyte mtDNA content, and by inference OXPHOS capacity during early embryogenesis, is a marker for female infertility. In this light, several groups have suggested that oocytes from women with premature ovarian failure or from older donors have a reduced ability to generate viable embryos, owing to a reduced mtDNA copy number. This argument could have some merit, not because of reduced copy number impairs OXPHOS capacity in the oocyte in an important way, but rather because reduced mtDNA copy number (and the associated reduced mitochondrial numbers) in the zygote would be predicted to aVect the distribution of mitochondria in the cells of the early embryo at a period when mitochondrial biogenesis is slowed or arrested. These studies have generated several estimates of mtDNA copy number in presumably healthy metaphase II human oocytes ranging from 50,000 to 1,500,000 copies (Barritt et al., 2002; Chan et al., 2005; May‐Panloup et al., 2005; Reynier et al., 2001; Steuerwald et al., 2000). The range in the estimates of mtDNA copy number (30‐fold) is at least tenfold greater than earlier measurements made by

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Southern blot in murine and bovine oocytes (Michaels et al., 1982; Piko and Taylor, 1987). There are a number of possible explanations for the variability in the estimates of mtDNA copy number in human oocytes as compared to those in other mammals. First, it may simply reflect the fact that the estimates from mice and cows are derived from animals with a more homogeneous nuclear genetic background than the human population. Second, the observed variability may reflect real biological variation in the sample of oocytes used in these studies. For obvious ethical reasons, it is seldom possible to harvest oocytes from normal healthy women for research purposes. Invariably, researchers must turn to in vitro fertilization (IVF) clinics to recuperate ooctyes from women who have given informed consent to donate their surplus oocytes. As a result, the oocyte quality and donor fertility are likely subject to stratification. Thus, it seems unlikely that oocytes harvested from this cohort accurately reflect random sampling from a healthy population. A third and more disconcerting cause of the inter‐ and intrastudy variation in mtDNA copy number variability may have to do with the use of the quantitative real‐time PCR platform that all the recent studies have used to estimate copy number. Real‐time PCR is a method of simultaneous DNA amplification and quantification. After each round of amplification, the DNA is quantified by measuring a fluorescent signal that is generated immediately following the synthesis of DNA, as it intercalates with a fluorescent dye such as SYBR Green. Tracking the fluorescence as the reaction progresses, it is possible to monitor the early cycles of the log–linear phase of amplification. In theory, there exists an inverse relationship between the quantity of starting material and the fluorescence generated during the log–linear phase of amplification. This is reported to the user as a Ct value, which represents the PCR cycle number at which the fluorescence is detectable above a predefined threshold. Using a standard curve generated from serial dilutions of a PCR product or plasmid of known concentration, real‐ time PCR machines can automatically extrapolate unknown starting concentrations from Ct values and provide the user with an absolute starting template number. Under the appropriate conditions, real‐time PCR quantitation of nucleic acid can be a powerful and sensitive platform, rivaling all other detection methods including hybridization techniques (i.e., Southern blots, Northern blots) and traditional endpoint PCR analyses. Nevertheless, real‐time PCR does has its own Achilles heel. Most calculations make the fundamental assumption that PCR eYciency of the amplicon of interest is constant over time and has the same value in all studied samples (i.e., that the amplification eYciency is equal to 2 such that each amplicon is doubled after every cycle). Largely ignored have been the reports that amplification eYciencies can vary over a range of 1.8–2.0 (Ramakers et al., 2003). When this variation in PCR eYciency is taken into account, a real

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tenfold diVerence can turn up as any value between 0.7 and 210 (Ramakers et al., 2003). Thus, without the proper postrun analyses that take into account variation in amplification eYciency, it becomes diYcult to accurately determine sample concentrations, even with a serially diluted standard curve, and hence derive meaningful conclusions from these data. It is not clear that any of the studies on single oocytes have analyzed their data with a rigor that would allow one to conclude that the data reflect real biological variation in the populations that have been sampled. Ramakers et al. (2003) designed an easily implemented, and since widely used, method for the assumption‐free analysis of quantitative real‐time PCR data that uses an empirically determined PCR eYciency in the calculation of starting sample concentration. Briefly, the program uses a sliding ‘‘window‐ of‐linearity’’ function to determine the linear phase of amplification and corresponding amplification eYciency for individual samples and performs a linear regression to determine the starting concentration of each sample. To accurately determine mtDNA content in single oocytes, we suggest the implementation of this program in conjunction with a complementary secondary technique like nucleic acid hybridization (slot blots) on pooled oocytes for the purposes of corroboration. Until such studies are done, we suggest that the reported variation in mtDNA copy number in human oocytes be treated with caution.

VII. Male Transmission of mtDNA There are only rare exceptions to the general rule of uniparental inheritance of organellar genomes, either mitochondrial or chloroplast (Birky, 1995), and it has been suggested that this nearly universal phenomenon may have evolved to limit the spread of selfish genetic elements (Hurst et al., 1996). The lack of a paternal contribution of mtDNA has been attributed to a variety of factors, including dilution and active elimination. That mtDNA content of the mammalian male gamete is estimated to be 3 orders of magnitude less than its female counterpart has caused some to invoke a simple dilution of paternal mtDNA in the oocyte at fertilization as the mechanism that makes transmission to the next generation unlikely. While this may be the most parsimonious explanation for the paucity of paternal mtDNA transmission in mammals, others have invoked a mechanism of active exclusion whereby active destruction of paternal mitochondria (which are present in the sperm midpiece that penetrates the oocyte during fertilization) in the zygote lead to uniparental inheritance of mitochondria and their genome (Sutovsky et al., 1999). The surveillance system that targets paternal mtDNA can apparently be bridged in interspecific crosses as shown in studies with mice (Gyllensten et al., 1991; Kaneda et al., 1995). Gyllensten et al. discovered that successive

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backcrossing between species of mice for up to 26 generations can yield animals harboring low levels of paternally derived mtDNA in all of their tissues. In a similar study using the same interspecific crosses between Mus musculus domesticus M. spretus, paternal mtDNA was detected by PCR analysis in early embryos and in about half of all oVspring generated from these crosses (Shitara et al., 1998). Unlike the previous report, the distribution of paternal mtDNA in tissues obtained from these positive oVspring was nonuniform. Less than 10% of hybrid females generated from this interspecific cross possessed paternal mtDNA in their ovaries, and none of the unfertilized oocytes from these animals were positive for the presence of paternal mtDNA. These observations lead to the notion that sperm from closely related species or subspecies may perchance escape the normal surveillance mechanisms put in place to ensure strict matrilineal transmission of mtDNA. Interestingly, no paternal mtDNA was detected in any of the N2 backcross animals (tested by examining embryos obtained by IVF), suggesting some barrier to transmission even in F1 hybrid animals, and that the genetic factors responsible act as either a dominant or codominant maternal trait. Other mammalian examples of paternal transmission (also referred to as paternal leakage) of mtDNA have been reported in sheep (Zhao et al., 2004) using similar PCR‐based methods. For instance, Zhao et al. demonstrated the occurrence of leakage in two related hybrid families (out of a total of 48) produced by Dorset crossed to Small‐Tail Han sheep. Taken together, these results suggest that the molecular targets for paternal mtDNA elimination in intraspecific crosses is a nuclear‐encoded molecule(s) associated with the sperm midpiece. It has been widely accepted that spermatozoa are translationally silent, but a study has challenged this dogma (Gur and Breitbart, 2006). Gur and Breitbart showed that labeled amino acid incorporation was completely blocked by mitochondrial translation inhibitors, but not by a cytoplasmic 80s ribosomal inhibitor. This suggests that mammalian sperm translates nuclear‐encoded proteins by mitochondrial‐type ribosomes that are important in the fertilization of the oocyte. Sutovsky et al. have suggested that mitochondria are earmarked for destruction by the addition of ubiquitin tags (Sutovsky, 2003; Sutovsky et al., 1999). The ubiquitination of sperm occurs in the male reproductive tract at the secondary spermatocyte/round spermatid stage, is masked during epididymal passage (possibly by disulfide bond cross‐linking), and reappears after fertilization (Sutovsky et al., 2000). Although markers of sperm mitochondria and anti‐ubiquitin antibodies colocalize by immunofluorescence in early embryos, there is no direct evidence that the mitochondria themselves are tagged. The destruction of sperm mitochondria could, however, be prevented by the injection of anti‐ubiquitin antibodies, or an antagonist of proteosome activity (Sutovsky et al., 2000), suggesting that ubiquitination is an important signal in the degradation pathway of paternal mitochondria. However, it is unclear

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how the nuclear genes products required for ubiquitin‐mediated protein degradation can be diVerentially recognized in closely related species by the oocyte proteosome. More recently, it was shown that the paternal mitochondrial genome in fish is actively removed prior to the purported degradation of sperm mitochondria. Using optical tweezers to remove intact sperm from within freshly fertilized fish eggs, Nishimura et al. (2006) showed that paternal mtDNA was in fact actively degraded well before the disappearance of the paternal mitochondria. While the elimination of paternal organellar DNA seems to be a conserved mechanism used to ensure matrilineal transmission (Nishimura et al., 2006), the nature of such a nucleolytic mechanisms remains a mystery. Currently, there are no data on how paternal mtDNA is actively eliminated from the mammalian oocyte. A single case of paternal transmission of a pathogenic mtDNA mutation was reported in an individual with a severe muscle myopathy (Schwartz and Vissing, 2002). This case prompted other investigators to search for paternal transmission retrospectively in a large group of patients (Filosto et al., 2003; Taylor et al., 2003). However, no evidence of paternal transmission was found in these studies, suggesting that it is likely to be an extremely rare event in human biology.

VIII. Clinical Perspectives Our current understanding of the transmission and segregation of pathogenic mtDNA mutations can be applied in the clinic in two major areas: counseling women who are carriers about recurrence risks and preventing the transmission of high proportions of mutant mtDNAs to their children. Although in principle, it ought to be possible to calculate recurrence risks for these diseases, there are a number of complicating issues. First, the distribution of mutant mtDNAs among tissues is rarely uniform, so it is generally not possible to infer the distribution in the germ cells from a blood or tissue sample. Second, the bottleneck ensures that there will be a range of heteroplasmy in the oocyte population, which is likely to be mutation dependent. Third, the threshold behavior of pathogenic mutations makes it diYcult to precisely predict the severity of the clinical phenotype, especially at intermediate levels of heteroplasmy. The Melbourne group has conducted the most thorough analysis of the prospects for genetic counseling in a large number of pedigrees segregating the ATP6 (8993) gene mutations associated with NARP (White et al., 1999). This analysis showed that the recurrence risk increased with the mutant load in the mother, as predicted from a stochastic model of transmission, but the confidence intervals on the risk estimate of having an aVected child were extremely large, demonstrating that the risk of

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recurrence is not insignificant at any level of heteroplasmy. This is a direct consequence of the genetic bottleneck. As mentioned previously, extreme skewing of heteroplasmy (suggesting a very narrow bottleneck) is often observed (but remains unexplained) in the germ line of individuals with mutations in the ATP6 gene in particular, so this might represent a worst case scenario for estimating recurrence risk. Although the experience is still limited, prenatal genetic diagnosis for these disorders seems promising. The fact that little, if any, segregation of mtDNA sequence variants occurs during fetal life (Jenuth et al., 1997), even those that are pathogenic (Harding et al., 1992; Matthews et al., 1994), suggests that sampling of any fetal tissue (chorionic villus, amniocytes) after establishment of pregnancy ought to provide a reliable indication of the overall mtDNA mutation load. Mosaicism for polymorphic mtDNA sequence variants has been reported in the placenta, but it was only apparent when very small tissue samples were analyzed (Marchington et al., 2006). It is also not known if all pathogenic mtDNA mutations will obey the same rules. A number of prenatal tests have been reported, mostly for the ATP6 mutation associated with NARP (reviewed in Jacobs et al., 2006). The utility of prenatal diagnosis for a broad spectrum of mtDNA mutations will likely only become apparent after years of clinical experience. The prospects for preimplantation genetic diagnosis are also bright. Because the individual’s mtDNA genotype is most likely determined by the relative proportions of wild‐type and mutant mtDNA in the oocyte, the mtDNA genotype of a blastomere sampled from an eight‐cell embryo (which should contain more than 10,000 copies of mtDNA), ought to be a random sample of the oocyte’s mtDNA complement and hence a good predictor of the mtDNA genotype of the fetus. A study using heteroplasmic mice has borne out this prediction (Dean et al., 2003), and confirmed earlier suggestions (Briggs et al., 2000) that the analysis can also be carried out on a polar body, without disturbing the embryo. There is at least one reported case where blastomere biopsy identified two homoplasmic wild‐type embryos and one homoplasmic mutant embryo from a mother carrying the ATP6 gene mutation associated with NARP (SteVann et al., 2006). Implantation of both wild‐type embryos resulted in the birth of a healthy baby. The introduction of new reproductive technologies (intracytoplasmic sperm injection, ICSI) has also given rise to concern that the normal surveillance mechanisms that target sperm for destruction may be abrogated and that some transmission of male mtDNA might occur. A number of studies have investigated this and to date no evidence for male transmission has been observed (Danan et al., 1999; Houshmand et al., 1997; Torroni et al., 1998). It must be said, however, that only blood has been examined in the individuals that have been born using this procedure, and questions remain about whether these studies had the sensitivity to detect paternal mtDNA at

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very low levels, so it may be premature to rule out the possibility of leakage of male mtDNA. Transfer of oocyte cytoplasm has been used in a number of instances in women with repeated implantation failure due to abnormal embryological development (Barritt et al., 2001). It is not, however, at all clear whether the introduction of exogenous mitochondria in these cases played any role in the establishment of a pregnancy. Some of the individuals that have been born as a result of this procedure are demonstrably heteroplasmic for the donor and recipient mtDNA haplotypes in blood and probably other tissues. This situation is analogous to male mtDNA leakage (although on a larger scale), and the consequences of such heteroplasmy are unknown. Although some surprising mtDNA segregation patterns have been observed in heteroplasmic mice (Jenuth et al., 1997), there is no indication that mixing two polymorphic mtDNA haplotypes can cause overt pathology (Battersby and Shoubridge, 2001).

IX. Conclusions and Future Prospects In mammals, all mitochondria and mtDNAs ultimately derive from those contained in an oocyte. Paternal mitochondria are actively eliminated from the early embryo by a process that remains ill‐defined, but which depends on unknown species‐specific targets recognized by components of the oocyte proteome. mtDNA haplotypes are thus propagated clonally along maternal lineages. The segregation of new germ line mtDNA mutations between generations is rapid and primarily stochastic, due to a genetic bottleneck for mtDNA in the PGCs. The eVect of this bottleneck is to rapidly fix neutral or near‐neutral polymorphisms in the population and to expose severe pathogenic mutations to selection. Investigating the mechanisms that produce the bottleneck, and possible variations in its size due to genetic modifiers or to OXPHOS defects, remain important future problems. In this context, it will be important to develop animal models that are segregating specific mtDNA point mutations. However, this will remain an extremely challenging problem until some robust method can be found to either transform mammalian mtDNA or identify and isolate cells containing naturally occurring mtDNA point mutations. Although oocytes contain a very large number of mitochondria, there is no evidence that this relates to a high demand for OXPHOS. The majority of mitochondria in an oocyte, early embryo, or fetus can harbor a pathogenic mutation without obviously aVecting prenatal development. The large number of mitochondria in the mature oocyte may in fact be a genetic, rather than a metabolic device, to ensure that mitochondria are apportioned to the PGCs before mitochondrial biogenesis restarts. The mechanisms that control mitochondrial biogenesis in the germ line and in early embryonic development

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remain completely unknown. The copy number of mtDNAs, and by association the number of mitochondria, might be an important determinant of oocyte quality because too small a number will result in maldistribution in the early embryo. It will be important to determine the independent eVects of varying mtDNA copy number and OXPHOS capacity on the development of the germ line, and the subsequent eVects on early embryonic development. The results of such investigations could have a major impact on our understanding of some causes of female infertility.

Acknowledgments Research in the author’s laboratory is supported by grants from the CIHR, NIH, Muscular Dystrophy Association (USA and Canada), and the March of Dimes. E.A.S. is a Senior Scientist of the CIHR and an International Scholar of the Howard Hughes Medical Institute. T.W. is supported by Canada Graduate Scholarship for Doctoral Research from the CIHR.

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Molyneaux, K. A., Stallock, J., Schaible, K., and Wylie, C. (2001). Time‐lapse analysis of living mouse germ cell migration. Dev. Biol. 240, 488–498. Nishimura, Y., Yoshinari, T., Naruse, K., Yamada, T., Sumi, K., Mitani, H., Higashiyama, T., and Kuroiwa, T. (2006). Active digestion of sperm mitochondrial DNA in single living sperm revealed by optical tweezers. Proc. Natl. Acad. Sci. USA 103, 1382–1387. O’Brien, T. W. (2002). Evolution of a protein‐rich mitochondrial ribosome: Implications for human genetic disease. Gene 286, 73–79. Ojala, D., Montoya, J., and Attardi, G. (1981). tRNA punctuation model of RNA processing in human mitochondria. Nature 290, 470–474. Perez, G. I., Trbovich, A. M., Gosden, R. G., and Tilly, J. L. (2000). Mitochondria and the death of oocytes. Nature 403, 500–501. Piko, L., and Taylor, K. D. (1987). Amounts of mitochondrial DNA and abundance of some mitochondrial gene transcripts in early mouse embryos. Dev. Biol. 123, 364–374. Ramakers, C., Ruijter, J. M., Deprez, R. H., and Moorman, A. F. (2003). Assumption‐free analysis of quantitative real‐time polymerase chain reaction (PCR) data. Neurosci. Lett. 339, 62–66. Reynier, P., May‐Panloup, P., Chretien, M. F., Morgan, C. J., Jean, M., Savagner, F., Barriere, P., and Malthiery, Y. (2001). Mitochondrial DNA content aVects the fertilizability of human oocytes. Mol. Hum. Reprod. 7, 425–429. Saitou, M., Barton, S. C., and Surani, M. A. (2002). A molecular programme for the specification of germ cell fate in mice. Nature 418, 293–300. Satoh, M., and Kuroiwa, T. (1991). Organization of multiple nucleoids and DNA molecules in mitochondria of a human cell. Exp. Cell Res. 196, 137–140. Schon, E. A., Rizzuto, R., Moraes, C. T., Nakase, H., Zeviani, M., and DiMauro, S. (1989). A direct repeat is a hotspot for large‐scale deletion of human mitochondrial DNA. Science 244, 346–349. Schwartz, M., and Vissing, J. (2002). Paternal inheritance of mitochondrial DNA. N. Engl. J. Med. 347, 576–580. Shadel, G. S., and Clayton, D. A. (1997). Mitochondrial DNA maintenance in vertebrates. Annu. Rev. Biochem. 66, 409–435. Shitara, H., Hayashi, J. I., Takahama, S., Kaneda, H., and Yonekawa, H. (1998). Maternal inheritance of mouse mtDNA in interspecific hybrids: Segregation of the leaked paternal mtDNA followed by the prevention of subsequent paternal leakage. Genetics 148, 851–857. Shutt, T. E., and Gray, M. W. (2006). Twinkle, the mitochondrial replicative DNA helicase, is widespread in the eukaryotic radiation and may also be the mitochondrial DNA primase in most eukaryotes. J. Mol. Evol. 62, 588–599. Snow, M. H. (1977). Gastrulation in the mouse: Growth and regionalization of the epiblast. J. Embryol. Exp. Morphol. 42, 293–303. Spremulli, L. L., Coursey, A., Navratil, T., and Hunter, S. E. (2004). Initiation and elongation factors in mammalian mitochondrial protein biosynthesis. Prog. Nucleic Acid Res. Mol. Biol. 77, 211–261. SteVann, J., Frydman, N., Gigarel, N., Burlet, P., Ray, P. F., Fanchin, R., Feyereisen, E., Kerbrat, V., Tachdjian, G., Bonnefont, J. P., Frydman, R., and Munnich, A. (2006). Analysis of mtDNA variant segregation during early human embryonic development: A tool for successful NARP preimplantation diagnosis. J. Med. Genet. 43, 244–247. Steuerwald, N., Barritt, J. A., Adler, R., Malter, H., Schimmel, T., Cohen, J., and Brenner, C. A. (2000). Quantification of mtDNA in single oocytes, polar bodies and subcellular components by real‐time rapid cycle fluorescence monitored PCR. Zygote 8, 209–215. Sutovsky, P. (2003). Ubiquitin‐dependent proteolysis in mammalian spermatogenesis, fertilization, and sperm quality control: Killing three birds with one stone. Microsc. Res. Tech. 61, 88–102.

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Sutovsky, P., Moreno, R. D., Ramalho‐Santos, J., Dominko, T., Simerly, C., and Schatten, G. (1999). Ubiquitin tag for sperm mitochondria. Nature 402, 371–372. Sutovsky, P., Moreno, R. D., Ramalho‐Santos, J., Dominko, T., Simerly, C., and Schatten, G. (2000). Ubiquitinated sperm mitochondria, selective proteolysis, and the regulation of mitochondrial inheritance in mammalian embryos. Biol. Reprod. 63, 582–590. Szabo, P. E., Hubner, K., Scholer, H., and Mann, J. R. (2002). Allele‐specific expression of imprinted genes in mouse migratory primordial germ cells. Mech. Dev. 115, 157–160. Tam, P. P., and Snow, M. H. (1981). Proliferation and migration of primordial germ cells during compensatory growth in mouse embryos. J. Embryol. Exp. Morphol. 64, 133–147. Tanaka, S. S., and Matsui, Y. (2002). Developmentally regulated expression of mil‐1 and mil‐2, mouse interferon‐induced transmembrane protein like genes, during formation and diVerentiation of primordial germ cells. Mech. Dev. 119(Suppl. 1), S261–S267. Tanaka, S. S., Yamaguchi, Y. L., Tsoi, B., Lickert, H., and Tam, P. P. (2005). IFITM/Mil/ fragilis family proteins IFITM1 and IFITM3 play distinct roles in mouse primordial germ cell homing and repulsion. Dev. Cell 9, 745–756. Taylor, R. W., McDonnell, M. T., Blakely, E. L., Chinnery, P. F., Taylor, G. A., Howell, N., Zeviani, M., Briem, E., Carrara, F., and Turnbull, D. M. (2003). Genotypes from patients indicate no paternal mitochondrial DNA contribution. Ann. Neurol. 54, 521–524. Thorburn, D. R. (2004). Mitochondrial disorders: Prevalence, myths and advances. J. Inherit. Metab. Dis. 27, 349–362. Thundathil, J., Filion, F., and Smith, L. C. (2005). Molecular control of mitochondrial function in preimplantation mouse embryos. Mol. Reprod. Dev. 71, 405–413. Torroni, A., D’Urbano, L., Rengo, C., Scozzari, R., Sbracia, M., Manna, C., Cavazzini, C., and Sellitto, D. (1998). Intracytoplasmic injection of spermatozoa does not appear to alter the mode of mitochondrial DNA inheritance. Hum. Reprod. 13, 1747–1749. Wartenberg, H., Hilscher, B., and Hilscher, W. (1998). Germ cell kinetics during early ovarian diVerentiation: An analysis of the oogonial cell cycle and the subsequent changes in oocyte development during the onset of meiosis in the rat. Microsc. Res. Tech. 40, 377–397. White, S. L., Collins, V. R., Wolfe, R., Cleary, M. A., Shanske, S., DiMauro, S., Dahl, H. H., and Thorburn, D. R. (1999). Genetic counseling and prenatal diagnosis for the mitochondrial DNA mutations at nucleotide 8993. Am. J. Hum. Genet. 65, 474–482. Wong, T. W., and Clayton, D. A. (1985). Isolation and characterization of a DNA primase from human mitochondria. J. Biol. Chem. 260, 11530–11535. Yang, M. Y., Bowmaker, M., Reyes, A., Vergani, L., Angeli, P., Gringeri, E., Jacobs, H. T., and Holt, I. J. (2002). Biased incorporation of ribonucleotides on the mitochondrial L‐strand accounts for apparent strand‐asymmetric DNA replication. Cell 111, 495–505. Ying, Y., Qi, X., and Zhao, G. Q. (2001). Induction of primordial germ cells from murine epiblasts by synergistic action of BMP4 and BMP8B signaling pathways. Proc. Natl. Acad. Sci. USA 98, 7858–7862. Yoshimizu, T., Sugiyama, N., De Felice, M., Yeom, Y. I., Ohbo, K., Masuko, K., Obinata, M., Abe, K., Scholer, H. R., and Matsui, Y. (1999). Germline‐specific expression of the Oct‐4/ green fluorescent protein (GFP) transgene in mice. Dev. Growth DiVer. 41, 675–684. Zhao, X., Li, N., Guo, W., Hu, X., Liu, Z., Gong, G., Wang, A., Feng, J., and Wu, C. (2004). Further evidence for paternal inheritance of mitochondrial DNA in the sheep (Ovis aries). Heredity 93, 399–403.

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Mitochondrial Disease—Its Impact, Etiology, and Pathology R. McFarland, R. W. Taylor, and D. M. Turnbull Mitochondrial Research Group, School of Neurology, Neurobiology, and Psychiatry, The Medical School, University of Newcastle upon Tyne Newcastle upon Tyne NE2 4HH, United Kingdom

I. Introduction A. Structure and Function of the Mitochondrion II. Mitochondrial Genetics A. Homoplasmy, Heteroplasmy, and Threshold B. Transmission, Segregation, and the Mitochondrial Genetic Bottleneck C. Transcription, Translation, and Replication of mtDNA III. Impact A. Epidemiology B. Economic Impact of Mitochondrial Disease C. Mitochondrial Involvement in Common Diseases IV. Etiology A. Making the Diagnosis B. What Is Causing the Problem? V. Pathology A. Organ‐Specific Mitochondrial Diseases B. Classical Mitochondrial Syndromes C. Childhood‐Specific Mitochondrial Syndromes VI. Conclusions Acknowledgments References

Mitochondria are ubiquitous organelles that are intimately involved in many cellular processes, but whose principal task is to provide the energy necessary for normal cell functioning and maintenance. Disruption of this energy supply can have devastating consequences for the cell, organ, and individual. Over the last two decades, mutations in both mitochondrial DNA (mtDNA) and nuclear DNA have been identified as causative in a number of well‐characterized clinical syndromes, although for mtDNA mutations in particular, this relationship between genotype and phenotype is often not straightforward. Despite this, a number of epidemiological studies have been undertaken to assess the prevalence of mtDNA mutations and these have highlighted the impact that mtDNA disease has on both the community and individual families. Although there has been considerable Current Topics in Developmental Biology, Vol. 77 Copyright 2007, Elsevier Inc. All rights reserved.

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improvement in the diagnosis of mitochondrial disorders, disappointingly this has not been matched by developments toward eVective treatment. Nevertheless, our understanding of mitochondrial biology is gathering pace and progress in this area will be crucial to devising future treatment strategies. In addition to mitochondrial disease, evidence for a central role of mitochondria in other processes, such as aging and neurodegeneration, is slowly accumulating, although their role in cancer remains controversial. In this chapter, we discuss these issues and oVer our own views based on our cumulative experience of investigating and managing these diseases over the last 20 years. ß 2007, Elsevier Inc.

I. Introduction A. Structure and Function of the Mitochondrion Mitochondria are small polymorphous organelles whose primary function of energy conversion is indispensable to every eukaryotic cell. They are primeval interlopers, primitive bacteria that fostered an endosymbiotic relationship with early eukaryotic cells and eventually became integral to the normal function of those cells as they evolved (Margulis, 1976). Mitochondria have themselves evolved and developed into highly specialized organelles with a key role in eukaryotic cell metabolism. Their capacity for generating adenosine triphosphate (ATP) by oxidative phosphorylation (OXPHOS) has cultivated a dependence on this energy source for normal eukaryotic cell function. The relationship is however reciprocal and mitochondria are critically reliant on importing molecules from the cytosol for a variety of specialized purposes (Lister et al., 2005). Indeed, this import of cytosolic proteins and the loss of the earliest bacterial pathways have been so extensive that only 14–16% of modern mitochondrial protein content (or proteome) can be traced back to the original bacterial endosymbiont (Gabaldon and Huynen, 2004). Structure and function of the mitochondrion are intimately related and two lipid membranes compartmentalize the organelle. The inner mitochondrial membrane houses the OXPHOS enzymes (the mitochondrial respiratory chain) and provides a highly eYcient barrier to ionic diVusion, a crucial factor in generating the proton gradient necessary to produce ATP (Fig. 1). This membrane also envelops the matrix, which contains the enzymes of the tricarboxylic acid (TCA) cycle and ‐oxidation, involved in the metabolism of carbohydrates and fats, respectively. An outer porous membrane allows passive diVusion of low molecular weight substances between the cytosol and the intermembrane space. Although often considered as a single entity, mitochondria vary substantially in their morphology across species, cell type, and even phase of the cell cycle. They are constantly striving to maintain structural integrity through a series of fission and fusion events

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OM H+

I

II

H+

H+

H+

III

IV

V

IM

Electron transfer ADP + Pi

7 41

4

ATP

1

3

2

10

10

12

Figure 1 The process of OXPHOS in mitochondria. A schematic representation of the process of OXPHOS. Complex I (NADH–ubiquinone oxidoreductase) accepts electrons from substrates, such as glutamate, pyruvate, and ‐hydroxybutyrate, while succinate donates an electron at complex II (succinate–ubiquinone oxidoreductase). Ubiquinone (Q) and electron transfer factor (ETF) then ‘‘shuttle’’ electrons to complex III (ubiquinol–cytochrome c reductase) where reduction of cytochrome c (Cu) enables transfer of electrons to complex IV (cytochrome c oxidase). In this way, electrons pass along the ‘‘chain’’ of complexes (I–IV) and, in doing so, provide suYcient energy to fuel proton pumping from the matrix across the membrane at complexes I, III, and IV. The electrochemical gradient generated by the extrusion of protons is then utilized by complex V, adenosine triphosphate (ATP) synthase, to generate ATP from the condensation of inorganic phosphate (Pi) and adenosine diphosphate (ADP). A circle below the complex indicates mtDNA‐ encoded subunits. Two chromosomes indicate nDNA‐encoded components. (IM, inner membrane; OM, outer membrane.)

that allow communication with other mitochondria (Margineantu et al., 2002). Mitochondrial OXPHOS ultimately results in the condensation of inorganic phosphate and adenosine diphosphate (ADP) to produce ATP, a readily utilizable and essential energy source for the eukaryotic cell (Fig. 1). This process is dependent on five multisubunit polypeptide complexes (I–V) located within the inner mitochondrial membrane, only one of which (complex II) is wholly encoded by the nuclear genome. The other complexes comprise subunits encoded by both the nuclear and mitochondrial genomes (Table I and Fig. 1). These polypeptide complexes utilize flavins, nicotinamides, cytochromes, iron–sulfur centers, and in the case of complex IV,

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Table I Subunit Composition of the Mitochondrial Respiratory Chain Complexes and Mitochondrial ATP Synthase Complex I II III IV V

Subunits

Nuclear‐Encoded

Mitochondrial‐Encoded

48 4 11 13 14

41 4 10 10 12

7:MTND1–6 and MTND4L 0 1:MTCYB 3:MTCO1–3 2:MTATP6 and MTATP8

copper ions to transfer electrons by a series of concomitant oxidation and reduction steps. Through this succession of oxidoreduction reactions, electrons pass along the ‘‘mitochondrial respiratory chain’’ of complexes (I–IV) and, in doing so, generate an electrochemical gradient by fueling the extrusion of protons from the matrix across the inner membrane at complexes I, III, and IV. ATP is then generated by the dissipation of this proton gradient through complex V (ATP synthase). This process of ATP production is essential for normal functioning of all eukaryotic cells, but mitochondria are particularly numerous in cells with high metabolic demands such as neurons and skeletal muscle. The exclusive niche that mitochondria occupy in the metabolism of the eukaryotic cell is central to their role in human disease. The cells dependence on ATP derived from OXPHOS makes it extremely vulnerable to mitochondrial malfunction and even small reductions in the eYciency of ATP production in some tissues may be suYcient to cause symptoms. The aim of this chapter is to discuss the molecular genetic mechanisms that impair the eYciency of ATP production, the variety of phenotypes that arise from such cellular energy crises and how these are diagnosed in patients.

II. Mitochondrial Genetics Mitochondria are exceptional in harboring the only extrachromosomal DNA within the cell and are thus under the dual genetic control of both the nuclear and mitochondrial genomes. In humans, the latter is a small (16,569 bp), circular, double‐stranded DNA molecule that encodes 13 polypeptides integral to the OXPHOS system (Fig. 2). It also encodes the RNA machinery (22 transfer ribonucleic acid (tRNAs) and 2 ribosomal ribonucleic acid (rRNAs)) necessary for the intramitochondrial translation of these polypeptides. All other proteins are imported from the cytosol, having been specifically targeted to the mitochondrion.

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5. Mitochondrial Disease—Its Impact, Etiology, and Pathology Several mutations − exercise intolerance, muscle weakness 14709T > C − myopathy, weakness, diabetes

1095T > C, 1555A > G − deafness

F

T

14484T > C - LHON 14459G > A - LHON/dystonia 14487T > C - Leigh/dystonia

D-loop 12S rRNA

V

P

Cyt b

E

16S rRNA

ND6 13513G > A, 13514A > G and others MELAS, Leigh, and overlap syndromes

3243A > G - MELAS/MIDD/CPEO 3271T > C − MELAS ND5

L1

3460G > A - LHON Other mutations - MELAS

L2 S2 H

ND1

I 4300A > G & others - cardiomyopathy

4977-bp deletion CPEO, KSS and Pearson's

Q

M

ND4

ND2

W

A N C Y

11777C > A - Leigh syndrome 11778G > A - LHON

ND 4L ND3 COI

S1

COIII COII

D 7445A > G 7472Cins and others deafness, myopathy

ATP ATP 6 8

R

G 10158T > C, 10191T > C Leigh/Leigh - like syndrome

K 8993T > G/C, 9176T > G/C NARP and MILS

8344A > G, 8356T > C MERRF

Figure 2 The mitochondrial genome and selected mutations. This figure illustrates the double‐ stranded circular nature of mtDNA. The genome is punctuated by mitochondrial tRNAs colored in this diagram according to their origin on the light (green) or heavy (gray) strand. Numerous point mutations and their associated phenotypes are shown to demonstrate the genetic and clinical heterogeneity of mtDNA disease. The arc of the ‘‘common’’ single large‐ scale mtDNA deletion is also shown.

There are a number of important diVerences between mitochondrial and Mendelian genetics that reflect the unusual evolutionary origins of the mitochondrion. Unlike the diploid nuclear genome, the mitochondrial genome is polyploid with multiple copies of mitochondrial DNA (mtDNA) within each mitochondrion and in some cells, such as oocytes, the copy number is in the region of 3 105 mtDNA molecules (Steuerwald et al., 2000). This polyploidy of mtDNA has important implications for the occurrence, transmission, and potential treatment of human mitochondrial disease and gives rise to the concepts of heteroplasmy, homoplasmy, and segregation that are unique to mitochondrial genetics. A. Homoplasmy, Heteroplasmy, and Threshold Homoplasmy refers to the situation where all copies of mtDNA are identical, whereas heteroplasmy describes the coexistence of two (or rarely more) stable mitochondrial genotypes within the same cell, tissue, or organism.

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Inherent and as yet undefined characteristics of some point mutations allow them to aVect all copies of mtDNA (homoplasmic mutations), while others only ever occur in a proportion of the mtDNA present (heteroplasmic mutations). These definitions are almost certainly oversimplifications of a complex situation, where the hostile environment of the mitochondrial matrix rarely permits either uniformity of genotype or establishment of true heteroplasmy. Available evidence suggests that mtDNA is constantly and randomly acquiring mutations, which undergo clonal expansion and subsequent extinction (Coller et al., 2001). These mutations and processes usually occur at low level and are not detectable by routine diagnostic sequencing methods in tissues such as muscle and blood, but they have been identified in colonic crypts where the anaerobically permissive environment is thought to have allowed extensive clonal expansion to occur (Taylor et al., 2003a). Although flawed, these conceptual ideas of homoplasmy and heteroplasmy are nevertheless valuable in considering mtDNA disease. Disease does not occur in everyone with an mtDNA mutation and it is apparent that for some mutations at least a ‘‘threshold’’ for mutated mtDNA must be exceeded before that tissue’s function is compromised and a clinical phenotype is evident (Chinnery et al., 1997). This threshold varies between individuals and from tissue to tissue. For homoplasmic mutations no threshold level exists and individuals may harbor 100% mutated mtDNA yet remain well (McFarland et al., 2002a, 2004b). Other nuclear or environmental factors must be influencing the phenotype in individuals with homoplasmic mutations and some of these will be discussed in this chapter.

B. Transmission, Segregation, and the Mitochondrial Genetic Bottleneck Until recently, it was thought that mtDNA was inherited exclusively through the maternal lineage. While for the purposes of genetic counseling, evolutionary and epidemiological studies this remains true, a single report of paternal inheritance has now been documented (Schwartz and Vissing, 2002). In this case, paternal mtDNA was confined to a single organ (muscle) and was identified through discrepancies in the mtDNA sequence between blood and muscle tissues. However, subsequent reanalysis of mtDNA from blood and muscle tissues from multiple cohorts of patients has failed to identify any other cases of paternal inheritance (Filosto et al., 2003; Taylor et al., 2003b). Mitochondrial disease caused by mtDNA defects can demonstrate remarkable phenotypic and genotypic variability between siblings, a phenomenon directly related to the ‘‘mitochondrial genetic bottleneck’’ (Poulton et al., 1998). Our understanding of this bottleneck has been greatly improved by studies on the transmission of heteroplasmic polymorphisms generated in

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mice by karyoplast transfer (Meirelles and Smith, 1997). For each female mouse, comparison of heteroplasmy levels in oVspring with those of oocytes at diVerent stages of development revealed that the bottleneck occurs early in oogenesis. In applying this finding to pathogenic mutations, consider a heteroplasmic female embryo developing in utero. The primordial germ cells (PGCs) of this embryo contain an mtDNA mutation at varying levels of heteroplasmy. During early oogenesis, these PGCs experience a significant reduction in the number of mtDNA molecules present (the bottleneck). This process is random, but can cause a ‘‘sampling eVect,’’ altering the level of mutant heteroplasmy (Fig. 3). Consequently, some primary oocytes harbor higher levels of mutated mtDNA and others significantly less. When this female embryo eventually reaches adult life only a small proportion of her primary oocytes will mature, but again the selection is thought to be random. If the primary oocyte selected for maturation contains a high level of mutated mtDNA, then the oVspring is

Figure 3 The mitochondrial genetic bottleneck. The primordial germ cells harbor a heteroplasmic mtDNA mutation. During early oogenesis, there is a substantial reduction in the amount of mtDNA present (the bottleneck). The process is random, but can cause a ‘‘sampling eVect’’ that alters the level of mutant heteroplasmy and consequently, some primary oocytes harbor higher levels of mutated mtDNA and others significantly less than the original primordial germ cell.

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likely to be aVected. Thus, it is entirely possible for an unaVected heteroplasmic female to have oVspring that are severely, mildly, or not aVected. In addition to the inherited mutant load from the oocyte, a number of other factors influence the development of disease including tissue‐specific levels of heteroplasmy and threshold. Following fertilization, a heteroplasmic mtDNA point mutation present in the oocyte will segregate to either of the two daughter cells. This segregation is random and one daughter cell may inherit significantly more mutated mtDNA than the other. As this process recurs during organogenesis, it can lead to quite diVerent levels of mutated mtDNA in various tissues (Lightowlers et al., 1997). If a particular tissue’s specific threshold for mutation is exceeded, then organ malfunction and clinical disease will ensue. It has been suggested that this process of mtDNA segregation is not a random event and there is some evidence to support this hypothesis. Shoubridge and coworkers successfully demonstrated that two distinct polymorphisms specific to two diVerent strains of mice are not randomly segregated to the various internal organs of the crossbred mouse (Battersby et al., 2003). How this relates to the situation with pathological mutations in humans is not clear, but it is tempting to speculate that some mtDNA mutations in humans are also actively segregated to particular tissues.

C. Transcription, Translation, and Replication of mtDNA Transcription, translation, and replication of mtDNA are important processes aVecting the generation, establishment, and consequences of mtDNA mutations. Transcription of mtDNA‐encoded genes is initiated at three major, functionally independent initiation sites on the light (L) and heavy (H) strands of the mtDNA molecule, respectively (Clayton, 1992; Shoubridge, 2002). The promoter elements for each strand, heavy (HSP) and light strand promoters (LSP), each include a short (15 bp) consensus sequence that is essential for transcription (Chang and Clayton, 1984). On the H‐strand, transcription most often begins at nucleotide 561 (ITH1) within the HSP adjacent to tRNAPhe, but a second less frequently used site also exists within tRNAPhe at nucleotide 638 (ITH2) (Parisi and Clayton, 1991). These dual initiation sites on the H‐strand are thought to permit more adaptable transcription, allowing independent synthesis of ribosomal RNA (rRNA) and messenger RNA (mRNA) (Montoya et al., 1982). Transcription of the light strand is initiated at nucleotide 407 (ITL1) within the LSP and produces a single polycistronic mRNA. The polycistronic transcripts generated by both strands are processed and subsequently matured to generate rRNA, tRNA, and mRNA molecules.

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Human mtDNA is transcribed by a dedicated mitochondrial RNA polymerase (POLRMT), and requires mitochondrial transcription factor A (TFAM), and either transcription factor B1 (TFB1M) or B2 (TFB2M) (Falkenberg et al., 2002; Fernandez‐Silva et al., 2003). TFAM binds to regulatory elements upstream of the LSP and HSP to form a protein–DNA complex (Fisher et al., 1987). Binding of TFAM induces a change in the conformation of LSP necessary for POLRMT‐dependent promoter recognition (Gaspari et al., 2004). In addition, in vitro studies have indicated that either TFB1M or TFB2M must also be present for transcription to occur (Falkenberg et al., 2002). In vivo studies performed in mice have demonstrated that TFAM also has a crucial role in regulating mtDNA copy number (Ekstrand et al., 2004). The mRNA sequences of mitochondrial transcripts are translated by mitochondrial ribosomal machinery housed within the matrix. The two mitochondrial‐ encoded rRNAs, 12S and 16S, are complemented by in excess of 85 ribosomal proteins imported from the cytosol (Matthes et al., 1982). Processing of the polycistronic transcripts is mediated by folding of the mt‐tRNAs that punctuate the transcribed mRNA (Ojala et al., 1981). mt‐mRNA is translated using a coding system that has several important diVerences from nuclear mRNA (Table II) and in accordance with the minimalist scheme of mitochondrial biology, the mt‐mRNA lacks both the 50 leader sequences that facilitate ribosome binding of nuclear RNAs (Montoya et al., 1981) and the 7‐methylguanylate cap structure, important in ribosome recognition (Grohmann et al., 1978). Pathological disruptions of translation commonly occur with mutations in mt‐tRNA (MTT) and rRNA (MTRNR) genes, but nuclear gene mutations encoding specific mitochondrial ribosomal proteins (MRPs) (Miller et al., 2004) and elongation factors (EFG1) (Coenen et al., 2004) have also been described in some families with generalized translation defects. Each mtDNA molecule appears to replicate individually and despite the high copy number, recombination is thought to occur only in extremely unusual circumstances (Sato et al., 2005; Zsurka et al., 2005). Nevertheless, the existence of significant recombination in humans with its implications for

Table II Codon Usage in Nuclear and Mitochondrial DNA Codon

Nuclear DNA

Mitochondrial DNA

AUA AGA AGG UGA

Isoleucine Arginine Arginine Termination

Methionine Termination Termination Tryptophan

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understanding human evolution, population genetics, and forensic analyses has been (and remains) controversial (Awadalla et al., 1999; D’Aurelio et al., 2004; Elson et al., 2001). The process of mtDNA replication appears to be temporally unrelated to the phase of the cell cycle and in contrast with the replication of nuclear DNA (nDNA) occurs many times per cell cycle (Bogenhagen and Clayton, 1977). Even though each of the mtDNA molecules is replicating independently of each other and of the cell cycle, the mtDNA copy number remains virtually constant—a situation described as ‘‘relaxed replication’’ (Chinnery and Samuels, 1999; Clayton, 1992). A small quorum of proteins known as the ‘‘replisome’’ is necessary for mtDNA replication and includes mtDNA polymerase (POLG), a helicase with 50 ‐30 activity known as twinkle (Korhonen et al., 2003) and the mitochondrial single‐stranded binding protein. Replisome dysfunction is viewed as an increasingly important cause of human disease and mutations have been identified in the catalytic (POLG1) (Naviaux and Nguyen, 2004) and accessory (POLG2) (Longley et al., 2006) subunits of POLG as well as in twinkle (Spelbrink et al., 2001). The precise mechanism of mtDNA replication remains elusive and somewhat controversial, with the two leading proponents advocating irreconcilably diVerent models. One theory advocated by Clayton is the strand displacement model, where a ‘‘leading’’ strand is synthesized for a distance from the origin of H‐strand replication (OH) before a ‘‘lagging’’ strand can be synthesized in the opposite direction from the displaced H‐strand template (Bogenhagen and Clayton, 2003a,b; Clayton, 1982). According to Clayton’s model, synthesis begins at the origin of OH, downstream of the LSP in the D‐loop and proceeds along the parental L‐strand to produce a new H‐strand. When the replication of the H‐strand reaches the origin of L‐strand replication (OL), some two‐thirds of the way around the genome from OH, the parental H‐strand is displaced. This displacement exposes the initiation site for L‐strand synthesis, enabling replication of this strand to occur in the opposite direction to that of the H‐strand. The two strands continue to grow until they form a pair of linked rings that are eventually decatenated to release two daughter molecules. In his development of this model, Clayton also proposed that short mitochondrial transcripts originating from ITL act as primers for the initiation of H‐strand synthesis (Chang and Clayton, 1985; Chang et al., 1985). However, Clayton could find no diVerences between the mechanisms initiating transcription rather than replication priming (Clayton, 1991), and it remains unclear how the two processes are distinguished. An alternative hypothesis endorsed by Holt proposes a synchronous strand‐ coupled synthesis with duplex replication intermediates (Holt et al., 2000). He identified two replication intermediates on two‐dimensional (2‐D) agarose gels, one of which proved resistant to single‐strand nuclease digestion

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and displayed the mobility characteristics expected of strand‐synchronous products. In subsequent experiments, this intermediate appeared to be more abundant, suggesting that coupled leading and lagging strand replication predominates (Yang et al., 2002). Holt’s group has further suggested that mtDNA replication initiates from multiple sites and may initially occur in both directions (Bowmaker et al., 2003). The interpretation of these 2‐D gels has been contested by Clayton and the issue remains unresolved (Brown et al., 2005). It is possible that under diVerent circumstances each model is correct, with for instance one form of replication predominant in mitotic tissue and the other in postmitotic cells.

III. Impact A. Epidemiology Disorders of mitochondrial OXPHOS present at any age and the ubiquitous tissue requirement for ATP can result in almost any organ being aVected (McFarland et al., 2002b). Consequently, patients are frequently seen by a variety of system‐based clinical specialists. This diverse clinical referral pattern means that few centers have amassed significant collective clinical experience of mitochondrial disease and the diagnosis may not be considered until a number of disparate organ systems are involved, for example, diabetes and deafness. Phenotypic diversity and highly variable clinical referral patterns are not helpful in establishing the prevalence of mitochondrial disease. In addition, accurate diagnosis may require biopsy of an aVected tissue, commonly muscle (Taylor et al., 2004). This facility may not be available, particularly for some patient groups such as children where an open biopsy under general anesthetic is preferred. These problems are further compounded by the ethical complexities of presymptomatic genetic testing in adults and children. Leber’s hereditary optic neuropathy (LHON) is characterized by subacute bilateral visual failure in young adults. It is the commonest mtDNA disorder (Chinnery et al., 2000a) with over 95% of patients harboring one of three, usually homoplasmic, point mutations in the MTND genes of complex I. In the Australian population, 2% of persons registered as blind harbor one of these three common LHON mutations (Mackey and Buttery, 1992). The minimum point prevalence of visual failure due to LHON has been recorded as 3.22 per 100,000 (95% CI: 2.47–3.97 per 100,000) in the population of northeast England (Man et al., 2003). The same study also demonstrated a minimum point prevalence for mtDNA LHON mutations of 11.82 per 100,000 (95% CI: 10.38–13.27 per 100,000) (Man et al., 2003). Thus, inheritance of the mutation alone is insuYcient to cause disease and other factors

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must be important in its clinical expression. One such factor is gender: 50% of men with the mutation develop visual failure, whereas 10% of women are clinically aVected. This sex bias stimulated the search for an X‐linked modifying gene and a locus has been identified (Hudson et al., 2005). Another common mtDNA defect is the m.3243A>G point mutation in the mt‐tRNALeu[UUR] gene (MTTL1). There has been great interest in the role of this particular mutation as a cause of diabetes mellitus. In the general diabetic population, the m.3243A>G mutation is found in approximately 1% of Caucasian and Japanese populations. Overall, the prevalence of diabetes in Western Europe is between 3% and 6% of the general population, and the prevalence of mitochondrial diabetes due to the m.3243A>G mutation is estimated at 0.06%, or 60 per 100,000 of the general population (Gerbitz et al., 1995). A number of population‐based studies have contributed to our understanding of the prevalence of mtDNA disease. In the first of these, Majamaa et al. (1998) studied 245,201 adults in northern Finland. They identified individuals with clinical features and family history suggestive of mitochondrial disease, and determined the frequency of the m.3243A>G mutation. Of the 615 patients identified on clinical grounds, 480 were screened for the m.3243A>G mutation and they detected 11 independent maternal pedigrees transmitting the m.3243A>G mutation, giving an overall point prevalence of 16.3 per 100,000 of the adult population (95% C.I.: 11.3–21.4 per 100,000). Subgroup analysis revealed a high prevalence of the m.3243A>G mutation in certain subgroups of the Finnish population. A second study performed in northeast England determined the point prevalence for adult mitochondrial disease for the midyear period of 1997 (Chinnery et al., 2000a). There were 104 cases of mtDNA disease identified with a further 161 maternal relatives at risk of inheriting the mtDNA defect, giving an estimated minimum prevalence of mtDNA defects of 12.48 per 100,000 for the population below retirement age. Following vigorous investigation of family trees from aVected individuals, further studies on the same population indicate that the prevalence is significantly higher, with 1 in 3500 adults at risk of mitochondrial disease (Schaefer et al., 2004). As regards children, Uusimaa et al. (2000), working in Finland, performed a study over a 7‐year period where they identified defects of OXPHOS in 26 children from a pediatric population of 146,482. Surprisingly, despite the high prevalence of the m.3243A>G mutation within the study region, only one of these children had an mtDNA defect (m.3243A>G). This might suggest that although disorders of OXPHOS are as common in children as in adults, they are infrequently due to mutations in mtDNA. However, several novel mtDNA mutations have been described in association with isolated defects of complex I (Kirby et al., 2004; McFarland et al., 2004a) and some of these mutations have occurred spontaneously, only being identified

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through whole‐genome sequencing. It is, therefore, quite possible that mtDNA mutations occur more frequently in the pediatric population than is currently recognized, but their prevalence remains low because they are more often fatal.

B. Economic Impact of Mitochondrial Disease Although mitochondrial disease is now recognized to be considerably more common than previously thought (Schaefer et al., 2004), large‐scale studies of the healthcare costs associated with these diseases are still lacking. The chronic nature and multisystem involvement of mitochondrial diseases means that patients often require many diVerent forms of support and care. For example, the patient with the m.3243A>G MELAS (mitochondrial encephalomyopathy, lactic acidosis, and stroke‐like episodes) mutation has an increased risk of developing diabetes (Damian et al., 1995), with its long‐ term complications of retinal degeneration, neuropathy, arteriopathy, and renal disease. In addition, they are independently at increased risk of deafness, seizures, myopathy, cardiomyopathy, depression, and stroke‐like episodes, all of which contribute to the burden of chronic disability they experience (Engelgau et al., 2004). Consequently, the most severely aVected of MELAS patients will utilize a wide range of health and social care services, including rehabilitation medicine and financial support. In contrast, patients with chronic progressive external ophthalmoplegia (CPEO) due to a single large‐scale mtDNA deletion often experience few symptoms that interfere significantly with their daily lives and may only develop major associated pathology (dysphagia, cardiac arrhythmias, myopathy) after many years of CPEO. Obscuration of vision by ptosis may become a problem, but surgical correction by an experienced ophthalmologist can produce excellent results (Wong et al., 2002). Thus, patients with this form of CPEO frequently do not represent a significant health or social burden during their working lives. It would, however, be misleading to suggest that this is the situation for the majority of patients with mitochondrial disease. While most do not experience the severe complications of MELAS described above, many are nevertheless commonly unable to work as a result of weakness, fatigue, cognitive impairment, or visual failure. Occasionally, a single feature of the disease, such as myopathy, impairs mobility to an extent where a motorized wheelchair and home adaptations are required. Children often experience severe forms of mitochondrial disease with developmental delay, seizures, and alterations of tone being common features. Parents are often the sole care providers for these children and this may in some respects be a ‘‘hidden’’ cost of mitochondrial disease. Although the impact of the health and social care costs of mitochondrial disease has not yet been quantified,

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the prevalence and chronic disability associated with these diseases would suggest that lifetime expenditure per patient is probably high.

C. Mitochondrial Involvement in Common Diseases The role of mitochondrial dysfunction in a number of common conditions including the process of aging has been hotly debated. Evidence is emerging to suggest that mitochondria play a key role in the etiology of a number of neurological disorders, although their contribution to the pathogenesis of other conditions such as cancer remains controversial.

1. Neurodegenerative Diseases Parkinson’s disease (PD) is a chronic neurodegenerative condition with an incidence in developed countries of between 12 and 20 per 100,000 population (Twelves et al., 2003). Mitochondrial dysfunction, and in particular oxidative stress, has been implicated in the pathogenesis of PD since the incidental exposure of humans to 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydopyridine (MPTP) was noted to produce parkinsonism in those exposed (Langston et al., 1983). Subsequent observation of complex I deficiency in some patients with parkinsonism and the transfer of this biochemical deficiency to cybrids [created by the fusion of enucleated patient cells with cells from a diVerent nuclear background that have no endogenous mtDNA ( cells)] fueled speculation that mutations in mtDNA were responsible for PD (Gu et al., 1998; Swerdlow et al., 1996). Some families with mtDNA mutations and parkinsonism have been identified, but exhaustive mitochondrial genome‐sequencing studies suggest that maternally inherited mtDNA mutations are not involved in the pathogenesis of PD (Vives‐Bauza et al., 2002). However, studies on postmortem substantia nigra cells do reveal high proportions of somatic mtDNA deletions in both aged controls and Parkinson’s disease patients (Bender et al., 2006; Kraytsberg et al., 2006). In some cells, there was evidence of mitochondrial respiratory chain compromise in the form of cytochrome c oxidase (COX) deficiency and in these cells the proportion of deleted mtDNA molecules was significantly higher. This finding suggests that these clonally expanded somatic mtDNA deletions are directly responsible for impaired cellular respiration and by implication involved in the pathogenesis of PD and the process of cell loss observed in normal aging. In addition to the mitochondrial genome, the products of more than 850 nuclear genes are also thought to be integral to normal mitochondrial function. Mutations in some of these genes such as PINK1, a gene encoding a mitochondrial protein kinase, have been identified in families with Mendelian inheritance

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of early‐onset PD (Valente et al., 2004). Other mutations aVect the ATP‐ dependent ubiquitin/proteasome system (UPS), which removes damaged or misfolded proteins from neurons (McNaught et al., 2001). Oxidative stress and mitochondrial dysfunction might be predicted to increase protein damage and limit ATP supply, thereby adversely aVecting the function of the UPS. Another approach has investigated the association of mtDNA polymorphisms with the risk of developing PD and Alzheimer’s disease (AD), another chronic neurodegenerative disease where mitochondrial dysfunction and oxidative stress have been considered important in the pathogenesis. Specific sets of single nucleotide polymorphisms scattered throughout the genome define haplogroups and reflect the evolution of a particular maternal lineage. Numerous studies have analyzed the relative risk of developing AD or PD by assessing the prevalence of these diseases in diVerent mitochondrial haplogroups (Chagnon et al., 1999; Ross et al., 2003; van der Walt et al., 2003, 2004). However, risk assessment for PD and AD by haplogroup association has proved diYcult and results are inconsistent. One particular problem faced in these studies has been the substructure that exists in what seem to be otherwise homogenous populations and the influence that this substructure exerts on the association of specific mutations with disease in cases and controls (Helgason et al., 2005). Nevertheless, one study has managed to overcome these population substructure diYculties and compared the frequency of haplogroups in both PD and AD with those in control patients. The UKJT haplogroup cluster was positively associated with a reduced risk of developing PD, but not AD, when compared with controls (Pyle et al., 2005).

2. Cancer An observation by Vogelstein and colleagues in 1998 (Polyak et al., 1998) triggered a flurry of papers exploring the potential link between mtDNA mutations and cancer. Vogelstein had originally noted that human colorectal carcinomas frequently harbored somatic (a mutation confined to a particular tissue) mtDNA mutations, some of which were homoplasmic. Others confirmed these findings in a variety of tumors and leukemias (Fliss et al., 2000; He et al., 2003; Jeronimo et al., 2001; Jones et al., 2001; Kirches et al., 2001). It has, however, proved diYcult to conclude that these mutations actively participate in the development of cancer and both modeling and molecular studies suggest that their accumulation to homoplasmic levels may occur even in the absence of selection (Coller et al., 2001; Taylor et al., 2003a). Although the link with tumor pathogenesis has not been established, the identification of such somatic mtDNA mutations may yet prove useful in the detection of tumor recurrence (Fliss et al., 2000).

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3. Diabetes and Hypertension Diabetes is one of the commonest conditions in the Western world with a prevalence rate in the United States of nearly 10% in those aged over 20 years. Several mitochondrial point mutations, particularly those in mt‐tRNA genes have been associated with the development of diabetes (MITOMAP, 2006), but in such a common disorder the contribution of mitochondrial disease to the overall morbidity of diabetes is small (Saker et al., 1997). However, it has been proposed that a common mtDNA variant (m.16189T>C) is associated with the development of Type 2 diabetes (DM2) and independently with cardiomyopathy, suggesting that mitochondrial polymorphisms may predispose to more familiar diseases (Khogali et al., 2001; Poulton et al., 2002). Again although, diYculty in controlling for population substructure with such a common mtDNA polymorphism undermines these studies and comparison of the poly‐C tract 16184–16193 in patients with DM2 and two independent control groups failed to detect any diVerences—a finding that was confirmed by a meta‐analysis of European studies (Chinnery et al., 2005). Nevertheless, mtDNA mutations continue to be linked with common conditions; most recently hypertension was associated with the matrilineal inheritance of a homoplasmic mt‐tRNA mutation, where family members also suVered from hypomagnesemia and hypercholesterolemia (Wilson et al., 2004). The impact of this finding on the etiology and treatment of hypertension and hypercholesterolemia is potentially huge, but to date no other families with this constellation of ‘‘metabolic’’ defects have been published.

4. Aging The mitochondrial theory of aging proposes that the progressive accumulation of somatic mutations in mtDNA during life leads to an inexorable decline in mitochondrial function (Harman, 1992). Reactive oxygen species (ROS) generated at very low levels during normal mitochondrial respiratory chain activity are believed to create a ‘‘hostile’’ environment in which spontaneous mtDNA mutations are more likely to occur. Some of these mutations will impair respiratory chain function, leading to increased ROS production and further mtDNA mutation. This process is reflected in the increased number of COX‐deficient cells and higher levels of mutated mtDNA observed in the rather unusual environment of aging colonic crypts (Taylor et al., 2003a). Positive feedback between mtDNA mutation and generation of ROS is thought to result in an exponential increase in oxidative damage during aging, with eventual loss of cellular and tissue functions through a combination of energy insuYciency, signaling defects, apoptosis, and replicative senescence.

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To test this hypothesis directly, knockin mice have been developed by two diVerent research groups (Kujoth et al., 2005; Trifunovic et al., 2004). These mice express a proofreading‐deficient version of PolgA, the nuclear‐encoded catalytic subunit of the mtDNA polymerase, and acquire mtDNA mutations at a much higher rate than normal. A point mutation introduced into the exonuclease domain of the PolgA gene does not aVect DNA synthesis capacity, but radically reduces its proofreading ability. Consequently, both PolgA mutator mice accumulate mtDNA point mutations in numerous tissues, reproducing the eVect of aging. The PolgA mutator mice have a phenotype consistent with premature aging, including osteoporosis, reduced activity, and severe weight loss with decreased muscle mass and lipoatrophy. Both models are supportive of somatic mtDNA mutations contributing to aging, yet further studies are required on human tissues to determine whether acquired mtDNA mutations, with subsequent clonal expansion, accumulate in human tissues to the degree found in the animal model.

IV. Etiology A. Making the Diagnosis 1. Clinical Assessment The diVerential diagnosis for patients with defects of OXPHOS is extraordinarily diverse. Furthermore, the investigation of presumed mitochondrial disease is made more diYcult because not only may the same genetic or biochemical defect present in a variety of diVerent ways, but also the same clinical syndrome may be due to a variety of diVerent biochemical or molecular defects (Taylor et al., 2004). A detailed history and examination of the patient with suspected mitochondrial disease is crucial to both initiation and interpretation of the relevant investigations. Common presenting symptoms include fatigue, myalgia, and weakness, but it is important to specifically address issues such as maternal health and obstetric history, family history of neonatal or childhood deaths, deafness, diabetes, cardiac symptoms, visual impairment, and developmental delay. Identification of optic atrophy, ophthalmoparesis, hearing impairment, cardiac enlargement, and the neurological signs associated with muscle, cerebellar, brainstem, and basal ganglia involvement are essential elements of the examination. 2. Clinical Investigation Simple blood tests may provide some supporting evidence for the clinical diagnosis and initial investigations and should include creatine kinase, resting

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blood lactate, full blood count, thyroid and liver function, bone chemistry, random blood glucose, and glycated hemoglobin (HbA1c). All patients should have an ECG to investigate possible conduction defects or cardiac hypertrophy (Santorelli et al., 2001). Chest radiography and echocardiography are then appropriate investigations in those individuals with clinical or ECG evidence of cardiorespiratory involvement. Lumbar puncture is a valuable investigation, particularly in children, as raised cerebrospinal fluid (CSF) lactate and mild protein elevation are consistent with mitochondrial dysfunction (Jackson et al., 1995). However, an increased CSF lactate following seizure activity or stroke, two common features of mitochondrial disease, should always be interpreted with caution. Electromyography may be normal, even in the presence of clinical myopathy and nerve conduction studies can demonstrate either an axonal or a mixed axonal‐demyelinating peripheral sensorimotor neuropathy. A pattern of generalized slow waves, indicative of a subacute encephalopathy, or subclinical seizure activity may be evident on electroencephalogram (EEG) (Canafoglia et al., 2001). Cognitive impairment, central neurological signs, movement disorder, or abnormal EEG all warrant some form of cerebral imaging. A variety of changes can be seen on computed tomography (CT) and magnetic resonance imaging (MRI), some of which are specific for certain mitochondrial disorders (e.g., the symmetrical hypodensities of brainstem, thalamus, and basal ganglia seen in Leigh syndrome), and may indeed be characteristic for a particular mutation (Farina et al., 2002). 3. Muscle Biopsy a. Histology and Histochemistry. Histological and histochemical analyses of muscle biopsy are fundamental to the detection of mitochondrial disease. Muscle biopsies obtained from quadriceps femoris are first orientated and then frozen in an isopentane bath (cooled to 160 C in liquid nitrogen) before being sectioned (8–10 mm sections). Various assays and stains can then be performed on these muscle sections. Ragged red fibers are pathological subsarcolemmal accumulations of mitochondria that have historically been identified using modified Gomori trichrome staining. Assessment of activity of succinate dehydrogenase (SDH), a specific mitochondrial enzyme and one that is unaVected by mutations in mtDNA, appears to be a more sensitive and is also useful in identifying disorders of complex II. It is also frequently used in conjunction with the COX reaction. The COX reaction is aVected by mutations in both mitochondrial and nDNA, as subunits of COX are encoded on both genomes. Significant variation is observed in COX reactivity between type I (oxidative) and type II (glycolytic) fibers; the former reacting strongly, resulting in a dark brown fiber. A mosaic pattern of COX activity is indicative of a heteroplasmic

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mtDNA mutation, the mosaic arising from the variation in mutation load between diVerent fibers (Taylor et al., 2004). In patients where only a very small number of fibers are COX deficient, sequential COX–SDH histochemistry is particularly useful for identifying abnormal fibers that might otherwise remain undetected; COX‐deficient fibers remain dark blue and are easily distinguishable from brown COX‐positive fibers. A global decrease in COX activity has previously been considered as evidence of an nDNA mutation, aVecting either COX subunits or one of the ancillary proteins involved in COX assembly such as SURF (surfeit)1 (Zhu et al., 1998), but homoplasmic tRNA mutations can also produce the same global pattern of COX deficiency (McFarland et al., 2002a). Although often informative, mitochondrial enzyme histochemistry should always be interpreted in the clinical context and with regard to other factors such as patient age and the results of biochemical respiratory chain analysis. Patients with defects involving complexes I, III, or V will have normal COX and SDH reactions and at present there are no histochemical methods of assessing activity of these enzymes. Consequently, patients with clearly defined mitochondrial diseases such as MELAS and CPEO can present with normal muscle histochemistry. In contrast, some elderly patients demonstrate low levels of COX‐deficient muscle fibers. This COX deficiency is not due to mitochondrial disease but rather to clonal expansion of acquired mtDNA deletions within individual fibers. Such focal deficiencies of COX can comprise up to 2% of all fibers in the muscle biopsy from an elderly patient and low levels of COX deficiency must therefore be interpreted with caution in this age group (Brierley et al., 1998).

b. Biochemical Analysis. Although the measurement of mitochondrial enzyme activities is an integral part of the diagnostic process, the protocols are not standardized between laboratories. The availability of fresh or frozen muscle biopsies and the choice of substrates and/or electron acceptors used in the assays complicate the development of universally accepted ‘‘normal ranges’’ for each of the enzyme activities. Rates of flux, substrate oxidation, and ATP production are measured using polarography or 14C‐labeled substrates. Often the most profound deficiencies are observed in children with recessive nuclear mutations and these tend to be isolated enzyme defects (Triepels et al., 2001). In contrast, patients with mtDNA disease have wide‐ ranging results, from normal enzyme activities, through isolated complex deficiency, to multiple enzyme defects involving complexes I, III, and IV. Measurement of each complex activity in isolation avoids some of the diYculties encountered with the linked spectrophotometric assays, where measurement of electron transfer through a section of the respiratory chain

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can obscure a partial defect, because the linked enzyme exerts a greater influence on electron flux through that section of the respiratory chain (Birch‐Machin and Turnbull, 2001). c. Molecular Genetics. The investigation of mitochondrial disease at the molecular level can be complex and should not be undertaken without first reviewing the available clinical, histochemical, and biochemical evidence. Information gleaned from these various sources will determine a rational approach to molecular investigation, as does an understanding of the genotype–phenotype relationship for specific mutations of both nuclear and mtDNA origin. Mitochondrial disease in the pediatric population is frequently the result of autosomal recessive mutations and children with isolated biochemical deficiencies in muscle should be investigated with this in mind (LoeVen et al., 2000). However, approximately 25% of pediatric presentations are due to mutations in mtDNA and may be misdiagnosed because of early‐onset disease and atypical (nonclassical) presentation (McFarland et al., 2004a). Blood is the tissue of choice for investigation of nuclear genetic defects but is generally less useful in this respect for mtDNA mutations. Nevertheless, it is possible to detect high levels of heteroplasmy for some mt‐tRNA point mutations (m.14709T>C and m.8344A>G) (Hammans et al., 1993; McFarland et al., 2004b) and although less reliable, single deletions or rearrangements can also be detected in early childhood. Muscle is the preferred tissue for molecular genetic analysis of mtDNA because it is often an aVected tissue (myopathy, rhabdomyolysis) and for some mutations, the levels of heteroplasmy in muscle parallel those in other aVected postmitotic tissues such as brain (Oldfors et al., 1995). Suspected rearrangements of mtDNA, including single deletions, duplications, and multiple mtDNA deletions, are usually investigated by Southern blot analysis. Cases of mtDNA depletion will also be detected if in addition to a mitochondrial probe, the Southern blot is hybridized simultaneously with a probe to detect a nuclear gene, usually 18S rRNA. It is important to recognize that although remaining the ‘‘gold standard’’ test, Southern blotting will occasionally miss low levels of multiple mtDNA deletions. For this reason, nonquantitative PCR‐based techniques such as long‐range PCR (LRPCR) are often employed where there is a strong clinical suspicion of multiple deletions (e.g., autosomal dominant CPEO phenotype) (He et al., 2002). Being PCR based, this technique preferentially amplifies smaller templates (deleted mtDNA molecules) over full‐length wild‐type mtDNA. These deleted mtDNA molecules can however be found at low levels in otherwise healthy elderly individuals and consequently the results of LRPCR must be interpreted with caution in this age group (Ozawa, 1995). Alternatively, the relative amount of deleted mtDNA can be detected in individual COX‐positive and

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COX‐deficient muscle fibers using real‐time fluorescence PCR (RTPCR). Common point mutations such as m.3243A>G, m.8344A>G, m.8993T>G/C, and m.9176T>G/C are frequently detected and quantified using radioactive (or fluorescent) ‘‘last cycle’’ PCR‐based restriction fragment length polymorphism (RFLP) analyses. Sequencing of the mitochondrial genome is useful for detecting novel mtDNA mutations and to a lesser extent for excluding mtDNA involvement prior to investigating candidate nuclear genes. The ease with which the whole mitochondrial genome can be sequenced has revealed its true diversity with numerous base variations identified (MITOMAP, 2005). DiYculties arise for the clinician in deciding which of these variants are polymorphisms and which are pathogenic. Again the results of previous investigations, and in particular biochemistry, are important in determining the biological plausibility of pathogenicity for any putative mutation. Other criteria for pathogenicity of an mtDNA mutation have been proposed (Table III) by DiMauro and Schon (2001), but these are not without exception and some well‐recognized heteroplasmic mutations fail to satisfy a number of these criteria, particularly with regard to evolutionary conservation. Homoplasmic mutations are an extreme example and cannot fulfill any criteria that specify segregation of mutated mtDNA, irrespective of whether this is at the level of the cell, tissue, individual, or family. Additional ‘‘functional’’ evidence is necessary to demonstrate pathogenicity of these homoplasmic mutations. The contribution that homoplasmic mutations make to the overall morbidity and mortality associated with mitochondrial disease is diYcult to estimate. Their frequency is likely to be underestimated by routinely employed

Table III Canonical Criteria for Pathogenicity After Schon and DiMauro Criteria Determining Pathogenicity of mtDNA Mutations Heteroplasmy Segregates within a family Segregates within an individual Segregates with biochemical defect Unique to aVected families Evolutionary conservation

The mutation must be heteroplasmic (coexistent wild‐type and mutant mtDNA) The proportion of mutated mtDNA should be higher in tissue from aVected individuals than in the same tissue from their unaVected relatives The proportion of mutated mtDNA should be higher in clinically aVected tissues The proportion of mutated mtDNA should segregate with a biochemical defect at the tissue and cellular level The mutation should be absent from healthy controls The mutation should occur at an evolutionary conserved site in the mitochondrial genome

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whole‐genome screening methods such as denaturing high performance liquid chromatography (DHPLC). This technique relies on the formation of heteroduplexes that only occur in the presence of heteroplasmy (Biggin et al., 2005; van Den Bosch et al., 2000). Thus, although whole genome sequencing or screening techniques are increasingly employed, it is important to interpret the results with caution and in the context of clinical, histochemical, and biochemical evidence.

B. What Is Causing the Problem? 1. A Variable Genome The physical proximity of mtDNA to the mitochondrial respiratory chain complexes and the free radicals they generate, together with its lack of protective histones and the absence of an eYcient repair system, encourages mutagenic events at a rate that far exceeds that observed in nDNA. Some benign mutations will quickly reach homoplasmic levels and be inherited through the maternal lineage. A small number of these will become fixed in the population and when found in association with other specific polymorphisms will come to define a particular haplogroup. This process has given rise to at least 18 major haplogroups of human mtDNA (Herrnstadt et al., 2002). Deleterious mutations also occur spontaneously and although they rarely reach homoplasmic levels, they can be inherited to varying degrees in the maternal lineage, causing disease in future generations. Some regions of the mitochondrial genome appear to be more vulnerable to mutation than others and the D‐loop in particular demonstrates considerable polymorphic variation (ExcoYer and Yang, 1999). Defects of mtDNA can have structural or functional consequences that result in impaired OXPHOS. Mutations in MTND or MTCO genes usually disrupt the structural integrity of the protein subunits they encode, leading to isolated biochemical deficiencies of complexes I and IV, respectively (McFarland et al., 2004c; Triepels et al., 2001). Multiple complex deficiencies commonly result from mutations in MTT genes where both structure (tertiary folding) and function (aminoacylation) of the mt‐tRNA molecule are impaired. Consequently, mutations in MTT genes have been associated with mtDNA translation defects that cause multiple deficiencies of complexes I, III, IV, and V in skeletal muscle (Boulet et al., 1992; Enriquez et al., 1995; Hanna et al., 1995; King et al., 1992; Schon et al., 1992). Some of these mutations, and in particular those in MTTK, MTTS1, and MTTL, have been shown to impair aminoacylation of the respective mt‐tRNA with a subsequent decrease in the rate or extent of protein synthesis (Chomyn et al., 2000;

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Hao et al., 2004; Park et al., 2003; Sissler et al., 2004; Toompuu et al., 2002). Such defects in protein translation with their associated multiple biochemical deficiencies are not confined to mutations in MTT genes and are also seen with nuclear gene mutations, including those in EFG1 (Coenen et al., 2004) and MPV17 (Spinazzola et al., 2006). To add further complexity, mutations in MTT genes can cause isolated or multiple respiratory chain deficiencies and although the mechanism for this variation is unclear, it appears to be unrelated to the proportion of mutated mt‐tRNA present in the tissue (Obermaier‐Kusser et al., 1991). Furthermore, although in vitro biochemical analysis may be within normal limits, the in vivo production of ATP may be severely aVected even when only very low levels of mutated mt‐tRNA are present (Chinnery et al., 2000b). 2. A Variable Phenotype The perplexing observation that a single mtDNA mutation can produce several diVerent phenotypes at similar levels of mutant heteroplasmy has been made for many diVerent mutations. Perhaps the best example of such phenotypic diversity is that which occurs with the MTTL1 gene mutation m.3243A>G. This mutation is responsible for MELAS (Goto et al., 1990), CPEO (Fang et al., 1993), and also the maternally inherited diabetes and deafness (MIDD) phenotypes (van den Ouweland et al., 1992). This clinical variation cannot be explained solely on the basis of heteroplasmy, as significant overlap in the levels of mutated mtDNA has been observed in these patients. Likewise, the same phenotype can be produced by a number of diVerent mutations. For instance, MELAS can result from several mutations in the MTTL gene (Goto et al., 1990, 1991, 1992, 1994) as well as other mtDNA mutations (Kirby et al., 2004; Liolitsa et al., 2003), while CPEO can be caused by point mutations (Campos et al., 2001; Hammans et al., 1995; Spinazzola et al., 2004), single large‐scale mtDNA deletions (Moraes et al., 1989), and multiple deletions of mtDNA (Zeviani et al., 1990). The factors responsible for determining the phenotype in patients with a particular genotype remain elusive, but the observation of genotypic diversity with the same phenotype probably reflects a common pathway of impaired OXPHOS in a specific set of tissues. Environmental exposure to aminoglycoside antibiotics is known to be important in the early onset of deafness in individuals with the homoplasmic m.1555A>G MTRNR1 mutation (Prezant et al., 1993), and it is likely that similar interactions with environmental factors determine clinical outcome in other mtDNA mutations. The deafness phenotype associated with this mutation is also modified by the nuclear background (Guan et al., 2001) and in particular by the mtRNA‐modifying enzymes MTO1 and GTPBP3 (Bykhovskaya et al., 2004). Thus, nonsyndromic deafness associated with

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the m.1555A>G MTRNR1 mutation is a paradigm for mitochondrial disease, with the corresponding nuclear, environmental, and mitochondrial influences determining the disease phenotype. 3. Nuclear Control The extent and complexity of nuclear–mitochondrial interactions is perhaps only just beginning to be appreciated, but already a number of nuclear genes that govern the maintenance and copy number of mtDNA have been identified, including POLG1 (Van Goethem et al., 2001), PEO1 (Spelbrink et al., 2001), ANT1 (Kaukonen et al., 2000), TP (Nishino et al., 1999), DGOUK (Mandel et al., 2001), and TK2 (Saada et al., 2001). Mutations in these genes aVect either the nucleotide pool available for mtDNA replication (ANT1, TP, DGOUK, TK2) or the enzymatic process itself (POLG1 and PEO1) and result in either multiple deletions or depletion of mtDNA. Most patients with autosomal dominant CPEO and multiple deletions are thought to have mutations in one of three genes, ANT1, PEO1, or POLG1 (Agostino et al., 2003). However, recessive mutations in POLG1 have also been identified in patients with CPEO, as well as other conditions such as sensory atactic neuropathy, dysarthria and ophthalmoplegia (SANDO) (Van Goethem et al., 2003), juvenile spinocerebellar ataxia epilepsy syndrome (SCAE) (Van Goethem et al., 2004), and Alpers–Huttenlocher’s disease, a condition aVecting young children with involvement of liver and brain (Ferrari et al., 2005; Naviaux and Nguyen, 2004). These mutations in POLG1 only explain a fraction of the depletion syndromes and the search for genes involved in regulating mtDNA copy number continues apace with the identification of a mutation in MPV17 as the cause of infantile hepatocerebral depletion (Spinazzola et al., 2006). In addition to these mutations aVecting mtDNA copy number or maintenance, other nuclear gene mutations have been implicated in the etiology of mitochondrial disease. Missense mutations in SURF‐1, a protein necessary for the formation of the COX complex have been shown to cause Leigh syndrome (Tiranti et al., 1998; Zhu et al., 1998) and mutations in SCO2 (synthesis of cytochrome c oxidase), a COX assembly gene located on chromosome 22, have been associated with fatal infantile cardiomyopathy and severe COX deficiency in heart and skeletal muscle (Jaksch et al., 2000; Papadopoulou et al., 1999). Other neurological diseases such as forms of hereditary spastic paraplegia (Casari et al., 1998), Friedreich’s ataxia (Calabrese et al., 2005; Koutnikova et al., 1997; Lodi et al., 1999), and Wilson’s disease (Lutsenko and Cooper, 1998) are not directly related to mtDNA maintenance, copy number, or complex assembly, but have all been ascribed to mutations in nuclear gene‐ encoding products essential to mitochondria. Thus, mutations in a disparate collection of nuclear genes have been directly implicated in disrupting normal

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mitochondrial function in a variety of tissues, including brain, heart, and liver. What remains to be seen is whether mutations in mtDNA have any bearing on the expression of nuclear genes.

V. Pathology Mitochondrial disease can present with a wide variety of symptoms and signs attributable to either single‐ or multiorgan failure (Taylor and Turnbull, 2005). Tissue‐specific disease may progress to involve other systems or may remain confined to a single organ. A number of ‘‘classical’’ syndromes have been described, many of them associated with, but not exclusive to, a specific mtDNA or nDNA defect. Inevitably though, a substantial group of patients with mitochondrial disease will either not conform to the clinical criteria for a particular syndrome or present with only a few of the features. Seizures, migraine, stroke‐like episodes, neuropathy, or movement disorder are common neurological manifestations, but mitochondrial disease is often only considered when these features occur in conjunction with other symptoms such as deafness, diabetes, or visual impairment. Common, nonspecific symptoms such as fatigue and myalgia may be dismissed and the patient not referred for specialist advice, yet these symptoms can be the most debilitating features of mitochondrial disease (Griggs and Karpati, 1999). Equally, some patients with cardinal signs of respiratory chain dysfunction undergo extensive, but ultimately fruitless investigation. Childhood presentations of mitochondrial disease tend to be more severe than those with their onset in adult life and frequently involve many diVerent organ systems. Hepatic dysfunction and hemopoietic stem cell failure (Pearson syndrome) are uncommon features of mitochondrial disease, but are seen more often in children than adults. Renal disease also appears to be a more prominent clinical feature of pediatric mitochondrial disorders, evident in both mitochondrial depletion syndrome and complex III deficiencies (BCS1L mutations) (de Lonlay et al., 2001; De Meirleir et al., 2003). A. Organ‐Specific Mitochondrial Diseases 1. Mitochondrial Cardiomyopathy Cardiovascular involvement can include cardiomyopathy and conduction defects, both of which can be life threatening and in some patients may be the presenting feature of mitochondrial disease. Familial forms of cardiomyopathy have been described in which the most prominent feature is biventricular hypertrophy (thickening) (Guenthard et al., 1995). Patients with this pattern of cardiac involvement warrant further investigation for mitochondrial disease.

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2. Mitochondrial Myopathy Myopathy is a prominent clinical feature of several ‘‘classical mitochondrial syndromes,’’ but occasionally muscle involvement is the only feature of disease. Mitochondrial myopathy typically involves the proximal muscles of the hip and shoulder girdle and is usually associated with a gradual deterioration in power. The disease is sometimes progressive and eventual involvement of other muscle groups including the diaphragm may require nocturnal ventilatory support and can be life‐limiting. Rhabdomyolysis (McFarland et al., 2004c; Saunier et al., 1995), muscle pain, and stiVness (Deschauer et al., 1999) are not uncommon features of mitochondrial myopathy and such patients should be given advice regarding overexertion and maintaining adequate hydration. 3. Leber’s Hereditary Optic Neuropathy The first mitochondrial disease to be ascribed to a point mutation in mtDNA, LHON is an acute or subacute, bilateral, painless, central visual loss, and the commonest cause of blindness in young men (Kleiner and Sherman, 1996). The clinical condition was described as a familial neuro‐ophthalmologic disease by Theodor Leber in 1871, but it was not until over a century later that Wallace (Wallace et al., 1988) demonstrated the majority of LHON families harbor the same mtDNA mutation (m.11778G>A). A number of mtDNA mutations have subsequently been described in association with LHON, but three mutations (m.11778G>A, m.3460G>A, and m.14484T>C) are present in at least 95% of families (Man et al., 2003). These mutations in the complex I (NADH: ubiquinone oxidoreductase) encoding genes MTND4, MTND1, and MTND6, respectively, are considered to be primary LHON mutations on the basis of their frequency, penetrance, and clinical severity. The excess of aVected males suVering from this condition is thought to be due to an X‐linked visual‐loss susceptibility locus (Hudson et al., 2005). Clinical examination reveals peripapillary telangiectasia, microangiopathy, disk pseudooedema, and tortuous retinal vessels (Huoponen, 2001). Onset of disease is commonest after 20 and before 40 years, with initial unilateral involvement being typical. The other eye is commonly involved within a 2‐month period, but the subsequent decline in visual acuity may be rather slowly progressive over a period of several years. Both the disease penetrance and the clinical course appear to be determined by the mutation responsible, with measurements of final visual acuity ranging from 20/60 to no light perception at all. Similarly, the extent of visual recovery also varies in relation to the mutation, with only 4% of m.11778G>A patients showing recovery (Stone et al., 1992), while there are reports of up to 71% of m.14484T>C patients recovering (Harding et al., 1995). Extraocular clinical features such as cardiac conduction defects (WolV–Parkinson–White and Lown–Ganong–Levine) are evident occasionally and minor neurological problems are also not uncommon. Specific mutations (m.14459G>A) can be associated with a severe,

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predominantly neurological, phenotype involving early‐onset dystonia accompanied by bilateral basal ganglial degeneration (Jun et al., 1994). 4. Sensorineural Deafness Deafness commonly occurs in conjunction with diabetes (neurological features) in families harboring the m.3243A>G mutation (Hammans et al., 1995) or other MTT gene mutations, including m.14709T>C (MTTE) (Rigoli et al., 2001), m.7511T>C (MTTS1) (Sue et al., 1999), m.7445A>G (MTTS1/ CO1) (Reid et al., 1994), and m.7472Cins (MTTS1) (Tiranti et al., 1995) (Fig. 3). Deafness has also been observed in patients with sporadic single large‐scale mtDNA deletions (Ballinger et al., 1992; Zupanc et al., 1991). In families with maternally inherited nonsyndromic deafness due to the m.1555A>G MTRNR1 mutation, sensorineural deafness occurs as the only symptom of mitochondrial disease (Prezant et al., 1993). Phenotypic expression of this mutation is subject to both environmental and nuclear genetic control (see above) (Bykhovskaya et al., 2004; Guan et al., 2001). Deafness, in association with dystonia, is also one of the defining features of Mohr– Tranebjaerg syndrome, a condition caused by a mutation in the deafness dystonia protein 1/translocase of mitochondrial inner membrane 8a (DDP1/ TIMM8a) (Jin et al., 1996). Thus, deafness is a common feature of mitochondrial disease and observed in patients with heteroplasmic and homoplasmic mtDNA mutations as well as nDNA mutations. B. Classical Mitochondrial Syndromes 1. Mitochondrial Encephalopathy, Lactic Acidosis, and Stroke‐Like Episodes This clinical syndrome is characterized by parietooccipital stroke‐like episodes, which frequently do not conform to a single recognized vascular territory. Other features of MELAS include intermittent encephalopathic episodes associated with elevated plasma and CSF lactate, vomiting, migraine, dementia, and focal or generalized epilepsy. Muscle biopsy most commonly reveals a complex I deficiency but histochemistry may reveal COX‐negative fibers and COX‐positive ragged red fibers. The m.3243A>G mutation in the MTTL1 gene was the first and most frequently described mtDNA mutation associated with this clinical phenotype (Goto et al., 1990; Morten et al., 1995; Sternberg et al., 2001). However, mutations in other MTT genes (Bataillard et al., 2001; de Coo et al., 1998) as well as MTND1 (Kirby et al., 2004) and MTND5 (Liolitsa et al., 2003) genes have been described in association with this presentation and the m.3243A>G mutation also causes other distinct clinical phenotypes such as diabetes and deafness (Hammans et al., 1995). Extraordinarily, such variations in phenotype can

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occur between individuals in the same family harboring identical mutations (Damian et al., 1995; Martinuzzi et al., 1992). 2. Myoclonus Epilepsy and Ragged Red Fibers Myoclonus epilepsy and ragged red fibers (MERRF) was initially reported in association with an A>G mutation in the gene encoding MTTK at position 8344, and although other mutations in the same tRNA have now been reported, this remains the most common cause of this disease (ShoVner et al., 1990). This severe neuromuscular disorder causes progressive cortical myoclonus, focal and generalized epilepsy, muscle weakness and wasting, hypertrophic cardiomyopathy, dementia, deafness, and cerebellar ataxia (Silvestri et al., 1993). Indeed the m.8344A>G mutation has been identified as the cause of the original hereditary syndrome described by Ekbom of cerebellar ataxia, photomyoclonus, skeletal deformities, and lipomata (TraV et al., 1995). 3. Neuropathy, Ataxia, and Retinitis Pigmentosa and Maternally Inherited Leigh Syndrome First described as a variable combination of developmental delay, retinitis pigmentosa, dementia, seizures, ataxia, proximal neurogenic muscle weakness, and sensory neuropathy in four members of a single family (Holt et al., 1990), the phenotype has now been expanded to include cardiomyopathy and maternally inherited Leigh syndrome (MILS). The original family had a heteroplasmic T>G transversion at nucleotide pair m.8993 in subunit 6 of mitochondrial ATPase (ATP6), probably impairing ATP synthesis. A T>C mutation at m.8993 has also subsequently been described with a generally milder clinical phenotype, but higher frequency of ataxia. The MILS phenotype and impaired ATP synthesis is also observed with other mutations in the ATP6 gene, including m.9176T>C (Thyagarajan et al., 1995) and m.9176T>G (Carrozzo et al., 2001). Mutations in the mitochondrial ATPase subunits do not aVect cytochrome oxidase activity and therefore neuropathy, ataxia, and retinitis pigmentosa (NARP) patients will have no evidence of mitochondrial myopathy on routine histochemical analysis. 4. Mitochondrial Neurogastrointestinal Encephalopathy (MNGIE) This multisystem disorder is of variable severity and characterized by onset of CPEO, ptosis, gastrointestinal dysmotility (pseudo‐obstruction), diVuse leukoencephalopathy, peripheral neuropathy, and myopathy (Hirano et al., 1994). The inheritance is autosomal recessive and the causative mutations have been identified in the thymidine phosphorylase (TP) gene (Nishino et al., 1999; Spinazzola et al., 2002).

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5. Chronic Progressive External Ophthalmoplegia CPEO is one of the commonest presentations in adults, this condition is defined by a slowly progressive paresis of eye musculature, bilateral ptosis, and occasionally cardiac conduction defects. Many patients first notice ptosis in their third or fourth decade, but age of onset is variable. As the disease progresses, there is often associated proximal muscle weakness and fatigue though rarely to a debilitating extent. A sporadic, single deletion of mtDNA (4977 bp) is a common cause of CPEO, but other single deletions of variable length and some MTT mutations result in an identical phenotype. Recessive and dominant families with CPEO are well recognized and are associated with multiple species of deleted mtDNA (Bohlega et al., 1996; Zeviani et al., 1989). In these patients, the ophthalmoparesis and ptosis demonstrate a slowly progressive course with a wide range in the age of onset. Other features can include fatigue, optic atrophy, cataracts, ataxia, peripheral neuropathy, deafness, cardiomyopathy, and depression. A number of causative nuclear gene mutations including ANT‐1, PEO1, and POLG1 have now been described in these families (Agostino et al., 2003). 6. Kearns–Sayre Syndrome The onset of ophthalmoparesis and pigmentary retinopathy before the age of 20 years is characteristic of Kearns–Sayre syndrome (KSS). This sporadic condition is usually the result of either a large‐scale single deletion or complex rearrangements of mtDNA (Poulton et al., 1995; Zeviani et al., 1988). Other clinical features include cerebellar ataxia, proximal myopathy, complete heart block, cardiomyopathy, endocrinopathies, short stature, deafness, and an elevated CSF protein. As might be predicted from the early onset of this multisystem disorder, life expectancy is considerably reduced. C. Childhood‐Specific Mitochondrial Syndromes 1. Depletion Syndromes The term depletion refers to the extremely low levels of mtDNA found in muscle, liver, and brain tissues in these disorders. These are severe disorders presenting in early infancy or childhood with profound weakness, encephalopathy, seizures, and liver failure. In one form of ‘‘hepatocerebral’’ depletion known as Alpers–Huttenlocher’s disease or progressive neuronal degeneration of childhood (PNDC), explosive onset of seizures, developmental delay, and spasticity are followed some variable time later by catastrophic liver failure (Huttenlocher et al., 1976). In the ‘‘myopathic’’ form of depletion, profound weakness impairs mobility and eventually involves respiratory muscles leading to severe diYculty in breathing. A number of genes have been

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associated with specific variations of the depletion syndromes: myopathic (TK2) (Saada et al., 2001), hepatocerebral (DGOUK, POLG1, MPV17) (Mandel et al., 2001; Naviaux and Nguyen, 2004; Spinazzola et al., 2006), and encephalomyopathic (SUCLA2) (Elpeleg et al., 2005). Although this currently has little bearing on treatment options, it does provide useful genetic information for prenatal diagnosis in future pregnancies.

2. Leigh Syndrome Leigh syndrome is a progressive neurodegenerative condition of infancy and childhood. The characteristic symmetric necrotic lesions distributed along the brainstem, diencephalon, and basal ganglia were first described on postmortem tissue (Leigh, 1951), but are now more often identified on MRI or CT scan. The clinical presentation and course vary considerably, but signs of brainstem or basal ganglia dysfunction such as respiratory abnormalities, nystagmus, ataxia, dystonia, hypotonia, and optic atrophy are frequently observed. Developmental delay and more particularly regression are prominent, but nonspecific clinical features of this disorder and their diagnostic usefulness are improved when they occur in conjunction with raised CSF lactate. Stepwise developmental deterioration with some recovery of skills between episodes of regression is usual, although a relentless neurological decline is observed in some individuals. A severe failure of oxidative metabolism due to a variety of biochemical and molecular defects, including nuclear and mtDNA mutations, have been described in Leigh syndrome (Rahman et al., 1996). Consequently, inheritance can be X‐linked recessive, autosomal recessive, or maternal depending on the genetic defect responsible.

3. Pearson Syndrome Pearson syndrome is a rare disorder similar to KSS in that it results from large‐scale rearrangements of mtDNA. In this instance, however, clinical features of sideroblastic anemia with pancytopenia and exocrine pancreatic dysfunction predominate in early life and frequently result in death during infancy (Morikawa et al., 1993). Survival through childhood leads to an improvement in anemia but patients then develop the characteristic features of KSS. For Pearson syndrome, KSS, and CPEO, the clinical severity appears to correlate with the tissue localization of mutated mtDNA. In Pearson syndrome (and to a lesser extent KSS), mutated mtDNA can be demonstrated in a wide variety of tissues, whereas in CPEO the defective mtDNA is confined to muscle.

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VI. Conclusions The publication of the human mitochondrial genome sequence (Cambridge Reference Sequence) in 1981 was a defining moment in the study of human mitochondrial disease. This spectacular achievement heralded the dawn of an era of intense interest in mitochondrial genetics, but even after 25 years of industrious and often ingenious research, much remains unknown about this extraordinary genome. The mechanisms governing fundamental processes such as replication, transcription, and translation are only slowly being unraveled, but should eventually improve our understanding of the pathogenesis of mtDNA disease. The role of nuclear genes in the control of normal mitochondrial function and interaction between the two genomes is an exciting area of flourishing research, which again also holds the prospect of elucidating pathogenic mechanisms. The development of a mouse model will permit a clearer understanding of the role that mitochondria play in the aging process and increasingly sophisticated imaging techniques such as magnetic resonance spectroscopy should improve diagnosis and monitoring of patients. There are of course frustrations, and the slow progress toward eVective treatment of mitochondrial disease is a major source of disappointment to patients, scientists, and clinicians. Nevertheless, some progress has been made in this area by employing strategies, such as preimplantation genetic diagnosis, so that transmission of these awful diseases through generations of aVected families can be avoided. Much has been done, but there remains much to do.

Acknowledgments R. M. is supported by an MRC Clinician Scientist Fellowship. We thank the Wellcome Trust, the Medical Research Council (UK), and the Newcastle upon Tyne Hospitals NHS Trust for their continued support. Thanks also to Dr. L. Greaves for assistance with Fig. 3.

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Cybrid Models of mtDNA Disease and Transmission, from Cells to Mice Ian A. Trounce* and Carl A. Pinkert{ *Center for Neuroscience, University of Melbourne, Victoria 3010, Australia { Department of Pathology and Laboratory Medicine, Center for Aging and Developmental Biology, University of Rochester Medical Center Rochester, New York 14642

I. What Can Be Learned from Cybrids About mtDNA Disease? A. Introduction B. The First Mammalian Cell Cybrids: CAPR and Other Selectable Markers C. 0 Cells D. Cybrid Modeling of Human Disease‐Linked mtDNA Mutations E. Complementation of Heteroplasmic mtDNA Mutants F. mtDNA Contributions to Polygenic Disorders and Tissue‐Specificity G. Unknown OXPHOS Functions H. Mouse Cell mtDNA Mutants and Cybrids I. Xenomitochondrial Cybrids II. Animal Modeling of mtDNA Transmission and Human Disease: From Transgenesis to Transmitochondrial Cybrid Models A. Modeling Rationale B. Spontaneous and Induced Models C. Modifying the Mitochondrial Genome D. Use of Transfected 0 Cells as Intermediate Mitochondrial Carriers in the Production of Mouse Models E. Transmitochondrial Mice F. Embryonic Cytoplast and Karyoplast Fusion G. Xenomitochondrial Mice III. Summary References

Oxidative phosphorylation (OXPHOS) is the only mammalian biochemical pathway dependent on the coordinated assembly of protein subunits encoded by both nuclear and mitochondrial DNA (mtDNA) genes. Cytoplasmic hybrid cells, cybrids, are created by introducing mtDNAs of interest into cells depleted of endogenous mtDNAs, and have been a central tool in unraveling eVects of disease‐linked mtDNA mutations. In this way, the nuclear genetic complement is held constant so that observed eVects on OXPHOS can be linked to the introduced mtDNA. Cybrid studies have confirmed such linkage for many defined, disease‐associated mutations. In general, a threshold principle is evident where OXPHOS defects are Current Topics in Developmental Biology, Vol. 77 Copyright 2007, Elsevier Inc. All rights reserved.

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expressed when the proportion of mutant mtDNA in a heteroplasmic cell is high. Cybrids have also been used where mtDNA mutations are not known, but are suspected, and have produced some support for mtDNA involvement in more common neurodegenerative diseases. Mouse modeling of mtDNA transmission and disease has recently taken advantage of cybrid approaches. By using cultured cells as intermediate carriers of mtDNAs, ES cell cybrids have been produced in several laboratories by pretreatment of the cells with rhodamine 6G before cytoplast fusion. Both homoplasmic and heteroplasmic mice have been produced, allowing modeling of mtDNA transmission through the mouse germ line. We also briefly review and compare other transgenic approaches to modeling mtDNA dynamics, including mitochondrial injection into oocytes or zygotes, and embryonic karyoplast transfer. When breakthrough technology for mtDNA transformation arrives, cybrids will remain valuable for allowing exchange of engineered mtDNAs between cells. ß 2007, Elsevier Inc.

I. What Can Be Learned from Cybrids About mtDNA Disease? A. Introduction Cybrids, or ‘‘cytoplasmic hybrids,’’ are cultured cells manipulated to contain introduced mitochondrial DNA (mtDNA). The cybrids are produced by first treating mitochondrial donor cells with cytochalasin B to weaken the cytoskeleton, before subjecting the cells to a centrifugal force, either as attached cells or in suspension. The dense nuclei are extruded, leaving plasma membrane‐bound ‘‘cytoplasts’’ containing cell cytoplasm and organelles, including mitochondria. These cytoplasts are then fused with a nuclear donor cell line (originally using Sendai virus, but now using either polyethylene glycol or electrofusion), and transformant clones (cybrids) selected either with drug‐resistance markers or by selecting for respiratory competence with 0 cell fusions as discussed below. Cybrids allow discrimination between eVects of transferred organelle genes and those of a controlled nuclear background in the mtDNA recipient cell. Mitochondria, and in particular the oxidative phosphorylation (OXPHOS) complexes, depend on the coordinated assembly of both nuclear‐ and mtDNA‐encoded proteins so that mutations in either genome can result in mitochondrial pathology (Chinnery and Schon, 2003; Smeitink et al., 2001; Wallace, 1999). Cybrid experiments have contributed greatly to our understanding of mtDNA mutations in human disease. Studies of mitochondrial pathology and biogenesis are increasingly focusing on polygenic diseases where an mtDNA contribution is suspected. For such studies, cybrids should continue to make important contributions, including

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exploration of yet to be defined pathogenic mechanisms, such as a greater understanding of the role of normal and abnormal OXPHOS complexes in programed cell death. Cybrid technology can also address important issues in reproductive biology. The formation of a zygote is eVectively a new cybrid, having mtDNA only from the oocyte, and a new diploid nucleus. Assisted reproductive technologies including karyoplast transfer and ooplasm replacement, as discussed later in this chapter, inevitably result in cells with altered mitochondrial genetic complements. We review here the application of cybrid technology in the study of mtDNA mutations linked to human diseases, before moving to review cybrid approaches, in comparison with other technologies, used in modeling mtDNA transmission in experimental animals. B. The First Mammalian Cell Cybrids: CAPR and Other Selectable Markers The first mammalian cultured cell phenotype identified to segregate with mtDNA was in human (HeLa) cells, where mtDNA imparted resistance to the antibiotic chloramphenicol (Spolsky and Eisenstadt, 1972). This group coined the term cybrid when they were the first to demonstrate cosegregation of this phenotype with mtDNA markers, implying mtDNA mutation(s) as the cause of the phenotype (Bunn et al., 1974; Wallace et al., 1975). Several other mtDNA‐linked drug‐resistant phenotypes were identified in mammalian cells in the 1970s and 1980s, including resistance to the complex III inhibitors antimycin and myxothiazol (Howell and Gilbert, 1988) and later, to the complex I inhibitor rotenone (Bai and Attardi, 1998; Bai et al., 2000). The development of robust DNA‐sequencing methods in the late 1970s led to the identification of single‐base substitutions in the 16S rRNA gene of the mtDNA of independently derived yeast, mouse, and human chloramphenicol‐resistant (CAPR) cell lines (Blanc et al., 1981a,b; Dujon, 1980; Kearsey and Craig, 1981). The field of mammalian mitochondrial genetics was thus born. These pioneering studies were in turn followed by identification of the first cytochrome b mutants (Howell and Gilbert, 1988) and more recently ND5, ND6, and COI mutants. C. r0 Cells King and Attardi (1989) described the isolation of a human cell line without mtDNA (called 0 cells from the convention followed in yeast studies where such mtDNA‐less cells were first described; Trounce and Pinkert, 2005). Employing an approach first used with yeast (Slonimski et al., 1968) and then with avian cells (Desjardins et al., 1985), cells were incubated with low levels

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of the drug ethidium bromide, which intercalates DNA. Low levels of the drug selectively inhibits the ‐DNA polymerase responsible for mtDNA replication, and with ongoing cell division, the mtDNAs are ‘‘diluted’’ to the point where clones can be isolated without detectable organelle genomes. King and Attardi (1989) also discovered the absolute requirement for pyruvate gained by these cells and confirmed the previous observation from Desjardins et al. (1985) that mtDNA‐less cells also required added uridine for growth. This allowed a selection regime to be used after cytoplast–0 cell fusion so that unfused 0 cells could be eliminated and cybrids selected with the use of an appropriate nuclear drug‐resistant marker (King and Attardi, 1989; Fig. 1). Apart from their value in cybrid experiments, such 0 cells represent a unique research tool in themselves. They are a surprising reminder that OXPHOS is dispensable, at least for some diVerentiated mammalian cell types. In this instance, ATP production is 100% from glycolysis, so the cells acidify culture media very rapidly by producing large quantities of lactate. They retain functional mitochondria (except lacking OXPHOS), which show

Figure 1 Schematic illustrating cybrid production. The mtDNA donor cell line (top left) may harbor an mtDNA mutation of interest, or in the case of xenomitochondrial transfer, may be from another mouse species. These cells are enucleated using centrifugal force against a Percoll cushion and fused with 0 cells using polyethylene glycol. Cybrids are then selected by incorporating BrdU in the culture medium to eliminate any TKþ donor cells since the 0 cells are TK. To create transmitochondrial embryonic stem cell (ES) cybrids, the new TK cybrids are in turn enucleated and fused to rhodamine 6G (R6G)‐treated female (XX) ES cells (top right). When cybrid ES cell clones are selected, expanded, genotyped, and frozen, these can be used for blastocyst injection and chimeric mouse production employing standard transgenic techniques.

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a transmembrane potential (probably from the electrogenic exchange of ATP for ADP) and can import the hundreds of other proteins needed for non‐OXPHOS functions. It is tempting to speculate that quiescent cells, including stem cells, may ‘‘switch oV ’’ OXPHOS to minimize reactive oxygen species (ROS) production. This remains an understudied area. In one study of hematopoietic stem cells, the mitochondrial density and activity of progenitor cells increased in inverse proportion to the CD34 commitment marker, suggesting that mitochondrial biogenesis is suppressed in progenitor cells, and engaged when such cells further diVerentiate (Piccoli et al., 2005). With the advent of 0 cells, researchers were able to introduce any mitochondria, including those from patients with putative mtDNA disease, without the need for selectable drug‐resistant markers. The pioneering cybrid work using the selectable markers was limited in the sense that endogenous mtDNAs were also present; that is, the cybrids were heteroplasmic. The 0 cell approach allowed creation of homoplasmic or heteroplasmic cells, depending on the mtDNA donor cell(s) used. As such, the emerging field of mitochondrial medicine was poised to dissect and characterize specific mitochondrial mutations.

D. Cybrid Modeling of Human Disease‐Linked mtDNA Mutations From 1988 to 1990, five diVerent mtDNA mutations associated with the major mitochondrial disease ‘‘syndromes’’ were discovered. Hence, single‐ base substitution mutations were identified which segregated with Leber’s hereditary optic neuropathy (LHON; Wallace et al., 1988), mitochondrial encephalomyopathy with lactic acidosis and stroke‐like episodes (MELAS; Goto et al., 1990), myoclonic epilepsy with ragged red fibers (MERRF; ShoVner et al., 1990), large heteroplasmic mtDNA deletions in chronic progressive external ophthalmoplegia (CPEO; Holt et al., 1988), and a single‐base substitution in the ATP6 gene in neurogenic weakness with ataxia and retinitis pigmentosa (NARP; Holt et al., 1990). These comprise the more commonly encountered mtDNA mutations, but the list has grown to several hundred (see Chapter 5 by McFarland et al., this volume). The 0 cell cybrid technique was applied to these and other mtDNA mutations, establishing OXPHOS phenotypes that were transferred with the diVerent mutations. The MERRF mutation, a single‐base substitution in the mtDNA tRNALys gene, was shown to segregate with a severe mitochondrial protein synthesis defect (Chomyn et al., 1991). Interestingly, this paper also showed that a heteroplasmic mtDNA donor sample from a patient could be used to produce cybrids with either normal or defective protein synthesis, segregating with wild‐type or mutant mtDNA cybrids, suggesting the mtDNA heteroplasmy was largely intercellular.

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This observation was repeated in several later studies with diVerent mtDNA mutants (Turner et al., 2005). However, other data illustrate largely intracellular heteroplasmy; this remains a controversial topic and is briefly addressed in the Section I.E on mtDNA complementation. Another controversy surrounds the pathogenic mechanism associated with the MELAS tRNALeu(UUR) mutation. In vitro studies suggested the mutation altered the ratio of rRNA to mRNA transcripts by altering the transcriptional termination sequence located in the tRNA gene (Hess et al., 1991). Cybrid studies did not support this, showing no alteration in the transcript ratios, but again a severe mitochondrial protein synthesis defect when the level of mutation was high (Chomyn et al., 1992; Hayashi et al., 1991; King et al., 1992). Another report found that in cybrids made by fusing diVerent 0 cells (i.e., diVerent nuclear backgrounds) and enucleated patient cells harboring the heteroplasmic MELAS mutation, that with passage of cells in culture diVerent subclones analyzed showed segregation to either the wild‐type or mutant mtDNAs, depending on the nucleus (Dunbar et al., 1995). Nonstochastic segregation of disease‐linked mtDNAs has also been described so that the mutant genome progressively increases as a proportion of total mtDNA with growth of cultured cybrids (Hayashi et al., 1993; Turner et al., 2005). The most commonly encountered missense mutations associated with mitochondrial disease are the LHON (at nucleotide 3460 in the ND1 gene, at 11778 in the ND4 gene, and at 14484 in the ND6 gene) and Leigh’s disease/NARP mutants most commonly found at nucleotide 8993 in the ATP6 gene. The nucleotide 8993 ATP6 mutation was shown in cybrid studies to segregate with a moderate respiration defect and a lowered ADP:O ratio, combining to produce a severe ATP synthesis defect (Trounce et al., 1994). The LHON mutants have been more diYcult to define or assign biochemical phenotypes in cybrid studies. Jun et al. (1996) found complex I activity to be moderately reduced when the 14459 mutation associated with LHON þ dystonia was transferred into lymphoblastoid cybrids. Others have found that nuclear background is again important, with diVerent cybrid nuclear backgrounds aVecting the expression of the complex I defect associated with the nucleotide 3460 mutation (Cock et al., 1998). Comparison of the three ‘‘primary’’ LHON mutants in the same cybrid nucleus also showed variable defects of complex I, with the 3460 mutant exhibiting a greater defect in respiration and enzyme assays than the nucleotides 11778 and 14484 mutants (Brown et al., 2000). E. Complementation of Heteroplasmic mtDNA Mutants An issue of relevance to mtDNA in development and transmission is whether mixing of mtDNA types occurs in organelles of heteroplasmic cells. Cybrid studies have addressed, but not resolved, this issue. Thus, Oliver

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and Wallace (1982) presented evidence that heteroplasmic mtDNAs could ‘‘complement’’ each other, implying fusion of organelles and mixing of mtDNAs and mitoribosomes from translation‐defective and competent mtDNAs. This conclusion was based on the observation that in heteroplasmic cybrids, a marker polypeptide linked to a defective 16S rRNA gene (carrying a 16S point mutation giving rise to CAPR) continued to be translated in the presence of CAP (Oliver and Wallace, 1982). Hayashi’s group have been a proponent of such intramitochondrial complementation, providing evidence in both a mouse model (Nakada et al., 2001) and in cultured cells (Hayashi et al., 1994; Nakada et al., 2001). In both cases, heteroplasmic mtDNA deletions were used as model systems. When present in cells in very high levels, the deleted mtDNA prevented any mitochondrial translation and cells were cytochrome oxidase (COX) negative. Newly created cybrids, or cells in mice carrying heteroplasmic deletions, always showed a uniform staining for COX, suggesting that single organelles had extensively fused (Hayashi et al., 1994; Nakada et al., 2001). Yet, another study does not support such extensive mixing. Using cybrids carrying two diVerent homoplasmic mtDNA point mutations, Enriquez et al. (2000) found a level of trans‐complementation of only around 1% of cybrids produced. Mitochondria exist as an interconnected reticulum in most cell types, but can exhibit dynamic morphology (see Chen and Chan, 2004 for a review). Fusion experiments using diVerently labeled mitochondria of 0 cells and respiratory competent cells showed that mitochondrial mixing is dependent on a membrane potential in the organelle, but not respiratory competence (Legros et al., 2002). This basic issue in mitochondrial genetics may have consequences for the expression of mtDNA mutants in tissues, and deserves further attention to better address whether diVerent cell types will alter the rate of mitochondrial fusion and mixing of mtDNAs.

F. mtDNA Contributions to Polygenic Disorders and Tissue‐Specificity In the classic mtDNA diseases, clear matrilineal inheritance, although often with variable penetrance, strongly supports a disease link. Claims have been made over the past decade that mtDNA variants may also be contributors to the commonest age‐related neurodegenerative disorders, especially Parkinson’s disease (PD) and Alzheimer’s disease (AD). Some support has been gained for the idea that at least a portion of PD patients may have mild, mtDNA‐encoded complex I defects that can be transferred in cybrids (Gu et al., 1998; Swerdlow et al., 1996). Adding to the controversy is the lack, to date, of identification of specific disease‐associated mtDNA variants in PD and AD.

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Such identification is extremely diYcult when faced with the large degree of polymorphism encountered in mtDNA from diVerent humans. As the number of fully sequenced individual mtDNAs grows, this big question may have begun to be answered. Wallace’s group has published the largest of such analyses thus far, which has provided some interesting clues (Ruiz‐Pesini et al., 2004). Populations from higher latitudes were found to exhibit higher rates of substitutions of more highly conserved amino acids, which may correlate with increased susceptibility to energetic diseases (Ruiz‐Pesini et al., 2004). Why diVerent tissues are adversely aVected by diVerent mtDNA mutations remains another enduring mystery of mitochondrial disease. One route to address this question has been to isolate 0 cells from diVerent cell types, including neuronal cell lines (Fukuyama et al., 2002; Sheehan et al., 1997; Swerdlow et al., 1996; Turner et al., 2005). Using 0 SH‐SY5Y human neuroblastoma cells, Sheehan et al. (1997) found that cybrids with mtDNAs from AD patients showed a 52% decreased COX activity, increased ROS, and decreased ability to buVer changes in cytosolic calcium. Onyango et al. (2005) used the same 0 SH‐SY5Y cells to produce cybrids with sporadic PD mtDNAs, and found evidence for increased activation of p38 and JNK oxidative stress pathways, and decreased viability. Neither of these studies compared cybrids made using another nonneuronal parent cell line, so no conclusions can be drawn regarding the utility of the neuron‐like lines in establishing such phenotypes. As mentioned above, these and the other cybrid studies investigating putative mtDNA‐linked eVects in PD and AD have not identified mtDNA candidate mutations. This does not mean that mtDNA‐linkage is unlikely. It is possible that combinations of mtDNA amino acid polymorphisms may act together to produce aberrant cellular phenotypes or act in concert with nuclear gene polymorphisms in subunits that interact (with mtDNA‐encoded subunits in OXPHOS complexes I, III, IV, or V). Further cybrid work in these and other sporadic neurodegenerative diseases is warranted. Large patient groups need to be studied to identify likely subgroupings or endophenotypes that may be mtDNA linked. Analysis of mtDNA sequences, when informed by such phenotype groupings, may then reveal patterns of amino acid substitutions with significant linkage to the disease. Conversely, putative nuclear gene alleles that may cause OXPHOS defects, when combined with an mtDNA type, could also be identified by cybrids. Primary patient fibroblasts, for example, can be treated with rhodamine 6G (R6G) so that endogenous mtDNA can be replaced with a mitochondrial genome of choice. This approach was shown to be feasible (Williams et al., 1999), but reports using such a system to tackle candidate diseases have not yet emerged.

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G. Unknown OXPHOS Functions The discovery of an alternate role of the respiratory chain electron carrier, cytochrome c, as a messenger in programed cell death signaling (Liu et al., 1996) was a reminder that ‘‘well‐established’’ biochemical pathways can continue to throw up major surprises. We look forward to many more unexpected findings, especially with regard to the role of the OXPHOS complexes in mediating aspects of the cell death program. It is likely that cybrid studies will continue to represent an important tool for defining pathogenic eVects of organellar genes as well as in ongoing modeling studies in mice. Even when transformation techniques finally catch up with mtDNA, cybrids will continue to have a role in allowing ease of transfer of mitochondria with altered mtDNAs between cells. Cybrid studies have greatly advanced our understanding of mtDNA disease genetics, but have also left major gaps to fill. Clues have been gained that nuclear gene eVects can be important modulators of OXPHOS phenotypes. As new information on the role of the mitochondrion in programed cell death pathways accumulates, new assays for cellular phenotypes will emerge beyond the simplistic assays of respiratory chain complex activity mostly used to date. While the OXPHOS assays will continue to be a vital part of such characterization, wider cellular eVects of mutations will need to be discovered before we can begin to understand the still baZing clinical features of human mitochondrial diseases. An example of such new cellular phenotypes, in addition to those of Sheehan et al. (1997) and Onyango et al. (2005) discussed above, may include some reports using LHON cybrids where the mtDNA mutant cells are shown to be more sensitized to enter programed cell death via mitochondrial pathways (Carrozzo et al., 2004; Danielson et al., 2002; Ghelli et al., 2003).

H. Mouse Cell mtDNA Mutants and Cybrids Only a dozen or so diVerent mouse cell mtDNA mutants have been described (Table I). These have contributed important insights into the function of the aVected OXPHOS complexes. As discussed later in this chapter, such mutants also represent a resource for production of transmitochondrial mice via a cybrid route. Where investigated, most of the mutants described have moderate to severe OXPHOS phenotypes, which may preclude their use in mouse‐modeling studies. The limited mouse models produced to date include the use of the CAPR mouse cell lines, which have moderate protein synthesis defects reflected in partial deficiencies of complexes I, III, and IV

Table I

Reported mtDNA Mutants in Cultured Mouse Cells

Locus

Nucleotide Change

16S

T2432C A2381T

ATP6/16S

T8563A C2380T G14830A G14251T G14563C C14578T A15020T G15263A 13879–13884 Cins

Val!Glu – Glu231!Asp Gly38!Val Gly142!Ala Thr147!Met Leu294!Phe E373K Frameshift

C12081A C6063A T6589C

Frameshift Leu246!Lle Val421!Ala

Cyt b

ND6 ND5 COI

a

NR, not reported.

Amino Acid Change

OXPHOS Phenotype

References

– –

Moderate complexes I, III, and IV defects

Blanc et al., 1981b; Howell and Lee, 1989; Howell and Nalty, 1987; Kearse and Craig, 1981 Slott et al., 1983

NRa NR

Howell and Gilbert, 1988; Howell et al., 1987

Severe Severe complex I defect

Acı´n‐Pe´rez et al., 2004 Acı´n‐Pe´rez et al., 2003; Bai and Attardi, 1998 Bai et al., 2000 Acı´n‐Pe´rez et al., 2003

Severe complex I defect 50% complex IV defect

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(Howell and Nalty, 1987; Sligh et al., 2000). Since the homoplasmic mice produced by Sligh et al. (2000) showed a neonatal lethal phenotype, our expectation is that only mutants with mild OXPHOS phenotypes will be viable and, therefore, most informative. The report of Sligh et al. (2000) is also significant for showing that mouse embryonic stem cell cybrids can maintain their pluripotency, even when treated with R6G prior to cytoplast fusion. The rhodamine treatment destroys the mitochondria, and it had been shown to reduce or prevent transmission of the mtDNAs of treated cells when fused to donor cytoplasts (Trounce and Wallace, 1996; Ziegler and Davidson, 1981). Unlike 0 cells, R6G‐treated cells cannot survive even in permissive media containing uridine and pyruvate, unless rescued by cytoplast fusion. This provides a means to eliminate endogenous mtDNA in recipient cells or to manipulate the degree of heteroplasmy. Remarkably, R6G treatment of mouse ES cells allowed cytoplast fusion with subsequent recovery of undiVerentiated, homoplasmic cybrid ES cells, in turn, opening the door for ES cell cybrid approaches to modeling mtDNA defects in mice (Kasahara et al., 2006; McKenzie et al., 2004; Sligh et al., 2000). There remains a complete lack of mouse mtDNA mutants with proven phenotypes identified in living mouse strains. In rats, mtDNA‐linkage has been described for the diabetes‐prone BHE/Cdb rat, which was found to harbor a pair of point mutations in the ATP6 gene compared with Sprague‐ Dawley rats (see Berdanier, 2001 and later in this chapter). A relative lack of rat 0 cell lines has meant this mutation has yet to be investigated in a cybrid system. Until breakthrough technologies allow mtDNA transformation (Khan and Bennett, 2004; Yoon and Koob, 2005), a greater eVort involving isolation of new mouse cell mtDNA mutants for use in modeling could be richly rewarded.

I. Xenomitochondrial Cybrids One approach to produce mtDNA‐based defects without mutants is by creating xenomitochondrial cybrids, defined as having mtDNAs from a diVerent species to the recipient cell. Interspecific transfer of mitochondria and mtDNAs has long been an important tool in plant genetics (see Gillham, 1994; Hagemann, 2000; Herrman et al., 2003 for reviews). Early cybrid and hybrid studies using interspecific crosses of mammalian cells determined that the mtDNAs segregated with the nuclear genes of the same species (Hayashi et al., 1980; Wallace et al., 1976). Studies with mice first explored the idea of introducing mtDNAs into interspecific nuclear backgrounds by backcrossing (Gyllensten et al., 1991; Nagao et al., 1998). Gyllensten et al. used this approach to examine putative

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paternal mtDNA transmission, finding extremely low levels using sensitive PCR detection. While limited to species where interbreeding results in fertile oVspring, these studies are interesting for demonstrating that such transfers can produce viable oVspring. Some success has been reported for manipulating the germ line mtDNAs in Drosophila, and subsequent breeding of heteroplasmic flies showed complete segregation of the endogenous mtDNA (Niki et al., 1989). Cybrids represent a cleaner experimental system, as the cross is achieved in a single generation, with the advantage that there is no doubt as to the nuclear gene complement. The use of 0 cells presented an opportunity to revisit interspecies mitochondrial transfer, since eVects of endogenous mtDNA in the nuclear donor cell could be removed. Kenyon and Moraes (1997) fused enucleated primate cells with human 0 cells, selected for respiratory competent cells and succeeded in producing xenocybrids. They showed that with increased evolutionary divergence a barrier was reached whereby the foreign mtDNA could no longer be maintained so that cybrids were not viable. Thus, chimpanzee and gorilla mtDNAs could be replicated and transcribed in human cells, but orangutan and more divergent primate mtDNAs could not. Interestingly, the OXPHOS of the cybrids showed defective complex I activity with the other respiratory chain complexes being relatively preserved (Kenyon and Moraes, 1997). In a similar way, mouse 0 cells were independently shown to be capable of maintaining other murid mtDNAs, including the closely related Mus spretus and the more distant mouse cousin the Norway rat (Rattus norvegicus) (Dey et al., 2000; McKenzie and Trounce, 2000; Yamaoka et al., 2000). While the M. spretus xenocybrid exhibited normal OXPHOS, the Rattus xenocybrid had severe combined respiratory chain defects. Murid rodents have more extant species than any other mammalian family, so that a large number of potential xenocybrids can be made, although the limited number of described cell lines necessitates creation or procurement of new primary cell lines. Using this approach, McKenzie et al. (2003, 2004) showed that species of intermediate divergence compared with Rattus could create graded respiratory chain defects in xenocybrids. Mild complex I and IV defects emerged in intermediate divergence cybrids, but a severe complex III defect was evident in the most divergent xenocybrids (McKenzie et al., 2003). Like the primate xenocybrids, mouse cells also display limits to the evolutionary divergence of mtDNA donor species. Hamster mtDNAs, divergent about 16 million years ago (mybp; million years before present), could not be maintained in cybrids with the mouse nucleus (McKenzie and Trounce, 2000). The 0 cell xenocybrids are selected by virtue of detectable recovery of respiratory chain function. These experiments cannot clearly distinguish whether the evolutionary barrier to xenocybrid creation results from total loss of respiratory chain function due to mismatching of

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nuclear and mtDNA subunits or from loss of mtDNA replication. This could be further addressed by transfection of key nuclear mtDNA replication factors (such as mitochondrial transcription factor A or Tfam) from the mtDNA donor species. From the mouse xenocybrid studies, it is nevertheless clear that OXPHOS dysfunction will precede such replicative barriers in interspecific crosses.

II. Animal Modeling of mtDNA Transmission and Human Disease: From Transgenesis to Transmitochondrial Cybrid Models A. Modeling Rationale The field of mitochondrial medicine is still in its infancy. The first published accounts of diseases caused by mutations of mtDNA were reported in 1988 and there are now a spectrum of genomic mitochondrial mutations that are known to be the underlying causes of various degenerative disorders (Clayton, 1991, 1992; Copeland, 2002; Lemasters and Nieminen, 2001; Pinkert and Trounce, 2002; Wallace, 2001; Wallace et al., 1988; see also MITOMAP at http://www.mitomap.org). Additionally, mutations that aVect mitochondrial function dramatically impact on tissues with high cellular energy requirements, including the central nervous system, cardiac and skeletal muscle, and various endocrine organs (Wallace, 2001). In mammals, mitochondrial genetics are inherited in matrilineal fashion. Furthermore, it was postulated that a developmental bottleneck exists in female germ cell development, whereby a small number of mitochondria give rise to approximately 105 mitochondria that populate the mature ovum (Cummins, 1998; Marchington et al., 1997; Piko and Matsumoto, 1976; see Chapter 3 by May‐Panloup et al., this volume). Through the late 1990s, in vivo animal models of mtDNA mutation and human disease were virtually nonexistent as techniques for modifying large numbers of mitochondria in early embryos were far more complex than comparable modifications of nuclear‐encoded genes. As we learn more about mitochondrial dysfunction and developmental consequences of mitochondrial mutation, creation of animal models will be of critical importance not only in the study of mitochondrial dynamics and function but in dissection of mtDNA‐based disease pathogenesis. Currently, the term ‘‘mitochondrial disease’’ is used most prominently to describe diseases caused by mutations in mtDNA. However, defining mitochondrial disease is probably more accurate in the broader context of recognizing that mitochondrial diseases are those with a mitochondrial component, genetically or environmentally influenced, that produce cell and organ dysfunction or failure under acute and chronic stress.

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B. Spontaneous and Induced Models Identified, spontaneous mtDNA mutants are extremely rare in mammalian models. The BHE rat was identified early on as a unique nonobese diabetic model that appeared inconsistent with expected disease phenotype (Berdanier, 1982, 1991; Lakshmanan et al., 1977). The salt‐sensitive BHE/ Cdb rat was first characterized with two homoplasmic and three heteroplasmic mutations in the mitochondrial ATP6 gene, causing impairment of OXPHOS and glucose intolerance; the specific mutations have since been studied extensively (Everts and Berdanier, 2002; Mathews et al., 1999). The BHE/Cdb rat and related models paved the way for current hypotheses related to nutritional regulation of mitochondrial function in mammals. Other early toxin‐induced models, identified spontaneous nuclear mitochondrial gene mutants, and nuclear mitochondrial transgenic models have been reviewed elsewhere (Pinkert and Trounce, 2005; Wallace, 2001). In the aggregate, data have accumulated over the years that readily reflect mitochondrial dysfunction in animal models. Many of these models have been molecularly characterized; yet, demonstration of methodology to tailor mitochondrial genetics and mitochondrial–nuclear dynamics was the next logical progression in eVorts to target mitochondrial mutations.

C. Modifying the Mitochondrial Genome With initial characterization of nuclear gene‐encoded models that impacted on mitochondrial function, our search for a greater understanding of mitochondrial biology would eventually lead us to develop methodology for mitochondria and mitochondrial gene transfer. As an initial endeavor, eYcient methods for introduction of either—(1) foreign or modified mtDNA or (2) intact foreign mitochondrial genomes—into somatic or germ cells would be needed. In order to employ genetically engineered mitochondria or mitochondrial genes for in vivo gene transfer or for gene therapy, eYcient methods to introduce foreign or altered mitochondrial genomes into somatic cells or germ cells must be identified. A primary focus for our work has included production of engineered mutations of mtDNA genomes and subsequent transduction or transfection into isolated mitochondria in vitro prior to transfer into mouse embryos. Methods for the delivery of complete foreign mtDNA genomes into intact mitochondria are presently nonexistent. The transfection of DNA into mitochondria presents some very formidable challenges. Although in vitro fusion of inner mitochondrial membrane vesicles was reported, there have been no reports of fusion of outer membrane vesicles or of whole mitochondria (Hackenbrock and Chazotte, 1986). Also, the basic mechanisms for

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mitochondrial membrane lipid addition have yet to be exposed (i.e., how newly synthesized lipid is targeted to mitochondrial membranes). Using mitochondrial‐targeting peptides covalently attached to DNA molecules, Seibel et al. (1995) demonstrated internalization of small DNA fragments into intact mitochondria via a protein import pathway. However, delivery of mitochondrial‐specific DNA sequences and subsequent integration into the host mitochondrial genome, while yet to be demonstrated, may not be possible if mechanisms for DNA recombination within mitochondria are lacking (Cannon et al., 2004; Pinkert and Trounce, 2005; Trounce and Pinkert, 2005).

D. Use of Transfected r0 Cells as Intermediate Mitochondrial Carriers in the Production of Mouse Models We and others have taken advantage of techniques in ES cell/knockout technologies and cybrid development to generate models of mitochondrial mutations. In developing in vitro models of human mitochondrial disorders, both immortalized human and mouse 0 cell lines were developed (Bai and Attardi, 1998; King and Attardi, 1989; Trounce et al., 2000). Thus, immortalized 0 cells would obviously provide an excellent vehicle for propagation of our mutated mtDNA containing mitochondria for developing mutant mitochondrial mouse models (Irwin et al., 2001; Pinkert and Trounce, 2002). With transfer of transfected mitochondria (either by microinjection or cytoplast fusion), selection pressure could be exerted by removal of uridine and pyruvate from the culture medium. An important advantage of this intermediate culture system is that heteroplasmic constructs can be produced at will by simply mixing cells of interest in the enucleation step, then fusing the mixed cytoplasts, in diVerent ratios if desired, and genotyping many cybrid clones to select heteroplasmic clones. These clones can then be used in fusion with the R6G‐treated ES cells as we have described (McKenzie et al., 2004; Trounce et al., 2004).

E. Transmitochondrial Mice 1. mtDNA Injection Versus ES Cell‐Derived Models Through the 1980s, paternal contribution to mtDNA was discounted in most mammalian species (Birky, 1995; Cummins, 1998; Giles et al., 1980; Gyllensten et al., 1991; Ingraham and Pinkert, 2003; Nishimura et al., 2006). While estimates vary widely, normal embryos prior to fertilization harbor over 100,000 mitochondria in comparison to the 75–150 contributed by

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sperm (Hauswirth and Laipis, 1985; Michaels et al., 1982). In the late 1990s, sperm mitochondria appeared preprogramed for destruction as illustrated by work where labeled‐mtDNA from spermatocytes microinjected into mouse zygotes could not be traced into subsequent embryos or young (Cummins, 1998; Cummins et al., 1999). Further studies reported active digestion of sperm mtDNA, and mitochondrial degradation initiated by ubiquitination (Nishimura et al., 2006; Sutovsky et al., 2000). Thus, generation of heteroplasmic mice by natural breeding is ineYcient and has limited utility in establishing mtDNA disease models (Irwin et al., 1999). In contrast, segregation of mtDNA was studied in heteroplasmic mice created by cytoplast fusion (Jenuth et al., 1996, 1997; see Chapter 4 by Shoubridge and Wai, this volume) and by karyoplast transplantation (Pinkert et al., 2005; Takeda et al., 2005). Microinjected murine mitochondria labeled with mitochondria‐specific TM MitoTracker (Invitrogen, Carlsbad, CA, USA) fluorophores allowed evaluation of subsequent viability and functionality of exogenous mitochondria populations in vivo. Double‐labeling experiments were performed to establish the fate and function of microinjection mitochondria (Ingraham and Pinkert, 2003). Embryos and isolated mitochondria were incubated with various TM TM combinations of MitoTracker Red (CM‐H2XRos) or MitoTracker TM Green. MitoTracker Red was used as a fluorescent probe that only adsorbs TM to viable mitochondria, while MitoTracker Green attaches to the mitochondrial membrane independent of viability or membrane potential (Molecular Probes, 2002). Endogenous mitochondria within embryos were labeled with TM MitoTracker Green and introduced/microinjected mitochondria were laTM beled with MitoTracker Red. Over time, introduced and labeled mitochondria became increasingly diVuse. Varying localization and fluorescence TM intensities were observed in MitoTracker Red labeled zygotes 2 hours postinjection (M. H. Irwin, L. W. Johnson, and C. A. Pinkert, unpublished data). Migration of fluorescence signaling throughout the cytoplasm was observed in zygotes surviving injection (Ingraham and Pinkert, 2003). In contrast, the zygotes that did not survive injection completely lost red fluorescence within 24 hours. Since the control green fluorescence remained, it was likely that the red fluorescence was lost concomitantly with a loss of mitochondrial viability. Mitochondrial research is progressing directly toward development of mitochondrial mutants with mtDNA or genomic modifications (Khan and Bennett, 2004; Pinkert and Trounce, 2002; Yoon and Koob, 2005), which illustrates the significance of maximizing the developmental fate of exogenously introduced mitochondria. Direct mitochondrial microinjection into the cytoplasm of embryos in interspecies transfer experiments demonstrated persistence of the introduced mitochondria in 100% of zygotes through the blastocyst stage of development (Pinkert et al., 1997). However, the

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persistence of introduced mitochondria diminished over time with only 10–15% of founder mice harboring detectable levels of foreign mitochondria at term (Irwin et al., 1999; Pinkert and Trounce, 2002). Exogenous mitochondria were clearly being eliminated and/or they were not replicating in our earlier studies. In addition to direct cytoplasmic injection of mitochondria, ES cell transfer technology, coupled with cytoplast fusion methods, can be used to introduce mutated mtDNA into 0 cell lines (Pinkert and Trounce, 2002; Trounce and Wallace, 1996). DiVerentiation between endogenous and exogenous populations of mitochondria can be established in addition to the viability of microinjected mitochondria. Cytoplasmic diVusion of the exogenous microinjected mitochondria within the zygote explores mitochondrial migration in vivo in relation to maintenance or elimination of heteroplasmy. However, the use of putatively specific dyes must be evaluated carefully in such studies. In other published studies, mitochondria were subjected to a specific membrane potential dye, tetramethylrhodamine methyl ester (TMRM), which may provide further insight into fate of microinjected mitochondria (Zhang et al., 2001). Our studies extend the finding that mitochondria relocate during oocyte maturation or fertilization in several species (Barnett et al., 1996; Bavister and Squirrell, 2000). The significance of mitochondrial migration through early cleavage stages remains unclear, although the process appears critical to normal embryonic development. It was further suggested that changes in mitochondrial organization might serve as predictors of preimplantation embryonic developmental competence (Bavister and Squirrell, 2000), and in relation to mitochondrial dysfunction and segregation in vivo.

F. Embryonic Cytoplast and Karyoplast Fusion Techniques to create heteroplasmic mice then progressed via cytoplast fusion (see Chapter 4 by Shoubridge and Wai, this volume; Jenuth et al., 1996, 1997; Laipis, 1996) and by embryonic karyoplast transplantation (Meirelles and Smith, 1997). In these experiments, rapid segregation of mtDNA was possible within maternal lineages; however, specific manipulations were not readily controllable or quantifiable in first‐generation animals. Generally, the recipient embryo appeared to dominate in terms of mitochondrial survival. Interestingly, early reports on development of cloned animals by nuclear transfer resulted in conflicting consequences when retrospective studies on mitochondrial transmission were reported (Evans et al., 1999; Hiendleder et al., 1999; Steinborn et al., 2000; Takeda et al., 1999). Similarly, using a

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human in vitro fertilization protocol, two heteroplasmic children may have been inadvertently created (Barritt et al., 2001; Cohen et al., 1997). As such, research independent of targeted mitochondrial genomic modifications may also help unlock mechanisms underlying the dynamics related to persistence of foreign mitochondria and maintenance of heteroplasmy in various cloning protocols. Indeed, specific culture‐related conditions that may influence the prevalence or development of heteroplasmy in these techniques provide one example to explain the range of results (heteroplasmy to homoplasmy) observed in nuclear transfer experimentation (Takeda et al., 2002). For in vivo modeling, a number of laboratories have reported on methodologies used to create transmitochondrial mouse models (Inoue et al., 2000; Irwin et al., 1999; Kasahara et al., 2006; Levy et al., 1999; Marchington et al., 1999; Pinkert et al., 1997; Sligh et al., 2000). These reports illustrate various technical and biological aspects of in vivo modeling of mitochondrial dynamics and human disease. Curiously, in reports where developmental consequences of the genetic manipulations were observed, either aberrant or unexpected phenotype has been regularly identified (Inoue et al., 2000; Sligh et al., 2000). The use of cybrid modeling in the context of whole animal biology will be most useful in addressing the biological bases of these findings. In 1999, Levy et al. and Marchington et al. independently demonstrated that it was possible to fuse cytoplasts prepared from a CAPR cell line with mouse ES cells and then introduce the mutant cells into mouse blastocysts. The CAPR mutation was used to target a respiratory deficiency in resultant animals, while not initially identified, anticipated phenotype and germ line competence in founders were later demonstrated (Sligh et al., 2000). An important innovation in this work was in the use of R6G to limit the transmission of endogenous ES cell mtDNAs, something that had been previously demonstrated only for somatic cells (Trounce and Wallace, 1996; Ziegler and Davidson, 1981). The advantage of this approach was illustrated by the failure of Marchington et al. (1999) to obtain homoplasmic ES cell cybrids or mice using chloramphenicol selection alone. In the absence of methods to produce targeted mtDNA mutants, the current limitation to the production of mtDNA mutant mice is the lack of suitable mouse cell mutants. A handful of mutants were described over the past 20 years, all produced by in vitro mutagenesis followed by selection of drug‐resistant mtDNA mutants (Table I). While some of these may prove interesting if successfully introduced into mice, the findings of Sligh et al. (2000) sound a warning. The relatively mild impairment of respiration consequent to the CAPR mutation as defined by polarographic measurements (Levy et al., 1999), surprisingly, resulted in the in utero or perinatal

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death of animals with high levels of the mutation. This suggested that mutations with mild respiratory chain impairment as measured in vitro would likely prove the most successful in modeling human disorders. Data on the respiratory chain function are lacking for many of the mouse cell mutants described, but when available suggest that these mutants are severe (Bai and Attardi, 1998; Howell, 1990).

G. Xenomitochondrial Mice From development of the first transmitochondrial animal models, xenocybrids were later created from several mouse species to evaluate a graded respiratory impairment in vivo in mouse models (McKenzie et al., 2004).

Figure 2 From cybrids cells to mice. (A) Phase contrast image of mouse LMTK 0 cells; (B) control cybrids produced by fusion of the 0 cells with Mus musculus cytoplasts. The morphology returns to the parental type, with stubby, fast‐growing cells in contrast to the elongate, granular, and slow‐growing 0 cells; (C) cybrids carrying mtDNAs from the Asian mouse M. terricolor (aka M. dunni), showing similar morphology to control cybrids; (D) our first germ line xenomouse carrying homoplasmic M. terricolor mtDNA, with her first litter. The mouse was produced by first transferring mtDNAs from the LMTK cybrids, shown above, into female ES cells that had been pretreated with R6G. The transmitochondrial ES cells were then used for blastocyst injection and chimera production (McKenzie et al., 2004; Fig. 1). (A–C) 200 magnification.

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Taking advantage of the enormous evolutionary diversity of Muridae species, one objective was to create a series of cybrids with increasing levels of respiratory chain impairment resulting from the presence of mismatched nuclear‐mtDNA subunits. Modeling of the defects was first done in a 0 cybrid system and then lineages where phenotype was observed (here, a mild respiratory impairment) could be readily transferred into mouse female ES cells for production of chimeric mice (Pinkert, 2002). Again, the use of cybrid technology formed the foundation for our continuing eVorts at generating in vivo respiratory deficiencies and a greater understanding of mitochondrial mutation and ensuing pathologies (Fig. 2).

III. Summary Through the early 1990s, various early attempts to create transmitochondrial mouse strains (and a number of other animal species) by introduction of foreign mitochondria into germ cells were largely unsuccessful. Novel in vivo methodologies, especially those associated with cybrid techniques, have since proven most encouraging and beneficial. Cybrids provide mitochondrial researchers with technologies comparable to those now taken for granted in producing nuclear genetic modifications (e.g., knockout and knock‐in nuclear gene modeling). With the use of cybrid cell lines, site‐ directed mutagenesis modeling is envisioned that is adaptable to now routine in vitro screening and selection techniques. With targeted mitochondrial genomic modifications just around the corner, such technologies will be instrumental in characterizing mechanisms underlying mitochondrial dynamics and disease.

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Section III The Use of Assisted Reproductive Technologies to Regulate mtDNA Disease

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7

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The Use of Micromanipulation Methods as a Tool to Prevention of Transmission of Mutated Mitochondrial DNA Helena Fulka* and Josef Fulka, Jr.{ *Institute of Experimental Medicine, Videnska 1083 CS‐142 20 Prague 10, Czech Republic { Center for Cell Therapy and Tissue Repair VUZV Laboratories, Pratelstvi 815 CS‐104 01 Prague 10 Czech Republic I. II. III. IV.

Biological Material: Oocytes and Zygotes Technical Equipment Germinal Vesicle Transfer Short‐ and Long‐Term Storage of Nuclear Material and Asynchronous GV to Cytoplast Transfer V. Chromosome Group Transfer VI. Transfer of Pronuclei VII. Conclusions Acknowledgments References

The introduction of diVerent micromanipulation techniques into reproductive and developmental biology has helped us not only to answer many essential biological questions but it is now evident that these techniques also have wide practical applications. In human‐assisted reproduction, the most commonly used approach is the injection of a donor sperm into the oocyte cytoplasm—intracytoplasmic sperm injection. It is, however, speculated that with these techniques it would be also possible to improve the oocyte developmental potential especially in those cases when the quality of the cytoplasm is rather poor and thus its function is compromised. Another important application would be the elimination of mutated mitochondrial DNA (mtDNA) by transferring the nuclear material from an abnormal oocyte into a healthy donor oocyte cytoplast. Some of these techniques were already successfully tested in experimental animals, but it is evident that before their introduction into human medicine many questions must be answered, and we must be sure that these approaches are absolutely or almost absolutely safe. In our contribution, we will be specifically oriented to the nuclear (nuclear material) replacement approaches that could be potentially used to prevent the transmission of mutated mtDNA from mother to oVspring. Because these techniques are very delicate, some Current Topics in Developmental Biology, Vol. 77 Copyright 2007, Elsevier Inc. All rights reserved.

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0070-2153/07 $35.00 DOI: 10.1016/S0070-2153(06)77007-7

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training with oocytes from other species other than human is strongly recommended. ß 2007, Elsevier Inc.

I. Biological Material: Oocytes and Zygotes The ovary contains essentially three basic populations of oocytes. The population of growing oocytes includes very small cells that are incompetent to initiate the final processes of maturation. This means that in these cells, germinal vesicle breakdown (GVBD, GV‐oocyte nucleus) does not occur either in vivo or in vitro when these oocytes are isolated from follicles and placed into a culture medium. The intermediate population contains larger oocytes that are able to initiate the process of maturation. In these oocytes, GVBD can be detected after their isolation from follicles and culture under appropriate conditions. The process of maturation is, however, not completed and is arrested at the metaphase I (MI) stage. The last population includes those oocytes that completed their growth and attained the full size (100–140 mm, depending on species origin). These oocytes respond to gonadotropin stimuli and initiate the process of maturation (Fulka et al., 1998). This means that they undergo GVBD and chromosome condensation. The chromosomes are then organized in the first meiotic metaphase that is followed by anaphase I to telophase I transition (AI–TI). Thereafter, the process of maturation is arrested in metaphase II (MII) with the first polar body (1PB) extruded. At this stage, the oocyte awaits the fertilizing spermatozoa (Chian, 2004; Trounson et al., 2001). Soon after fertilization the oocyte chromosomes complete the process of meiosis and after anaphase II to telophase II transition the oocyte as well as sperm chromosomes decondense and form the female pronucleus (FPN) and male pronucleus (MPN). Both pronuclei (PNs) increase their size and approximately after 6–7 hours postfertilization DNA replication begins. The pronuclei then closely appose each other and after the completion of DNA replication their chromosomes condense and are arranged in the first mitotic metaphase. Typically, this can be detected approximately 18–24 hours postfertilization. The oocyte (zygote) has two basic components—the nucleus (germinal vesicle, pronuclei) and the cytoplasm. It is absolutely essential for both of these to function perfectly, not only for the final phases of oocyte maturation but especially later on after the oocyte is fertilized by sperm and the embryo develops (Eichenlaub‐Ritter and Peschke, 2002). Oocytes or zygotes can be manipulated at any stage mentioned above. An excellent example is Kaguya—the parthenogenetic mouse—where germinal vesicle DNA from very small oocytes was combined with DNA from oocytes that had already completed their growth (Kono et al., 2004).

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In this chapter, we will however concentrate on fully grown oocytes and zygotes that still contain fully developed pronuclei (PNs). Here, their manipulation is simpler than at other stages. Moreover these micromanipulation schemes can be theoretically used in assisted human reproduction. We will also discuss enucleation and nuclear transfer in immature fully grown oocytes (germinal vesicle stage), maturing oocytes (metaphase I stage), mature oocytes (metaphase II stage) as well as in zygotes that contain fully developed pronuclei. Finally, we will discuss some potential advantages and disadvantages for each developmental stage mentioned above as well as some potential risks. The basic procedural steps are essentially the same for all the above stages. The most prominent diVerence is only in the diameter of the enucleation pipette opening that is considerably wider for the manipulation of germinal vesicle oocytes and pronuclear stage embryos.

II. Technical Equipment Mammalian oocytes or zygotes are rather small and can be easily damaged when improperly manipulated. Although certain manipulations can be done simply just by hand (Karnikova et al., 1998), it is recommended to use high quality equipment, commercially produced culture and manipulation media, and high quality chemicals. The most essential is an inverted microscope (Olympus, Leica, Nikon) with diVerential contrast (Nomarski or HoVman) optics. These microscopes must be equipped with precise micromanipulators (Narishige, Leica) and injectors (Eppendorf, Narishige) that allow us to perform all the necessary steps, including the oocyte or zygote enucleation and transfer of nuclear material under the zona pellucida (ZP) of the recipient cytoplast. A very useful instrument that was introduced into reproductive and developmental biology relatively recently is the Piezo injector either from PrimeTech (www. primetech‐jp.com) or Burleigh (www.burleigh.com). These injectors allow very gentle manipulations, especially safe penetration through the zona pellucida. According to our opinion and in the context with those methods that will be further discussed, this is the most diYcult step as the oocyte or zygote can be easily damaged. It is also beneficial if the microscope is equipped with fluorescence. Thus, success at enucleation can be checked after the nuclear material is stained with some vital DNA stains (Hoechst 33258, bisBenzimide). This may be important when whole chromosome groups are transferred between oocytes. Moreover, mitochondria can be easily labeled with some vital stains that can be purchased from Molecular Probes (www.probes.com). In this case, the fluorescence microscope is absolutely necessary.

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A high standard of quality is absolutely essential for enucleation and transfer pipettes. The holding pipettes can be purchased from diVerent companies. According to our experience, it is much better if enucleation or injection pipettes are made by the person who directly manipulates the oocytes or zygotes. These pipettes can be prepared relatively easily with the help of some instruments that are also commercially available (Sutter, Narishige). The nuclei or chromosome groups are almost exclusively isolated from donor oocytes or zygotes in the form of ‘‘karyoplasts.’’ This means that they are enclosed within the cell membrane and thus these karyoplasts also contain a certain volume of cytoplasm. The cytoplasm without the nuclear material is called ‘‘the cytoplast.’’ The karyoplasts are not directly injected into the recipient cell cytoplasm (cytoplast) because they are too large and after their direct injection one can be almost absolutely certain that the reconstructed cell will be destroyed. Thus, the method of induced cell‐to‐cell fusion is commonly used for the transfer of the nucleus (chromosomes) from the karyoplast into the cytoplast. This is typically performed by electrofusion with the help of some commercially available equipment (BTX, Eppendorf). Potentially, several alternative fusion methods can be used: polyethylene glycol (PEG)‐induced fusion, inactivated Sendai virus, or the method developed by Tesarik et al. (2000) where the intracytoplasmic sperm injection (ICSI) pipette penetrates through both the karyoplasts and cytoplasts and this leads to a subsequent fusion between them. The culture of reconstructed cells is also very important. All of the above steps are described in detail in several manuals and books and for this reason they are described here only briefly (Hogan et al., 1994; Monk, 1987; Wassarman and DePamphilis, 1993). As mentioned, the transfer of nuclear material isolated from a defective cytoplasm into a healthy cytoplasm can be carried out at diVerent stages of oocyte maturation or in zygotes with distinct pronuclei. Next, we will discuss the methodological approaches and biological aspects that are typical for given stages. We will, however, concentrate only on those cells where we can expect the potential application of a given approach in human medicine.

III. Germinal Vesicle Transfer Enucleation and nucleus transfer can be performed relatively easily in those cells that contain a nucleus—this means in immature oocytes with germinal vesicles (or in zygotes with pronuclei). The immature, fully grown oocytes can be isolated from nonstimulated or stimulated ovaries from mid or large antral follicles. These oocytes are enclosed with tightly packed follicular

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cells that must be removed from the oocyte (zona pellucida) surface before manipulation. Without this step the manipulation is impossible as the oocyte cytoplasm and germinal vesicle are invisible. The germinal vesicle can be easily observed, for example, in mouse, rat, and human oocytes. On the other hand, the location of germinal vesicles is more complicated in bovine and especially porcine oocytes because their cytoplasm is very dark. In cattle, germinal vesicle can be detected as a lighter area below the vitelline membrane. In the pig, the centrifugation of oocytes is usually necessary. The oocytes without cumulus cells are first incubated in the manipulation medium with cytochalasin B (5 mg/ml) and thereafter centrifuged for 10 min at about 9000 rpm. After centrifugation, the lipid particles are located in one hemisphere while the other hemisphere becomes lighter. The germinal vesicle is almost always located in the light hemisphere (Fulka et al., 2003). In general, before enucleation, the oocytes are first incubated in the manipulation medium droplets (approximate volume 20 ml) with cytochalasin B (5 mg/ml) under paraYn oil for at least 10 min. The position of the manipulated oocyte (zygote) is stabilized by a holding pipette. Then, the zona pellucida must be penetrated with the enucleation pipette. This step depends on the origin of oocytes and the equipment the given laboratory has. The zona pellucida opening can be made easily with a laser beam or by applying several piezo pulses from a piezo injector. Here, care must be taken to avoid damage to the vitelline membrane. This means that the perivitelline space must be suYciently large and the intensity of pulses must be checked before. If not, we recommend increasing the osmolarity of the manipulation media, for example, with sucrose supplementation. If the laboratory is not equipped with the above instruments, alternative approaches exist. The opening can be made by acid Tyrode solution that is released from the narrow injection pipette tip in the close vicinity of the zona pellucida. However, in this case, for subsequent enucleation, another wider pipette must be used. Alternatively, the enucleation pipette can be beveled and sharpened. This enables the penetration of zona pellucida without damage to the vitelline membrane (cytoplast). Beveled and sharp pipettes are commonly used for the enucleation of ungulate (cattle, sheep, goat, pig) oocytes. The enucleation of mouse oocytes is more complicated even if a beveled pipette is used because their zonae pellucidae are rather soft and resistant to penetration, so the oocyte can be damaged very easily. Thus, a slit must be first made in the zona pellucida with a sharp thin glass needle and subsequently the enucleation pipette is introduced under the zona through this slit. For simplicity, the use of a laser or piezo injector is strongly recommended. After opening the zona pellucida, the enucleation pipette is introduced under it and positioned in close vicinity to the germinal vesicle. Thereafter, a very slow suction is applied and the germinal vesicle is slowly aspirated into

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a pipette. As soon as the whole germinal vesicle is inside, the pipette is slowly withdrawn from the oocyte cytoplasm. As mentioned above, the isolated germinal vesicle is enclosed with a certain volume of the cytoplasm and the cytoplasmic (vitelline) membrane (karyoplast). The diameter of the enucleation pipette opening is extremely important. If too wide, it is very diYcult to introduce it under the zona pellucida and if piezo pulses are applied to make a hole in the zona, the oocyte can be easily damaged and destroyed. On the other hand, if too small, the nucleolus can be released from the germinal vesicle and translocated into the cytoplasm—so called ‘‘enucleolation’’ (Fulka et al., 2003). In general, the recommended diameter is approximately 20 mm. Next, the germinal vesicle karyoplast must be transferred into the perivitelline space of another oocyte that was previously enucleated in the same way, as described above. The tip of the enucleation pipette penetrates through the opening in the zona that was already made before and the germinal vesicle karyoplast is then slowly released into the perivitelline space of a donor cytoplast. The germinal vesicle karyoplast is rather fragile and can be easily damaged and destroyed. Thus, it is highly recommended to transfer it directly, after its removal from the original oocyte, to just under the zona pellucida of a recipient oocyte. It is better not to release it into manipulation medium with subsequent aspiration into a pipette and transfer later on. If this is necessary the aspiration into the pipette must be very gentle and slow or the diameter of a transfer (enucleation) pipette must be larger. The germinal vesicle transfer method is demonstrated in Fig. 1a–i. The cytoplast/karyoplast constructs are then removed from the manipulation medium, washed several times in the medium without cytochalasin, and cultured in a standard culture medium for at least 30 min (5% CO2, 37–38 C) before their fusion is induced. In those species where the process of GVBD occurs very rapidly, the manipulation and culture media must be supplemented with some drugs inhibiting the onset of maturation. For example, in the mouse where GVBD occurs within 30–60 min, the media are usually supplemented with dibutyryl cyclic AMP (100–150 mg/ml). This is not necessary when human (GVBD, 10–15 hours), bovine (6–7 hours), or porcine immature oocytes (16–18 hours) are used. After this short preculture interval, the cytoplast/karyoplast constructs are fused. As mentioned above, several approaches can be used. Here, we will however mention only the electrofusion method because of its simplicity and safety. The cytoplast/karyoplast constructs are removed from the culture medium and washed briefly in fusion medium (isotonic glucose solution or 0.30 M mannitol containing 100 mM CaCl2 and 100 mM MgCl2) before being transferred into the same medium in a fusion chamber consisting of two electrodes (gap between them is usually 100–200 mm).

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Figure 1 (a–i) Enucleation and germinal vesicle transfer in immature mouse oocytes. (a) The immature mouse oocyte just before enucleation. The oocyte is fixed in a holding pipette (H), the enucleation pipette is on the right (E). ZP, zona pellucida; GV, germinal vesicle; N, nucleolus. Interference contrast, 350. (b) Parallel picture where the oocyte germinal vesicle DNA is stained with Hoechst (arrow). Fluorescence, 350. (c) Parallel picture showing the mitochondria labeled with MitoTracker Green FM (M). Fluorescence, 350. (d) The GV vesicle is aspirated into the enucleation pipette and forms the karyoplast (K). The enucleated oocyte is called ‘‘the cytoplast (C).’’ Interference contrast, 350. (e) Parallel staining with Hoechst demonstrates that all of the nuclear DNA is located in the karyoplast (K). Fluorescence, 350. (f) Parallel picture showing the oocyte mitochondria that were labeled before enucleation. Note the mitochondria‐rich cytoplast (C), while in the karyoplast (K), mitochondria are less abundant. Fluorescence, 350. (g) The isolated germinal vesicle (karyoplast, K) transferred under the zona pellucida (ZP) of another previously enucleated oocyte (C). The karyoplast is subsequently fused to the cytoplast (C). Interference contrast, 350. (h) Parallel picture showing the absence of nuclear DNA in the cytoplast (C), on the other hand, GV DNA is positively labeled (arrow). Fluorescence, 350. (i) Mitochondria in the original karyoplast donor oocyte were labeled with MitoTracker FM Green, thus the isolated karyoplast (K) exhibits the positive signal. The cytoplast (C) from the recipient oocyte was not labeled and is without fluorescence. Fluorescence, 350.

Prior to pulsing, the cytoplast/karyoplast constructs must be aligned in a chamber so that membranes to be fused are parallel to the electrodes. This can be done either manually or by applying an AC field (usually 3 V for 10 s). The alignment is followed by one or two fusion DC pulses (1–2 kV/cm for

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30–50 ms each). The treated constructs are then washed two to three times in culture medium and incubated in it (5% CO2, 37–38 C) for 30–60 min before being checked for signs of fusion. The above fusion parameters must be modified for each laboratory, species, and type of biological material used. It is our experience that human, porcine, and bovine oocytes can be fused very easily with a high eYciency (fusion rate more than 75%). On the other hand, the fusion of mouse material is sometimes more complicated and needs to be tested before. For example, GV karyoplast to cytoplast fusion can be diYcult; on the other hand, karyoplasts with pronuclei in this species can be easily fused to corresponding cytoplasts.

IV. Short‐ and Long‐Term Storage of Nuclear Material and Asynchronous GV to Cytoplast Transfer Whilst with laboratory and domestic animals the supply of biological material is usually quite suYcient, it can be the case in humans that we have the patient’s oocytes but we do not have recipient’s cytoplasts and vice versa. Consequently, we have oocytes that are at diVerent stages of maturation. Karnikova et al. (1998) found in the mouse that isolated germinal vesicle karyoplasts remained intact when cultured in vitro whilst in nonmanipulated oocytes the process of GVBD occurred within the same culture interval. These observations prompted us to study this phenomenon in more details. We have found that if GV karyoplasts are very small, they remain intact even after a prolonged culture (more than 2 days). These karyoplasts can then be used for fusion to freshly isolated cytoplasts with a high rate of maturation in reconstructed cells (more than 80% of oocytes in MII; Fulka, Jr., unpublished data). When cytoplasts were stored under the same condition, they could not be used for transfer because the reconstructed cell lysed or died in almost all cases. In large germinal vesicle karyoplasts, the process of GVBD occurs and chromosomes condense, but they never reach the metaphase II stage. Small karyoplasts can be even frozen by vitrification. In our laboratory, we transfer them into evacuated porcine or mouse zonae pellucidae (Fig. 2) in which they are frozen and stored for further use. After thawing, they can be fused to freshly isolated cytoplasts, again with a suYciently high rate of maturation. Almost all karyoplasts survive the vitrification procedure (Kren et al., 2005). We suppose that the inability of small karyoplasts to undergo GVBD and the possibility of their low‐temperature storage would be extremely important in humans where the immediate supply of recipient oocytes may sometimes be diYcult.

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Figure 2 The isolated germinal vesicles’ karyoplasts can be frozen by vitrification and stored in liquid nitrogen. The picture shows a single mouse GV karyoplast (K) in an evacuated mouse zona pellucida (ZP). Up to 10 karyoplasts can be stored in a single evacuated zona. Interference contrast, 200.

It can also be the case that the patient oocytes are in the appropriate stage (immature, germinal vesicle) and the donor oocytes are more advanced in the process of maturation—for example, already undergoing GVBD. Can we use these more advanced oocytes as a source of cytoplasts for germinal vesicle transfer? Here, the results obtained are very variable and rather inconsistent. We have transferred GV karyoplasts into cytoplasts that were obtained from oocytes just undergoing GVBD (1) and also into cytoplasts from metaphase I (2) or metaphase II (3) oocytes. Consistent results were only obtained in combination (1) where the reconstructed oocytes regularly completed the process of maturation and were arrested in metaphase II. The other two combinations yielded rather inconsistent results where in some cases oocytes proceeded up to metaphase II whilst in others they remained arrested at the metaphase I stage (Fulka et al., 2002). Basically, the same results were reported by Cheng et al. (2003a) who showed that only the combination: GV karyoplasts GVBD (MI) cytoplasts yielded satisfactory results. Clearly, some additional experiments are necessary before a definitive conclusion can be made. We, however, suppose that these asynchronous transfer experiments are extremely important from the point of view of practical use. It is well known that the period of maturation from the intact germinal vesicle stage up to shortly after GVBD includes the most important processes that are essential for further embryonic development. We may thus suppose that the quality of cytoplasts that are obtained from those oocytes having completed GVBD is significantly higher than in those obtained from completely immature oocytes (see below). The feasibility of the GV transfer method has been documented in several papers where mouse oocytes almost exclusively served as a model (Tsai et al., 2000). All these papers demonstrate that a very high proportion of mouse oocytes can be successfully enucleated (80%). The fusion eYciency is also satisfactory (70%) and the same proportion of reconstructed oocytes matured up to metaphase II (Fulka et al., 2002, 2005; Palermo et al., 2002;

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Takeuchi et al., 2001). The reconstructed and matured oocytes can be fertilized by ICSI and approximately half of them form both the MPN and FPN. The further development of these embryos depends on many factors and diVers between laboratories. The most critical factors are the origin of oocytes: that is, oocytes from stimulated females pregnant mare serum gonadotropin (PMSG) are better than those from nonstimulated females, also culture conditions and media have a significant influence (Liu et al., 1999, 2003a; Takeuchi et al., 2001, 2004). Nevertheless, if the method of GV transfer is perfected, live oVspring can be obtained as demonstrated by Takeuchi et al. (2004, 2005a). In other species, the GV transfer method has been used in the cattle where Kuwayama (2003) reported the birth of a calf by transferring a germinal vesicle from the oocyte of an animal that was assessed as infertile due to its advanced age into a donor enucleated oocyte that was obtained from a fertile female. The potential practical use is, however, clearly evident when human oocytes would be used. This approach may, in theory, serve for the elimination of mutated mitochondrial DNA (mtDNA), the elimination of aneuploidies that result from the insuYcient cytoplasmic function (low number of mitochondria, unknown factors), and eventually for the treatment of certain oocyte maturation defects that are diYcult to explain (Mrazek and Fulka, 2003; Palermo et al., 2002; Tsai et al., 2000). Logically, the development of an eYcient GV transfer scheme in humans is very complicated not only because of the limited accessibility to convenient biological material (oocytes) but also from the ethical point of view. We must bear in mind that there are evident physiological diVerences and diVerent culture requirements between model animals (mouse) and human oocytes. For example, in the mouse, GVBD occurs within 1 hour and the process of maturation is completed approximately after 10–12 hours of culture. There is no evident diVerence in the speed of GVBD between fully grown oocytes obtained either from stimulated or nonstimulated animals. On the other hand in humans, the source of oocytes significantly influences the speed of GVBD and the whole process of maturation. The time course of GVBD is diVerent between oocytes from nonstimulated and stimulated cycles. The oocytes from a nonstimulated cycle still contain intact germinal vesicles after 12 hours of culture, whilst the oocytes from stimulated cycles already contain condensed chromosomes at this time point. Similarly, the speed of maturation diVers between these two groups. The oocytes from stimulated cycles reach metaphase II after 30 hours in culture. On the other hand, the oocytes from nonstimulated cycles need an additional 12 hours to reach this stage (for review see Chian, 2004; Trounson et al., 2001). From the technical point of view, enucleation as well as GV transfer in immature human‐staged oocytes (as they are rather resistant to damage) can be performed relatively easily with high eYciency (Palermo et al., 2002;

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Takeuchi et al., 2001). Moreover, the maturation of reconstructed oocytes seems to also be satisfactory (60% MII) and so is their fertilization by ICSI (50%). The further early development of fertilized embryos seems to be also good (Takeuchi et al., 2001). However, many questions remain. If these techniques should be introduced in humans, one must be absolutely sure of their safety. The results obtained especially in farm animals, where the length of the process of oocyte maturation is considerably longer than in the mouse, clearly document that we must be cautious when we propose these eventual applications in humans. As mentioned above, the process of GVBD is of crucial importance and influences oocyte quality. The oocytes collected from follicles soon after GVBD have basically the same developmental potential as those that completed the process of maturation inside the follicle (Motlik and Fulka, 1981). Gioia et al. (2005) clearly demonstrated in the pig that the degree of paternal genome demethylation in MPNs diVers between oocytes that were matured completely in vitro, in vivo, or if they were collected from follicles after GVBD and then cultured in vitro. In the first case, in 40% of zygotes, demethylation of the paternal chromatin was clearly evident; on the other hand in ‘‘in vivo’’ matured oocytes, this demethylation was detectable in 80%. In those oocytes where GVBD occurred in vivo, 73% of zygotes showed the correct demethylation pattern. It remains to be determined if the same applies to human oocytes. The evaluation of methylation–demethylation processes in human zygotes showed rather inconsistent results (Beaujean et al., 2004; Fulka et al., 2004) and further analysis is clearly necessary. Moreover, the presence of follicular cells seems to be essential for the appropriate completion of oocyte maturation (Staigmiller and Moor, 1984). For germinal vesicle transfer, cumulus cells must be removed from the oocyte surface. For example, in the mouse, the maturation of denuded oocytes is associated with aberrant expression of Oct‐4 (Chang et al., 2005). It is not known if the coculture of reconstructed oocytes with follicular cells would be suYcient for the elimination of some negative eVects. That certain negative eVects can be expected has been demonstrated by Takeuchi et al. (2005a) who produced a few mouse oVspring from GV transfer oocytes. There were evident diVerences in the weights of fetuses and placentas between control and experimental oVspring born. Also, significant differences were detected in expression of Igf2 and H19 in corresponding placentas. It was also clearly documented that not all oocytes isolated from stimulated follicles are in the same developmental stage. Some of them are more advanced while others are slightly behind (for review see Fulka et al., 1998; Moor et al., 1998). It is beyond the scope of this chapter to discuss these diVerences as they are comprehensively discussed in the above‐mentioned reviews. We will give just three examples, H19 gene expression showed evident diVerence in about one‐quarter of human oocytes collected from

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stimulated cycles (Borghol et al., 2006). These results indicate that some oocytes evidently did not reach a developmentally optimal situation. It is not known if this slightly aberrant expression will have some negative long‐term eVects. We have analyzed the transcriptional activity in mouse oocytes after they were labeled against phosphorylated Pol II. The oocytes were isolated from PMSG‐stimulated follicles and very carefully selected. Even in this case, the results imply that in about 25% of these oocytes the Pol II transcription shutdown has not taken place yet. Again, this indicates that some oocytes even from stimulated cycles are developmentally less advanced and perhaps not at the optimal stage (Helena Fulka, in preparation). It remains to be determined if a prolonged culture could overcome this asynchrony. Also, some recent articles analyzing the process of maturation in vitro in human oocytes indicate certain diVerences when compared to in vivo matured oocytes. These experiments concentrated on the spindle morphology, which was evidently diVerent between in vitro and in vivo maturing oocytes. These diVerences consequently resulted in a higher frequency of chromosomal abnormalities in ‘‘in vitro’’ matured oocytes (Li et al., 2006). In conclusion, we suppose that the above overview clearly documents the feasibility of germinal vesicle transfer in mammalian oocytes from a technical point of view. On the other hand, from a safety point of view, many questions remain unanswered and evidently some additional experiments are clearly necessary.

V. Chromosome Group Transfer Concomitantly with GVBD, chromosomes condense and form a single group. Interestingly, even very shortly after GVBD, chromosomes are relatively compact and can be easily removed from the oocyte as a whole group without loosing one or more of them. Later on, the metaphase I plate of chromosomes is formed with a clearly visible spindle. This stage is followed by a short anaphase I to telophase I transition. Thereafter, the chromosomes are arranged as the metaphase II plate. Chromosomes can be removed from the oocyte at any stage mentioned above (Fig. 3a–i). It is our experience that the probability of either a single or multiple chromosomes remaining in the oocyte whilst the rest are removed is minimal (Fulka, Jr., unpublished data). When compared to germinal vesicle stage oocytes, the manipulation of maturing or mature oocytes has also some advantages as well as disadvantages. Among the advantages, the maturing and mature oocytes are more resistant to damage, so their manipulation is easier. Moreover, the metaphase plate covers a smaller area in the oocyte cytoplasm and the diameter of the enucleation pipette can thus be smaller (15 mm). As mentioned above,

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Figure 3 (a–i) Enucleation and metaphase group transfer in maturing oocytes. (a) The maturing mouse oocyte just before the enucleation. The oocyte is fixed in a holding pipette (H), the enucleation pipette is on the right. The metaphase group of chromosomes is located in the oocyte center (arrow). ZP, zona pellucida; C, cytoplasm. Interference contrast, 350. (b) Parallel picture where the oocyte chromosomes were stained with Hoechst (arrow). Fluorescence, 350. (c) Parallel picture showing the mitochondria labeled with MitoTracker Green FM. Fluorescence, 350. (d) The metaphase chromosome group (arrow) is then slowly aspirated into the enucleation pipette. Interference contrast, 350. (e) Parallel picture where chromosomes are stained with Hoechst (arrow). Fluorescence, 350. (f) The same oocyte with mitochondria labeled by MitoTracker Green FM. Fluorescence, 350. (g) The metaphase group as the karyoplast (K) is then completely detached from the oocyte and subsequently used for transfer into another, previously enucleated oocyte. Interference contrast, 350. (h) Parallel picture demonstrating the absence of nuclear DNA in the cytoplast (C), the positive staining of chromosomes is evident only in the karyoplast (arrow). Fluorescence, 350. (i) Labeling of mitochondria showed that they are very abundant in the cytoplast (C), while in the karyoplast (arrow) less intensive signal can be detected. Fluorescence, 350.

the period of GVBD is of crucial importance from the oocyte quality point of view. Thus, when we manipulate oocytes with condensed chromosomes, we can suppose that they had already overcome this essential period and their quality will be higher when compared with manipulated immature oocytes. The manipulation of maturing and mature oocytes has also some disadvantages. The most crucial one is the detection of the chromosome groups.

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The chromosomes can easily be seen in the living oocyte in the mouse if the quality of the microscope optics is suYcient. In some other species, where the cytoplasm is rather dark, diVerent strategies must be used. The most common is the staining of chromosomal DNA with some vital stains like Hoechst (bisBenzimide). The chromosomes can then be seen in the oocyte cytoplasm under UV light either as individual spots or as a bright area. Another possibility is the use of the Pol‐scope microscope. This microscope incorporates liquid crystals with electrooptical control and software and thus employs circular polarized light to image birefringent structures. The successful use of the Pol‐ scope for spindle imaging has been demonstrated even in oocytes with high lipid content (sheep, bovine) as well as in human oocytes (Liu et al., 2000). When metaphase II oocytes are used, the chromosome group is usually located underneath the 1PB. Thus, it is possible to aspirate this part of the oocyte. Eventually, some additional noninvasive approaches could be used (Fulka et al., 2004). These approaches are used for the preparation of cytoplasts for subsequent nuclear transfer—cloning—the metaphase chromosome groups are discarded after the enucleation. Because we are discussing, in this chapter, the possible approaches with the eventual application in human medicine, where we suppose further use of chromosomes, we recommend the enucleation of oocytes either under UV light or with the Pol‐scope microscope. From a safety point of view, the Pol‐scope microscope seems to be at present the ideal equipment to use. Although technically feasible, only few papers reported the exchange of chromosome groups between oocytes. Moreover, these experiments were done exclusively in the mouse. Basically, this approach does not diVer significantly from the one where immature (germinal vesicle stage) oocytes are manipulated. The only diVerence is in the size of the enucleation pipette opening which is smaller. Cheng et al. (2003b) enucleated metaphase I mouse oocytes matured in vitro and transferred the isolated karyoplasts by electrofusion into corresponding cytoplasts. Although the fusion rate was rather lower, some oocytes were able to complete the process of maturation and reached the metaphase II stage. These oocytes were then fertilized in vitro and developed at least to the morula stage. An interesting observation has been made during this experiment. When isolated karyoplasts were cultured in vitro, they cleaved into two equal halves, each containing a clear metaphase spindle with attached chromosomes. Moreover, the distribution of chromosomes between these two halves was not random. Typically, each half contained 20 univalent chromosomes (metaphase II). Thus, in theory, it would be possible to separate these halves and use them individually for transfer into metaphase II enucleated oocytes. Then, we could expect that these oocytes would be identical. These observations were confirmed in our laboratory where we enucleated mouse oocytes

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soon after GVBD or later on approximately 1–2 hours before the expected exit from metaphase I (Fulka, Jr., unpublished data). The karyoplasts from more advanced oocytes also cleaved into two halves, though those karyoplasts from less advanced oocytes did not. (Fig. 4a–d). We believe that the observations describing the behavior of germinal vesicles as well as chromosome groups in karyoplasts are important from a practical point of view. Thus, the time interval between the karyoplast isolation from immature or early‐maturating oocytes can be relatively longer because the nuclear material remains intact. On the other hand, when more advanced metaphase I karyoplasts are isolated, they must be transferred into a host metaphase I cytoplast soon, that is, before they cleave. If they have already cleaved into two halves, it is impossible to transfer them into MI cytoplasts and they can be transferred only into metaphase II cytoplasts. Moreover, we assume that the transfer of the metaphase I group has another potential advantage. It is speculated that the aberrant function of the cytoplasm (mitochondria) directly leads to a disproportional segregation of chromosomes after the exit from metaphase I and consequently to aneuploidies in the embryo (Hassold and Hunt, 2001). It must be determined

Figure 4 (a) When karyoplasts isolated from advanced maturing oocytes are not immediately fused with cytoplasts, they cleave into two equal halves (arrows). Interference contrast, 200. (b) The parallel picture demonstrates that in cleaved karyoplasts, each half contains a group of chromosomes (arrows). Fluorescence, 200. (c) On the other hand, karyoplasts isolated from oocytes soon after germinal vesicle breakdown remain intact and do not cleave even after prolonged culture (arrow). Interference contrast, 200. (d) The parallel picture after staining with Hoechst shows a single chromosome group (arrow) in this type of karyoplast. Fluorescence, 200.

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whether the frequency of these chromosomal abnormalities will be reduced if metaphases are isolated from a defective cytoplasm and transferred into a normal cytoplasm in which the maturation process will be completed. Some results, however, showed that the transfer of germinal vesicles from aged mice into cytoplasts from young mice does not prevent age‐associated chromosome misalignment (Cui et al., 2005). The procedure where metaphase II oocytes are enucleated is basically the same as for oocytes in metaphase I. As already mentioned, this approach is commonly used for the preparation of cytoplasts that are used for nuclear transfer. In the context of this chapter, we must assume that karyoplasts are not discarded and will be used for transfer into a healthy cytoplasm. If ovulated oocytes are used, their developmental potential is theoretically very high. This will not be the case when the number of mtDNA copies is under an optimal level (Cummins, 2001), where for example oocytes originated from aged donors (Palermo et al., 2002) or in those specific cases that are diYcult to explain (Mrazek and Fulka, 2003). Here, in theory, chromosomal abnormalities can be expected; consequently, the developmental potential of these oocytes after fertilization will be low. The mature oocyte is naturally activated by sperm. Similarly, the MII cytoplast is also activated when the somatic cell is fused to it. This means that there is a potential danger of activation when isolated MII karyoplasts will be fused to recipient cytoplasts by electric pulses. Surprisingly, this has not been reported in experiments studying the exchange of metaphase II chromosome groups between oocytes. Moreover, in vitro fertilization of mouse oocytes reconstructed by nuclear transfer of metaphase II chromosome groups resulted in the birth of normal oVspring (Wang et al., 2001). Metaphase II chromosome transfer may however solve another problem, that is, the quality of the cytoplasm. In vitro parthenogenetic development is compromised when metaphase II in vitro matured oocytes, isolated from follicles that were grown in culture, were activated. On the other hand, a high rate of parthenogenetic development was reported for in vivo matured oocytes. The exchange of metaphase II groups between these two types of oocytes demonstrated that the developmental inability of cultured oocytes is of cytoplasmic origin (Liu et al., 2003b). This was confirmed in the mouse by Liu et al. (2003a) who transferred metaphase II groups of chromosomes from oocytes produced by germinal vesicle transfer into naturally ovulated and subsequently enucleated oocytes and reported the birth of several oVspring. The authors conclude that the process of oocyte maturation has two parts—cytoplasmic and nuclear and the relevant quality of both of them is essential for further embryonic development.

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VI. Transfer of Pronuclei The method enabling the exchange of pronuclei between mouse zygotes was in fact the first successful and repeatable technique that formed a firm basis for subsequent nuclear transfer and nucleus replacement experiments. Essentially, all those experiments mentioned above are based on the papers describing this method (McGrath and Solter, 1983, 1986). Moreover, with this approach some essential biological questions were also answered—imprinting, parthenogenesis, andro‐ and gynogenesis in mammals, nucleocytoplasmic interactions, and so on. The approach is relatively very simple, the zygote that was preincubated in a medium with cytochalasin is stabilized by the holding pipette in a position where both pronuclei are clearly visible. Then the enucleation pipette penetrates the zona pellucida and through this opening it is slowly pushed to the close vicinity of one pronucleus. The pipette does not penetrate into the cytoplasm. Very gentle suction is then applied and this aspirates a given pronucleus into the pipette. Thereafter, the opening of the pipette is positioned near the second pronucleus that is also aspirated into the pipette. As soon as both pronuclei are inside, the pipette is withdrawn (Fig. 5a–f). The cytoplasmic bridge that connects the karyoplast (containing both pronuclei and a small volume of the zygote cytoplasm) with the cytoplast is gradually stretched and forms a fine thread that is finally broken with the continued withdrawal of the pipette and the membranes in both the karyoplast and cytoplast seal. The karyoplast is then transferred to a position under the zona pellucida of another previously enucleated zygote. After a short incubation in a medium without cytochalasin, both pronuclei are fused with the cytoplast. This can be done either by inactivated Sendai virus or by electric pulse‐induced fusion. Polyethylene glycol‐induced fusion is not commonly used. The fusion rate as well as the development of reconstructed embryos is typically very high. Moreover, the karyoplasts can be successfully frozen and used for transplantation after thawing later on (He et al., 2003). The above approach has been almost exclusively used in the mouse but with some occasional reports in other species. Moreover, some species like pig and cattle do not contain clearly visible pronuclei in living zygotes. Thus, the centrifugation of oocytes before they are manipulated is necessary. The morphology of human zygotes is, however, completely diVerent because pronuclei are clearly visible in living zygotes (Fulka et al., 2004). In fact, the correction of polyploidy, when one pronucleus is removed from tripronuclear zygotes, uses some of the steps described above (Takeuchi et al., 2005b). Thus, in theory, we can expect that the pronuclear exchange

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Figure 5 (a–f) The enucleation and pronuclear transfer in mouse zygotes. (a) The zygote with male pronucleus (MPN) and female pronucleus (FPN) just before enucleation. The zygote is fixed in a holding pipette (H), the enucleation pipette (E) is on the right. 2PB, second polar body. Interference contrast, 350. (b) Parallel picture where the zygote DNA was stained with Hoechst. Note the positive staining in both pronuclei (PNs) as well as in the second polar body (arrow). Fluorescence, 350. (c) Parallel picture showing the distribution of mitochondria after their labeling with MitoTracker Green FM. Fluorescence, 350. (d) The isolated karyoplast (K) with both pronuclei is transferred under the zona pellucida (ZP) of a previously enucleated zygote and subsequently fused to it. Interference contrast, 350. (e) In a parallel picture, the staining with Hoechst demonstrates the presence of both pronuclei (arrows) in the karyoplast. Fluorescence, 350. (f) The karyoplast (K) with labeled mitochondria (arrows) can then be fused to the cytoplast in which mitochondria were not labeled (C). Fluorescence, 350.

between zygotes could be successfully used also in humans for the elimination of certain detrimental cytoplasmic defects. From a general point of view, we can suppose that the developmental potential of zygotes will be in a normal range or slightly lower in manipulated cells. This will not be the situation when the original oocyte cytoplasm is abnormal, for example, when it contains dysfunctional mitochondria or their number is low. It has been assumed that mitochondrial dysfunction may lead to an incorrect segregation of chromosomes especially during the transition from metaphase I to metaphase II. Then, after fertilization the embryonic chromosome complement will be also abnormal. Theoretically, pronuclear exchange would be the most convenient approach for the elimination of mutated mtDNA. Due to the mitochondria bottleneck segregation when primordial germ cells multiply, oocytes can contain a diVerent amount of mutated mtDNA—from low to very high number of copies (Taylor and Turnbull, 2005). The presence and quantity

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of mutated mtDNA can be assessed by preimplantation genetic diagnosis when isolated 2PBs are used (Briggs et al., 2000). They must be, however, isolated from fertilized eggs very soon after they are extruded and the analysed results must be made available whilst the zygote still contains visible pronuclei. If not, the zygotes or cleaving embryos with defected mtDNA should be discarded. We do believe that from the above‐described possibilities of nucleus replacement approaches, it is clear that these techniques can be in theory used at diVerent stages of maturation in fully grown oocytes as well as in zygotes with pronuclei. The choice of the most convenient stage depends on many factors. If we suppose the use in humans, the choice must be influenced by the problem we want to solve. For example, for the elimination of mutated mtDNA, the best would be the exchange of pronuclei between the patient and the donor zygote (Taylor and Turnbull, 2005). If the oocyte contains a suboptimal number of mtDNA copies (Reynier et al., 2001; Wilding et al., 2001) or oocytes are collected from patients in advanced age, the preferred alternative will be the exchange of germinal vesicles or metaphase I groups. Essentially, for every problem or disease, a very careful calculation must be performed before the most convenient alternative is selected. Ethical questions must be also taken into consideration. While the exchange of pronuclei in animals is without any problems, the use of the same approach is problematic in humans because the zygote that is used as a donor of the cytoplasm is in fact destroyed. This can be eventually avoided if parthenogenetic embryos are used as donors of cytoplasts. However, many biological questions resulting from the use of nucleus replacement techniques remain to be elucidated or are not yet fully answered (St. John et al., 2004). Even very small karyoplasts contain a certain amount of mitochondria that are introduced into the cytoplast and the resulting reconstructed embryo will then contain the mixed population of mitochondria—this situation is called ‘‘heteroplasmy.’’ It has been demonstrated basically for every oocyte or zygote stage that mitochondria are more abundant around nuclei (germinal vesicles, pronuclei) or chromosome groups (Van Blerkom et al., 2000). Thus, in theory, a relatively high amount of mitochondria will be introduced into the reconstructed oocyte or embryo. However, as shown in Fig. 3i, when mitochondria were labeled with MitoTracker Green FM, the fluorescence in karyoplasts was considerably lower than in corresponding cytoplasts. It cannot be excluded that cytochalasin treatment also disrupts some cytoskeletal components inside the oocyte or zygote and this disruption leads to a decreased number of mitochondria in isolated karyoplasts. These visual observations must be confirmed by detailed and precise analysis.

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The distribution of mitochondria from the karyoplast in reconstructed cells will also be essential. We have labeled isolated mouse GV karyoplasts with MitoTracker Green FM. These karyoplasts were then fused to unlabeled cytoplasts and the reconstructed oocytes were subsequently cultured in vitro up to the metaphase II stage. Our results showed that labeled mitochondria were uniformly distributed in mature oocytes (Fulka, 2004). Interestingly, the speed of mitochondrial distribution was cell‐cycle dependent, that is, rapid in interphase and slow in M‐phase cells (Fulka, 2004). When labeled mitochondria were injected directly into oocytes or zygotes slightly diVerent results were obtained (Barritt et al., 2001; Van Blerkom et al., 1998). Here, the injected population of mitochondria dispersed more slowly. The uniform distribution of mitochondria introduced is essential. Van Blerkom et al. (2000) showed that in developing human embryos their distribution is not uniform; with some blastomeres containing a very rich population of mitochondria while some were almost without these organelles. Thus, the best solution would be a complete elimination or destruction of the karyoplast’s mitochondria. These attempts were reported in several papers. Palermo et al. (2002) and Takeuchi et al. (2005a) labeled mitochondria in immature mouse oocytes with chloromethyl‐X‐rosamine (CMXRos) and subsequently induced their damage by UV light photoirradiation with a Texas Red filter for 10 s. This photosensitization inhibited GVBD and oocyte maturation in almost all cases. When germinal vesicles from these oocytes were transferred into nontreated cytoplasts, more than 70% of reconstructed cells completed maturation and reached metaphase II. These oocytes were then injected with spermatozoa and about 21% of successfully fertilized cells reached blastocyst stage. The transfer of 132 two‐cell stage embryos into oviducts of pseudo‐pregnant recipients resulted in 17 healthy oVspring. Another approach used the treatment of mouse zygotes with H2O2 that induced mitochondrial dysfunction and embryonic developmental arrest. However, when pronuclei from the treated zygotes were transferred into a healthy cytoplasm, the development of reconstructed embryos was only partly rescued. This indicates that this treatment also damages pronuclei but the cytoplasm is more sensitive (Liu and Keefe, 2000). There are some other possibilities for destroying mitochondria (e.g., with some potent mitochondrial toxins like chloramphenicol). The above examples, however, indicate that the evaluation of these treatments must be very exact. The oocyte or embryo represents a very sensitive complex system where the precise coordination between the nucleus and the cytoplasm plays an essential role not only during the embryonic development but also later on, and some abnormalities and defects can appear at any stage during a lifetime (Hawes et al., 2002; Hiendleder et al., 2005; St. John, 2002; St. John et al., 2004).

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VII. Conclusions It was not our intention to give, in this chapter, a comprehensive review of those approaches that can be used for the rescue of nuclei from a cytoplasm harboring defects. It is, however, clear that nuclear replacement can be performed basically in all stages of oocyte maturation as well as in pronuclear stage zygotes—that is, in cells that are currently used in human‐assisted reproduction. From the technical point of view, in experienced and skilled hands, these techniques are rather simple and trivial. As it has been clearly demonstrated, the number and especially the function of mitochondria influences many processes, including oocyte maturation, the proper segregation of chromosomes, fertilization outcome as well as embryonic development (Almeida Santos et al., 2006; El Shourbagy et al., 2006). Moreover, mutated mtDNA that is exclusively maternally inherited (or almost exclusively) may lead to certain devastating diseases in children (Wallace, 1999). Some other cytoplasmic defects that are not yet fully understood could be also eliminated by nucleus replacement techniques (Mrazek and Fulka, 2003). We have shown that the replacement of nuclei can be done relatively easily and with high eYciency. Many questions do however remain. One such question is the importance of the compatibility between the nucleus and the cytoplasm. Mitochondria do not represent an independent functional unit and also some nuclear factors, like mitochondrial transcription factor A, play a vital role too (Van Blerkom, 2004). Very important will be the elimination of the residual karyoplast’s mitochondria and thus the prevention of heteroplasmy in the reconstructed cells. The injection of mitochondria (together with the sperm) from a donor oocyte into a patient’s defective oocyte improves developmental potential of the embryo and several children were born after this treatment. The slight increase in the frequency of certain abnormalities in these children clearly indicates that some additional experiments with careful evaluation must be done (Spikings et al., 2006). However, the nuclear transfer experiment in mito‐mice that carry pathogenic mtDNA clearly demonstrates the feasibility and perspective of nucleus replacement techniques (Sato et al., 2005).

Acknowledgments We would like to dedicate this chapter to J. Fulka, Sr., the founder of modern reproductive biology in the Czech Republic, who will be 80 in 2007 and to our teachers and friends Bob Moor who will be 70 in the same year and Jan Motlik who is 60 in 2006. H. F. and J. F., Jr. are supported from ESF GACR STE/05/E004 and MZE0002701401.

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Gioia, L., Barboni, B., Turriani, M., Capacchietti, G., Pistilli, M. G., Berardinelli, P., and Mattioli, M. (2005). The capability of reprogramming the male chromatin after fertilization is dependent on the quality of oocyte maturation. Reproduction 130, 29–39. Hassold, T., and Hunt, P. (2001). To err (meiotically) is human: The genesis of human aneuploidy. Nat. Rev. Genet. 2, 280–291. Hawes, S. M., Sapienza, C., and Latham, K. E. (2002). Ooplasmic donation in humans. The potential for epigenetic modifications. Hum. Reprod. 17, 850–852. He, Z., Liu, H. C., and Rosenwaks, Z. (2003). Cryopreservation of nuclear material as a potential method of fertility preservation. Fert. Steril. 79, 347–354. Hiendleder, S., Zakhartchenko, V., and Wolf, E. (2005). Mitochondria and the success of somatic cell nuclear transfer cloning: From nuclear‐mitochondrial interactions to mitochondrial complementation and mitochondrial DNA recombination. Reprod. Fert. Dev. 17, 69–83. Hogan, B., Beddington, R., Constantini, F., and Lacy, E. (1994). ‘‘Manipulating the Mouse Embryo: A Laboratory Manual,’’ 2nd edn., p. 497. Cold Spring Harbor Laboratory Press, New York. Karnikova, L., Urban, F., Moor, R. M., and Fulka, J., Jr. (1998). Mouse oocyte maturation: The eVect of modified nucleo‐cytoplasmic ratio. Reprod. Nutr. Dev. 38, 665–670. Kono, T., Obata, Y., Wu, Q., Niwa, K., Ono, Y., Yamamoto, Y., Park, E. S., Seo, J. S., and Ogawa, H. (2004). Birth of parthenogenetic mice that can develop to adulthood. Nature 428, 860–864. Kren, R., Fulka, J., Jr., and Fulka, H. (2005). Cryopreservation of isolated mouse germinal vesicles. J. Reprod. Dev. 51, 289–292. Kuwayama, M. (2003). Birth of normal calf after transfer of the defective oocyte from old infertile cattle after germinal vesicle transfer. Theriogenology 59, 268(Abstract). Li, Y., Feng, H. L., Cao, Y. J., Zheng, G. J., Yang, Y., Mullen, S., Critser, J. K., and Chen, Z. J. (2006). Confocal microscopic analysis of the spindle and chromosome configurations of human oocytes matured in vitro. Fert. Steril. 85, 827–832. Liu, H., Wang, C. W., Grifo, J. A., Krey, L. C., and Zhang, J. (1999). Reconstruction of mouse oocytes by germinal vesicle transfer: Maturity of host oocyte cytoplasm determines meiosis, Hum. Reprod. 14, 2357–2361. Liu, H., Chang, H. C., Zhang, J., Grifo, J., and Krey, L. C. (2003a). Metaphase II nuclei generated by germinal vesicle transfer in mouse oocytes support embryonic development to term. Hum. Reprod. 18, 1903–1907. Liu, J., Van der Elst, J., and Dhont, M. (2003b). In vitro parthenogenetic development of mouse oocytes following reciprocal transfer of the chromosomes between in vivo‐matured oocytes and in vitro‐matured oocytes. Biol. Reprod. 68, 186–189. Liu, L., and Keefe, D. L. (2000). Cytoplasm mediates both development and oxidation‐induced apoptotic cell death in mouse zygotes. Biol. Reprod. 62, 1828–1834. Liu, L., Oldenbourg, R., Trimarchi, J., and Keefe, D. L. (2000). A reliable, noninvasive technique for spindle imaging and enucleation of mammalian oocytes. Nature Biotechnol. 18, 223–225. McGrath, J., and Solter, D. (1983). Nuclear transplantation in the mouse embryo by microsurgery and cell fusion. Science 220, 1300–1302. McGrath, J., and Solter, D. (1986). Nuclear and cytoplasmic transfer in mammalian embryos. In ‘‘Developmental Biology’’ (R. B. L. Gwatkin, Ed.), Vol. 4, pp. 37–55. Plenum Press, New York. Monk, M. (Ed.) (1987). ‘‘Mammalian Development: A Practical Approach,’’ p. 313. IRL Press, Oxford, Washington. Moor, R. M., Dai, Y., Lee, C., and Fulka, J., Jr. (1998). Oocyte maturation and embryonic failure. Hum. Reprod. Update 4, 223–236.

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Motlik, J., and Fulka, J. (1981). Fertilization of rabbit oocytes cocultured with granulosa cells. J. Reprod. Fert. 63, 425–429. Mrazek, M., and Fulka, J., Jr. (2003). Failure of oocyte maturation: Possible mechanisms for oocyte maturation arrest. Hum. Reprod. 18, 2249–2252. Palermo, G. D., Takeuchi, T., and Rosenwaks, Z. (2002). Technical approaches to correction of oocyte aneuploidy. Hum. Reprod. 17, 2165–2173. Reynier, P., May‐Panloup, P., Chretien, M. F., Morgan, C. J., Jean, M., Savanger, F., Barriere, P., and Malthiery, Y. (2001). Mitochondrila DNA content aVects the fertilizability of human oocytes. Mol. Hum. Reprod. 7, 425–429. Sato, A., Kono, T., Nakada, K., Ishikawa, K., Inoue, S. I., Yonekawa, H., and Hayashi, J. I. (2005). Gene therapy for progeny of mito‐mice carrying pathogenic mtDNA by nuclear transplantation. Proc. Natl. Acad. Sci. USA 102, 16765–16770. Spikings, E. C., Alderson, J., and St. John, J. C. (2006). Transmission of mitochondrial DNA following assisted reproduction and nuclear transfer. Hum. Reprod. Update 12(4), 401–415; doi: 10.1093/humupd/dm1011. St. John, J. C. (2002). Ooplasm donation in humans. The need to investigate the transmission of mitochondrial DNA following cytoplasmic transfer. Hum. Reprod. 17, 1954–1958. St. John, J. C., Lloyd, R. E. I., Bowles, E. J., Thomas, E. C., and El Shourbagy, S. (2004). The consequence of nuclear transfer for mammalian foetal development and oVspring survival. A mitochondrial DNA perspective. Reproduction 127, 631–641. Staigmiller, R. B., and Moor, R. M. (1984). EVect of follicle cells on the maturation and developmental competence of ovine oocytes matured outside the follicle. Gamete Res. 9, 221–229. Takeuchi, T., Gong, J., Veeck, L. L., Rosenwaks, Z., and Palermo, G. D. (2001). Preliminary findings in germinal vesicle transplantation of immature human oocytes. Hum. Reprod. 16, 730–736. Takeuchi, T., Rosenwaks, Z., and Palermo, G. D. (2004). A successful model to assess embryo development after transplantation of prophase nuclei. Hum. Reprod. 19, 975–981. Takeuchi, T., Neri, Q. V., Katagiri, Y., Rosenwaks, Y., and Palermo, G. D. (2005a). EVect of treating induced mitochondrial damage on embryonic development and epigenesis. Biol. Reprod. 72, 584–592. Takeuchi, T., Neri, Q. V., and Palermo, G. D. (2005b). Construction and fertilization of reconstituted human oocytes. RBM Online 11, 309–318. Taylor, R. W., and Turnbull, D. M. (2005). Mitochondrial DNA mutations in human disease. Nature Rev. Genet. 6, 389–402. Tesarik, J., Nagy, Z. P., Mendoza, C., and Greco, E. (2000). Chemically and mechanically induced membrane fusion: Non‐activating method for nuclear transfer in mature human oocytes. Hum. Reprod. 15, 1149–1154. Trounson, A. O., Anderiesz, C., and Jones, G. (2001). Maturation of human oocytes in vitro and their developmental competence. Reproduction 121, 51–75. Tsai, M. C., Takeuchi, T., Bedford, J. M., Reis, M., Rosenwaks, Z., and Palermo, G. D. (2000). Alternative sources of gametes: Reality or science fiction? Hum. Reprod. 15, 988–998. Van Blerkom, J. (2004). Mitochondria in human oogenesis and preimplantation embryogenesis: Engines of metabolism, ionic regulation and developmental competence. Reproduction 128, 269–280. Van Blerkom, J., Sinclair, J., and Davis, P. (1998). Mitochondrial transfer between oocytes: Potential applications of mitochondrial donation and the issue of heteroplasmy. Hum. Reprod. 13, 2857–2868. Van Blerkom, J., Davis, P., and Alexander, S. (2000). DiVerential mitochondrial distribution in human pronuclear embryos leads to disproportionate inheritance between blastomeres:

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Difficulties and Possible Solutions in the Genetic Management of mtDNA Disease in the Preimplantation Embryo J. Poulton,* P. Oakeshott,{ and S. Kennedy * *NuYeld Department of Obstetrics and Gynaecology, The Women’s Center University of Oxford, Oxford OX3 9DU, United Kingdom { Community Health Sciences, St. George’s University of London London SW17 0RE, United Kingdom

I. Heteroplasmy of mtDNA Presents Problems for Chorionic Villus Sampling A. Consequences of Heteroplasmy: Dosage and Thresholds B. Is mtDNA Exclusively Maternally Inherited? C. Variable Distribution of mtDNA in Heteroplasmic Embryos: Implications for CVS D. Routine CVS for mtDNA Disease II. The Mitochondrial Bottleneck A. Physiological Basis of the Bottleneck: Animal Models of the Bottleneck Using Polymorphic mtDNA Variants B. Mathematical Models of the Bottleneck III. Management Options Beyond CVS A. Preconception Counseling Based on Analysis of Oocytes and Pedigree Analysis B. Preimplantation Genetic Diagnosis C. Nuclear Cell Transfer References

Families who have had a child die of a severe, maternally inherited mitochondrial DNA (mtDNA) disease are usually desperate to avoid having further aVected children. Here we discuss the problems of applying classical genetic management to mtDNA diseases (Poulton and Turnbull, 2000) and the biology underlying these problems. We explain why these disorders have lagged so far behind the genetics revolution. We then outline the directions in which management is likely to develop, including the use of preimplantation genetic diagnosis (PGD). ß 2007, Elsevier Inc.

Current Topics in Developmental Biology, Vol. 77 Copyright 2007, Elsevier Inc. All rights reserved.

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0070-2153/07 $35.00 DOI: 10.1016/S0070-2153(06)77008-9

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I. Heteroplasmy of mtDNA Presents Problems for Chorionic Villus Sampling A. Consequences of Heteroplasmy: Dosage and Thresholds Mitochondrial diseases manifest with a range of clinical presentations from severe liver disease, which may be fatal from the neonatal period, to adult onset myopathy, cardiomyopathy, or acute loss of vision. The conditions may be caused by mutations in either mitochondrial or nuclear genes and are hence associated with both maternal and Mendelian (dominant or recessive) patterns of inheritance. Prenatal diagnosis using chorionic villus sampling (CVS) is available for mitochondrial diseases caused by mutations in nuclear genes that are inherited as Mendelian traits. However, CVS is unreliable for maternally inherited diseases because of the problem of heteroplasmy. Thousands of mtDNA copies are present in every nucleated cell and normal individuals are homoplasmic, that is, virtually all their mtDNA copies are identical. However, aVected individuals are usually heteroplasmic, that is, most of their tissues or cells have both normal (wild type) and mutant mtDNAs, with proportions varying from 0% to 100%. Heteroplasmy may cause the variable clinical severity of mtDNA disorders because, in some tissues, the mutant levels change over time: falling, for example, in blood (Rahman et al., 2001) and accumulating in nondividing cells such as muscle (Poulton and Morten, 1993). There is also a threshold eVect (tissues function normally unless the proportion of mutant mtDNA rises above a particular level) in most diseases whose progression is explained by preferential accumulation of mutant mtDNAs in aVected tissues (Poulton et al., 1995; Weber et al., 1997). B. Is mtDNA Exclusively Maternally Inherited? In humans the term mitochondrial inheritance is synonymous with maternal inheritance. At fertilization, the sperm contributes about 700–1200 mtDNAs (Diez‐Sanchez et al., 2003) or approximately 0.5% of the zygote’s total DNA. Studies of paternal transmission in intraspecific mouse crosses show that paternal mtDNA is lost very early in embryogenesis (Kaneda et al., 1995). Paternal mitochondria labeled with vital dyes prior to in vitro fertilization (IVF) are degraded by the early pronucleus stage apparently via ubiquitinylation (Sutovsky et al., 2000). However, in mouse interspecific crosses, paternal mtDNA is detectable from the stage of single cell embryos through to neonates albeit at low levels (<1%) (Kaneda et al., 1995). However, the paternal mtDNA is restricted to certain tissues and is not transmitted to oVspring; hence, it does not enter the germ line. Transmission of paternal mtDNA in mouse thus appears to be unlikely (Shitara et al., 1998). In man, the possibility of paternal transmission has become an issue because

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of advances in treatment of male infertility due to deficient sperm function and/or low sperm numbers. It is now possible to generate viable human embryos by microinjection of single sperm into unfertilized human ooctyes (so‐called ICSI for intracytoplasmic sperm injection) (Palermo et al., 1992). Mitochondria in the sperm tail are not specifically excluded from the embryo during natural fertilization (Ankel‐Simons and Cummins, 1996) but rapidly destroyed (Sutovsky et al., 2000), the ICSI procedure might introduce more paternal mtDNA than normal. This is an important issue with potentially long‐term clinical consequences because evidence is accumulating that mitochondrial dysfunction may contribute to male infertility (Rovio et al., 2001; Ruiz‐Pesini et al., 2000). The procedure may therefore increase the likelihood of injecting mtDNA carrying pathogenic mutants. However, several studies suggest that no detectable paternal contribution results from this procedure (Marchington et al., 2002). Just occasionally paternal mtDNA may be important. There is a case report of a unique patient with mitochondrial disease arising from a mutation in paternal mtDNA (Schwartz and Vissing, 2002). No wild‐type (or normal) paternal mtDNA was detectable in any tissue, and the mutant paternal mtDNA was detectable only in muscle where it composed 90% of mtDNA. It would therefore appear that the mtDNA mutation conferred an exceptional advantage to the mutant mtDNA, but only in the myogenic lineage. In another case report, the mtDNA haplotype of a boy with Klinefelter’s syndrome, extensively investigated for medicolegal reasons, was substantially diVerent from that of his mother (Oikawa et al., 2002). It is possible that elimination of nonmaternal mtDNA may be incomplete in abnormal embryos. This is suggested by apparent persistence of paternal mtDNA in arrested human embryos (St. John et al., 2000) and by loss of donor mtDNA in preference to recipient in mouse chimeras derived from aneuploid embryonic stem cells (Marchington et al., 1999). While it is conceivable that this might indicate persistence and proliferation of paternal mtDNA (Schwartz and Vissing, 2002) in the presence of paternal nondysjunction (Oikawa et al., 2002), critics have suggested that this paper is flawed and that sequencing errors are the correct explanation (Bandelt et al., 2005). It is widely believed that these are exceptional cases and that paternal transmission of mtDNA is rare; hence, it will be discounted for the remainder of this chapter.

C. Variable Distribution of mtDNA in Heteroplasmic Embryos: Implications for CVS If the fate of an 11‐week fetus is to be determined by analysis of the load of mutant mtDNA in a single CVS, its distribution should be uniform throughout the embryo and extraembryonic tissues. Data exist on the distribution of pathogenic mtDNA mutants in diVerent neonatal tissues, but not in

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multiple simultaneous placental samples from a known female carrier. There is one report of prenatal diagnoses for A3243G in five families, in which CVS followed by two amniocenteses revealed 6–8% variation in level of mutant mtDNA (Bouchet et al., 2006) that could represent either local variation or sequential change. Few carrier/aVected fetuses have been analyzed for mutant mtDNA load in diVerent tissues. However, the limited existing data from studies on human fetuses or neonates with pathogenic mtDNA mutations also indicate that mutant mtDNAs do not segregate greatly during embryogenesis (Harding et al., 1992b; Matthews et al., 1994; Suomalainen et al., 1993). In two of the animal models of heteroplasmic mtDNA segregation, the proportion of each mtDNA variant was uniform in all tissues of the fetuses analyzed. However, in two of these experiments there was tissue specific, directional selection for diVerent mtDNA genotypes in the same animal (Jenuth et al., 1997; White, 1999). These studies used nonpathogenic mtDNA variants but are consistent with the limited studies that have been performed on pathogenic variants, indicating that mtDNA probably does not segregate greatly during embryogenesis. We used naturally occurring mtDNA variants to investigate mtDNA segregation in human placenta (Marchington et al., 2006). Using large samples of control placenta, we demonstrated that the level of polymorphic heteroplasmic mtDNA variants is very similar in mother, cord blood, and placenta. However, where placental samples were very small [<10 mg, comparable in size to normal CVS (10–30 mg)] there was clear evidence of variation in the distribution of mtDNA polymorphic variants. This is entirely consistent with previous data showing that neighboring regions of placental tissue are arranged in ‘‘patches’’ or clones, arising from diVerent cells in the early embryo (Lau et al., 1997). The larger 50–100 mg fragments are presumably more likely to sample two or more placental patches than the smaller fragments. Our data suggest that sample variation attributable to patch size is more marked in the term placenta, where patches will presumably have grown larger than they are in the 7‐week placenta. In practice, the magnitude of the variation that we found in placentas at 7 weeks gestation may be too small to have much clinical impact (coeYcient of variation <0.18). We and others suggest that accurate prenatal diagnosis for some mtDNA diseases may require at least two separate samples (Bouchet et al., 2006; Marchington et al., 2006). While it is not clear whether and by how much taking two samples would impair fetal viability, this might increase the miscarriage rate (Rhoads et al., 1989).

D. Routine CVS for mtDNA Disease CVS has been successfully carried out in several women carrying the neuropathy, ataxia, and retinitis pigmentosa (NARP) mutation (Harding et al., 1992a;

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White et al., 1999a,b). In all cases, the mutant load was either high or low, enabling accurate predictions. However, caution is needed for other mtDNA disorders in which the correlation between severity and mutant load is less precise (Poulton and Turnbull, 2000). Patients must be carefully selected to ensure that the procedure will be helpful, as CVS may be uninformative where the mutant load has an intermediate level that is poorly predictive of outcome (Bouchet et al., 2006). As this requirement excludes a large proportion of the women in families carrying pathogenic mtDNA mutants from seeking help, some groups take the risk of an uncertain outcome (Bouchet et al., 2006). Hence, classic CVS is diYcult to apply to mtDNA disease without additional measures. However, our knowledge of mtDNA during gametogenesis and preimplantation development is rudimentary. In particular, the mtDNA bottleneck has a major impact on the feasibility and time window for genetic management.

II. The Mitochondrial Bottleneck Accurate estimates of the risk of recurrence are diYcult because of the mitochondrial ‘‘bottleneck’’ in transmission of mtDNA. Furthermore, the feasibility of prenatal diagnosis is critically dependent on the nature and timing of this bottleneck. This is a unique consequence of heteroplasmy (Marchington et al., 1998). At its most extreme, there may be complete switching of mtDNA type in a single transmission so that all the copies of mtDNA in a mother diVer from those of her oVspring by a single nucleotide. Because oocytes contain approximately 100,000 mtDNAs and yet the mutation probably only occurs once, a single mtDNA molecule must have been the mitochondrial founder for the child in this extreme example. This restriction in mtDNA numbers followed by amplification is described as a genetic ‘‘bottleneck’’ (Marchington et al., 1997). It has been studied in patients with and without mtDNA disease. We investigated the mitochondrial bottleneck in normal oocytes from couples in our IVF clinic referred for male infertility, using naturally occurring length variation in the large noncoding region of mtDNA. This region of mtDNA deviates from the rule that homoplasmy is the norm, as some individuals are heteroplasmic for diVerent length variants that are apparently neutral polymorphisms. The modal length variant diVered between oocytes from a single control individual, showing that segregation of founder mtDNA molecules probably occurs well before the oocytes are mature (Marchington et al., 1997). However, in most cases, the segregation was less extreme and more comparable to situations seen in clinical practice. For instance, a woman carrying 30% pathogenic 8344A>G mutant mtDNA may have children with anything from 0% mutant (hence healthy) to 80% mutant (severely aVected) (Larsson et al., 1992). In one previous report (Blok

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et al., 1997), oocytes from a mother with three severely aVected children (the mother carrying 50% and the children 87–98% 8993T>G mutant mtDNA) were obtained by conventional IVF methods, that is, ovarian stimulation and transvaginal oocyte aspiration. Seven oocytes were retrieved and analyzed of which one carried 0% and six carried >95% mutant. In addition, we have examined oocytes from a patient with Kearn–Sayre syndrome caused by mtDNA rearrangements (Marchington et al., 1998). Significant levels of rearranged mtDNA were detectable in the majority of the patient’s oocytes, by use of multiplex PCR, with wide variation in the levels of mutant and wild‐ type molecules between individual oocytes. We also used length variation in a homopolymeric C tract, which is often heteroplasmic in normal controls, to identify founder subpopulations of mtDNAs in this patient’s oocytes. The number of segregating units (n) was 3–5 orders of magnitude less than the number of mitochondria in the human female oocyte (Marchington et al., 1998). A similar study was carried out on oocytes from a woman with the 3243A>G mutation (Brown et al., 2001). In conclusion, the bottleneck appears to contribute a major component of the variable transmission from mother to oocyte. That this bottleneck had occurred by the time that oocytes were mature advances the prospects for prenatal diagnosis of mtDNA diseases (Marchington et al., 1997). These examples suggest that a small number of mtDNAs (as few as one in the latter case) become the mtDNA founder for each oVspring.

A. Physiological Basis of the Bottleneck: Animal Models of the Bottleneck Using Polymorphic mtDNA Variants The precise timing of the bottleneck is a critical determinant of the genetic management options available. To study this in detail, four groups constructed heteroplasmic mouse models of mtDNA segregation (Jenuth et al., 1996; Laipis, 1996; Meirelles and Smith, 1997). Heteroplasmy for two common polymorphic mtDNA types was generated by introducing donor cytoplasm into a fertilized recipient mouse egg. Analysis of developing female germ cells demonstrated that the major component of the bottleneck occurs between the primordial germ cell and primary oocyte stage. As in humans, these data imply that the major component of the bottleneck has occurred by the time oocytes are mature. Blastomeres were analyzed from preimplantation embryos derived from these mouse strains. All blastomeres from a single embryo contained very similar loads of mutant mtDNA (Dean et al., 2003; Jenuth et al., 1996). Furthermore, the polar body of an unfertilized oocyte or early embryo was representative of the level in the oocyte or embryo as a whole. Reliable results were obtained from both polar bodies and blastomeres, but the eYciency of diagnosis was greater with blastomeres.

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In two of the animal models of heteroplasmic mtDNA segregation, the proportion of each mtDNA variant was uniform in all the fetal tissues analyzed. However, in two of these experiments there was tissue‐specific, directional selection for diVerent mtDNA genotypes in the same animal after birth (Jenuth et al., 1997; White, 1999). When this type of segregation occurs in human diseases it is generally believed to indicate a functional diVerence between mtDNA types (Lightowlers et al., 1997). This suggests that the experiments are, in fact, modeling pathogenic human mtDNA mutants, and if so, are generally reassuring for prenatal diagnosis. Taken together, these studies suggest that a major bottleneck occurs during oogenesis and that mtDNA segregate little during prenatal development, even in mutants that are only mildly detrimental.

B. Mathematical Models of the Bottleneck Two types of mathematical model have been used to describe the mitochondrial bottleneck, diVering in whether they consider a single or multiple selection (Poulton et al., 2003). The multiple selection model is the most widely used, and it assumes that the bottleneck occurs as a result of mtDNA segregation over 30 identical cell divisions, during which the mtDNA content of the cell partitions stochastically. The model is a poor fit in biological terms because reality does not fit these assumptions: the mtDNA content of the oocyte varies 50‐ to 100‐fold. In practice, both single and multiple selection models generate frequency distributions for heteroplasmic oocytes/ oVspring of a similar shape, even though the bottleneck size (or number of segregating units, n) is radically diVerent. The advantage of the single selection model is that it can be used to model recurrence risks ‘‘on the back of an envelope’’ in the clinic for counseling patients. As new data emerge on mtDNA synthesis and segregation in early embryos, it will be possible to generate better models. Some authors have asserted that selection in favor of certain pathogenic mtDNA mutants (Wong et al., 2002) and others that random genetic drift determine the level of mutant mtDNA in human primary oocytes (Brown et al., 2001; Jenuth et al., 1996). Such studies are problematic both because of ascertainment bias and because they are largely based on measurements of mutant loads in blood, and this generally falls with time. They tend to underestimate the level of mutant in the older generation (Wong et al., 2002). A closer look at the data (Poulton et al., 2003) suggests alternatives. Selection against detrimental mtDNA mutants during oocyte maturation oVers a simple explanation of the data on the human and mouse germ line. In severe pathogenic mutants, this results in a trend toward a lower proportion of mutant mtDNA in the oVspring/oocytes and a larger apparent bottleneck.

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In milder/polymorphic mtDNA variants, the apparent bottleneck size is smaller and consequently swings between generations can be much larger. The assumption of random genetic drift that has been made by previous investigators is not able to explain the apparent diVerences in bottleneck sizes (Blok et al., 1997; Brown et al., 2001; Chinnery et al., 2000).

III. Management Options Beyond CVS A. Preconception Counseling Based on Analysis of Oocytes and Pedigree Analysis Given the uncertainties of CVS, families who have had a child with a severe, maternally inherited mtDNA disease often seek preconceptual counseling, but providing accurate information about risk recurrence is diYcult. Consequently, many families reluctantly decide not to have more children. Thankfully, alternative approaches are now available such as oocyte sampling, which involves measuring the mutant load in 6–10 unfertilized oocytes obtained by conventional IVF methods. The first attempt to use oocyte sampling for estimating recurrence risk of severe maternally inherited mtDNA disease in humans was conducted in 2002. In this case, the happy outcome was a normal baby for one family. CVS would be advisable after oocyte sampling, even though the concerns about intermediate levels would still exist (Marchington et al., submitted for publication). There are other reasons why genetic counseling in mtDNA diseases is diYcult. One can obtain a mother’s oocytes to estimate likely mutant levels in her unborn children, but is it appropriate to take tissue samples from her existing children? The British Society for Human Genetics recommends that presymptomatic children should only undergo genetic testing where there is a clear benefit to early diagnosis (http://www.bshg.org.uk/documents/ oYcial_docs/testchil.htm 1994). Suppose an asymptomatic sibling is found to have an intermediate or even a high mutant level, does this mean that (s)he will develop symptoms? Without eVective treatments, is the label an advantage or disadvantage? If oocytes are obtained, will they be typical of those that might produce the next child?

B. Preimplantation Genetic Diagnosis Although not yet available in the United Kingdom for mtDNA diseases, preimplantation genetic diagnosis (PGD) could be the best option for patients with a high risk of transmission. This involves analyzing embryos produced by IVF and selecting only those at very low risk for transfer to the

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uterus. A single blastomere is removed from the cultured embryo around the 16‐cell stage. Only embryos with low mutant loads that are well below the threshold for aVecting cell function (see above) are placed into the uterus to implant. As in the case of CVS, the accuracy of this technique relies on the load of mutant mtDNA in the cell sample, reflecting the load in the rest of the embryo. While the distribution of polymorphic (neutral) variants in both early human (SteVann et al., 2006 and J. P., unpublished data) and mouse (Dean et al., 2003) embryos suggests that this is likely, there is very little or published data on pathogenic mtDNA mutations (SteVan et al., 2005). PGD could reduce the likelihood of later obtaining an intermediate mutant level at CVS by selecting only those embryos with low levels. However, the ‘‘take home baby’’ rate after PGD is not high (Platteau et al., 2006) and women with normal fertility might be reluctant to undergo an invasive treatment with known risks. Oocyte donation would avoid all these problems, but there is a shortage of oocyte donors and first‐degree relatives are obviously unsuitable (donors in the maternal lineage are potential carriers of mutant mtDNA). The polar body of an unfertilized oocyte or from a single blastomere of a mouse embryo is representative of the level in the embryo as a whole (Dean et al., 2003). Reliable results were obtained from both polar bodies and blastomeres, but the eYciency of diagnosis was greater with blastomeres (Dean et al., 2003). In contexts where PGD is not ethically acceptable, it might also be possible to choose oocytes for fertilization by analysis of the first polar body (Dean et al., 2003). However, in view of the paucity of data on human embryos with pathogenic mutations, concordance testing of two blastomeres for embryos to be implanted in the uterus might be necessary. This implies that dual sampling must be considered for PGD for mtDNA disease, the second sample being taken either before implantation or as a CVS.

C. Nuclear Cell Transfer What about nuclear cell transfer (Hwang et al., 2005)? Despite the technical challenges (Hwang et al., 2005) and recent bad publicity (Kennedy, 2006), this approach is making significant strides. The development will facilitate treatment not only of diseases requiring stem cell therapy but also of mtDNA diseases. Since Dolly, the sheep, was created by fusing a cell nucleus from an adult somatic cell with a recipient‐enucleated oocyte, producing in Dolly mtDNA inherited not from the somatic cell donor but the recipient oocyte (Campbell et al., 1996), researchers have realized it might be possible to alter the mitochondrial population of a human embryo using nuclear transfer. Hence, a group in Newcastle (United Kingdom) has recently applied to the Human Fertility and Embryology Authority for permission to attempt nuclear transfer to prevent recurrence of maternally inherited mtDNA disease.

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Is it likely that the Newcastle team’s approach of pioneering nuclear transfer will be a viable alternative strategy to more conventional genetic management? It sounds simple yet problems have been reported. These problems can be reduced by transferring zygotic, instead of somatic cell, nuclei. As yet there are very many unknowns. Most of the animal oVspring generated by nuclear transfer experiments seem normal, but malformations (Sun et al., 2005) and growth abnormalities (Hiendleder et al., 2004) have been reported. mtDNA only has a small number of genes, the vast majority of the thousands of mitochondrial proteins are encoded by the nucleus. There are therefore a large number of important interactions between the products of mitochondrial and nuclear genes that could go wrong. In a normal embryo, half the nuclear genes are inherited with the mtDNA from the mother, which means they must interact appropriately with the mitochondria. In embryos derived by nuclear transfer, the DNA originates from three unrelated parents (two providing the nucleus from the zygote and one the mtDNA in the oocyte), but none of these nucleo‐mitochondrial interactions has been tested. Furthermore, genetic inheritance of mtDNA may be even more complex than this. Nonhuman primate Macaca mulatta oVspring born by nuclear transfer have mtDNA from three sources: (1) maternal mtDNA from the recipient egg, (2) maternal mtDNA from the egg contributing to the donor blastomere, and (3) paternal mtDNA from the sperm that fertilized the egg from which the donor blastomere was isolated (St. John and Schatten, 2004). Hence, the Newcastle team will be tackling formidable unknowns. In conclusion, the many ethical, scientific, and pragmatic problems have been a major impediment to the genetic management of mtDNA diseases. Despite this, the most ethical course of action may be to weigh up the uncertainties and use new approaches in an attempt to help these distressed families.

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Key points It is very diYcult to predict recurrence risk in maternally inherited

mtDNA disease due to heteroplasmy. This means genetic counselling of aVected families is extremely

diYcult. CVS may be unreliable. The major bottleneck in transmission of mtDNA occurs during

oogenesis. Analysis of oocytes may give an estimate of risk recurrence. PGD may also be used in future.

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Section IV The Effects of Invasive Assisted Reproductive Technologies on mtDNA Transmission

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9

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Impact of Assisted Reproductive Technologies: A Mitochondrial Perspective of Cytoplasmic Transplantation A. J. Harvey,*,{ T. C. Gibson,* T. M. Quebedeaux,* and C. A. Brenner *,{ *Department of Biology, University of New Orleans New Orleans, Louisiana 70148 { Pennington Biomedical Research Center Baton Rouge, Louisiana 70808

I. Why Are Invasive Assisted Reproductive Technology Procedures Important? II. Cytoplasmic Transfer: Animal Models That May Assist Human Infertility Therapeutics III. The Mitochondrial Genome and Mitochondrial Biogenesis A. Mitochondrial Replication B. Mitochondrial Transcription IV. Molecular Control of Mitochondrial Function in Preimplantation Embryos V. Mitochondrial Copy Number, Deletions, and Mutations VI. Oocyte and Embryo Mitochondrial Localization Patterns VII. Implications of Mitochondrial Heteroplasmy in Infertility, Mitochondrial Disease, and Stem Cell Therapeutics References

Many of the assisted reproductive techniques associated with maternal aging, disease states, or implantation failure aim to correct poor developmental capacity. These techniques are highly invasive and require the exchange of nuclear or cytoplasmic material from a donor oocyte to compensate for deficiencies inherent in the aVected individual. These techniques are based on the assumption that the cytoplasm of the donor oocyte can eVectively substitute the necessary component(s) to enable development to proceed. Several studies have attempted to inject cytoplasm from ‘‘normal’’ (young) donors, into aged eggs, again assuming that beneficial components of the cytoplasm are transferred to restore developmental capacity. These invasive assisted reproduction technology (ART) procedures aim to eliminate chromosomal abnormalities, improve the quality of oocytes deficient in some important cytoplasmic factors necessary for maturation and/or subsequent development, and eliminate maternally inherited diseases (particularly mitochondrial myopathies). However, in order to develop such ART, understanding the processes involving mitochondrial DNA replication and transcription is imperative, as asynchrony between mitochondrial and nuclear genomes may cause problems in mitochondrial function, localization, and biogenesis. ß 2007, Elsevier Inc. Current Topics in Developmental Biology, Vol. 77 Copyright 2007, Elsevier Inc. All rights reserved.

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I. Why Are Invasive Assisted Reproductive Technology Procedures Important? Cytoplasmic transfer was initially developed to treat infertility patients exhibiting persistent poor embryonic development and recurrent implantation failure after in vitro fertilization (IVF). The technique was based on the assumption that the ooplasm of eggs, particularly from older women, could be rescued by the introduction of cytoplasm from eggs of younger donors. The procedure involved microinjection of 5–15% of the ooplasm from a presumptively young fertile donor oocyte into a putative defective recipient oocyte (Cohen et al., 1998). While the technique resulted in successful births, the transfer of small amounts of donor cytoplasm was accompanied by the introduction of proteins, mRNAs, small molecules, and organelles, particularly mitochondria, from the donor into the recipient oocyte (Van Blerkom et al., 1998). A number of questions arise regarding the outcomes of such techniques, particularly since these procedures have become more invasive in order to overcome (or prevent inheritance of) a range of diseases. Indeed, the normal pattern of presumably maternal (homoplasmic) inheritance of mitochondria, and any existing cross talk between the nuclear and mitochondrial genomes, may be compromised with unknown effects on the resulting child. Furthermore, there is evidence that infants resulting from ‘‘ooplasm transfer’’ exhibit heteroplasmy. Mitochondrial dysfunction is suggested as a prime cause of low oocyte and embryo quality and hence the poor eYciency of human ART. Not surprisingly, attention has been drawn to possible eVects of current invasive ARTs on mitochondrial function and inheritance. The more invasive the ART procedure, the more opportunities there are to disrupt the normal pattern of mitochondrial inheritance. If the mechanisms for maintaining uniparental mitochondrial DNA (mtDNA) inheritance fail following these techniques, oVspring may harbor various forms of mtDNA heteroplasmy. It is possible that wild‐type mtDNA can coexist with donor mtDNA, either mtDNA can be preferentially replicated or two wild‐type mtDNAs may persist in the reconstructed oocyte. It is apparent that mtDNA introduced into the oocyte can be transmitted following the use of these invasive techniques. To date, over 30 ooplasmic transplantation cases have been completed at the Institute of Reproductive Medicine and Science at Saint Barnabas. In these cases, previous IVF attempts on these patients’ embryos had failed because of poor embryo development; thus, cytoplasmic transfer was performed with 27 couples’ consent. The resulting 30 cases had normal fertilization and pregnancy rates with the following outcomes: 1 miscarriage, 11 singletons, 1 twin, and 1 quadruplet birth (Barritt et al., 2000; Brenner et al., 2000). The first trimester spontaneous miscarriage was diagnosed as 45, XO karyotype

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(Barritt et al., 2001a,b), which is the most common aneuploidy associated with intracytoplasmic sperm injection (ICSI). Additionally, the twin pregnancy resulted in a female diagnosed as normal 46, XX with a chromosomally abnormal (45, XO) twin, who was also diagnosed with a form of autism called pervasive disorder. While the incidence rate of chromosomal abnormalities (1/17 or 5.9%) was in the upper range of that normally reported in the New York region (1–6%), an increased aneuploidy rate is commonly observed following ICSI and/or with advanced maternal age. Obviously the sample size was too small to draw any concrete conclusions. DNA analysis indicated the presence of donor and recipient mitochondria in embryos, amniocytes, fetal tissues, and blood of oVspring following cytoplasmic transfer (Barritt et al., 2000, 2001a,b; Brenner et al., 2000). Donor mtDNA was detected in samples with the following frequencies: embryos (6/13), amniocytes (1/4), and fetal cord blood (2/4). mtDNA fingerprinting analysis performed on blood samples from two children following ooplasmic transfer detected mitochondrial polymorphisms in which both alleles were present in the hypervariable region of the mitochondrial genome. Nickerson et al. (1997) had previously estimated that, in order for the allele to be detected on the DNA chromatograph, the allele must be present in at least 30% of the sample, thereby being highly represented in blood samples from the oVspring. Since the mitochondrial fingerprinting technique only qualitatively evaluated the presence of mtDNA heteroplasmy in these children, sensitive quantitative molecular beacon analysis was also employed to document these changes accurately (Barritt et al., 2000; Brenner et al., 2000). Molecular beacons were designed to include an identified single nucleotide variation between donor and recipient mtDNA, each labeled with a diVerent fluorophore. The percentages of donor mtDNA in the cytoplasmic transfer samples are given in Table I (Brenner et al., 2004). With the exception of

Table I Percentage Donor Mitochondria in Fetal Tissues and Infant Blood Following Cytoplasmic Transfer (Brenner et al., 2004)a Sample Type Amniocytes Placenta Fetal cord blood 9‐m.o. blood 12‐m.o. blood 21‐m.o. blood

Case 1

Case 2

Case 3

N/A 28 4 32 N/A 36

N/A 81 28 N/A 43 N/A

63 58 N/A 70 N/A N/A

a Each value represents the percentage of donor mitochondria quantified by the mean of three to five experiments with eight replicates for each sample. m.o., month old; N/A, not analyzed.

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fetal cord blood in Case 1, the proportion of donor mtDNA far exceeded 10–15%, which would have been equivalent to the proportion of ooplasm that had initially been transferred. The placenta from Case 2 contained a surprisingly high proportion (81%) of donor mitochondria. Significant variability in donor mtDNA between tissues was also found in Case 1 where donor mtDNA constituted 28.6% in the placenta, 4.0% in fetal cord blood, and 31.8% and 36.1% in the 9‐ and 24‐month old, respectively. Surprisingly, the third infant’s blood preferentially replicated the donor population’s contribution (Brenner et al., 2004). It is still unknown whether injection of donor cytoplasm can improve the quality of the resulting oocyte/embryo, and in the long term the child, despite the birth of a number of children. Notably, children resulting from IVF of these reconstructed oocytes carried mitochondria from both the donor and recipient in varying proportions in diVerent tissues. However, as this procedure is no longer oVered in infertility clinics (while other techniques are still being considered), these children require further monitoring of mitochondrial heteroplasmy levels, yet these children are not being monitored. This is unfortunate since cytoplasmic transfer was an experimental infertility treatment, and therefore, it is impossible to determine its safety and eYcacy, when there are only a few individuals that were created in this manner.

II. Cytoplasmic Transfer: Animal Models That May Assist Human Infertility Therapeutics There have been various animal models that have involved the direct transfer of cytoplasm (in the form of cytoplasts) between diVerent developmental stages and across separate strains and subspecies (Levron et al., 1996; Meirelles and Smith, 1997, 1998). While a complete evaluation of the technique was not a primary goal of these experiments, their results indicate that cytoplasmic transfer has no detrimental eVect on mouse development. Normal, healthy mice have been eYciently produced from such cytoplast transfer experiments and maintained over 15 generations (Meirelles and Smith, 1997; Takeda et al., 2000). Preliminary work has demonstrated that the transfer of cytoplasts between metaphase II (MII) mouse eggs and zygotes has led to preimplantation development rates that were identical to those of control embryos (Levron et al., 1996; Meirelles and Smith, 1998). However, these experimental results have not clearly demonstrated that cytoplasmic transfer has any beneficial eVects on defective oocyte or embryos. At the present time, the only models that will demonstrate eYcacy and safety for novel infertility therapeutics is the non‐human primate.

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III. The Mitochondrial Genome and Mitochondrial Biogenesis It is important to consider the critical role that mitochondria play in order to understand the potential eVects of heteroplasmy on cell function and oVspring health. It is generally accepted, and believed essential for human evolution, that mtDNA inheritance is strictly maternal. Although the sperm carries paternal mitochondria into the oocyte at fertilization, the mitochondria are normally eliminated in early embryonic development by highly specific proteolysis set in place by ubiquitin tagging of the surface protein prohibitin during spermiogenesis (reviewed by Cummins, 2000). Mitochondria play a key role in the physiology of eukaryotic cells, including the mammalian oocyte, during preimplantation embryo development, and in nuclear transplantation and stem cell‐based therapeutics (Bavister et al., 2005; Cummins, 2001; Hiendleder and Wolf, 2003). This is in part due to the fundamental contribution of mitochondria to energy metabolism, as well as calcium homeostasis, signal transduction, and apoptosis. Mitochondria also play a role in metabolic pathways such as those involved in the biosynthesis of heme, lipids, amino acids, and nucleotides. These mitochondrial functions are therefore likely to be critical determinants of early embryonic development at various levels, including spindle organization, chromosomal segregation, cell cycle regulation, and morphodynamic processes such as compaction, cavitation, and blastocyst hatching (Van Blerkom, 2004). Preexisting oocyte components are critical during the interval between fertilization and embryonic genome activation (EGA) when the transcriptional activity of the embryonic genome becomes fully functional. During this period of development, the embryo is supported by maternal mRNAs, proteins, and organelles stored in the ooplasm. Indeed, embryonic transcription and the degradation of maternal mRNA are gradual processes (Schultz, 2005). When the embryo reaches the 8‐ to 16‐cell stage, EGA occurs, marking the major activation of the embryonic transcription with a sharp increase in the level of mRNA at the blastocyst stage (Schultz, 2005). Throughout the preimplantation period, the gene expression pattern is not constant, but varies according to the gene considered. Despite the central contribution of mitochondria to cell function and oocyte/embryo development, our understanding of mitochondrial biogenesis is rudimentary. Thus, it is important to elucidate the mechanism(s) that coordinate mitochondrial replication and transcription (Clayton, 2000). A feature of eukaryotic cells is that they contain both nuclear and mitochondrial genomes, sequestered into distinct subcellular compartments. The mitochondrial compartment is composed of an approximately 16.5‐kb double‐stranded circular DNA genome (mtDNA), as well as enzymes required for its transcription and replication, and the protein synthesis machinery

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necessary for the translation of mitochondrial mRNAs (reviewed by Garesse and Vallejo, 2001). These mRNAs account for the total protein‐coding capacity of mtDNA, encoding subunits of respiratory complexes I, III, IV, and V, and RNAs required for translation (2 ribosomal, 22 transfer RNAs). Consequently, nuclear‐encoded genes contribute significantly to mitochondrial metabolic systems and molecular architecture (Garesse and Vallejo, 2001; Table II). Proper expression, and/or replication of these genes, is dependent on nuclear factors interacting with the control region of the mitochondrial genome. The control region is approximately 1.1 kb in length and within it lies the hypervariable displacement loop (D‐loop), several conserved sequence blocks (CSB) I–III, and the heavy strand origin of replication (OH; Shadel and Clayton, 1997). Because of the limited coding capacity of the mtDNA, mitochondrial function is dependent on the exchange of information between the cytoplasm and the nucleus. The nuclear contribution to mitochondria includes catalytic and auxiliary proteins, gene‐encoding proteins, import and assembly factors, and proteins required for replication and expression of the mitochondrial genome (such as nucleic acid polymerases, RNA processing enzymes, and transcription and replication factors). Indeed, more that 200 genes required for mitochondrial function reside in the nuclear genome of the cell (Cummins, 2002). Thus, the program regulating mitochondrial biogenesis involves the coordinate actions of nuclear and mitochondrial genes. The active transcription of the mitochondrial genome starts at diVerent developmental stages depending on the species. In mice, mtDNA transcription occurs in the late 2‐cell stage, whereas it occurs in the 4‐ to 8‐cell stage in

Table II

Proteins Involved in Mitochondrial Replication, Repair, and Transcriptiona

Protein

Size (kDa)

Role

POLG Catalytic subunit

140

Accessory subunit Twinkle mtSSB

55 77 15

Polymerase and 30 –50 exonuclease activity during mt replication Enhances processitivity of catalytic subunit Unwinding of mtDNA during mt replication Prevents reannealing of single‐stranded mtDNA during mt replication Produces RNA copy number of mtDNA sequence Unwinding of mtDNA for mt transcription Involved in mt transcription Involved in mt transcription

PolRmt TFAM TFB1 TFB2

150 24 39 45

mt, mitochondrial; POLG, DNA polymerase ; Twinkle, helicase; mtSSB, mt single‐ stranded binding protein; PolRmt, mtRNA polymerase; TFAM, mt transcription factor A; TFB1 and TFB2, mt transcription factors B1 and B2, respectively. a

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humans and in the 8‐ to 16‐cell stage in cattle (May‐Panloup et al., 2005b; Thundathil et al., 2005). The molecular mechanisms responsible for this transcriptional activation of mtDNA during early embryogenesis are not well understood (Thundathil et al., 2005). Moreover, as mtDNA transcription and replication, and therefore cellular ATP generating capacity, rely heavily on proteins encoded by the nuclear genome, mutations in these transcription factors may result in mtDNA depletion syndromes (reviewed by Suomalainen and Kaukonen, 2001), which can be transmitted in a Mendelian fashion. The localization of these factors, in association with mitochondria, and their exact roles in the oocyte and embryo remain to be determined.

A. Mitochondrial Replication It is generally accepted that several nuclear factors must be transported into the mitochondria for replication. The key mtDNA replication factors include, but are not limited to, polymerase (POLG), Twinkle DNA helicase, and mitochondrial single‐stranded DNA‐binding protein (mtSSB; refer to Table I). Together these proteins form a replisome that interacts with mtDNA. POLG is the recognized polymerase responsible for mammalian mtDNA replication and repair. The overall structure of POLG can be divided into two subunits: the catalytic and the accessory subunits. The larger catalytic subunit contains DNA polymerase and exonuclease domains that are conserved across several species (Graziewicz et al., 2006). The smaller accessory subunit enhances the processing capacity and realignment of the catalytic subunit for eYcient mtDNA replication (Longley et al., 2001). The association between both subunits became further apparent after Chan et al. (2005b) reported reduced polymerase activity in patients exhibiting a common A467T mutation within the human POLG gene. This mutation causes dissociation between the two subunits of POLG, and generates large‐ scale mtDNA deletions associated with a wide range of mitochondrial disorders. To date, numerous other studies have investigated the role of POLG in mtDNA maintenance because of mtDNA’s susceptibility to damage from oxidants produced by the electron transport chain. While most of these studies are not directly linked to this chapter, it is important to note that POLG exonuclease activity is an irreplaceable component of mtDNA replication, and without it mitochondrial myopathies may develop. Twinkle, an mtDNA helicase, appears to be the rate‐limiting step in mtDNA replication initiation (Korhonen et al., 2003; Spelbrink et al., 2001; Tyynismaa et al., 2004). Mutations in Twinkle are associated with multiple mtDNA deletions and inhibition of Twinkle expression in cultured cells results in rapid mtDNA depletion, suggesting that Twinkle is required

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for mtDNA maintenance (Tyynismaa et al., 2004). In contrast, overexpression of Twinkle in transgenic mice results in increased mtDNA copy number (Tyynismaa et al., 2004). Furthermore, Twinkle helicase activity is stimulated by mtSSB (Korhonen et al., 2003). Together with POLG, these proteins form a minimal replisome (Fig. 1A) capable of synthesizing 16 kb lengths of DNA in cell‐free systems (Kaguni, 2004; Korhonen et al., 2004),

Figure 1 Schematic representation of mitochondrial transcription (B) and replication (A). (A) mtDNA is replicated by DNA polymerase (PolG), which in humans consists of two subunits: a 140‐kDa catalytic subunit possessing DNA polymerase and exonuclease proofreading activities, and a 55‐kDa subunit (PolG2) that promotes tight binding to DNA. The primer for initiation of mtDNA replication at OH is generated by processing the transcript starting at light strand promoter (LSP). PolG initiates H‐strand synthesis by extending the RNA primer. Mitochondrial single‐stranded DNA‐binding protein (mtSSB) enhances helix destabilization for Twinkle helicase and PolG to support mtDNA replication, recombination, and repair processes. Mitochondria transcription factor (TFAM, also known as mtTFA and mtTF‐1) is a High‐Mobility‐Group (HMG) box protein involved in mtDNA replication. TFAM binds conserved regulatory sequences within the D‐loop of mtDNA, recruiting other replication factors to the D‐loop. (B) mtDNA transcription complex containing mitochondrial RNA polymerase (PolRmt), TFAM, and mitochondrial transcription factor B (TFB1 and TFB2) are depicted. Transcription begins from two promoters, the LSP and the heavy strand promoter (HSP), within the control region (D‐Loop). TFAM binds to LSP, then recruits PolRmt and TFB1 and TFB2, with transcription initiated toward the origin of H‐strand replication (OH).

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since POLG alone is unable to support DNA synthesis. Twinkle also colocalizes with mitochondrial transcription factor A (TFAM) and the mtSSB, acting as a stabilizer of mitochondrial chromosome and nucleoid structure.

B. Mitochondrial Transcription Transcription of mtDNA takes place following interaction between the nuclear‐encoded regulatory proteins and regions within the mitochondrial D‐loop (Fig. 1B). Transcription requires mitochondrial RNA polymerase (PolRmt; Tiranti et al., 1997), TFAM (Fisher and Clayton, 1985, 1988), and at least one of the transcription factors TFB1M or TFB2M (Falkenberg et al., 2002). Several studies suggest that transcription is initiated bidirectionally at two promoters, the heavy strand promoter (HSP) and the light strand promoter (LSP), within the D‐loop regulatory region (Clayton, 2000; Shadel and Clayton, 1997). Light strand transcription commences from the LSP to the second conserved sequence block within the D‐loop region, generating a complementary RNA sequence (Xu and Clayton, 1995). Binding of the mtDNA‐specific polymerase to the resulting sequence enables replication of the heavy strand to initiate within the D‐Loop. The D‐loop is the longest noncoding region in vertebrate mtDNA and contains the H‐strand replication origin (OH; Fig. 1). Both promoters share a critical upstream enhancer that serves as the recognition site for TFAM, a high mobility group (HMG)‐ box protein that stimulates transcription through specific binding to upstream enhancers. TFAM can bend and unwind DNA and is linked to its ability to stimulate transcription on binding DNA (Fisher et al., 1992). Both TFB1 and TFB2 proteins work together with TFAM and PolRmt to direct initiation from HSP and LSP. Knockdown of the TFB2 isoform results in reduced mtDNA transcription and copy number, while knockdown of TFB1 has no eVect on mtDNA transcription or replication (Matsushima et al., 2005). Termination of mitochondrial transcription is associated with the specific initiation site for H‐strand transcription, via the mitochondrial transcription termination factor (mTERF; Martinez‐Azorin, 2005).

IV. Molecular Control of Mitochondrial Function in Preimplantation Embryos It is essential to understand the mechanisms controlling mitochondrial activity during early embryogenesis to determine their role in developmental outcome. Our laboratory and others have begun to study the molecular

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control of mitochondrial transcription and mtDNA replication in mouse, bovine, and primate preimplantation development. The expression patterns of two mitochondrial genes and several nuclear genes that encode mitochondrial transcription and replication factors have been investigated in mouse preimplantation development. A significant increase in the amount of mRNA for nuclear genes encoding mtDNA transcription factors, TFAM, TFB1, Polg1, Polg2, NRF‐1, RNase mitochondrial RNA processing (MRP) were observed in eight‐cell stage mouse embryos (Thundathil et al., 2005). Although a similar increase in the mRNA levels of nuclear genes encoding mtDNA replication factors was observed in morula and blastocyst stage embryos, the mtDNA copy number remains stable during preimplantation stages, suggesting that nuclear‐encoded mitochondrial transcription factors are involved in the regulation of mtDNA transcription during early development. Unlike findings reported in mouse embryos, the mtDNA content during early bovine embryogenesis was not constant (May‐Panloup et al., 2005b). They found a 60% decrease in mtDNA content between the two‐ and four/ eight‐cell stages. COX1 mRNA was constant until the morula stage after which it increased dramatically. TFAM mRNA was undetectable in oocytes and remained so until the 8/16‐cell stage; it began to appear only at the morula stage, suggesting de novo synthesis. In contrast, nuclear respiratory factor (NRF)‐1 mRNA was detectable in oocytes and the quantity remained constant until the morula stage. The results reveal a reduction of mtDNA content in early bovine embryos and a de novo upregulation of TFAM expression associated with mitochondrial biogenesis activation. High levels of NRF‐1 mRNA from the oocyte stage onward argue for the essential function of these factors during the first steps of bovine embryogenesis (May‐Panloup et al., 2005b). Recent knockout studies have demonstrated the importance of nuclear‐ encoded mitochondrial transcription and replication factors during embryo development. TFAM heterozygous knockout mice exhibit reduced mtDNA copy number and respiratory chain deficiency. Homozygous knockout embryos exhibit severe mtDNA depletion, with abolished oxidative phosphorylation (OXPHOS). Mutant embryos proceed through implantation and gastrulation but die prior to embryonic day (E) 10.5 (Larsson et al., 1998). TFAM levels correlate with increased mtDNA in ragged red muscle fibers and decreased mtDNA levels in mtDNA‐depleted cells (Larsson et al., 1994; Poulton et al., 1994). Hance et al. (2005) reported embryonic lethality in POLG knockouts. POLG deficiency in mouse embryos caused early developmental arrest between E7.5 and E8.5, associated with severe mtDNA depletion. Heterozygous knockout mice had half the wild‐type levels of POLG transcripts with a slight reduction in mtDNA levels but developed normally. Interestingly, POLG transcripts in heterozygous knockout mice were increased in response to

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artificially elevated mtDNA copy number, suggesting a possible regulatory link between mtDNA maintenance and POLGA expression. These mice (mtDNA mutator mice) expressed an error‐prone version of POLG and exhibited a dramatic decrease in exonuclease activity, whereas POLG polymerase activity is preserved. Abolished exonuclease activity leads to a three‐ to fivefold increase in somatic mtDNA mutations that, in turn, cause a progressive respiratory chain deficiency and premature aging phenotypes. These results oVer the first direct genetic evidence that the collective eVect of a variety of somatic mtDNA mutations can cause aging. Multiple mtDNA deletions accumulate in the tissues of mice expressing a mutant form of Twinkle, resulting in progressive respiratory dysfunction and chronic late‐onset mitochondrial disease around 1 year of age (Tyynismaa et al., 2005). DNA‐binding mutants of mtSSB in Drosophila embryos are defective in stimulation of DNA synthesis by POLG. RNAi knockdown of mtSSB reduces expression to <5% of its normal level, resulting in growth defects and in the depletion of mtDNA. Moreover, overexpression of mtSSB restores cell growth rate and mtDNA copy number (Farr et al., 2004). NRF‐1 and NRF‐2 are key transcription factors that regulate the expression of TFB1M and TFB2M promoters thus are implicated in mitochondrial biogenesis. Embryos homozygous for NRF‐1 die between E3.5 and E6.5. While the majority of wild‐type and heterozygous NRF‐1(þ/) blastocysts develop further in vitro, NRF‐1(/) blastocysts arrest, despite having a normal morphological appearance (Huo and Scarpulla, 2001). In an earlier study, homozygous NRF‐1 mutant mice exhibited anemia due to a noncell autonomous defect in definitive erythropoiesis, also dying in utero (Chan et al., 1998). Homozygous NRF‐1 blastocysts also exhibited markedly reduced levels of mtDNA, consistent with a requirement for NRF‐1 in the maintenance of mtDNA and respiratory chain function during preimplantation embryo development (Huo and Scarpulla, 2001). Homologous recombination of NRF‐1 by a ‐galactosidase‐neomycin cassette, such that ‐galactosidase expression was under the control of the NRF‐1 promoter, established positive staining in growing oocytes within the ovary and in day 2.5 eight‐cell embryos and day 3.5 blastocysts. These results demonstrate that the NRF‐1 gene is expressed during oogenesis and early stages of preimplantation development (Huo and Scarpulla, 2001).

V. Mitochondrial Copy Number, Deletions, and Mutations While most mammalian cells contain between 103 and 104 copies of mtDNA, the mature mammalian oocyte contains approximately 105 copies (Cummins, 1998). Within each mitochondrial organelle, there are generally 2–10 copies

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of mtDNA. It has been postulated that a massive increase of mitochondria occurs in the oocyte during early follicular development. In the human, premigratory primordial germ cells contain approximately 10 mitochondria per cell (Jansen and De Boer, 1998), while later in folliculogenesis, in the primary oocyte, mtDNA copy number ranges from 10,000 to 700,000 (Almeida‐Santos et al., 2006). It has historically been assumed that all of the mitochondria in the mature MII oocyte arise from the clonal expansion of an extremely small number of mitochondria during oogenesis (Van Blerkom, 2004). While the number of mitochondrial organelles increases, it is believed that the mtDNA copy number remains constant, thereby producing more mitochondria within which there are fewer copies of mtDNA. Due to the increase in mitochondria and the conserved number of mtDNA copies, the relative amount of mtDNA copy numbers may be extremely important in oogenesis and preimplantation embryo development. Oocytes with higher copy numbers have been shown to have higher fertilization rates (Spikings et al., 2006) and ovarian insuYciency has been associated with low mtDNA content (May‐Panloup et al., 2005a). It has also been shown that there is a significantly lower mtDNA copy number in arrested two‐cell embryos compared with developing embryos (Almeida‐Santos et al., 2006). There appears to be a minimum number of mtDNA molecules required for embryo development beyond the mature unfertilized oocyte; approximately 100,000 copies in the mouse and the pig (El Shourbagy et al., 2006; Piko and Taylor, 1987). Over 100 mitochondrial mutation‐associated diseases have been identified and are associated with the accumulation of mtDNA deletions found in brain, cardiac and skeletal muscle, liver, oocytes, and embryos. In order for mtDNA mutations to actually lead to problems, the mutant mtDNA copy number must reach a critical threshold level, or percentage of the overall mtDNA volume, thus causing a reduction of OXPHOS and subsequent ATP production. It is postulated that mouse preimplantation embryos are dependent on energy produced by oocyte‐inherited mitochondria due to the lack of an mtDNA replication event occurring before the blastocyst stage (Piko and Taylor, 1987). Contrary to the dogma, it has been suggested that mitochondrial replication may also occur during a very short period from the one‐ to two‐cell stage in the mouse embryo after fertilization (McConnell and Petrie, 2004). Additionally, data have shown that oocyte ATP content increases significantly during both bovine and porcine oocyte in vitro maturation (Brevini et al., 2005; Stojkovic et al., 2001). Further data also suggest the possibility that mtDNA replication occurs in the monkey during oocyte maturation from the germinal vesicle stage (GV) to MII of meiosis (Gibson et al., 2005). In the human, large scale mtDNA deletions have been shown to be responsible for specific neuromuscular diseases such as Kearns–Sayre syndrome

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(KSS) and chronic progressive external ophthalmoplegia (CPEO) syndrome. Both of these mtDNA diseases are characterized by a 4977‐bp deletion, now referred to as the ‘‘common’’ deletion, as this specific mutation has been shown to be present in a myriad of tissues. The presence of the ‘‘common’’ deletion in human oocytes and embryos from IVF clinics has been well established. Barritt et al. (1999) found that 34% of oocytes and 21% of embryos from IVF clinics contained the human ‘‘common’’ deletion and that half of oocytes and one‐third of embryos harbored single mtDNA rearrangements. Certain laboratories have associated an increase in the ‘‘common’’ deletion with an increase in age (Chan et al., 2005a; Keefe et al., 1995), while others were unable to detect a correlation (Barritt et al., 1999; Brenner et al., 1998; Chen et al., 1995; Hsieh et al., 2002). It is also well established that in slow and nondividing tissues, such as muscle and brain, mtDNA rearrangements accumulate with age (reviewed by Barritt et al., 1999). In the Rhesus macaque, a ‘‘common’’ mutation, similar to that found in humans, accumulates with age in skeletal muscle and contributes to age‐related fiber loss (Gokey et al., 2004; Schwarze et al., 1995). In a subsequent study, Gibson et al. (2005) analyzed gonadotrophin‐stimulated, mature oocytes, IVF embryos, and unstimulated, immature oocytes for the Rhesus mitochondrial ‘‘common’’ deletion. They found that 77.3% of gonadotrophin‐stimulated MII oocytes and 67.7% embryos harbored the mutation compared to 21.3% of immature GV oocytes from necropsied ovaries. The specific mechanisms responsible for this increase remain unknown. Significantly, these studies suggest (1) that there is an mtDNA replication event that may occur during oocyte recruitment or during the resumption of meiosis and (2) that the reason for the high frequency of mtDNA defects in oocytes may be from gonadotrophin stimulation. In humans and nonhuman primates, the administration of high doses of exogenous follicle stimulating hormone (FSH) dramatically increases the number of follicles recruited as compared to a natural cycle, and causes numerous defective oocytes, destined for atresia, to be collected with normal oocytes. Also, the administration of high doses of human chorionic gonadotrophin (hCG) may have additional eVects on oocyte maturation and the resumption of meiosis. Furthermore, repeated ovarian stimulations in the mouse have been shown to cause multiple mtDNA rearrangements, oxidative damage, and declined ovarian competence (Chao et al., 2005). Results in the mouse and both human and nonhuman primates suggest that the administration of gonadotrophin stimulation may actually cause mtDNA defects in oocytes. Further experiments are in progress in our laboratory to determine how many mitochondrial replication events occur during oogenesis and embryo development in the nonhuman primate and whether hormonal stimulation increases the copy number of mtDNA deletions.

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VI. Oocyte and Embryo Mitochondrial Localization Patterns The distribution of mitochondrial networks throughout the cell varies with diVerent stages of the cell cycle (Margineantu et al., 2002) and may play an important regulatory role in nuclear–mitochondrial signaling. Immature and mature oocytes also display distinct patterns of mitochondrial distribution (Nishi et al., 2003; Stojkovic et al., 2001). Staining of mitochondria has shown that during the cleavage stages of embryo development they cluster around the nucleus of blastomeres (Bavister and Squirrell, 2000). Mitochondria are asymmetrically localized in mouse oocytes and become homogeneously distributed during fertilization (Calarco, 1995; Muggleton‐ Harris and Brown, 1988; Van Blerkom and Runner, 1984). Likewise, rat oocyte mitochondria exhibit a perinuclear aggregation, which subsequently disperses (Zernicka‐Goetz and Maro, 1993). Mitochondria in hamster embryos redistribute from a homogeneous profile in the oocyte and early pronucleate stage embryo to a distinct perinuclear organization in the late pronucleate stage and at the two‐ and four‐cell stages (Barnett et al., 1996). In contrast, mitochondria exhibit a range of organizations in the bovine oocyte from a uniform distribution to restricted localization around the cortex. Regardless of oocyte organization, following fertilization, blastomeres exhibit an organelle‐free zone around the cortex (Van Blerkom et al., 1990). During porcine oocyte maturation, the majority of mitochondria are initially localized cortically (Sun et al., 2001) and subsequently show perinuclear accumulation in both the oocyte and early embryo (Hyttel and Niemann, 1990; Sun et al., 2001). A pronuclear accumulation of mitochondria has also been described in fertilized monkey (Squirrell et al., 2003) and human oocytes (Noto et al., 1993; Van Blerkom et al., 2000). Significantly, several studies indicate that mitochondrial organization is associated with developmental competence in rodents (Barnett et al., 1997; Lane and Bavister, 1998; Muggleton‐Harris and Brown, 1988; Squirrell et al., 2001), domestic species (Hyttel and Niemann, 1990; Van Blerkom et al., 1990), and monkeys (Squirrell et al., 2003). Mitochondria become dispersed from the nuclei under culture conditions that disrupt development in the hamster (Barnett et al., 1997; Lane and Bavister, 1998; Ludwig et al., 2001; Squirrell et al., 2001). Developmentally competent mouse embryos exhibit a homogeneous distribution of mitochondria in their cytoplasm, while impaired embryos show a more perinuclear organization (Muggleton‐Harris and Brown, 1988). Bovine oocytes with the highest developmental potential display a homogenous mitochondrial distribution pattern, while those with lower developmental potential exhibit vacuoles surrounded by mitochondria (Stojkovic et al., 2001). Similarly, a disruption of mitochondria organization that clumps around empty vesicles has been described for developmentally retarded pig embryos (Luoh and Wu, 1996).

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The mechanism by which the mitochondria alter their localization in early mammalian embryos has not yet been entirely elucidated. It is likely that the cytoskeleton, particularly microtubules, participate in the translocation of mitochondria in these embryos. The importance of the cytoskeleton to the movement of subcellular components has been shown in the embryos of a number of species (Bement et al., 1992). At present, no study has examined whether mitochondrial localization follows the pattern of microtubule distribution. Reorganizations in mitochondria may be providing localized energy and/or metabolites (Barnett et al., 1996; Van Blerkom et al., 2000) and may be related to changes in ionic balance (Lane and Bavister, 1998; Squirrell et al., 2001). Close association is required for nuclear–mitochondrial communication, particularly for mitochondrial replication, or for providing energy required for cell division processes such as spindle formation. Perinuclear clustering of mitochondria during cell division allows more equal transmission of mtDNA molecules to daughter blastomeres. Mitochondria are also known to be involved in chromosome organization and movement. Although elucidating the direct connection between cytoplasmic organization (including the localization of mitochondria) and subsequent embryo development is diYcult, understanding this relationship is important for furthering our understanding of embryo physiology and how it is aVected by culture conditions. It has been suggested that preferential replication of mtDNA closer to the nucleus occurs (Davis and Clayton, 1996) potentially due to faster diVusion of nuclear‐encoded regulatory factors. Therefore, the position of mitochondria in the early embryo may play a role in determining which mtDNA molecules are selected for amplification. This information is of paramount importance for determining the possible detrimental consequences of physically invasive reproductive technologies.

VII. Implications of Mitochondrial Heteroplasmy in Infertility, Mitochondrial Disease, and Stem Cell Therapeutics It has been proposed that invasive ART be used to prevent the transmission of mitochondrial disease in patients who wish to transfer their own genetic material into an enucleated donor oocyte (Cummins et al., 1997; Trounson and Pera, 2001). However, these techniques will be associated with similar heteroplasmic mitochondrial inheritance issues. The transferred germinal vesicle or pronuclear stage nucleus is accompanied by small amounts of cytoplasm, and as such the nuclear material remains surrounded by mitochondria which will also be transferred into the donor ooplasm. It is likely that these mitochondria remain close to the reconstructed oocyte nucleus,

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but are then dispersed throughout the cytoplasm as maturation proceeds (Fulka et al., 2005). Since the children resulting from cytoplasmic transfer carried mitochondria from both the donor and recipient in varying proportions in diVerent tissues, it is not clear what would happen with the proposed invasive ART to prevent the transmission of mitochondrial disease in the children of these patients. Cytoplasmic and nuclear transfer techniques provide an opportunity where karyoplast mitochondria may be present in all cell types of the resulting oVspring. Therefore, any donor or diseased mitochondria transferred may continue to be transmitted and replicated. Since mitochondrial disease is based on mutational load, these children may eventually show symptoms of these mitochondrial myopathies later in life. In studies on cloned sheep, cattle (St. John et al., 2004), and monkeys (St. John and Schatten, 2004), mitochondrial heteroplasmy may also result from nuclear transfer, that is, the fusion of somatic cells with enucleated oocytes (somatic cell nuclear transfer, SCNT). All eVorts to produce cloned (identical) monkeys by SCNT for biomedical research have so far failed, but any success in this approach will have to contend with the problem of mitochondrial heteroplasmy. It may be that techniques such as cytoplasmic transfer, nuclear transfer, and therapeutic cloning involving mitochondrial heteroplasmy may perturb the normal interchanges between nuclear and mitochondrial genomes required for proper mtDNA replication and transcription. This could account for the low success of these technologies. Nevertheless, if any of these invasive ARTs for therapeutics are to ever gain acceptance for treatment of human infertility or embryonic stem cell therapeutics, it seems that their safety as well as their eYcacy must first be confirmed in monkey models. Understanding the processes involving mtDNA replication and transcription is imperative, as asynchrony between mitochondrial and nuclear genomes may be the source of problems in mitochondrial function, localization, and biogenesis.

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Korhonen, J. A., Pham, X. H., Pellegrini, M., and Falkenberg, M. (2004). Reconstitution of a minimal mtDNA replisome in vitro. EMBO J. 23(12), 2423–2429. Lane, M., and Bavister, B. D. (1998). Calcium homeostasis in early hamster preimplantation embryos. Biol. Reprod. 59, 1000–1007. Larsson, N. G., Oldfors, A., Holme, E., and Clayton, D. A. (1994). Low levels of mitochondrial transcription factor A in mitochondrial DNA depletion. Biochem. Biophys. Res. Commun. 200, 1374–1381. Larsson, N. G., Wang, J., Wilhelmsson, H., Oldfors, A., Rustin, P., Lewandoski, M., Barsh, G. S., and Clayton, D. A. (1998). Mitochondrial transcription factor A is necessary for mtDNA maintenance and embryogenesis in mice. Nat. Genet. 18(3), 231–236. Levron, J., Willadsen, S., Bertoli, M., and Cohen, J. (1996). The development of mouse zygotes after fusion with synchronous and asynchronous cytoplasm. Hum. Reprod. 11, 1287–1292. Longley, M. J., Nguyen, D., Kunkel, T. A., and Copeland, W. C. (2001). The fidelity of human DNA polymerase gamma with and without exonucleolytic proofreading and the p55 accessory subunit. J. Biol. Chem. 276, 38555–38562. Ludwig, T. E., Squirrell, J. M., Palmenberg, A. C., and Bavister, B. D. (2001). Relationship between development, metabolism, and mitochondrial organization in 2‐cell hamster embryos in the presence of low levels of phosphate. Biol. Reprod. 65, 1648–1654. Luoh, Y., and Wu, M. (1996). Mitochondrial distribution of swine owa and embryos analyzed by rhodamine 123 fluorescent assay. J. Chin. Soc. Anim. Sci. 25, 53–66. Margineantu, D. H., Gregory Cox, W., Sundell, L., Sherwood, S. W., Beechem, J. M., and Capaldi, R. A. (2002). Cell cycle dependent morphology changes and associated mitochondrial DNA redistribution in mitochondria of human cell lines. Mitochondrion 1, 425–435. Martinez‐Azorin, F. (2005). The mitochondrial ribomotor hypothesis. IUBMB Life 57, 27–30. Matsushima, Y., Adan, C., Garesse, R., and Kaguni, L. S. (2005). Drosophila mitochondrial transcription factor B1 modulates mitochondrial translation but not transcription or DNA copy number in Schneider cells. J. Biol. Chem. 280(17), 16815–16820. May‐Panloup, P., Chretien, M. F., Jacques, C., Vasseur, C., Malthiery, Y., and Reynier, P. (2005a). Low oocyte mitochondrial DNA content in ovarian insuYciency. Hum. Reprod. 20, 593–597. May‐Panloup, P., Vignon, X., Chretien, M. F., Heyman, Y., Tamassia, M., Malthiery, Y., and Reynier, P. (2005b). Increase of mitochondrial DNA content and transcripts in early bovine embryogenesis associated with upregulation of mtTFA and NRF1 transcription factors. Reprod. Biol. Endocrinol. 3, 65–72. McConnell, J. M., and Petrie, L. (2004). Mitochondrial DNA turnover occurs during preimplantation development and can be modulated by environmental factors. Reprod. Biomed. Online 9, 418–424. Meirelles, F., and Smith, L. (1997). Mitochondrial genotype segregation in a mouse heteroplasmic lineage produced by embryonic karyoplast transplantation. Genetics 145, 445–451. Meirelles, F., and Smith, L. (1998). Mitochondrial genotype segregation during preimplantation development in mouse heteroplasmic embryos. Genetics 148, 877–884. Muggleton‐Harris, A. L., and Brown, J. J. (1988). Cytoplasmic factors influence mitochondrial reorganization and resumption of cleavage during culture of early mouse embryos. Hum. Reprod. 8, 1020–1028. Nickerson, D. A., Tobe, V. O., and Taylor, S. L. (1997). PolyPhred: Automating the detection and genotyping of single nucleotide substitutions using fluorescence‐based resequencing. Nucleic Acids Res. 25, 2745–2751. Nishi, Y., Takeshita, T., Sato, K., and Araki, T. (2003). Change of the mitochondrial distribution in mouse ooplasm during in vitro maturation. J. Nippon Med. Sch. 70, 408–415. Noto, V., Campo, R., Roziers, P., Swinnen, K., Vercruyssen, M., and Gordts, S. (1993). Mitochondrial distribution after fast embryo freezing. Hum. Reprod. 8, 2115–2118.

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Nuclear Transfer: Preservation of a Nuclear Genome at the Expense of Its Associated mtDNA Genome(s) Emma J. Bowles,* Keith H. S. Campbell,{ and Justin C. St. John* *The Mitochondrial and Reproductive Genetics Group The Division of Medical Sciences, The Medical School The University of Birmingham Birmingham B15 2TT, United Kingdom { Animal Development and Biotechnology Group School of Biosciences, University of Nottingham Sutton Bonington, Loughborough Leics LE12 5RD, United Kingdom

I. II. III. IV. V.

Introduction Adaptations to NT Applications of NT Health Problems Associated with NT Possible Causes of NT Failure A. Cell Cycle B. Epigenetics C. Pluripotent and ‘‘Epigenetic Memory’’ Instability D. mtDNA Transmission Following NT E. Haplotype and Genetic Divergence F. mtDNA Copy Number G. mtDNA Transcription and Replication VI. Are All the Potential Reasons for Low Success Rates in NT Interlinked? VII. Conclusions References

Nuclear transfer technology has uses across theoretical and applied applications, but advances are restricted by continued poor success rates and health problems associated with live offspring. Development of reconstructed embryos is dependent upon numerous interlinking factors relating both to the donor cell and the recipient oocyte. For example, abnormalities in gene expression following somatic cell nuclear transfer (SCNT) have been linked with an inability of the oocyte cytoplasm to sufficiently epigenetically reprogram the nucleus. Furthermore, influences on the propagation of mitochondria and mitochondrial DNA (mtDNA) could be of great importance in determining the early developmental potential of NT embryos and contributing to their genetic identity. mtDNA encodes some of the subunits of the electron transfer chain, responsible for cellular ATP production. The remaining subunits and those factors required Current Topics in Developmental Biology, Vol. 77 Copyright 2007, Elsevier Inc. All rights reserved.

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for mtDNA replication, transcription and translation are encoded by the nucleus, necessitating precise intergenomic communication. Additionally, regulation of mtDNA copy number, via the processes of mtDNA transcription and replication, is essential for normal preimplantation embryo development and diVerentiation. Unimaternal transmission following natural fertilization usually results in the presence of a single identical population of mtDNA, homoplasmy. Heteroplasmy can result if mixed populations of mtDNA genomes co‐exist. Many abnormalities observed in NT embryos, fetuses, and oVspring may be caused by deficiencies in OXPHOS, perhaps resulting in part from heteroplasmic mtDNA populations. Additionally, incompatibilities between the somatic nucleus and the cytoplast may be exacerbated by increased genetic divergence between the two genomes. It is important to ensure that the nucleus is capable of suYciently regulating mtDNA, requiring a level of compatibility between the two genomes, which may be a function of evolutionary distance. We suggest that abnormal expression of factors such as TFAM and POLG in NT embryos will prematurely drive mtDNA replication, hence impacting on early development. ß 2007, Elsevier Inc.

I. Introduction Nuclear transfer (NT), often termed ‘‘cloning,’’ involves the transfer of a donor cell nucleus into a recipient enucleated oocyte (Figs. 1 and 2). Subsequent development of the reconstructed embryo in utero gives rise to oVspring genetically identical to the donor nucleus (a clone; Campbell et al., 1996a). Monozygotic twins are examples of naturally occurring clones as they are produced by blastomere separation or embryo splitting and result in two genetically identical oVspring. A variety of viable oVspring have been produced using NT. These include sheep (Wilmut et al., 1997), nonhuman primates (Meng et al., 1997), cattle (Cibelli et al., 1998), mice (Wakayama and Yanagimachi, 1999), goats (Baguisi et al., 1999), pigs (Polejaeva et al., 2000), cats (Shin et al., 2002), rabbits (Chesne et al., 2002), mules (Woods et al., 2003), rats (Zhou et al., 2003), horses (Galli et al., 2003), dogs (Lee et al., 2005), and ferrets (Li et al., 2006a). The donor cell can arise from (1) an embryonic cell (EC) derived from a preimplantation embryo, (2) a fetal cell such as an embryonic fibroblast, or (3) an adult cell.

II. Adaptations to NT Since its inception, a variety of NT techniques have been developed that result in distinct outcomes. For instance, single NT can be performed by injecting a cell’s nucleus (Lacham‐Kaplan et al., 2000; Trounson et al., 1998)

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MII oocyte

Enucleation Chromosomes

Oocyte/cell couplet

Figure 1 Nuclear transfer (NT). The process of NT involves the selection of a mature MII oocyte. This is held in place using a holding pipette while the nuclear material is removed with a needle. Prestaining the nucleus with Hoechst 33258 enables the chromosomal material to be easily identified to ensure its complete removal. Following enucleation, a donor cell is injected, usually into the perivitelline space, and the oocyte/cell couplet is fused. Subsequent activation and in vitro maturation will initiate development of the reconstructed embryo.

or a whole cell directly into the cytoplasm of an enucleated oocyte (Lee et al., 2003) rather than into the perivitelline space prior to fusion. In all cases, the reconstructed oocytes are then activated to mediate cellular division. A further modification of this technique, double NT, requires the donor cell to be fused to an enucleated oocyte and then transferred into an enucleated zygote (Polejaeva et al., 2000). Serial NT describes the transfer of a newly replicated metaphase nucleus from an early‐stage embryo into an enucleated recipient oocyte followed by subsequent transfer of the individual ‘‘pronuclei‐ like nuclei’’ into two enucleated recipient oocytes (Kwon and Kono, 1996). The intermittent period of time spent in the enucleated oocyte mediates reprograming of the donor cell before it is regulated by the cellular processes of the zygote. Alternatively, an oocyte can be stripped of its zona pellucida and dissected to eliminate its haploid nucleus and then fused to a donor cell, a technique referred to as handmade cloning (HMC; Peura et al., 1998; Trounson et al., 1998). Here, a single or multiple cytoplasts can be fused to the donor cell. This approach has been successfully utilized to increase inner cell mass (ICM) cell number at the blastocyst with the aim of increasing developmental potential (Tecirlioglu et al., 2005). Additionally, aggregation of multiple four‐cell mouse NT embryos has been performed in order to increase cell numbers at blastocyst (Boiani et al., 2003).

Figure 2 Gene targeting in somatic cell nuclear transfer (SCNT). Donor cells for SCNT are obtained from a live animal or fetus, plated and cultured in vitro. Homologous recombination technology allows specific genes to be targeted and knocked in or knocked out. Cells that have correctly incorporated the transgene can be cultured and selected for use in NT. These donor cells are injected into the perivitelline space of an oocyte usually from a diVerent animal. Oocytes are fused, activated, and allowed to divide in vitro until the blastocyst stage, when they are transferred into a recipient animal’s uterus. The resulting clone will express the altered version of the targeted gene.

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III. Applications of NT This ever‐improving technology has great potential in numerous areas of research. NT could act as a valuable tool in embryological studies (McGrath and Solter, 1983) and may oVer a way of determining the genetic and nongenetic aspects of complex disease (Schafer and Hawkins, 1998). It certainly provides great potential for understanding epigenetic disorders and aberrant patterns of mtDNA transmission when compared to more conventional forms of embryo production such as in vitro fertilization and coitus. Indeed, cloning can also be considered as a basic research tool to study biological phenomena such as cell diVerentiation, cell aging, and rejuvenation (Wakayama and Yanagimachi, 1999). Consequently, any developments that may improve oVspring viability in NT and other associated technologies are incredibly important. Somatic cells can also be subjected to genetic manipulation prior to their use as nuclear donors for NT (Fig. 2). For example, ovine fetal fibroblast cells have been transfected with a human coagulation factor IX genomic construct under the control of a mammary‐specific promoter and NT performed to produce live oVspring. The resulting expression of that gene in sheep milk has been used as a mechanism for commercially producing this factor for clinical purposes (Schnieke et al., 1997). The use of homologous recombination in nuclear donor cells followed by their use in NT has provided a method for the precise genetic modification of mammals other than the mouse and provides numerous opportunities in agricultural and medical research. This was first demonstrated in the sheep by creating a knockout of a single allele at the ‐1 (I) procollagen (COL1A1) locus (McCreath et al., 2000). The technique can also be used for gene knock‐in strategies to a suitable locus in order to produce high expression of biopharmaceutical proteins, such as alpha‐1‐antitrypsin (AAT) in cattle for the treatment of cystic fibrosis (Jang et al., 2006). In pigs, the ‐1,3‐galactosyltransferase gene has been successfully targeted and knocked out in cloned animals (Dai et al., 2002; Lai et al., 2002). ‐1,3‐ Galactosyltransferase is responsible for hyper acute rejection of transplants when pig organs are transferred to humans, as humans naturally possess antibodies against this protein (Sandrin et al., 1993). The production of pigs through NT that do not express this protein therefore provides a partial solution to porcine transplant rejection and increases the possibility of xenotransplantation of both organs and tissues (Prather et al., 2003). This ability to produce specific genetic modifications in species other than the mouse oVers a range of other opportunities. Thousands of transgenic mouse lines are already used for gene function and disease research, but NT technology allows such studies to be conducted in species more physiologically similar to the human. In addition, the ability to generate a series of genetically identical

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oVspring would therefore be of considerable value to the pharmaceutical industry for drug screening and functional studies as many variables arising from diVerent genetic individuals will be removed. Agricultural applications of cloning may include creating pigs that produce pork faster or more eYciently, or creating animals that are resistant to specific diseases (Prather et al., 2003). Through NT it would be possible to produce uniform stocks of superior genotypes in the animal breeding industry (Hiendleder et al., 1999). Similarly, the cattle industry is keen to preserve certain rare breeds to conserve genetic diversity. Those lineages of bulls which have good semen parameters and good reproductive histories are also often sought (Fitzpatrick et al., 2002; Holroyd et al., 2002). Again, NT oVers opportunities for these approaches to be realizable. The horse industry may also, in the long term, regard NT as an appropriate tool to preserve lineages of known sporting ability (Galli et al., 2003). NT can also assist in saving endangered species and maintaining endangered populations (Gomez et al., 2004; Lanza et al., 2000; Loi et al., 2001). This allows endangered species to be cloned using adult somatic cells from the species itself and oocytes from related species where they are in abundance. Success achieved to date includes the birth of African wildcat clones using domestic cat oocytes (Gomez et al., 2003, 2004) and cloning of the Banteng (Bos javanicus) using B. taurus or B. indicus oocytes (Sansinena et al., 2005). Recent interest in therapeutic cloning highlights the potential for using NT as a method of producing embryonic stem cells (ESCs). Here, the aim is to generate tailor‐made ESCs specific to the patient’s requirements. In this instance, a somatic cell from a patient would be used to produce an embryo by NT. ESCs would be derived from this embryo and used for cell therapies, once homologous recombination had been performed to correct the patient‐ specific gene defects prior to transplantation. Indeed, homologous recombination is currently more successful in ESCs than somatic cells (Templeton et al., 1997). This form of treatment is highly applicable to single‐gene disorders such as Parkinson’s disease, cardiomyopathy, sickle cell anemia, and Tay–Sachs disease (see Trounson, 2005 for review). ESCs can be directed in culture to diVerentiate into a vast array of cell types, including insulin‐ producing cells (Assady et al., 2001), cardiomyocytes (He et al., 2003), neuronal cells (Bibel et al., 2004; ReubinoV et al., 2001), and smooth muscle cells (Sinha et al., 2006), amongst others. The advantage of using such a sophisticated approach is that the resultant ESCs will be immune‐specific to the patient who will therefore not have to endure ongoing administration of immunosuppressives. Success has been achieved in isolating ESCs through NT in the mouse (Munsie et al., 2000; Wakayama et al., 2001). Additionally, significant steps forward were later reported where the Rag2 mutation was repaired in NT‐derived ESCs through homologous recombination (Rideout et al., 2002). These mice were subsequently successfully treated for severe

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immune deficiency, clearly demonstrating the feasibility of such an approach. However, so far, there are no published reports of hESCs derived from human NT embryos, although a blastocyst has been generated using such technology (Stojkovic et al., 2005). The potential to use cross‐species NT is perhaps even more vital considering the shortage of human oocytes available due to medical, legal, and ethical reasons (Holden, 2005). Bovine oocyte cytoplasm has been shown to support early development of somatic cell nuclei from cattle, sheep, pig, monkey, and rat with a number of embryos of each species reaching the blastocyst (Dominko et al., 1999). This suggests that mechanisms regulating early development may be conserved among mammalian species. Cross‐ species experiments have also demonstrated the rabbit oocyte to be capable of supporting the development of transplanted human nuclei (Chen et al., 2003). From the resulting embryos, ES cells were isolated that were capable of self‐renewal and diVerentiation into all three germ layers. These studies demonstrate the possibility of using nonhuman mammalian oocytes to sufficiently reprogram the human somatic cell nucleus and support development to blastocyst. Consequently, either excellent programs of human oocyte recruitment need to be founded or a cross‐species methodology needs to be fully developed.

IV. Health Problems Associated with NT Despite the increasing success of NT across a range of species, survival rates at all developmental stages are low and the health of many oVspring is poor. Successful development of reconstructed embryos varies between laboratories due to species and technical diVerences, but on average only 2–3% of reconstituted eggs develop into oVspring (Wilmut, 2002) with numbers as low as 0.1–1% in tadpoles (Gurdon and Colman, 1999). Major health problems associated with cloned animals include diabetes (Hill et al., 2000), pulmonary hypertension (Lanza et al., 2001), obesity (Wakayama and Yanagimachi, 2001), placental abnormalities (De Sousa et al., 2001; Hill et al., 2001; Wells et al., 1999), respiratory failure (Wakayama and Yanagimachi, 1999), and kidney, liver, and brain defects (Denning et al., 2001; McCreath et al., 2000). The successful development of embryos reconstructed by NT is dependent on a wide range of known factors, including oocyte quality, the reconstruction process, the culture conditions, and the cell cycle stages of both donor and recipient cells (see Campbell, 1999; Campbell et al., 1996b for review). In addition, many other factors are likely to be involved at all stages of development, including the source of donor genetic material, genetic imprinting, and the largely ignored influences of mtDNA.

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V. Possible Causes of NT Failure A. Cell Cycle Ovine studies in which live oVspring have been produced by NT from embryonic (Campbell et al., 1996a), fetal (Wells et al., 1997), and adult (Wilmut et al., 1997) donor tissues utilized cells which had been induced to enter quiescence prior to NT. This is commonly achieved through either serum starvation or by allowing the cells to reach confluency. Coordination of the cell cycle in the donor somatic cell with that of the recipient cytoplast is important to prevent DNA damage and/or uncoordinated DNA replication from taking place (Campbell, 1999; Campbell et al., 1996b). The stage of the cell cycle for the donor cell must also be compatible with the high levels of maturation‐promoting factor (MPF) found in a metaphase II (MII) oocyte, that is, G0, G1, or M (Campbell et al., 1993). When a cell is in quiescence the chromatin becomes more condensed, there is a reduction in transcription and translation, and mRNAs are actively degraded. The chromatin of quiescent cells may be more amenable to structural changes following NT and it has been suggested that nuclei in G0 are the best candidates for NT (Campbell et al., 1996b). Indeed, a significant eVect of G0 donor cells on the condensation of chromosomes and the formation of nuclei in the reconstituted embryos has been reported (Tao et al., 1999). B. Epigenetics ‘‘Epigenetic’’ changes in gene expression are those that occur in addition to alterations in genetic sequence (Weinhold, 2006) and include methylation, acetylation, phosphorylation, ubiquitylation, and sumolyation. These processes can result in changes in chromatin structure and hence the accessibility of DNA, thereby altering gene expression (Nguyen et al., 2001). An important control during development is genetic imprinting, a mechanism by which a proportion of genes is repressed on one of the chromosomes according to the parental origin of the gene, thought to be especially important for regulating fetal growth (Latham et al., 1994). An important regulator of ‘‘imprinting’’ is methylation of DNA at CpG dinucleotides. In general, DNA methylation is associated with transcriptional repression, although there are some exceptions to this rule (Zhang et al., 2005). It has been established that two genome‐wide methylation reprograming events occur during development: the first during germ cell development and the second during preimplantation development (reviewed in Reik et al., 2001). The first of these events results in highly methylated primordial germ cells rapidly losing their methylation and the subsequent resetting of allele‐specific imprinting (Tucker et al., 1996). Remethylation occurs in

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postimplantation stages in male germ cells (Davis et al., 2000; Ueda et al., 2000) and after birth in females during oocyte growth (Obata et al., 1998). By the time these cells have formed mature gametes of either sex, they are highly methylated again. The second round of methylation reprograming occurs between fertilization and the blastocyst stage. This time, methylation is asymmetric with a rapid loss of paternal‐specific methylation (Dean et al., 2003). De novo methylation is initiated to coincide with the first diVerentiation event and gives rise to tissue‐specific methylation patterns. The ICM becomes hypermethylated, while the trophectoderm is undermethylated (Dean et al., 2001; Santos et al., 2002). This pattern is maintained throughout development where the somatic tissues are highly methylated in contrast to the hypomethylated placenta (Dean et al., 2001). However, these changes in methylation that occur on fertilization and during preimplantation development are not conserved across all species, with sheep embryos exhibiting little demethylation prior to the blastocyst stage (Beaujean et al., 2004). It is evident that the mechanisms of imprinting and gene expression during preimplantation development are highly regulated events. As such, they provide an obstacle for somatic cell NT (SCNT) which involves the introduction of a highly methylated cell into an oocyte (Wade and Kikyo, 2002). This raises a number of questions. To what extent can the oocyte cytoplasm reprogram the somatic cell nucleus? Could incomplete reprograming contribute to the poor success rate of SCNT? How can donor cells be better reprogramed for use in SCNT? The status of genes in diVerentiated somatic cells must be ‘‘reprogramed to the zygotic state’’ and failure of cloned fetuses to reach term could result from incomplete reprograming of genes in the transferred somatic cell nuclei (Tamashiro et al., 2000). To this extent, higher cloning success rates were achieved when using nuclei from fetal, ovarian, and testicular cells than adult somatic cells, though success rates were still very low (Wakayama and Yanagimachi, 2001). This suggests that nuclear reprograming may be easier to achieve in less diVerentiated cell types, a hypothesis supported by the higher rates of development observed when embryonic blastomeres are used directly as nuclear donors (Wakayama and Yanagimachi, 2001). It has indeed been demonstrated that abnormal imprinting can severely compromise NT, often with lethal consequences (Lee et al., 2002). There is increasing evidence for abnormal methylation patterns and gene expression in cloned embryos (Kang et al., 2001; Li et al., 2005, 2006b; Ohgane et al., 2001; Yang et al., 2005), which may in part explain the abnormal symptoms observed. In one study where phenotypic and epigenetic characteristics of NT mice were compared with normal mice, a significant suppression of imprinted and nonimprinted placental genes was observed. This resulted in placental overgrowth where NT mice exhibited two‐ to threefold larger placentas than control mice (Ogura et al., 2002). Aberrant expression of

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genes essential for implantation and placental diVerentiation has been reported in cloned bovine blastocysts and linked with implantation failure (Hall et al., 2005). In order to investigate the extent of abnormalities in gene expression following NT, microarray analysis was utilized to detect and compare levels of over 10,000 mRNAs in tissues from mouse clones produced from both ESCs and cumulus cells, with control embryos (Humpherys et al., 2002). This study found approximately 4% genes, including some imprinted genes, in NT placentas to be abnormal. Interestingly, while most diVerences were common to both types of clones, other abnormalities were confined to clones made from a particular donor cell type, indicating cell‐specific eVects on epigenetic reprograming.

C. Pluripotent and ‘‘Epigenetic Memory’’ Instability The importance of epigenetics has been discussed in relation to cloning, with NT embryos exhibiting abnormal patterns of methylation. This leads us to question whether pluripotency may be compromised in SCNT embryos produced by insertion of a highly diVerentiated donor nucleus, with established methylation patterns, into an enucleated oocyte. Indeed, studies have demonstrated an incomplete reactivation of Oct4 in mouse SCNT embryos (Bortvin et al., 2003). Oct4 protein is required for embryonic development and maintenance of a pluripotent state in ESCs so in the absence of Oct4, embryos fail to produce viable blastocysts (Nichols et al., 1998). Studies in mouse show that when somatic cells, in which Oct4 is no longer expressed, are used in NT, the Oct4 gene is incompletely reactivated through epigenetic changes in 38% embryos analyzed (Bortvin et al., 2003) or exhibits aberrant expression in blastocysts (Boiani et al., 2002). Additionally, other markers of pluripotency such as Dppa and Pramel‐like genes were also incompletely reactivated in cloned embryos (Bortvin et al., 2003). Conversely, it has also been shown in studies on Xenopus clones that transcripts expressed by the donor somatic cells are prematurely expressed in tissues of NT embryos (Ng and Gurdon, 2005). The authors suggest that ‘‘epigenetic memory’’ is established in these somatic cells during diVerentiation, and are not completely reprogramed on insertion into the recipient oocyte. Additionally, de novo methylation is initiated to coincide with the first diVerentiation event in embryos (Dean et al., 2003). Could the bypassing of normal methylation events that occurs in SCNT therefore contribute to diVerentiation being initiated at the incorrect time and consequently would this impact on mitochondrial number, mitochondrial DNA (mtDNA) replication, and mtDNA copy number that would be disadvantageous to the cell, tissue, or oVspring (St. John et al., 2004)?

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D. mtDNA Transmission Following NT As donor cells for NT contain their own mtDNA population, there are implications for the unimaternal transmission of mtDNA that takes place in oVspring derived from natural fertilization (Birky, 1995, 2001; Giles et al., 1980). Consequently, using a somatic cell for NT could result in the oVspring exhibiting one of three potential patterns of mtDNA inheritance: (1) homoplasmic for donor somatic cell mtDNA, (2) homoplasmic for recipient oocyte mtDNA, and (3) heteroplasmic due to coexistence of donor and recipient mtDNA. Indeed, both homoplasmy for the recipient oocyte mtDNA (Evans et al., 1999) and heteroplasmy (Hiendleder et al., 1999; Meirelles et al., 2001; Steinborn et al., 1998a) have been reported in NT‐derived animals (Table I) with donor mtDNA transmission ranging from 0 to 59% (Takeda et al., 2003). For example, Dolly and nine other NT‐derived sheep were all homoplasmic for their respective recipient oocytes, although analysis was of only blood, skeletal muscle, placenta, and milk samples (Evans et al., 1999). This potential homoplasmic transmission may have resulted from the failure of the donor mitochondria to be taken up by the ooplasm following electrofusion (Evans et al., 1999). Alternatively, the donor mitochondria could have been actively destroyed by a mechanism similar to that which destroys sperm‐derived mitochondria in the ooplasm (Ankel‐Simons and Cummins, 1996; Sutovsky et al., 1999). A number of studies on cattle clones have detected heteroplasmic mtDNA populations where donor mtDNA was shown to persist and even proliferate in embryos and tissues from oVspring (Hiendleder et al., 1999; Steinborn et al., 1998a; Takeda et al., 2003). Interestingly, levels of donor mtDNA have been found to vary greatly in diVerent tissues of the same animal, showing that it has been segregated in a nonuniform manner during development (Takeda et al., 2003). Although the existence of heteroplasmy in healthy cloned animals (Han et al., 2004; Steinborn et al., 2002) could suggest a level of compatibility between the two mtDNA populations, there may be unknown consequences or undetected deficiencies in these oVspring. Importantly, it has been demonstrated in parthenogenetically activated bovine oocytes that the addition of somatic cell mtDNA was detrimental to development (Takeda et al., 2005). Cloned oVspring with heteroplasmic mtDNA populations may suffer from similar abnormalities associated with ooplasmic/cytoplasmic transfer (CT). In the human, the transfer of healthy donor ooplasm, along with a sperm, into mature oocytes from patients with repeated implantation failure has successfully led to births of healthy babies (Cohen et al., 1997). This transfer of cytoplasm can, but does not necessarily, involve the transmission of mitochondria and therefore mtDNA from the donor, resulting in heteroplasmy (Brenner et al., 2000). This raises an important question: Does heteroplasmy cause an increased risk of defects in CT oVspring?

Table I

Transmission of Donor mtDNA in Intraspecific NT Crosses

Species

Nuclear Donor Source

Percentage Donor mtDNA in Material Analyzed

References

B. taurus (cattle)

Blastomere

Steinborn et al., 1998a

B. taurus Ovis aries (sheep)

Blastomere Somatic

B. B. B. B.

Blastomere Blastomere Somatic Somatic

0–18% in blood from oVspring. Levels were dependent on mtDNA contribution of donor blastomeres 13% at one cell, 13% at morula, and 9% at blastocyst 0% in all tissues analyzed (blood, skeletal muscle, placenta, and skin) of 10 live oVspring 2–57% in oVspring 1/27 oVspring possessed donor mtDNA 0.6–2.8% several tissues from oVspring Present in 1‐, 2‐, 4‐, 8‐, 16‐cell, morula, and blastocyst stage embryos 0–59% in tissues and blood from 11 oVspring and fetuses 0–12.7% in 11 tissues of from oVspring 24/25 mice contained donor mtDNA in all tissues analyzed; tissue‐specific patterns: mean of 0.6% in brain, 1.7% in liver Two oVspring contained mtDNA from the blastomere and the sperm originally used to fertilize, in addition to that of the oocyte used in NT 0% donor mtDNA in blood from oVspring. Double NT resulted in heteroplasmy as mtDNA from oocyte and zygote 0.1–1% in blood and hair root from oVspring 0–44% in tissues of progeny 0–8.72% in preimplantation stage embryos. 0–0.08% in embryos made from donor cells with only residual mtDNA levels

taurus taurus taurus taurus

B. taurus B. taurus Mus musculus (mouse) Macaca mulatta (macaque)

Somatic Somatic Somatic Blastomere

Sus scrofa (pig)

Somatic

Sus scrofa

Somatic

Ovis aries

Somatic

Steinborn et al., 1998b Evans et al., 1999 Hiendleder et al., 1999 Takeda et al., 1999 Steinborn et al., 2002 Do et al., 2002 Takeda et al., 2003 Hiendleder et al., 2003 Inoue et al., 2004 St. John et al., 2004

St. John et al., 2005 Takeda et al., 2006 Lloyd et al., 2006

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This question remains to be resolved. However, analysis of outcomes related to CT suggests a sense of caution should be taken. Reports have included two cases of Turner syndrome, one of which was electively aborted while the other was spontaneously aborted, and one case of pervasive development disorder (Barritt et al., 2001). A heteroplasmic mtDNA population may result in impaired or suboptimal respiratory function of the electron transfer chain (ETC) and could even increase the incidence of mitochondrial diseases in the NT oVspring. This could arise due to diVerences in mtDNA sequences from the two fusion partners giving rise to altered amino acid sequence and consequently protein structure (St. John et al., 2005; Steinborn et al., 2002). This may result in inadequate interaction between the separate subunits of the electron transport chain with reduced capacity for energy production, as described in cybrid studies (Trounce and Pinkert, 2007). Furthermore, foreign mtDNA, whether donor or recipient, may be present in the oVspring but not actively replicated or transcribed. Therefore, a heteroplasmic NT oVspring may only have 40% plus functioning mtDNA genomes, based on the range of 0–59% donor mtDNA molecules being present in offspring (Takeda et al., 2003). Many mitochondrial diseases result from oxidative phosphorylation (OXPHOS) deficiencies (McFarland et al., 2007). The severity of an OXPHOS defect that results from an mtDNA mutation is dependent on the proportion of mutated mtDNAs (Wallace et al., 1992). A threshold level exists where in some cases up to 85–90% mutant mtDNA needs to be present for a biochemical or clinical disease phenotype to be expressed (Boulet et al., 1992). Other reports suggest that the presence of >60% mutant mtDNA is characteristic of phenotypic onset in LHON (Chinnery et al., 2001) with some longitudinal variability in mutant load (Howell et al., 2000). Consequently, some NT oVspring may be phenotypically normal, despite having considerable levels of mtDNA diversity present in their genomes whilst others will have reached the threshold for phenotypic onset. Although heteroplasmy has been reported in a number of cattle clones, the reduction or elimination of donor mtDNAs was observed later in development in both NT embryos and oVspring (Steinborn et al., 1998a; Takeda et al., 1999). This could be explained by the oocyte contribution at reconstruction being far more significant than the donor cells. Bovine oocytes tend to have a mean mtDNA content of 2.6 105 copies, while somatic cells contain between 103 and 104 copies (Michaels et al., 1982). Paradoxically, oocyte mitochondria contain only a single mtDNA copy whereas somatic cell mitochondria each possess between 1 and 15 copies (Satoh and Kuroiwa, 1991), reflecting the cell’s specific requirement for OXPHOS (Moyes and Battersby, 1998). However, the greater persistence of oocyte mtDNA following NT could also be explained by recent reports, suggesting that a threshold of about 100,000 mtDNA copy numbers must be exceeded for fertilization to ensue in mouse (Piko and Taylor, 1987), human (Almeida Santos et al.,

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2006; Reynier et al., 2001), and pig (El Shourbagy et al., 2006). Interestingly, it has been proposed that by using more advanced preimplantation ECs, lower levels of donor mtDNA would be introduced and fewer of these mtDNA molecules would be transmitted to oVspring (Steinborn et al., 1998a). Two bovine oVspring derived from 92‐cell morula blastomeres as nuclear donors indeed harbored less donor mtDNA than two oVspring derived from 24‐cell morula blastomeres (Steinborn et al., 1998b). However, two oVspring derived from 52‐cell morula blastomeres harbored less donor mtDNA than the oVspring derived from 92‐cell morula blastomeres. Analysis of the only two nonhuman primate oVspring generated through ECs detected the persistence of both donor cell sources of mtDNA, namely the oocyte and sperm contributions which gave rise to these ECs (St. John and Schatten, 2004). This persistence of sperm mtDNA, which would have been introduced at very low levels, was most likely due to the inter‐specific nature of the donor cell which was generated using an Indian Rhesus macaque oocyte fertilized by a Chinese Rhesus macaque sperm, resulting in its substantial replication. In a more recent study, donor mtDNA from the somatic cells depleted to only residual levels, approximately 0.02% of their original mtDNA content, still persisted to blastocyst (Lloyd et al., 2006). Consequently, in some cases, there appears to be a tendency for the donor cell to rescue its own mtDNA during early embryonic development. Preservation of donor mtDNA following NT might occur to a greater extent when the donor nucleus and recipient oocyte are from more diverse genetic backgrounds. B. taurus oocytes injected with nuclei from the closely related B. indicus have been found capable of supporting development and subsequent production of live oVspring. mtDNA analysis showed the occurrence of genetic drift in early development that reduced or completely eliminated B. indicus (donor) mtDNA (Meirelles et al., 2001). Another study revealed the coexistence of B. taurus and B. indicus mtDNA in healthy clones (Steinborn et al., 2002). Interspecies NT, however, leads to further potential problems for the ETC generation of ATP. In this instance, the nuclear‐ encoded genes of the ETC from one species are required to interact with mtDNA‐encoded genes from another species. The persistence of donor mtDNA following interspecies NT has produced varying results (Table II). For example, the fusion of a C. ibex donor cell with a C. hircus recipient oocyte resulted in the donor mtDNA persisting to the two‐cell stage (Jiang et al., 2004). The fusion of chicken blastodermal cells with rabbit oocytes resulted in 9.7% of fused embryos, developing to the blastocyst stage where both mtDNA populations were detectable (Liu et al., 2004). Likewise, the fusion of a human nucleus into a bovine oocyte demonstrated that human mtDNA persisted up to the 16‐cell stage (Chang et al., 2003), while the rabbit has proven to be a successful vehicle for the transmission of human DNA in the form of human ESCs (Chen et al., 2003) although no mtDNA

Table II

Donor mtDNA Transmission in Interspecific and Cross‐species NT Crosses

Species (Donor; Recipient)

Nuclear Donor Source

B. indicus

Blastomere

B. taurus B. indicus; B. Taurus

Somatic

Ailuropoda melanoleuca (panda); Oryctolagus cuniculus (rabbit) Homo sapiens (human); B. taurus Macaca mulatta; Oryctolagus cuniculus Gallus domesticus (chicken); Oryctolagus cuniculus C. ibex; C. hircus (goat) C. ibex; Oryctolagus cuniculus

Percentage Donor mtDNA in Material Analyzed

References

Somatic Somatic

0–63% in morula/blastocyst. 0.6–3.6% in several tissues from a fetus 0% in Several tissues from oVspring 0–28% in blood, muscle, skin, oocytes, follicular cells Detected in blastocyst stage embryos but not in fetuses Present up to 16‐cell stage 2% at one‐cell stage, 0.011% at blastocyst stage

Chang et al., 2003 Yang et al., 2003, 2004

Embryonic

Present up to blastocyst stage

Liu et al., 2004

Somatic Somatic

Detected in one‐ and two‐cell embryos but not later Detected in preimplantation embryos

Jiang et al., 2004 Jiang et al., 2005

Somatic

Meirelles et al., 2001 Meirelles et al., 2001 Steinborn et al., 2002 Chen et al., 2003

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analysis was conducted. In similar studies, macaque donor cells transferred into rabbit oocytes, resulted in the persistence of 0.011% donor mtDNA at blastocyst (Yang et al., 2003, 2004). It is evident from these NT experiments that the cytoplasm of one species is capable of supporting some preimplantation development with nuclei of other species. However, outcomes from these studies, both in terms of developmental stage reached and persistence of donor mtDNA, are very variable. Consequently, in this context, it is also important to consider the diVerent techniques employed to quantify levels of donor mtDNA present, following NT to determine how stringent some forms of analysis really are. For example, PCR followed by restriction enzyme digest is capable of detecting heteroplasmy at a proportion of >2% (Hiendleder et al., 2003). Alternatively, allele‐specific real‐time PCR assays can detect heteroplasmy to a sensitivity of <0.1% (Lloyd et al., 2006; Steinborn et al., 2002). The many uncertainties in NT outcome highlight the importance of mtDNA analysis in cloned embryos, ESCs, and oVspring. First, the detection of recipient oocyte mtDNA, alongside nuclear genome analysis, will verify that reconstruction has taken place. Basic techniques such as PCR and restriction enzyme digests can be performed post‐ reconstruction for this purpose. However, to study the transmission of donor mtDNA we need quantified data where the assay is sensitive. We propose 0.1% as a minimum assay threshold, which is similar to the level of sperm mtDNA found in eggs. With more uniform analyses being performed, it would be easier and more appropriate for drawing comparisons between the vast numbers of NT experiments in increasingly diverse species in order to determine the true impact that donor mtDNA and heteroplasmy have on NT outcome.

E. Haplotype and Genetic Divergence It is important to consider the influence of mtDNA haplotype on developmental potential for NT. mtDNA haplotype influences milk quality in Holstein cows (Schutz et al., 1994), growth and physical performance in mice (Nagao et al., 1998), and fertility in beef cattle (Sutarno et al., 2002) and pigs (El Shourbagy et al., 2006). Interestingly, a comparison between two cattle haplotypes has shown increased preimplantation development following IVF for one haplotype with distinct diVerences in their ATP consumption despite both sets of embryos possessing similar levels of mtDNA (Tamassia et al., 2004). However, a reversed outcome was observed for the same haplotypes following NT (BruggerhoV et al., 2002). Consequently, the matching of a strong or desired chromosomal genome with an appropriately functional mtDNA could confer a developmental advantage as long as the power of the mtDNA haplotype had been determined. For the livestock industry, super‐oVspring or founders could be established from

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which breeding programs could be driven. However, such speculative invention could be limited by the extent to which the evolutionary distance between the mtDNA haplotype of the donor cell and the recipient oocyte will allow faithful mtDNA replication, transcription, and translation (McKenzie et al., 2003). F. mtDNA Copy Number Due to the importance of mitochondria in terms of mtDNA copy number, ATP production, and cellular distribution during oocyte maturation, it is important to select appropriate oocytes for use in IVF and NT procedures. A threshold level of approximately 100,000 mitochondria is required for fertilization and for subsequent embryo development to take place (El Shourbagy et al., 2006). This may arise due to sufficient levels of ATP being required for cell division (Brenner et al., 2000), or to mediate mitochondrial dynamics during cell division (Hsieh et al., 2002; Van Blerkom et al., 1998). In addition, oocyte maturation, fertilization rates, and development of subsequent embryos were all influenced by changes in active mitochondrial distribution, mitochondrial reorganization, and ATP levels in pig oocytes (Stojkovic et al., 2001; Sun et al., 2001). Furthermore, supplementation of both mouse and human oocytes with metabolically active mitochondria resulted in a net increase in ATP production (Van Blerkom et al., 1998). Similarly, developmentally less competent pig oocytes had lower mtDNA copy number than more competent oocytes but supplementation with donor mitochondria improved fertilization rates to normal levels (El Shourbagy et al., 2006), suggesting that preselection techniques could improve NT outcome. As mtDNA copy number may influence embryo development, it is important to consider how the reconstruction process may influence the resulting mtDNA populations (Fig. 3). For example, if a large amount of cytoplasm is removed from the oocyte at enucleation, then mtDNA levels may drop below the threshold level and result in a nonviable embryo. Conversely, the addition of donor mtDNA at reconstruction could increase mtDNA levels above the threshold, hence allowing development to commence, given that the additional mtDNA is functional, though as previously indicated, this could be detrimental (Takeda et al., 2005). G. mtDNA Transcription and Replication In order to make predictions about compatibilities between nuclear and mitochondrial genomes, we must have a clear understanding of the interactions that are involved between these genomes during the processes of mtDNA transcription and replication. To date, the evidence for the onset of mtDNA

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Gastrulation

mtDNA copy number per cell

Critical threshold

Oocyte maturation

Preimplantation development

Cell differentiation

Figure 3 mtDNA replication prior to and during preimplantation development. mtDNA copy number is cell type specific and related to specific energy requirements. The primordial germ cell contains very few mtDNA copies, having passed through a restriction event. Throughout oocyte maturation, mtDNA copy number increases considerably as the cytoplasm expands. Copy number should normally exceed a threshold level for fertilization to take place. Those oocytes that fall below this threshold will be developmentally incompetent and will not fertilize (red line). Those oocytes that lie above the threshold can undergo fertilization and subsequent development (green line). As mtDNA replication is inactive during preimplantation development, the large number of cell divisions results in few mtDNA copies being present in each cell. During later development as cells diVerentiate down particular lineages, mtDNA copy number is increased again to match each cell’s requirement for OXPHOS (green lines). The blue dotted line predicts outcomes following NT. If mtDNA replication occurs prematurely during preimplantation development, mtDNA copy numbers are likely to be higher than normal.

replication and transcription during embryogenesis has been conflicting. Initial studies suggested that there are high levels of mtDNA transcription during early embryonic development (Piko and Taylor, 1987) but that mtDNA replication does not occur until after the blastocyst stage (Huo and Scarpulla, 2001; Larsson et al., 1998; Piko and Chase, 1973). However, mitochondria are dynamic organelles, being generally dispersed throughout the cell’s cytoplasm and then moving close to the nucleus in readiness to provide energy for cell division processes such as spindle formation (Stojkovic et al., 2001; Sun et al., 2001; Wilding et al., 2001a,b). This evidence suggests that the mitochondria are active prior to the blastocyst stage and asks us to question whether the mtDNA molecule needs to be replenished at this stage of development to support these activities. Interestingly, a small burst of mtDNA replication at the two‐cell stage in mouse embryos has been reported (McConnell and Petrie, 2004), perhaps coinciding with embryonic genome activation (Bolton et al., 1984). This switch from maternal to zygotic transcription occurs later in others species being at the four‐cell stage in pig (Jarrell et al., 1991) and at 8‐ to 16‐cell stage in cattle

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(Camous et al., 1986) and sheep (Calarco and McLaren, 1976). This led others to reconsider whether mtDNA replication is restricted to post‐implantation stages (May‐Panloup et al., 2005; Thundathil et al., 2005). However, as replication is dependent on transcription, we will briefly describe the process of mtDNA transcription first, followed by replication and then consider two factors we believe are important in regulating mtDNA replication during preimplantation development and how they could be aberrantly regulated following NT. 1. Transcription of mtDNA The enzymes and factors responsible for regulating mtDNA transcription are encoded in the nucleus. These include mitochondrial transcription factor A (TFAM; Fisher and Clayton, 1985, 1988) and either mitochondrial transcription factor B1 (TFB1M) or B2 (TFB2M; Falkenberg et al., 2002; McCulloch et al., 2002) which interact with the mitochondrial‐specific RNA polymerase (mtRNApol; Tiranti et al., 1997). The mtDNA genome consists of a heavy (H)‐ and a light (L)‐strand, both of which are completely transcribed from initiation sites located in the D‐loop: H1 and H2 for the H‐strand and L for the L‐strand (Bogenhagen et al., 1984; Montoya et al., 1982, 1983; Fig. 4). The H1 and L transcription initiation sites are located within promoter regions, named the heavy strand promoter (HSP) and the light strand promoter (LSP), respectively. TFAM recognizes and binds to specific sequences upstream of HSP and LSP and stimulates transcription (Dairaghi et al., 1995a; Parisi and Clayton, 1991). 2. Replication of mtDNA As RNA primers produced by transcription from the LSP are required for mtDNA replication, initiation is dependent on transcription having first taken place. Two separate origins of replication exist, one for each strand (Fig. 4). The origin of H‐strand replication, OH, is located within the displacement (D)‐loop region, and OL is located two‐thirds round the genome, clockwise, from the D‐loop (Anderson et al., 1981). Replication begins when the primer precursor transcribed from the LSP hybridizes to a region upstream of OH and forms a loop structure. Processing of the precursor by mitochondrial RNA‐ processing endoribonuclease (RNaseMP) produces the mature primers for replication, as demonstrated in Fig. 4 (Shadel and Clayton, 1997). The mitochondrial‐specific DNA polymerase (polymerase gamma; POLG) starts replication from the RNA primer and extends along the template (Lee and Clayton, 1996; Shadel and Clayton, 1997; Xu and Clayton, 1996). Replication of mtDNA begins by initiation of H‐strand synthesis from the parental L‐strand and on reaching OL the parental H‐strand is displaced, exposing the L‐strand initiation site, and L‐strand synthesis then begins in the opposite direction (Shadel and Clayton, 1997). Following complete replication of both

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RNA

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CSBs

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Figure 4 Schematic representation for the initiation of mitochondrial DNA transcription and replication. Transcription proceeds from two initiation sites on the heavy (H)‐strand and one on the light (L)‐strand. Initiation from H1 and L requires binding of TFAM (shown in red) to an enhancer element which allows mtRNApol (shown in yellow) to begin transcription in the presence of TFB1M or TFB2M (shown in blue). Transcription from H1 produces tRNAphe, the 12S and 16S rRNAs, and tRNAval. H2 initiation occurs 10 times more frequently, does not require TFAM binding and produces a polycistronic molecule containing most of the H‐strand. Transcription from L produces a polycistronic molecule for the L‐strand and a primer required for mtDNA replication. After processing of the primer by RNase MRP (shown in green), mtDNA replication proceeds from OH by action of POLG (shown in purple). The transition from the RNA primer to the newly synthesized DNA (shown in orange) takes place over a series of conserved sequence blocks (CSBs; shown in black). Once H‐strand replication reaches and exposes OL, L‐strand synthesis begins in the opposite direction.

strands, the RNA primers are removed, the gaps are filled and ligated, and mtDNA adopts its tertiary structure (Lecrenier and Foury, 2000). A second mechanism for mtDNA replication has also been described, involving ‘‘coupled leading‐ and lagging‐strand synthesis,’’ where both H and L strands are replicated at the same time from the same initiation site (Holt et al., 2000). This mechanism is proposed to occur in cells in addition to the mechanism described above, but is thought to be particularly active in partially depleted cells, suggesting that alternative modes of mtDNA replication can occur in cells under diVerent conditions according to the cell’s

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need to regulate mtDNA copy number. Since changes in mtDNA copy number will alter ratios between mtDNA and factors such as TFAM and POLG, such relationships could contribute to the initiation of this alternative mtDNA replication mechanism (Holt et al., 2000). These authors have suggested that mtDNA replication may occur primarily, or indeed, solely by their proposed mechanism and that it may initiate from multiple origins near to OH (Bowmaker et al., 2003; Yang et al., 2002).

a. Mitochondrial Transcription Factor A. TFAM is encoded on chromosome 10q21 and is made up of 204 amino acids and largely comprises two regions of sequence similar to nuclear High Mobility Group (HMG) box proteins and a mitochondrial‐targeting sequence (Parisi and Clayton, 1991). These boxes, termed HMG1 and HMG2, are the important structural domains of the protein concerned with DNA binding and stimulation of transcription (Clayton, 1992). In addition, the protein has a 27 amino acid linker domain between these two HMG‐box domains and a 25 amino acid carboxyl tail, which are important for the protein’s ability to initiate transcription (Dairaghi et al., 1995b). TFAM plays an important role in the diVerential regulation of gene expression (Attardi and Schatz, 1988), as it regulates mtDNA copy number in vivo and is essential for mitochondrial biogenesis and embryonic development (Larsson et al., 1998). An increased rate of transcription can be brought about by increasing the intramitochondrial level of TFAM, suggesting that it is a regulator of mitochondrial gene expression in vivo (Gensler et al., 2001). Reduced levels of TFAM are associated with mtDNA depletion and a number of severe mtDNA diseases, including mitochondrial myopathy and encephalomyopathy disorders (Larsson et al., 1994; Poulton et al., 1994; Siciliano et al., 2000; Spelbrink et al., 1998). The importance of TFAM in regulating mtDNA copy number has been demonstrated in TFAM knockout mice generated through LoxP‐Cre‐mediated excision. Heterozygous knockouts exhibited reduced mtDNA copy number and respiratory chain deficiency in heart while homozygous mice exhibited severe mtDNA depletion and abolished OXPHOS (Larsson et al., 1998). However, homozygous mutant embryos did survive through implantation and gastrulation. This could suggest that OXPHOS is not required up to that stage. Despite reduced mtDNA copy number, levels of mitochondrial transcripts and respiratory chain subunits did not significantly decrease in some tissues (Larsson et al., 1998). Furthermore, tissue‐specific TFAM knockouts had diVerent survival abilities, with most mice targeted with the heart‐specific disruption of TFAM dying in the neonatal period (Wang et al., 1999). Consequently, post‐gastrulation, the timing of initiation of transcription

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and replication of the mtDNA genome appears to be critical for embryo survival and depends on the expression of TFAM. However, other studies have demonstrated that there is not always a direct relationship between mtDNA copy number and TFAM levels. To this extent, TFAM and mtRNApol were shown to be down regulated when mtDNA copy number was reduced perhaps preventing the accumulation of these proteins in order to maintain the remaining genomes in an actively replicating state (Seidel‐Rogol and Shadel, 2002). Indeed, excess TFAM can bind and saturate mtDNA template, resulting in complete inhibition of transcription from LSP in vitro (Dairaghi et al., 1995a). However, when partially mtDNA‐depleted cells were cultured in the absence of their depleting agent, TFAM levels recovered to normal at a significantly slower rate than mtDNA, as did mtRNApol (Seidel‐Rogol and Shadel, 2002). The observed discrepancies in the relationship between mtDNA and TFAM may arise due to varying levels of accessory and regulatory factors being present thus mediating differential regulation of TFAM activity. TFAM function is also dependent on two mitochondrial transcription factors, TFB1M and TFB2M, which have mitochondrial‐targeting sequences (Falkenberg et al., 2002). These have been shown to have homologues in the mouse, Tfb1m and Tfb2m. These two factors are ubiquitously expressed and can both independently support promoter‐specific initiation in the presence of TFAM and mtRNApol, but levels of transcription with mtRNApol‐TFB1M are at least tenfold lower than for mtRNApol‐TFB2M (Falkenberg et al., 2002). TFB1M interacts with the C terminus of TFAM, a region necessary for transcriptional activation (McCulloch et al., 2002). These factors regulate transcription of mtDNA and therefore mtDNA gene expression, and are necessary for basal transcription of mtDNA (Falkenberg et al., 2002). Activity of TFAM is also highly dependent on nuclear respiratory factors 1 (NRF‐1) and 2 (NRF‐2) which have recognition sites on TFAM’s proximal promoter (Evans and Scarpulla, 1989; Virbasius et al., 1993).

b. Polymerase Gamma. The POLG gene (PolG) has been mapped to human chromosome 15q24–15q26 (Ropp and Copeland, 1996; Walker et al., 1997; Zullo et al., 1997), while the mouse homologue (Polg) is located on murine chromosome 7 (Zullo et al., 1997). POLG is made up of two protein subunits, a 140‐kDa catalytic subunit (POLGA) and a smaller 54‐kDa accessory subunit (POLGB; Gray and Wong, 1992). PolGA encodes the catalytic unit which is responsible for both the polymerase and 50 ‐30 exonuclease activity, while PolGB encodes the accessory unit that enhances the enzyme’s DNA‐binding ability, processivity, and fidelity (Graves et al., 1998;

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Lim et al., 1999; Longley et al., 2001). The accessory subunit is also thought to play a role in the recognition of RNA primers in mtDNA replication, helping recruitment of POLG to the primer–template junction (Fan et al., 1999). Mutations in the gene‐encoding POLG can aVect its targeting, transport, or proofreading function and are likely to result in impaired OXPHOS and therefore ATP production (Ropp and Copeland, 1996). For example, three PolG missense mutations have been associated with progressive external opthalmoplegia (PEO) or mitochondrial neurogastrointestinal encephalomyopathy (Van Goethem et al., 2001, 2003). In addition, patients receiving treatment with nucleoside analogue reverse transcriptase inhibitors (NRTIs) for HIV infection have been shown to develop varying degrees of myopathy and neuropathy, resembling mitochondrial disease resulting from inhibition of POLG (Dalakas et al., 2001). In the human, POLG also possesses a series of CAG repeats that encode a polyglutamine tract (Ropp and Copeland, 1996). Heterozygosity for CAG repeats has been implicated in some cases of idiopathic male subfertility (Jensen et al., 2004; Rovio et al., 2001, 2004) and testicular cancer (Nowak et al., 2005). The precise role of POLG in regulating mitochondrial biogenesis is unclear. Overexpression of Polg in mouse cell lines did not result in a corresponding increase in mtDNA copy number (Schultz et al., 1998). However, knockin mice that express an error‐prone version of Polg show a decrease in exonuclease activity with a corresponding increase in mtDNA mutations and a reduction in mtDNA copy number to 70% of the wild‐type levels (Trifunovic et al., 2004). The generation of a transgenic mouse model lacking PolgA through LoxP‐Cre homologous recombination has produced interesting outcomes (Hance et al., 2005). PolgA heterozygous oVspring were crossed to produce homozygotes which died between E7.5 and E8.5. They were also smaller and much less developed than the wild type and exhibited severe respiratory chain dysfunction, shown by an absence of COX activity. Although no diVerences in mtDNA levels were observed between wild type and heterozygotes, homozygous embryos contained barely detectable levels of mtDNA. Interestingly, mtDNA levels in adult tissue from heterozygous and wild‐type mice were comparable (15% decrease in heterozygotes) despite transcript levels for PolgA being half that of the wild‐type. This shows that mtDNA levels and PolgA expression are not directly related. Additionally, when crossed with TFAM overexpressor mice, heterozygous mutants were equally capable of increasing mtDNA levels as were their wild‐type counterparts (Hance et al., 2005). However, POLG is stable in the absence of mtDNA due to there being no feedback mechanism to alter its levels (Davis et al., 1996). It is also not upregulated by stimuli that increase mitochondrial biogenesis but is consistently expressed in all tissues (Schultz et al., 1998).

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VI. Are All the Potential Reasons for Low Success Rates in NT Interlinked? Many of the issues we have discussed are unlikely to act independently in determining NT success. Rather, as in any biological system, we would expect them to be interlinked (Fig. 5). For example, the ability of an enucleated oocyte to support development of a donor nucleus is dependent on there being a level of nucleo‐mitochondrial compatibility. This is determined by the cell cycle phases of both the oocyte and donor cell, which in turn could influence mtDNA turnover and hence copy number. The regulation of mtDNA replication is dependent on the appropriate time‐dependent expression of those

Epigenetic reprograming

Correct state of pluripotency

Sufficient oocyte mtDNA copy number

Evolutionary distance

Nuclear transfer outcome

↑ Blastocyst rate ↑ Cells per blastocyst mtDNA homoplasmy Appropriate respiratory capacity Healthy, functional offspring

Nuclear regulation of mtDNA replication and transcription

Reduction in donor mtDNA transmission

Cell cycle stage coordination of donor cell and oocyte

Figure 5 The factors combining to influence NT outcomes and subsequent mtDNA replication. Successful NT and the production of healthy oVspring are dependent on a number of interlinking factors. It has been determined that the cell cycles of the donor cell and recipient oocyte must be compatible for development to commence and that the somatic donor nucleus must be epigenetically ‘‘reprogramed.’’ This will ensure the correct state of pluripotency and correct patterns of gene expression. The mtDNA content is also critical as total copy number must exceed a threshold value to provide enough ATP for development. In addition, when introducing foreign mtDNA into the oocyte from the donor cell, a state of heteroplasmy exists that could negatively impact on OXPHOS. It is also important that the processes of mtDNA transcription and replication are eYciently regulated as these will determine whether mtDNA populations can proliferate. These factors could be influenced by the evolutionary distance between the donor cell and recipient oocyte as increased genetic diversity could cause a breakdown in communication between the nucleus and mitochondria. mtDNA replication could equally be influenced by the potential epigenetic status of the mtDNA‐encoded transcription and replication factors and the pluripotent status of the cell.

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nuclear‐encoded replication factors and on somatic cell reprograming and ‘‘epigenetic memory.’’ The evolutionary distance between donor cell and recipient oocyte could provide a critical constraint on development. For example, crosses within or between more closely related species might result in successful development, as seen with B. taurus and B. indicus (Meirelles et al., 2001), but as evolutionary distance increases, developmental potential would decrease as might be the case in, for example, goat–sheep crosses. This would result not only from the inability of the nucleus to mediate mtDNA replication but also the ability to undergo epigenetic changes essential for reprograming. Increases in evolutionary distance have also been noticed in respect of sperm mtDNA transmission. In this situation, we see elimination of sperm mtDNA in intraspecific crosses by the eight‐cell stage (Gyllensten et al., 1991; Shitara et al., 1998), but persistence in interspecific crosses (Cummins et al., 1997; Kaneda et al., 1995; Sutovsky et al., 1996) and following NT (St. John and Schatten, 2004). Here, normal processes of sperm mtDNA elimination by ubiquitination (Sutovsky et al., 1999) are bypassed due to interspecies diVerences. Consequently, with an acceptable range of evolutionary distance, one might expect to see donor mtDNA replicated following NT and successful development. As mtDNA replication is reliant on the interactions of a large number of different nuclear encoded proteins, there are numerous possible sources of error when the nucleus from one species is required to regulate the mitochondria of another. For example, specific binding sites for TFAM exist in the mtDNA D‐loop (Fisher and Clayton, 1988; Fisher et al., 1987). Base pair diVerences exist in functionally important sections of the mtDNA D‐loop between mtDNA molecules of diVerent haplotypes from the same species (Hiendleder et al., 1998; Steinborn et al., 1998c). Hence, even greater diVerences are likely between those of diVerent species. This is important when thinking about cross‐ species NT experiments, especially as heteroplasmic mtDNA populations are likely to result in suboptimal functioning of the ETC and increased risk of mtDNA‐related diseases of OXPHOS (St. John et al., 2004). Consequently, the existence of mixed mtDNA populations, together with abnormal expression of factors that drive mtDNA replication and transcription, are likely to have a major impact on the development of cloned embryos. Pronuclear transfer (PNT) oVers some explanation as to the power of donor mtDNA replication during pre‐ and postimplantation development. The transferred PN is still surrounded by mitochondria, which tend to remain close to the center of the reconstruction. They then disperse throughout the cytoplasm (Fulka, 2004). It is therefore most likely that, as the mitochondria are initially in close proximity to the nucleus, the donor mtDNA would be preferentially replicated in the reconstructions. Indeed, it has been demonstrated that exogenous mtDNA accompanying the murine zygotic karyoplast persists at higher levels once the embryo has reached blastocyst than mtDNA from transferred

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cytoplasts (Meirelles and Smith, 1998). Furthermore, oVspring derived from zygotic karyoplasts being transferred into recipient zygotes harbored varying amounts of donor mtDNA (0–69%; Meirelles and Smith, 1997), which are very similar to outcomes associated with NT. Consequently, if this technique was to be used as a form of treatment for those women with mtDNA‐type disease, then it would be very diYcult to predict how safe this technique would actually be. Indeed, recent data suggest that outcomes might be species‐specific and, those species with longer gestational periods would be more prone to transmitting greater amounts of the mtDNA accompanying the karyoplast (Sato et al., 2005). For example, in humans, the gestational period is 13 times longer than in the mouse and this coincides with the window in which mutant mtDNA replication is most proliferative (Sato et al., 2005). For NT, this would suggest that for those larger mammals such as cattle, pigs, sheep, goats, and monkeys, the longer gestational periods would give rise to a greater propensity of donor mtDNA being transmitted and that the levels of mtDNA variation between two alleles could be close to those typical for the onset of disease. Initial work has demonstrated that somatic cells used as nuclear donors are still programed to drive mtDNA replication through their continued expression of PolG and TFAM (Lloyd et al., 2006). These factors also appear to be unregulated in cloned embryos unlike their IVF counterparts (Lloyd et al., 2006). The abnormal expression of TFAM and POLG results in the loss of the tight regulation of mtDNA replication with the potential to initiate premature mtDNA replication (Fig. 2). This is highlighted by the persistence of donor mtDNA present at residual levels prior to NT which, consequently, has not been simply diluted out (Lloyd et al., 2006) or has even been preferentially replicated. This could therefore result in the unequal segregation of donor mtDNA into individual blastomeres and explain the huge variation seen in tissues of the same individual (Takeda et al., 2003). As epigenetics can contribute to repression of transcription by the methylation of CpG sites within promoter regions, so selective demethylation of promoter DNA may be a required mechanism for reprograming somatic cell nuclei (Simonsson and Gurdon, 2004). It has been proposed that TFAM packages mtDNA due to its high aYnity to the mtDNA molecule (Alam et al., 2003). The promoter regions of TFAM have been characterized in human, mouse, and rat (Choi et al., 2002; Larsson et al., 1997a,b; Tominaga et al., 1992). The proximal region of the human sequence contains binding sites for Sp1, NRF‐1, and NRF‐2 to which binding is a major determinant in TFAM promoter activity (Virbasius and Scarpulla, 1994). The activation of the TFAM promoter is completely abolished by mutating the NRF‐1‐binding site and mutating the NRF‐2‐ or Sp1‐binding site reduces activity, but to a much smaller extent (Gleyzer et al., 2005). Methylation of these sites interferes with binding and hence suppresses transcription (Choi et al., 2004). Furthermore, an Sp1‐binding site has also been detected at about 90 bp

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upstream from the start site of the mouse POLG sequence, in the promoter region (Mott et al., 2000). We have previously hypothesized that promoter methylation is a likely mechanism responsible for regulating mtDNA replication (St. John et al., 2004). Therefore, the incomplete reprograming of the somatic donor cell would result in methylation patterns similar to those of somatic cells that are driving mtDNA replication. Consequently, the epigenetic regulation of mtDNA replication may be indirectly organized by its nuclear‐encoded transcription and replication factors.

VII. Conclusions It is becoming increasingly evident that the replication, transcription, and transmission of mtDNA are aVected by NT. This, in turn, may influence the OXPHOS function of the oVspring and the ESCs derived. It is perhaps pertinent that we do not just consider matters such as epigenetic organization of gene expression and pluripotency of the reconstructed oocyte when attempting to improve NT outcome. mtDNA haplotype, nucleo‐mitochondrial interaction, nucleo‐mitochondrial compatibility, and mtDNA copy number require appropriate regulation. The interesting proposition to determine is whether the factors established as the key problems hindering NT success are those that also regulate mtDNA replication, transcription, and transmission. Such an outcome would therefore warrant a holistic approach.

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Index

A AAT. See -1-antitrypsin AD. See Alzheimer’s disease Adenosine diphosphate (ADP), 52, 115 Adenosine trisphosphate (ATP), 3, 8, 12, 21–22, 31, 52, 70, 73, 123, 127 production, 4, 6, 10–11, 32, 35–36, 88, 114 in mitochondria, 7, 9, 33–34, 37–38, 52–53, 63, 65–66, 116, 131, 135, 140 ADP. See Adenosine diphosphate A3243G, 215–216 Aging mitochondrial involvement, 128–129 oocyte, 24, 40, 98–99 AIF. See Apotosis-inducing factor Alzheimer’s disease (AD), 127, 163–164 Aneuploidies elimination of, 196 in embryo, 201 Animal modeling of mtDNA transmission and human disease, from transgenesis to transmitochondrial cybrid models embryonic cytoplast and karyoplast fusion, 173–175 mitochondrial genome, modifying, 170–171 modeling rationale, 169 spontaneous and induced models ATPase6 gene, 170 BHE/Cdb rat, salt-sensitive, 170 transfected 0 cells use as intermediate mitochondrial carriers, mouse models production and, 171 transmitochondrial mice mtDNA injection versus ES cell-derived models, 171–173 xenomitochondrial mice and, 175–176 Animal models of bottleneck, polymorphic mtDNA variants and, 218–219 Antimycin A, 7 -1-antitrypsin (AAT), 255 Apoptosis, 71 in ascidians, 41 in embryo, 22 in blastocyst, 33

hyperglycemia, 41 mitochondria, 24, 40–41 in oocytes, 38, 40, 59 in Xenopus, 24, 41 in zebrafish embryos, 24 Apoptosis-inducing factor (AIF), 40 Ascidians, 24, 27–28, 30, 33–35, 38–39, 41 embryo, 23, 30 Assisted reproductive techniques (ART), importance of cytoplasmic transfer and, 230–232 fetal cord blood and, 231–232 ICSI and, 231 IVF and, 230, 232 mitochondrial dysfunction and, 230 mitochondrial fingerprinting technique, 231 mitochondrial heteroplasmy, implications of, 243–244 molecular beacon analysis and, 231 A467T, 235 ATP. See Adenosine trisphosphate ATPase, 9–10, 70, 140 ATP6 gene NARP and, 161 spontaneous and induced models, 170 ATP synthase, 7, 34, 53, 88, 116 Atresia, 59, 88 in oocyte, 98–99 Autofluorescence, in mitochondria, in mouse/sea urchin eggs, 30–31

B Banteng (Bos javanicus), 256 Biochemical analysis, in muscle biopsy, 131–132 Bos indicus, 275 mtDNA transmission, NT and, 264–265 NT applications and, 256 Bos javanicus. See Banteng Bos taurus, 275 mtDNA transmission, NT and, 262, 264–265 NT applications and, 256 291

292 Bottleneck theory, in oogenesis, 58, 72 Bovine embryos, 26, 69 4977-bp (common deletion) Rhesus mitochondrial, 241

C Ca2þ dynamics and mitochondria ascidian and mouse fertilization, 24 ATP supply and demand, 34 homeostasis in eggs/embryos, 33–34 in sea urchins, 34 somatic cells, signaling of, 24 Cancer, mitochondrial involvement, 127 CAPR. See Chloramphenicol-resistant cell lines Capra hircus, 264–265 Capra ibex, 264–265 Cell cycle, NT failure and, 258 0 cells, 159 mtDNA-less cells, 160 OXPHOS and, 160–161 xenomitochondrial cybrids and, 168 Childhood-specific mitochondrial syndromes depletion syndromes, 141–142 Leigh Syndrome, 142 Pearson Syndrome, 142 Chimeric mice, xenomitochondrial cybrids and, 176 Chloramphenicol-resistant (CAPR) cell lines in human HeLa cells, 159 mouse cell mtDNA mutants and, 165 mutation, 174 Chloromethyl-X-rosamine (CMXRos), 206 Chorionic villus sampling (CVS), heteroplasmy of mtDNA and consequences of Mendelian traits and, 214 threshold eVect and, 214 mtDNA disease and, 216–217 distribution, heteroplasmic embryos and, 215–216 maternal inheritance and, 214–215 Chromosome group transfer aneuploidies, 201 GVBD and, 198–201 IVF, mouse oocytes and, 202 metaphase stages and, 198–202 parthenogenetic development and, 202

Index Chronic progressive external ophthalmoplegia (CPEO), 125, 140–141 heteroplasmic mtDNA deletions in, 161 syndrome, 241 Citrate metabolism, in mitochondrial embryos, 36 Classical mitochondrial syndromes CPEO, 140–141 KSS, 141 MELAS, 139 MERRF, 140 MNGIE, 140 NARP and MILS, 140 Clayton’s model, mtDNA replication, 122 Clinical applications, mtDNA in counseling women, 103–104 ICSI, 104 mutant mtDNAs, preventing transmission of, 103–104 oocyte cytoplasm, transfer of, 105 preimplantation genetic diagnosis, 104 prenatal genetic diagnosis, 104 Clinical assessment of, mitochondrial disease, 129 Clinical investigation of, mitochondrial disease, 129–130 Cloning. See also Nuclear transfer agricultural applications of, 256 monozygotic twins and, 252 CMXRos. See Chloromethyl-X-rosamine Coupled leading-and lagging-strand synthesis, 270 COX. See Cytochrome oxidase COX. See Cytochrome c oxidase CPEO. See Chronic progressive external ophthalmoplegia CtBP. See C-terminal-binding proteins C-terminal-binding proteins (CtBP), 39 CVS. See Chorionic villus sampling CVS, management options and nuclear cell transfer, 221–222 oocytes and pedigree analysis, preconception counseling and, 220 preimplantation genetic diagnosis, 220–221 Cybrid modeling of human disease-linked mtDNA mutations CPEO and, 161 LHON and, 161–162 MELAS and, 161–162 MERRF and, 161

Index NARP and, 161 Leigh’s disease, 162 Cybrids, mtDNA disease and CAPR, first mammalian cell cybrids and, 159 cytochalasin B, 158 0 cells, 159–161 heteroplasmic mtDNA mutants, complementation of, 162–163 modeling of, 161–162 mouse cell mtDNA mutants and, 165–167 xenomitochondrial cybrids, 167–169 OXPHOS and complexes, 158–159 functions, 165 polygenic disorders and tissue-specificity, mtDNA contributions to, 163–164 Sendai virus and, 158 Cytochalasin B, 191 Cytochrome c oxidase (COX), 126 in spermiogenesis, 5 Cytochrome oxidase (COX), 163 decreased activity of, 164 Cytoplasmic transfer, human infertility therapeutics and, 232 D De novo methylation, 259 Diabetes and hypertension, mitochondria involvement, 128 Dibutyryl cyclic AMP, GVBD and, 192 Displacement loop (D-loop), 234, 269 D-loop. See Displacement loop Drosophila embryos, mtSSB in, 239 xenomitochondrial cybrids and, 168 E EGA. See Embryonic genome activation Eggs Ca2þ homeostasis and mitochondria, 33–34 GFP, 28 mitochondrial autofluoresence, in mouse/sea urchins, 30–31 mitochondrial study in, 27–28 mitocondrial generation and distribution, 25–27 Ejaculated spermatozoa, survival of, 13–14 Electron transport chain (ETC) in OXPHOS, 7

293 respiratory complexes antimycin A, 7 gossypol, 7 NO, 7 oligomycin, 7 rotenone, 7 Embryogenesis, mtDNA and blastocyst, 65–66 in bovine embryos, 67–70 cell divisions, 65–70 in mouse embryos, 66–70 NRF1mRNA, 70 TFAM mRNA, 69–70 Embryonic cytoplast and karyoplast fusion, 173–175 CAPR mutation, 174 R6G and, 174 Embryonic genome activation (EGA), 233 Embryonic stem cell (ESC), 256–257 Embryos apoptosis and mitochondria, 24, 40–41 apoptosis in zebrafish/Xenopus embryos, 24 bovine embryos, mtDNA copy number, 26 Ca2þ homeostasis and mitochondria, 33–34 energy production and mitochondria, 31–33 GFP, in mouse, 28–30 glutathione (GSH/GSSG), 35 homoplasmy, 23 implantation, 65–66 mitochondrial impact CtBP, 39 oncosis, 38 S-glutathionylation, 39–40 sirtuins, 39 transcription factors, 39 mitochondrial study in, 27–28 mutations in, oocytes and, 71–74 NADH/NADþ and NADPH/NADPþ, 35 oxidative metabolism in mitochondria, 24 preimplantation, in mammals, 31–33, 36 redox metabolism and mitochondria, 34 citrate metabolism, 36 enzymes in mouse, 36 oxidant action in sea urchin, 38 Encephalomyopathy disorders, TFAM and, 271 Endoplasmic reticulum (ER), 24

294 Endosymbiosis in mitochondria, 114 in -proteobacteria, mtDNA in, 54 Epigenetic memory instability and NT failure, 260 Epigenetics CpG dinucleotides and, 258 disorders, NT applications and, 255 ICM and, 259 methylation and, 258–259 SCNT and, 259 trophectoderm and, 259 ER. See Endoplasmic reticulum ESC. See Embryonic stem cell ETC. See Electron transport chain Ethidium bromide, 272 F Female pronucleus (FPN), 188 Female reproductive tract, substrates in diabetic women, 13 glucose level, 11–13 lactate level, 11–13 oxygen level, 11–13 Flagellum implantation, defects, 4–5 Flavine adenine dinucleotide (FADH2), 53 Follicle cells, in oogenesis, 57–58 FPN. See Female pronucleus Free radical theory of aging, mutations in, 71 G -Galactosidase-neomycin cassette, 239 -1,3-Galactosyltransferase gene, 255 GAPDH. See Glyceraldehyde 3-phosphate dehydrogenase Gene knockin strategies, NT applications, 255 Germ cells mitochondrial location, 26–27 mitochondrial production, 25 mtDNA in, 87 segregation of mitochondria, 23 Germinal vesicle breakdown (GVBD), 188, 194 chromosome group transfer and, 198–201 germinal vesicle transfer and, 192 Germinal vesicle (GV), mitochondrial copy number and, 241–242 Germinal vesicle transfer antral follicles and, 190 cytochalasin B and, 191

Index enucleation and, 191–193 GVBD and, 192 GV karyoplast and, 194 piezo injector and, 191–192 vitelline membrane and, 191–192 zona pellucida and, 190–193 Germ line in females, mtDNA transmission bottleneck concept, 95–96 homoplasmy, 95 mouse, 95–96 point mutations, 97 replication of, 95 segregation of, 96–97 stochastic process, 97–98 mutations, in oogenesis, 98–99 origin and development of follicles, 93 formation of gonads, 93–95 Fragilis and Stella, 92 IFITM2, 92 IFITM3, 92 mitochondrial number, 94 in mouse embryo, 92 Oct4, 92 oogonia, 93 PGC, 92–95 GFP. See Green fluorescent protein Gluconeogenesis, in sperm motility, 10–11 Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) glycolysis, 9 in mouse, 9 Glyceraldehyde 3-phosphate (G3P), 10 Glycolysis, in sperm motility ATP production, 9–11 GADPH/, 9–10 gluconeogenesis, 10–11 G3P, 10 Gossypol, 7 G3P. See Glyceraldehyde 3-phosphate Green fluorescent protein (GFP), 22 mitochondria in eggs and embryos, 28–31, 93 GVBD. See Germinal vesicle breakdown H Handmade cloning (HMC), 253 Haplotype and genetic diversity, as NT failure, 266–267 IVF and, 266

295

Index Heavy strand promoter (HSP), 56, 120–121 Heteroplasmic embryos, mtDNA distribution and A3243G and, 216 CVS, implications for, 215–216 Heteroplasmic mtDNA mutants, complementation of, 162 COX and, 163 intramitochondrial, 163 Heteroplasmy mitochondrial genetics and, 117–118 polymorphism, in mouse, 118–120 in mtDNA mutations, 71 Heteroplasmy of mtDNA, chorionic villus sampling (CVS) and consequences of, dosage and thresholds and Mendelian traits and, 214 threshold eVect and, 214 mtDNA disease and, 216–217 distribution, heteroplasmic embryos and, 215–216 maternal inheritance and, 214–215 H19 gene, 197 High Mobility Group (HMG), 271 Histochemistry, in muscle biopsy, 130–131 HMC. See Handmade cloning HMG. See High Mobility Group Homoplasmy, mitochondrial genetics and, 117–118 HSP. See Heavy strand promoter Human infertility therapeutics, cytoplasmic transfer and, 232 Hyperglycemia, induction of apoptosis, 41 Hyperpolarized electrical potential, in mitochondria, 27–28

I ICM. See Inner cell mass ICSI. See Intracytoplasmic sperm injection Inner cell mass (ICM) epigenetics and, 259 NT applications and, 253 Inter-oocyte cytoplasmic transfer, in human fertility, 61–63

Intracellular redox potential (IRP), in mitochondria, 22, 24 redox metabolism in embryo, 34–35 transcription factors, 39 Intracytoplasmic sperm injection (ICSI), 60, 104 ART and, 231 oocytes fertilization and, 196–197 pipette of, technical equipment, micromanipulation methods and, 190 procedure, 215 In vitro fertilization (IVF), 59, 100, 266 ART and, 230, 232 chromosome group transfer and, 202 mitochondrial bottleneck and, 217–218 mouse spermatozoa and, 8 mtDNA, maternal inheritance and, 214 NT applications and, 255 oocytes and pedigree analysis and, 220 OXPHOS, sperm motility, 7 IRP. See Intracellular redox potential IVF. See In vitro fertilization K Kaguya, parthenogenetic mouse, 188 Kearns-Sayre syndrome (KSS), 97 mitochondrial bottleneck and, 218 mitochondrial copy number and, 241 Klinefelter’s syndrome, 215 Krebs cycle intermediates, synthesis of, in mitochondria, 36 KSS. See Kearns-Sayre syndrome

L Lactate dehydrogenase C4 (LDH-C4), in spermatogenic activity, 13 Lactate level, respiratory substrate in female reproductive tract, 11–13 in mouse uterine/oviduct/follicular fluids, 13 Leber’s hereditary optic neuropathy (LHON), 97, 123, 138–139 cybrid modeling and, 161–162 OXPHOS functions and, 165 Leigh’s disease, 162 LHON. See Leber’s hereditary optic neuropathy Light strand promoter (LSP), 56

296 Low success rates in NT, potential reasons for mtDNA D-loop and, 275 NRF-1 and, 276 nucleo-mitochondrial compatibility and, 274 TFAM and, 275–276 LoxP-Cre-mediated excision, TFAM and, 271 LSP. See Light strand promoter M Macaca mulatta, 222 Male pronucleus (MPN), 188 paternal genome demethylation in, degree of, 197 Male transmission, of mtDNA fish eggs, 103 mouse, 101–103 sperm ubiquitination, 102–103 uniparental inheritance, 101–103 Mammalian mtDNA homoplasmy, 95 point mutations, 97 structure/organization/ replication/expression heavy (H) strand, 89–91 light (L) strand, 89–91 mtTFA, 89–91 strand-displacement model, 89–91 transcription of, 91 Twinkle helicase, 89, 91 Mammalian oocyte atresia in, 98–99 mtDNA copy number, 89, 99–101 PCR eVciency, determination of, 101 real-time PCR, 100–101 mtDNA deletion, 97–98 mtDNA in, 87, 98–101 NARP mutation, 97 OXPHOS, 98–99 Mammals energy production in embryos preimplantation embryos, 31–33 redox and energetic metabolism, 31–33 yolk proteins, 31 mitochondrial activity in embryos, 23 mitochondrial-dependent apoptosis, 41 morpholino approach, for mitochondrial function, 26 mtDNA copy number, 26

Index Maternally inherited Leigh syndrome (MILS), 90, 140 Maternal-embryo transition (MET), 66 Maternal transmission, in mtDNA, 57 Mathematical models of bottleneck, 219–220 MELAS. See Mitochondrial encephalomyopathy with lactic acidosis and stroke-like episodes MERRF. See Mitochondrial encephalomyopathy, and ragged red fibers; Myoclonus epilepsy and ragged red fibers Messenger RNA (mRNA), 120 MET. See Maternal-embryo transition Micromanipulation methods use as prevention tool, transmission of mutated mitochondrial DNA and chromosome group transfer, 198–202 germinal vesicle transfer and, 190–194 nuclear material and asynchronous GV to cytoplast transfer, storage of, 194–198 oocytes and zygotes, 188–189 pronuclei, transfer of, 202–206 technical equipment, 189–190 MILS. See Maternally inherited Leigh syndrome Mitochondria in ascidian embryos, 23 ATP production, 52 autofluorescence, in mouse/sea urchin eggs, 30–31 behavior of, 21 biogenesis, 26 Ca2þ oscillations in ascidian and mouse fertilization, 24, 33–34 Ca2þ signaling of somatic cells, 24, 33 cell survival, role in, 53 cellular energetic metabolism, 52–53 cellular processes, role in, 22 in eggs Ca2þ homeostasis, 33–34 generation/distribution in, 25–27 GFP, 28 heterogenic distribution, 23 in sea urchins, 27 study of, 27–31 embryonic development apoptosis, 24 in mammals, 31 regulation of, 24

Index in embryos Ca2þ homeostasis, 33–34 citrate metabolism, 36 energy production, in mammals, 31–33 generation/distribution in, 25–27 GFP, 28 homoplasmy, 23 mtDNA, 26 oogenesis, 23, 26 oxidant action in sea urchin, 38 oxidative metabolism, 24 preimplantation, 31, 36 redox metabolism, 34 study of, 27–31 ER Ca2þ release, activity of, 24 FADH2, 53 functions in eucaryotic cells, 52 hyperpolarized electrical potential, 27–28 IRP, 22 Krebs cycle intermediates, synthesis of, 35 location, in germ plasms, 26–27 in mammalian embryos, 23 MitoTrackers, 28 NADH-dehydrogenase complex, 53 nutrient catabolism, 53 OXPHOS, 52 patterning in mouse, 27 pleiotropic properties, 53 potentiometric dyes, use of, 27–28 production during oogenesis, 25 production in germ cells, 25 proteins, 26 ROS, 22, 53 segregation in germ cells, 23 succinate-dehydrogenase complex, 52–53 Mitochondria, in sperm function metabolic pathways glycolytic process, 9–11 OXPHOS system, 6–9 m, 6–7 Mitochondria, structure and function of ADP, 115 ATP production and OXPHOS process, 114–116 endosymbiosis, 114 in eukaryotic cells, 114 oxidoreduction reactions, 115–116 Mitochondrial ATP, 9, 33–34, 37–38, 52, 115

297 Mitochondrial biogenesis, 87 in fertility, role of, 60 spermatogenesis, 63 Mitochondrial bottleneck animal models of, polymorphic mtDNA variants and, 218–219 heteroplasmic consequence and, 217 IVF and, 217–218 Kearn–Sayre syndrome and, 218 mathematical models of, 219–220 neutral polymorphisms and, 217 in normal oocytes, 217 Mitochondrial cardiomyopathy, 137 Mitochondrial copy number, deletions, and mutations 4977-bp deletion (common deletion) and Rhesus macaque, 241 CPEO syndrome and, 241 folliculogenesis and, 240 gonadotrophin stimulation and, 241 GV and, 241–242 Kearns–Sayre syndrome (KSS) and, 241 mammalian oocyte and, 239 OXPHOS and, 240 Mitochondrial diseases, 113–114 aging, in mouse, 128–129 cancer, 127 diabetes and hypertension, 124, 128 epidemiology. See Mitochondrial diseases, impact of etiology. See Mitochondrial diseases, etiology impact of. See Mitochondrial diseases, impact of mitochondrial genetics bottleneck concept, 118–120 heteroplasmy, 117–118 homoplasmy, 117–118 mtDNA replication, 120–123 segregation, 118–120 transcription, 120–123 translation, 120–123 transmission, 118–120 paternal inheritance, 118 polyploidy, 116–117 threshold, 118 neurodegenerative diseases AD, 127 PD, 126

298 Mitochondrial diseases (cont.) organ-specific Leber’s hereditary optic neuropathy, 138–139 mitochondrial cardiomyopathy, 137 mitochondrial myopathy, 138 sensorineural deafness, 139 Mitochondrial diseases, etiology causes of problems nuclear control, 136–137 variable genome, 134–135 variable phenotype, 135–136 diagnosis of clinical assessment, 129 clinical investigation, 129–130 muscle biopsy, 130–134 pathology childhood-specific mitochondrial syndromes, 141–142 classical mitochondrial syndromes, 139–141 organ-specific mitochondrial diseases, 137–139 Mitochondrial diseases, impact of economic CPEO, 125 m.3243A>G MELAS, 125 epidemiology diabetes mellitus, 124 LHON, 123 m.3243A>G, 124 mitochondrial involvement, 126–129 aging, 128–129 cancer, 127 diabetes and hypertension, 128 neurodegenerative diseases, 126–127 Mitochondrial DNA (mtDNA), 22, 54, 87, 113 proteobacteria, endosymbiosis, 54 bottleneck theory, in oogenesis, 58 clinical application of counseling women, 103–104 ICSI, 104 mutant mtDNAs, 103–104 oocyte cytoplasm, transfer of, 105 preimplantation genetic diagnosis, 104 prenatal genetic diagnosis, 104 content variability in cohorts, 60–61 oocyte, maturity of, 61 and early embryogenesis, transcripts blastocyst, 65–66

Index in bovine embryos, 67–70 cell divisions, 65–70 embryo implantation, 65–66 MET, 66 in mouse embryos, 66–70 NRF1mRNA, 70 TFAM mRNA, 69–70 female germ line, transmission of bottleneck concept, 95–96 homoplasmy, 95 mouse, 96 point mutations, 97 replication, 96 stochastic process, 97–98 germ cells, 87 heavy (H) strand, 89–91 light (L) strand, 89–91 maintenance and expression PGC, 57 replication of, 55 respiratory chain, 56–57 TFAM, 56 transcription of, 56 male transmission of fish eggs, 103 mouse, 101–103 sperm ubiquitination, 102–103 uniparental inheritance, 101–103 mammalian oocyte, 99–101 in mammals structure, organization, replication, expression, 89–92 maternal lineage, 6 maternal transmission of, 57 MELAS syndrome, 8 mitoribosomes, 91 in mouse oocytes, 59 mtTFA, 89 mutations, in oocytes and embryos cell aging, 71–74 free radical theory, 71 haplogroups, 73 heteroplasmy, 71–74 in mouse, 71 mtDNA mutations, 72 POLG, 71–72 in women, 72 nucleoid, 89 oocyte quality, 65 in oogenesis, restriction/amplification of content, 57–59 ovarian insuYciency, 63–65

Index in OXPHOS, 6 POLG, 8–9 polymorphism, 73 replication of Clayton’s model, 122 nDNA, 122 relaxed, 122 replisome, 122 strand displacement model, 122 segregation of, 95–96 in spermiogenesis, 5–6 strand-displacement model, 89–91 structure of, 54 transcription of, 91 polycistronic transcripts, 120–121 translation of, 120–123 Mitochondrial dysfunctions in diabetic women, 25 in mouse eggs, 41 mtDNA mutations, 72 oxidative stress in embryo, 24–25 Mitochondrial encephalomyopathy with lactic acidosis and stroke-like episodes (MELAS), cybrid modeling and, 97, 161–162 Mitochondrial encephalopathy and ragged red fibers (MERRF), 97 Mitochondrial fingerprinting technique, 231 Mitochondrial genetic bottleneck, transmission/segregation and, 118–120 Mitochondrial genetics bottleneck concept, 118–120 heteroplasmy polymorphism in mouse, 118–120 homoplasmy, 117–118 mtDNA replication Clayton’s model, 122 nDNA, 122 relaxed, 122 replisome, 122 strand displacement model, 122 mtDNA segregation, 118–120 mtDNA transcription mouse, 121 polycistronic transcripts, 120–121 TFAM, 121 mtDNA translation, 120–123 mtDNA transmission, 118–120 paternal inheritance, 118

299 polyploidy, 116–117 threshold, 118 Mitochondrial genome, 54 expression of, 55–56 in oocyte, 60 replication of, 55–56 Mitochondrial genome, modifying gene therapy and, 170 protein import pathway and, 171 Mitochondrial genome and biogenesis EGA and, 233 heteroplasmy, potential eVects of, 233 mRNAs, 233–234 nuclear genome and, 234–235 preimplantation embryo development and, 233 replication, 235–237 transcription, 237 Mitochondrial heteroplasmy, implications of ART and, 243–244 in infertility, 243–244 in mitochondrial disease, 244 SCNT and, 244 in stem cell therapeutics, 244 Mitochondrial localization patterns, oocyte and embryo microtubule distribution and, 243 nuclear-encoded regulatory factors, diVusion of, 243 nuclear-mitochondrial signaling and, 242 pronuclear accumulation and, 242 Mitochondrial membrane potential ( m) in OXPHOS, sperm motility and, 7 Mitochondrial myopathy, 138–139 Mitochondrial neurogastrointestinal encephalopathy (MNGIE) mitochondrial nuclear respiratory factors NRF1 and NRF2, 56–57, 140 Mitochondrial number, PGC, 94 Mitochondrial replication A467T and, 235 POLG and, 235–237 Twinkle, DNA helicase and, 235–237 Mitochondrial ribosomal proteins (MRP), 121 Mitochondrial single-stranded DNA-binding protein (mtSSB) Drosophila embryos and, 239 mitochondrial replication and, 235–237

300 Mitochondrial transcription D-loop and, 237 heavy strand promoter (HSP) and, 236–237 light strand promoter (LSP) and, 236–237 TFAM and, 236–237 TFB1M and, 236–237 TFB2M and, 236–237 Mitochondrial transcription factor A (TFAM), 56, 121, 238, 275–276 encephalomyopathy disorders and, 271 ethidium bromide and, 272 HMG1, 271 HMG2, 271 LoxP-Cre-mediated excision and, 271 mitochondrial replication and, 236–237 mitochondrial transcription and, 236–237 in mouse, 70 mtDNA transcription and replication, 269–271 NRF-1 and, 272 TFB1M, 272 TFB2M, 272 Mitoribosomes, 91 MitoTrackerTM Green, 172, 205–206 in mitochondria, 28 Red (CM-H2XRos), 172 MNGIE. See Mitochondrial neurogastrointestinal encephalopathy Molecular beacon analysis, 231 Molecular genetics, in muscle biopsy, 132–134 Monozygotic twins, 252 Morpholino approach, in mitochondrial function, mammals, 26 Mouse aging, mitochondrial disease, 121 Ca2þ dynamics in eggs, 34 heteroplasmic polymorphism, 118–120 male transmission, of mtDNA, 101–103 mitochondrial autofluorescence in eggs, 30–31 mitochondrial dysfunction in eggs, 41 mitochondrial enzymes, 36 mitochondrial GFP, 28 mitochondrial patterning, 27

Index mtDNA, 26 content and embryogenesis, 66–70 content in oocytes, 59 copy number, 25–26 mutations, 71 transcription, 121 transmission, female germ line, 96 oogonia, 93 ovarian stimulation, 73 PGC, in embryos, 92 preimplantation embryos, 36 preimplantation genetic diagnosis, 104 redox metabolism, 37 TFAM, 56> Mouse cell mtDNA mutants and cybrids CAPR and, 165 OXPHOS complexes and, 165–167 R6G treatment and, 167 xenomitochondrial cybrids, 167–169 Drosophila and, 168 0 cells and, 168 PCR detection and, 168 Mouse spermatozoa glycolysis, GAPDH, 9 IVF assays, 8 OXPHOS, sperm motility in POLG, 6–9 POLG, 8–9 MPN. See Male pronucleus mRNA. See Messenger RNA MRP. See Mitochondrial ribosomal proteins mtDNA. See Mitochondrial DNA mtDNA, maternal inheritance and ICSI procedure and, 215 IVF and, 214 Klinefelter’s syndrome and, 215 myogenic lineage and, 215 mtDNA copy number, 89, 96 in bovine embryos, 26 mammalian oocytes PCR eYciency, determination of, 101 real-time PCR, 100–101 in mammals, 26 in mouse, 25–26 mtDNA copy number, NT failure and ATP and, 267 fertilization rates and, 267 mtDNA disease, CVS and, 216–217

Index mtDNA disease in preimplantation embryo, genetic management of chorionic villus sampling (CVS), heteroplasmy of mtDNA and consequences of, dosage and thresholds and, 214 mtDNA disease and, 216–217 distribution, heteroplasmic embryos and, 215–216 maternal inheritance and, 214–215 CVS, management options and nuclear cell transfer, 221–222 oocytes and pedigree analysis, preconception counseling and, 220 preimplantation genetic diagnosis, 220–221 mitochondrial bottleneck, 217 animal models of, polymorphic mtDNA variants and, 218–219 mathematical models of, 219–220 mtDNA distribution, heteroplasmic embryos and A3243G and, 216 CVS, implications for, 215–216 mtDNA homoplasmy, in mammals, 95 mtDNA injection versus ES cell-derived models, 171 heteroplasmic mice, generation of, 172 MitoTrackerTM Green, 172 Red (CM-H2XRos), 172 TMRM and, 173 in vivo. Doublelabeling experiments and, 172 mtDNA replication and NT failure, 267 blastocyst stage and, 268 coupled leading-and lagging-strand synthesis, 270–271 D-loop and, 269 POLG, 269–273 mtDNA segregation bottleneck concept, 95–96 population genetic model, 96 mtDNA transcription and NT failure, 267 spindle formation and, 268 TFAM and, 269–272 TFB1M, 269

301 mtDNA transmission female germ line bottleneck concept, 95–96 homoplasmy, 95 in mouse, 96 replication, 96 segregation, 96–97 stochastic process, 97–98 male in fish eggs, 101–103 in mouse, 101–103 sperm ubiquitination, 102–103 uniparental inheritance, 101–103 mtDNA transmission, animal modeling of, from transgenesis to transmitochondrial cybrid models and human disease embryonic cytoplast and karyoplast fusion, 173–175 mitochondrial genome, modifying, 170–171 modeling rationale, 169 spontaneous and induced models ATPase6 gene, 170 BHE/Cdb rat, salt-sensitive, 170 transfected 0 cells use as intermediate mitochondrial carriers, mouse models production and, 171 transmitochondrial mice mtDNA injection versus ES cell-derived models, 171–173 xenomitochondrial mice and, 175–176 mtDNA transmission and NT failure cytoplasmic transfer (CT) and, 261–263 heteroplasmic, 261–266 homoplasmic, 261 OXPHOS and, 263 PCR and, 266 variability in, 261 mtDNA variation, early embryogenesis, transcripts in, 65–70 mtSSB. See Mitochondrial single-stranded DNA-binding proten mtTFA, 89 Muscle biopsy biochemical analysis, 131–132 histology and histochemistry, 130–131 molecular genetics, 132–134 Mus spretus, 168

302 Mutations, in mtDNA, oocytes/embryos, 71–74 cell aging, 71–74 free radical theory of aging, 71 haplogroups, 73 heteroplasmy, 71 in mouse, 71 ovarian stimulation, 72 POLG, 71–72 Myoclonic epilepsy with ragged red fibers (MERRF), 140 cybrid modeling and, 161

N NADPH. See Nicotinamide dinucleotide NARP. See Neurogenic weakness, ataxia, and retinitis pigmentosa nDNA. See Nuclear DNA Neurodegenerative diseases, mitochondria involvement AD, 127 PD, 126 Neurogenic weakness, ataxia, and retinitis pigmentosa (NARP), 97, 140, 216 ATP6 gene in, 161 Leigh’s disease, 162 Nicotinamide dinucleotide (NADH), in mitochondria, 52–53 NO, respiratory complex, in sperm motility, 7 Normozoospermic individuals, 6 Norway rat (Rattus norvegicus), 168 NRF-1. See Nuclear respiratory factor NRF1mRNA, 70 NT. See Nuclear transfer NT failure, possible causes of cell cycle, 258 epigenetics, 258–260 haplotype and genetic diversity, 266–267 mtDNA copy number, 267 transcription and replication, 267–273 transmission, 261–266 pluripotent and epigenetic memory instability, 260 Nuclear cell transfer Macaca mulatta and, 222 stem cell therapy and, 221

Index Nuclear DNA (nDNA), 122 Nuclear material and asynchronous GV to cytoplast transfer, storage of FPN and, 196 GVBD and, 194–197 GV karyoplasts and, 194–195 ICSI and, 196 MPN and, 196 paternal genome demethylation in, degree of, 197 PMSG and, 196–197 thylation-demethylation processes, 197 vitrification and, 194–195 Nuclear respiratory factors 1 (NRF-1), 238–239, 272, 276 Nuclear transfer (NT). See also Cloning adaptations to, 252–254 applications of agricultural applications and, 256 cattle industry and, 256 embryological studies and, 255 epigenetic disorders and, 255 ESC and, 256–257 gene knock in strategies and, 255 genetic manipulation and, 255 IVF and, 255 xenotransplantation and, 255 failure, possible causes of cell cycle, 258 epigenetics, 258–260 haplotype and genetic diversity, 266–267 mtDNA copy number, 267 mtDNA transcription and replication, 267–273 mtDNA transmission and, 261–266 pluripotent and epigenetic memory instability, 260 health problems associated with, 257 HMC and, 253 ICM and, 253 low success rates in, potential reasons for, 274–277 monozygotic twins and, 252 SCNT and, 254 Nucleoids, in mtDNA, 54, 89 O Oct4, 260 Oligomycin, respiratory complex, in sperm motility, 7

303

Index Oligozoospermic individuals, 6 Oncosis, in embryonic development, 38 Oocytes aging, 24, 40, 98–99 atresia, 59, 88, 98–99 fertilization and mtDNA, mitochondrial content in spermatogenesis, 63 women, 61–63 in mammals atresia, 98–99 germ line, 93–94 NARP mutation, 97 OXPHOS in, 98–99 real-time PCR, 100–101 maturation of, 61 mitochondrial genomes, 60 mtDNA, in mammals copy number, 99–101 deletion, 97–98 mtDNA content, 54 largest cellular content of organism, 59–60 quality of, 65 variability in, 60–61 mutations in, embryos and women, 72 oogenesis, restriction/amplification of content, in mtDNA apoptosis, 59 bottleneck theory, 58 follicle cells, 57–58 in germ cells, 57–59 oocytic atresia, 59 point mutations, 58 ovarian insuYciency, mtDNA depletion ovarian dystrophy, 63–64 POLG1, 64–65 quality, 98–99 Oocytes and pedigree analysis, preconception counseling and asymptomatic siblings and, 220 IVF methods and, 220 Oocytes and zygotes, biological material gonadotropin stimuli and, 188 GVBD and, 188 GV-oocyte nucleus and, 188 Kaguya, parthenogenetic mouse and, 188 metaphase stages and, 188–189 methylation–demethylation processes in, 197

Oocytic atresia, in oogenesis, 59 Oogenesis apoptosis, 40–41 germ line mutations, 98–99 mitochondrial production, 25–26 in oocytes bottleneck theory, 58 oocytic atresia, 59 point mutations, elimination of, 58 oocytic atresia, 59 Oogonia, 57–58, 93–96 mouse, 93 Organ-specific mitochondrial diseases Leber’s hereditary optic neuropathy, 138–139 mitochondrial cardiomyopathy, 137 mitochondrial myopathy, 138–139 sensorineural deafness, 139 Ovarian dystrophy, 63–64 Ovarian insuYciency, mtDNA depletion, in oocytes POLG1, 64–65 in three groups of women, 63–65 Ovarian stimulation, in mouse, 73 Oxidative phosphorylation (OXPHOS), 3, 52, 88, 157, 238, 240 ATP production, 9 biochemical analysis, 7 biogenesis, 6 complexes Cybrids, mtDNA disease and, 158–159 mouse cell mtDNA mutants and cybrids, 165–167 0 cells and, 160–161 functions, LHON cybrids and, 165 in mitochondria, 114–116 mtDNA transmission and, 263 in oocyte, 98–99 respiratory complexes, 7 spermiogenesis, 5 sperm mitochondria, 6–9 sperm motility, supply of energy in mouse, 7 normozoospermic, 6 oligozoospermic, 6 m, 7 sperm quality, 8 subunits cytochrome c, 5 VIb-2, 5

304 Oxidoreduction reactions, in mitochondria, 115–116 OXPHOS. See Oxidative phosphorylation P Parkinson’s disease (PD), 126, 163–164, 256 Pathology, 137–142 PD. See Parkinson’s disease PDH. See Pyruvate dehydrogenase PEO. See Progressive external opthalmoplegia PGC. See Primordial germ cells Pgc-1-related coactivator (PRC), 57 PGD. See Preimplantation genetic diagnosis Piezo injector, 189, 191 Pluripotent instability and NT failure, 260 PMSG, 196, 198 Point mutations in mammalian oocytes A3243G (MELAS), 97 A8344G (MERRF), 97 G3460A (LHON), 97 G11778A (LHON), 97 T8993C (NARP), 97 T8993G (NARP), 97 mtDNA in, 71–74 oogenesis, elimination of, 58 POLG, 71–72. See also Polymerase ; Proofreading-deficient version of polymerase gamma PolgA, 17, 128–129 Pol-scope microscope, 200 Polyethylene glycol (PEG)-induced fusion, 190, 203 Polygenic disorders and tissue-specificity mtDNA contributions to Alzheimer’s disease (AD), 163–164 COX, decreased activity of, 164 disease-associated mtDNA variants, identification of, 163–164 0 SH-SY5Y cells, 164 Parkinson’s disease (PD), 163–164 rhodamine 6G (R6G), 164 Polymerase (POLG), 269–272 mitochondrial replication and, 235–237 PEO and, 273 preimplantation embryos, molecular control of mitochondrial function in, 238–239

Index subunits of, 235 transgenic mouse model and, 273 Polymorphic mtDNA variants, animal models of bottleneck and, 218–219 Polymorphism, mtDNA in, 73 Polyploidy, of mtDNA, 116–117 Potentiometric dyes, in mitochondrial imaging, use of, 27–28 PRC. See Pgc-1-related coactivator Preimplantation embryos in mammals, 31–33 molecular control of mitochondrial function in, 237 -galactosidase-neomycin cassette, 239 mtSSB in Drosophila embryos and, 239 NRF-1 and, 238–239 OXPHOS and, 238 POLG and, 238–239 TFAM and, 238 TFB1 and, 238–239 Preimplantation genetic diagnosis (PGD) blastomeres and, 221 CVS and, 221 IVF and, 220 Primordial germ cells (PGC), 92, 119 mitochondrial number, 94 Progressive external opthalmoplegia (PEO), 97, 273 Pronuclei transfer, mouse zygotes and cytochalasin and, 203 cytoplasmic bridge and, 203 heteroplasmy and, 205 mitochondrial dysfunction and, 204, 206 MitoTracker Green FM and, 205–206 mutated mtDNA, elimination of, 205 Sendai virus and, 203 Proofreading-deficient version of polymerase gamma (POLG), in mtDNA, 6–9, 64–65, 71–72 Pyruvate dehydrogenase (PDH), in sperm motility, 13

R Rag2 mutation, 256 Rattus norvegicus. See Norway rat Reactive oxygen species (ROS), 53, 128 in mitochondria, 22 Real-time PCR, in mammalian oocytes, 99–101

Index Redox metabolism, in embryo citrate metabolism, 36 glutathione (GSH/GSSG), 35 IRP, 35 Krebs cycle intermediates, synthesis of, 36 in mouse, 37 NADH/NADþ and NADPH/NADPþ, 35 oxidant action in sea urchin, 38 oxidative metabolism, 37 pyruvate metabolism, 36–37 Relaxed replication, in mtDNA, 122 Replisome, in mtDNA replication, 122 Respiratory complexes, in ETC, OXPHOS system antimycin A, 7 gossypol, 7 NO, 7 oligomycin, 7 rotenone, 7 Respiratory substrate, lactate in female vagina, 12–13 in mouse, 13 R6G. See Rhodamine 6G Rhesus macaque, 264 Rhodamine 6G (R6G), 158, 167 embryonic cytoplast and karyoplast fusion, 174 polygenic disorders and tissue-specificity, 164 Ribosomal RNA (rRNA), 120 ROS. See Reactive oxygen species Rotenone, respiratory complex, in sperm motility, 7 rRNA. See Ribosomal RNA

S SCNT. See Somatic cell nuclear transfer Sea urchins mitochondrial autofluoresence, 30–31 mitochondrial Ca2þ dynamics, 34 mitochondrial density, 27 oxidant action, in mitochondria, 38 Sendai virus, 190 cybrids, mtDNA disease, 158 pronuclei transfer and, 203 Sensorineural deafness, 139 S-glutathionylation, in embryonic development, 39–40 Single nucleotide polymorphism (SNP), 73 Single-strand-binding protein (SSB), 70 SNP. See Single nuceotide polymorphism

305 Somatic cell nuclear transfer (SCNT), 244, 254 embryos pluripotent instability and NT failure, 260 epigenetics, 259 Sperm quality, in OXPHOS, 8 structure, 4–5 ubiquitination, 102–103 Spermatids, substrates in, sperm motility and, 12 Spermatogenesis, mitochondrial biogenesis in, 63 Spermiogenesis COX activity, 5 mtDNA, 5–6 OXPHOS cytochrome c, 5 VIb-2, 5 sperm mitochondria, 5–6 sperm structure, 4–5 Sperm mitochondria, in functional purposes glycolysis, 9–11 lactate dehydrogenase C4 (LDH-C4), 13 OXPHOS, 6–9 spermiogenesis, 5–6 Sperm motility gluconeogenesis, 10–11 glycolysis GAPDH/, 9–10 G3P, 10 in mouse, 9–11 glycolytic susbtrates, 11–13 lactate, 12–13 LDH-C4 and, 13 malate, 13 mtDNA mutations, 8 OXPHOS biogenesis, 6 in mouse spermatozoa, 7 oxygen consumption and, 7 respiratory complexes, 7 m, 7 PDH, 13 respiratory susbtrates, 11–13 spermatids, 12 SSB. See Single-strand-binding protein Strand displacement model, mtDNA, 89–91 replication, 122

306 Substrates, energy suppliers, in sperm motility in female reproductive tract, 11–13 glycolytic substrates malate, 13 spermatids, 12 ketones, 13 oxygen levels, 11–13 respiratory substrates lactate, 12–13 T Tay-Sachs disease, 256 Technical equipment, micromanipulation methods and cytoplast and, 189–190 enucleation and, 189–190 ICSI pipette and, 190 inverted microscope, 189 karyoplasts and, 190 piezo injector, 189 polyethylene glycol (PEG)-induced fusion, 190 zona pellucida and, 189 Tetramethylrhodamine methyl ester (TMRM), 173 TFAM. See Mitochondrial transcription factor A TFAM mRNA, 69–70 TFB1M, 237, 239, 269–270, 272

Index TFB2M, 237, 239, 269–270, 272 Tissue-nonspecific alkaline phosphatase (TNAP), 92 TMRM. See Tetramethylrhodamine methyl ester TNAP. See Tissue-nonspecific alkaline phosphatase Turner syndrome, 263 Twinkle, DNA helicase, 239 mitochondrial replication and, 235–237 mtDNA copy number and, 236 Tyrode solution, acidified, 191

U Untranslated regions (UTR), 54 UTR. See Untranslated regions

X Xenomitochondrial cybrids, 167–169 chimeric mice and, 176 Drosophila and, 168 0 cells and, 168 muridae species and, 176 PCR detection and, 168 transmitochondrial animal models and, 175 Xenotransplantation, NT applications and, 255

Contents of Previous Volumes Volume 47 1 Early Events of Somitogenesis in Higher Vertebrates: Allocation of Precursor Cells during Gastrulation and the Organization of a Moristic Pattern in the Paraxial Mesoderm Patrick P. L. Tam, Devorah Goldman, Anne Camus, and Gary C. Shoenwolf

2 Retrospective Tracing of the Developmental Lineage of the Mouse Myotome Sophie Eloy-Trinquet, Luc Mathis, and Jean-Franc¸ois Nicolas

3 Segmentation of the Paraxial Mesoderm and Vertebrate Somitogenesis Olivier Pourqule´

4 Segmentation: A View from the Border Claudio D. Stern and Daniel Vasiliauskas

5 Genetic Regulation of Somite Formation Alan Rawls, Jeanne Wilson-Rawls, and Eric N. Olsen

6 Hox Genes and the Global Patterning of the Somitic Mesoderm Ann Campbell Burke

7 The Origin and Morphogenesis of Amphibian Somites Ray Keller

8 Somitogenesis in Zebrafish Scott A. Halley and Christiana Nu¨sslain-Volhard

9 Rostrocaudal Differences within the Somites Confer Segmental Pattern to Trunk Neural Crest Migration Marianne Bronner-Fraser

307

308

Contents of Previous Volumes

Volume 48 1 Evolution and Development of Distinct Cell Lineages Derived from Somites Beate Brand-Saberi and Bodo Christ

2 Duality of Molecular Signaling Involved in Vertebral Chondrogenesis Anne-He´le`ne Monsoro-Burq and Nicole Le Douarin

3 Sclerotome Induction and Differentiation Jennifer L. Docker

4 Genetics of Muscle Determination and Development Hans-Henning Arnold and Thomas Braun

5 Multiple Tissue Interactions and Signal Transduction Pathways Control Somite Myogenesis Anne-Gae¨lle Borycki and Charles P. Emerson, Jr.

6 The Birth of Muscle Progenitor Cells in the Mouse: Spatiotemporal Considerations Shahragim Tajbakhsh and Margaret Buckingham

7 Mouse–Chick Chimera: An Experimental System for Study of Somite Development Josiane Fontaine-Pe´rus

8 Transcriptional Regulation during Somitogenesis Dennis Summerbell and Peter W. J. Rigby

9 Determination and Morphogenesis in Myogenic Progenitor Cells: An Experimental Embryological Approach Charles P. Ordahl, Brian A. Williams, and Wilfred Denetclaw

Volume 49 1 The Centrosome and Parthenogenesis Thomas Ku¨ntziger and Michel Bornens

2 g-Tubulin Berl R. Oakley

Contents of Previous Volumes

309

3 g-Tubulin Complexes and Their Role in Microtubule Nucleation Ruwanthi N. Gunawardane, Sofia B. Lizarraga, Christiane Wiese, Andrew Wilde, and Yixian Zheng

4 g-Tubulin of Budding Yeast Jackie Vogel and Michael Snyder

5 The Spindle Pole Body of Saccharomyces cerevisiae: Architecture and Assembly of the Core Components Susan E. Francis and Trisha N. Davis

6 The Microtubule Organizing Centers of Schizosaccharomyces pombe Iain M. Hagan and Janni Petersen

7 Comparative Structural, Molecular, and Functional Aspects of the Dictyostelium discoideum Centrosome Ralph Gra¨f, Nicole Brusis, Christine Daunderer, Ursula Euteneuer, Andrea Hestermann, Manfred Schliwa, and Masahiro Ueda

8 Are There Nucleic Acids in the Centrosome? Wallace F. Marshall and Joel L. Rosenbaum

9 Basal Bodies and Centrioles: Their Function and Structure Andrea M. Preble, Thomas M. Giddings, Jr., and Susan K. Dutcher

10 Centriole Duplication and Maturation in Animal Cells B. M. H. Lange, A. J. Faragher, P. March, and K. Gull

11 Centrosome Replication in Somatic Cells: The Significance of the G1 Phase Ron Balczon

12 The Coordination of Centrosome Reproduction with Nuclear Events during the Cell Cycle Greenfield Sluder and Edward H. Hinchcliffe

13 Regulating Centrosomes by Protein Phosphorylation Andrew M. Fry, Thibault Mayor, and Erich A. Nigg

14 The Role of the Centrosome in the Development of Malignant Tumors Wilma L. Lingle and Jeffrey L. Salisbury

15 The Centrosome-Associated Aurora/IpI-like Kinase Family T. M. Goepfert and B. R. Brinkley

310

Contents of Previous Volumes

16 Centrosome Reduction during Mammalian Spermiogenesis G. Manandhar, C. Simerly, and G. Schatten

17 The Centrosome of the Early C. elegans Embryo: Inheritance, Assembly, Replication, and Developmental Roles Kevin F. O’Connell

18 The Centrosome in Drosophila Oocyte Development Timothy L. Megraw and Thomas C. Kaufman

19 The Centrosome in Early Drosophila Embryogenesis W. F. Rothwell and W. Sullivan

20 Centrosome Maturation Robert E. Palazzo, Jacalyn M. Vogel, Bradley J. Schnackenberg, Dawn R. Hull, and Xingyong Wu

Volume 50 1 Patterning the Early Sea Urchin Embryo Charles A. Ettensohn and Hyla C. Sweet

2 Turning Mesoderm into Blood: The Formation of Hematopoietic Stem Cells during Embryogenesis Alan J. Davidson and Leonard I. Zon

3 Mechanisms of Plant Embryo Development Shunong Bai, Lingjing Chen, Mary Alice Yund, and Zinmay Rence Sung

4 Sperm-Mediated Gene Transfer Anthony W. S. Chan, C. Marc Luetjens, and Gerald P. Schatten

5 Gonocyte–Sertoli Cell Interactions during Development of the Neonatal Rodent Testis Joanne M. Orth, William F. Jester, Ling-Hong Li, and Andrew L. Laslett

6 Attributes and Dynamics of the Endoplasmic Reticulum in Mammalian Eggs Douglas Kline

7 Germ Plasm and Molecular Determinants of Germ Cell Fate Douglas W. Houston and Mary Lou King

Contents of Previous Volumes

311

Volume 51 1 Patterning and Lineage Specification in the Amphibian Embryo Agnes P. Chan and Laurence D. Etkin

2 Transcriptional Programs Regulating Vascular Smooth Muscle Cell Development and Differentiation Michael S. Parmacek

3 Myofibroblasts: Molecular Crossdressers Gennyne A. Walker, Ivan A. Guerrero, and Leslie A. Leinwand

4 Checkpoint and DNA-Repair Proteins Are Associated with the Cores of Mammalian Meiotic Chromosomes Madalena Tarsounas and Peter B. Moens

5 Cytoskeletal and Ca2+ Regulation of Hyphal Tip Growth and Initiation Sara Torralba and I. Brent Heath

6 Pattern Formation during C. elegans Vulval Induction Minqin Wang and Paul W. Sternberg

7 A Molecular Clock Involved in Somite Segmentation Miguel Maroto and Olivier Pourquie´

Volume 52 1 Mechanism and Control of Meiotic Recombination Initiation Scott Keeney

2 Osmoregulation and Cell Volume Regulation in the Preimplantation Embryo Jay M. Baltz

3 Cell–Cell Interactions in Vascular Development Diane C. Darland and Patricia A. D’Amore

4 Genetic Regulation of Preimplantation Embryo Survival Carol M. Warner and Carol A. Brenner

312

Contents of Previous Volumes

Volume 53 1 Developmental Roles and Clinical Significance of Hedgehog Signaling Andrew P. McMahon, Philip W. Ingham, and Clifford J. Tabin

2 Genomic Imprinting: Could the Chromatin Structure Be the Driving Force? Andras Paldi

3 Ontogeny of Hematopoiesis: Examining the Emergence of Hematopoietic Cells in the Vertebrate Embryo Jenna L. Galloway and Leonard I. Zon

4 Patterning the Sea Urchin Embryo: Gene Regulatory Networks, Signaling Pathways, and Cellular Interactions Lynne M. Angerer and Robert C. Angerer

Volume 54 1 Membrane Type-Matrix Metalloproteinases (MT-MMP) Stanley Zucker, Duanqing Pei, Jian Cao, and Carlos Lopez-Otin

2 Surface Association of Secreted Matrix Metalloproteinases Rafael Fridman

3 Biochemical Properties and Functions of Membrane-Anchored Metalloprotease-Disintegrin Proteins (ADAMs) J. David Becherer and Carl P. Blobel

4 Shedding of Plasma Membrane Proteins Joaquı´n Arribas and Anna Merlos-Sua´rez

5 Expression of Meprins in Health and Disease Lourdes P. Norman, Gail L. Matters, Jacqueline M. Crisman, and Judith S. Bond

6 Type II Transmembrane Serine Proteases Qingyu Wu

7 DPPIV, Seprase, and Related Serine Peptidases in Multiple Cellular Functions Wen-Tien Chen, Thomas Kelly, and Giulio Ghersi

Contents of Previous Volumes

313

8 The Secretases of Alzheimer’s Disease Michael S. Wolfe

9 Plasminogen Activation at the Cell Surface Vincent Ellis

10 Cell-Surface Cathepsin B: Understanding Its Functional Significance Dora Cavallo-Medved and Bonnie F. Sloane

11 Protease-Activated Receptors Wadie F. Bahou

12 Emmprin (CD147), a Cell Surface Regulator of Matrix Metalloproteinase Production and Function Bryan P. Toole

13 The Evolving Roles of Cell Surface Proteases in Health and Disease: Implications for Developmental, Adaptive, Inflammatory, and Neoplastic Processes Joseph A. Madri

14 Shed Membrane Vesicles and Clustering of Membrane-Bound Proteolytic Enzymes M. Letizia Vittorelli

Volume 55 1 The Dynamics of Chromosome Replication in Yeast Isabelle A. Lucas and M. K. Raghuraman

2 Micromechanical Studies of Mitotic Chromosomes M. G. Poirier and John F. Marko

3 Patterning of the Zebrafish Embryo by Nodal Signals Jennifer O. Liang and Amy L. Rubinstein

4 Folding Chromosomes in Bacteria: Examining the Role of Csp Proteins and Other Small Nucleic Acid-Binding Proteins Nancy Trun and Danielle Johnston

314

Contents of Previous Volumes

Volume 56 1 Selfishness in Moderation: Evolutionary Success of the Yeast Plasmid Soundarapandian Velmurugan, Shwetal Mehta, and Makkuni Jayaram

2 Nongenomic Actions of Androgen in Sertoli Cells William H. Walker

3 Regulation of Chromatin Structure and Gene Activity by Poly(ADP-Ribose) Polymerases Alexei Tulin, Yurli Chinenov, and Allan Spradling

4 Centrosomes and Kinetochores, Who needs ‘Em? The Role of Noncentromeric Chromatin in Spindle Assembly Priya Prakash Budde and Rebecca Heald

5 Modeling Cardiogenesis: The Challenges and Promises of 3D Reconstruction Jeffrey O. Penetcost, Claudio Silva, Maurice Pesticelli, Jr., and Kent L. Thornburg

6 Plasmid and Chromosome Traffic Control: How ParA and ParB Drive Partition Jennifer A. Surtees and Barbara E. Funnell

Volume 57 1 Molecular Conservation and Novelties in Vertebrate Ear Development B. Fritzsch and K. W. Beisel

2 Use of Mouse Genetics for Studying Inner Ear Development Elizabeth Quint and Karen P. Steel

3 Formation of the Outer and Middle Ear, Molecular Mechanisms Moise´s Mallo

4 Molecular Basis of Inner Ear Induction Stephen T. Brown, Kareen Martin, and Andrew K. Groves

5 Molecular Basis of Otic Commitment and Morphogenesis: A Role for Homeodomain-Containing Transcription Factors and Signaling Molecules Eva Bober, Silke Rinkwitz, and Heike Herbrand

Contents of Previous Volumes

315

6 Growth Factors and Early Development of Otic Neurons: Interactions between Intrinsic and Extrinsic Signals Berta Alsina, Fernando Giraldez, and Isabel Varela-Nieto

7 Neurotrophic Factors during Inner Ear Development Ulla Pirvola and Jukka Ylikoski

8 FGF Signaling in Ear Development and Innervation Tracy J. Wright and Suzanne L. Mansour

9 The Roles of Retinoic Acid during Inner Ear Development Raymond Romand

10 Hair Cell Development in Higher Vertebrates Wei-Qiang Gao

11 Cell Adhesion Molecules during Inner Ear and Hair Cell Development, Including Notch and Its Ligands Matthew W. Kelley

12 Genes Controlling the Development of the Zebrafish Inner Ear and Hair Cells Bruce B. Riley

13 Functional Development of Hair Cells Ruth Anne Eatock and Karen M. Hurley

14 The Cell Cycle and the Development and Regeneration of Hair Cells Allen F. Ryan

Volume 58 1 A Role for Endogenous Electric Fields in Wound Healing Richard Nuccitelli

2 The Role of Mitotic Checkpoint in Maintaining Genomic Stability Song-Tao Liu, Jan M. van Deursen, and Tim J. Yen

3 The Regulation of Oocyte Maturation Ekaterina Voronina and Gary M. Wessel

4 Stem Cells: A Promising Source of Pancreatic Islets for Transplantation in Type 1 Diabetes Cale N. Street, Ray V. Rajotte, and Gregory S. Korbutt

316

Contents of Previous Volumes

5 Differentiation Potential of Adipose Derived Adult Stem (ADAS) Cells Jeffrey M. Gimble and Farshid Guilak

Volume 59 1 The Balbiani Body and Germ Cell Determinants: 150 Years Later Malgorzata Kloc, Szczepan Bilinski, and Laurence D. Etkin

2 Fetal–Maternal Interactions: Prenatal Psychobiological Precursors to Adaptive Infant Development Matthew F. S. X. Novak

3 Paradoxical Role of Methyl-CpG-Binding Protein 2 in Rett Syndrome Janine M. LaSalle

4 Genetic Approaches to Analyzing Mitochondrial Outer Membrane Permeability Brett H. Graham and William J. Craigen

5 Mitochondrial Dynamics in Mammals Hsiuchen Chen and David C. Chan

6 Histone Modification in Corepressor Functions Judith K. Davie and Sharon Y. R. Dent

7 Death by Abl: A Matter of Location Jiangyu Zhu and Jean Y. J. Wang

Volume 60 1 Therapeutic Cloning and Tissue Engineering Chester J. Koh and Anthony Atala

2 a-Synuclein: Normal Function and Role in Neurodegenerative Diseases Erin H. Norris, Benoit I. Giasson, and Virginia M.-Y. Lee

3 Structure and Function of Eukaryotic DNA Methyltransferases Taiping Chen and En Li

4 Mechanical Signals as Regulators of Stem Cell Fate Bradley T. Estes, Jeffrey M. Gimble, and Farshid Guilak

Contents of Previous Volumes

317

5 Origins of Mammalian Hematopoiesis: In Vivo Paradigms and In Vitro Models M. William Lensch and George Q. Daley

6 Regulation of Gene Activity and Repression: A Consideration of Unifying Themes Anne C. Ferguson-Smith, Shau-Ping Lin, and Neil Youngson

7 Molecular Basis for the Chloride Channel Activity of Cystic Fibrosis Transmembrane Conductance Regulator and the Consequences of Disease-Causing Mutations Jackie F. Kidd, Ilana Kogan, and Christine E. Bear

Volume 61 1 Hepatic Oval Cells: Helping Redefine a Paradigm in Stem Cell Biology P. N. Newsome, M. A. Hussain, and N. D. Theise

2 Meiotic DNA Replication Randy Strich

3 Pollen Tube Guidance: The Role of Adhesion and Chemotropic Molecules Sunran Kim, Juan Dong, and Elizabeth M. Lord

4 The Biology and Diagnostic Applications of Fetal DNA and RNA in Maternal Plasma Rossa W. K. Chiu and Y. M. Dennis Lo

5 Advances in Tissue Engineering Shulamit Levenberg and Robert Langer

6 Directions in Cell Migration Along the Rostral Migratory Stream: The Pathway for Migration in the Brain Shin-ichi Murase and Alan F. Horwitz

7 Retinoids in Lung Development and Regeneration Malcolm Maden

8 Structural Organization and Functions of the Nucleus in Development, Aging, and Disease Leslie Mounkes and Colin L. Stewart

318

Contents of Previous Volumes

Volume 62 1 Blood Vessel Signals During Development and Beyond Ondine Cleaver

2 HIFs, Hypoxia, and Vascular Development Kelly L. Covello and M. Celeste Simon

3 Blood Vessel Patterning at the Embryonic Midline Kelly A. Hogan and Victoria L. Bautch

4 Wiring the Vascular Circuitry: From Growth Factors to Guidance Cues Lisa D. Urness and Dean Y. Li

5 Vascular Endothelial Growth Factor and Its Receptors in Embryonic Zebrafish Blood Vessel Development Katsutoshi Goishi and Michael Klagsbrun

6 Vascular Extracellular Matrix and Aortic Development Cassandra M. Kelleher, Sean E. McLean, and Robert P. Mecham

7 Genetics in Zebrafish, Mice, and Humans to Dissect Congenital Heart Disease: Insights in the Role of VEGF Diether Lambrechts and Peter Carmeliet

8 Development of Coronary Vessels Mark W. Majesky

9 Identifying Early Vascular Genes Through Gene Trapping in Mouse Embryonic Stem Cells Frank Kuhnert and Heidi Stuhlmann

Volume 63 1 Early Events in the DNA Damage Response Irene Ward and Junjie Chen

2 Afrotherian Origins and Interrelationships: New Views and Future Prospects Terence J. Robinson and Erik R. Seiffert

3 The Role of Antisense Transcription in the Regulation of X-Inactivation Claire Rougeulle and Philip Avner

Contents of Previous Volumes

319

4 The Genetics of Hiding the Corpse: Engulfment and Degradation of Apoptotic Cells in C. elegans and D. melanogaster Zheng Zhou, Paolo M. Mangahas, and Xiaomeng Yu

5 Beginning and Ending an Actin Filament: Control at the Barbed End Sally H. Zigmond

6 Life Extension in the Dwarf Mouse Andrzej Bartke and Holly Brown-Borg

Volume 64 1 Stem/Progenitor Cells in Lung Morphogenesis, Repair, and Regeneration David Warburton, Mary Anne Berberich, and Barbara Driscoll

2 Lessons from a Canine Model of Compensatory Lung Growth Connie C. W. Hsia

3 Airway Glandular Development and Stem Cells Xiaoming Liu, Ryan R. Driskell, and John F. Engelhardt

4 Gene Expression Studies in Lung Development and Lung Stem Cell Biology Thomas J. Mariani and Naftali Kaminski

5 Mechanisms and Regulation of Lung Vascular Development Michelle Haynes Pauling and Thiennu H. Vu

6 The Engineering of Tissues Using Progenitor Cells Nancy L. Parenteau, Lawrence Rosenberg, and Janet Hardin-Young

7 Adult Bone Marrow-Derived Hemangioblasts, Endothelial Cell Progenitors, and EPCs Gina C. Schatteman

8 Synthetic Extracellular Matrices for Tissue Engineering and Regeneration Eduardo A. Silva and David J. Mooney

9 Integrins and Angiogenesis D. G. Stupack and D. A. Cheresh

320

Contents of Previous Volumes

Volume 65 1 Tales of Cannibalism, Suicide, and Murder: Programmed Cell Death in C. elegans Jason M. Kinchen and Michael O. Hengartner

2 From Guts to Brains: Using Zebrafish Genetics to Understand the Innards of Organogenesis Carsten Stuckenholz, Paul E. Ulanch, and Nathan Bahary

3 Synaptic Vesicle Docking: A Putative Role for the Munc18/Sec1 Protein Family Robby M. Weimer and Janet E. Richmond

4 ATP-Dependent Chromatin Remodeling Corey L. Smith and Craig L. Peterson

5 Self-Destruct Programs in the Processes of Developing Neurons David Shepherd and V. Hugh Perry

6 Multiple Roles of Vascular Endothelial Growth Factor (VEGF) in Skeletal Development, Growth, and Repair Elazar Zelzer and Bjorn R. Olsen

7 G-Protein Coupled Receptors and Calcium Signaling in Development Geoffrey E. Woodard and Juan A. Rosado

8 Differential Functions of 14-3-3 Isoforms in Vertebrate Development Anthony J. Muslin and Jeffrey M. C. Lau

9 Zebrafish Notochordal Basement Membrane: Signaling and Structure Annabelle Scott and Derek L. Stemple

10 Sonic Hedgehog Signaling and the Developing Tooth Martyn T. Cobourne and Paul T. Sharpe

Volume 66 1 Stepwise Commitment from Embryonic Stem to Hematopoietic and Endothelial Cells Changwon Park, Jesse J. Lugus, and Kyunghee Choi

Contents of Previous Volumes

321

2 Fibroblast Growth Factor Signaling and the Function and Assembly of Basement Membranes Peter Lonai

3 TGF- Superfamily and Mouse Craniofacial Development: Interplay of Morphogenetic Proteins and Receptor Signaling Controls Normal Formation of the Face Marek Dudas and Vesa Kaartinen

4 The Colors of Autumn Leaves as Symptoms of Cellular Recycling and Defenses Against Environmental Stresses Helen J. Ougham, Phillip Morris, and Howard Thomas

5 Extracellular Proteases: Biological and Behavioral Roles in the Mammalian Central Nervous System Yan Zhang, Kostas Pothakos, and Styliana-Anna (Stella) Tsirka

6 The Genetic Architecture of House Fly Mating Behavior Lisa M. Meffert and Kara L. Hagenbuch

7 Phototropins, Other Photoreceptors, and Associated Signaling: The Lead and Supporting Cast in the Control of Plant Movement Responses Bethany B. Stone, C. Alex Esmon, and Emmanuel Liscum

8 Evolving Concepts in Bone Tissue Engineering Catherine M. Cowan, Chia Soo, Kang Ting, and Benjamin Wu

9 Cranial Suture Biology Kelly A Lenton, Randall P. Nacamuli, Derrick C. Wan, Jill A. Helms, and Michael T. Longaker

Volume 67 1 Deer Antlers as a Model of Mammalian Regeneration Joanna Price, Corrine Faucheux, and Steve Allen

2 The Molecular and Genetic Control of Leaf Senescence and Longevity in Arabidopsis Pyung Ok Lim and Hong Gil Nam

3 Cripto-1: An Oncofetal Gene with Many Faces Caterina Bianco, Luigi Strizzi, Nicola Normanno, Nadia Khan, and David S. Salomon

322

Contents of Previous Volumes

4 Programmed Cell Death in Plant Embryogenesis Peter V. Bozhkov, Lada H. Filonova, and Maria F. Suarez

5 Physiological Roles of Aquaporins in the Choroid Plexus Daniela Boassa and Andrea J. Yool

6 Control of Food Intake Through Regulation of cAMP Allan Z. Zhao

7 Factors Affecting Male Song Evolution in Drosophila montana Anneli Hoikkala, Kirsten Klappert, and Dominique Mazzi

8 Prostanoids and Phosphodiesterase Inhibitors in Experimental Pulmonary Hypertension Ralph Theo Schermuly, Hossein Ardeschir Ghofrani, and Norbert Weissmann

9 14-3-3 Protein Signaling in Development and Growth Factor Responses Daniel Thomas, Mark Guthridge, Jo Woodcock, and Angel Lopez

10 Skeletal Stem Cells in Regenerative Medicine Wataru Sonoyama, Carolyn Coppe, Stan Gronthos, and Songtao Shi

Volume 68 1 Prolactin and Growth Hormone Signaling Beverly Chilton and Aveline Hewetson

2 Alterations in cAMP-Mediated Signaling and Their Role in the Pathophysiology of Dilated Cardiomyopathy Matthew A. Movsesian and Michael R. Bristow

3 Corpus Luteum Development: Lessons from Genetic Models in Mice Anne Bachelot and Nadine Binart

4 Comparative Developmental Biology of the Mammalian Uterus Thomas E. Spencer, Kanako Hayashi, Jianbo Hu, and Karen D. Carpenter

5 Sarcopenia of Aging and Its Metabolic Impact Helen Karakelides and K. Sreekumaran Nair

6 Chemokine Receptor CXCR3: An Unexpected Enigma Liping Liu, Melissa K. Callahan, DeRen Huang, and Richard M. Ransohoff

Contents of Previous Volumes

323

7 Assembly and Signaling of Adhesion Complexes Jorge L. Sepulveda, Vasiliki Gkretsi, and Chuanyue Wu

8 Signaling Mechanisms of Higher Plant Photoreceptors: A Structure-Function Perspective Haiyang Wang

9 Initial Failure in Myoblast Transplantation Therapy Has Led the Way Toward the Isolation of Muscle Stem Cells: Potential for Tissue Regeneration Kenneth Urish, Yasunari Kanda, and Johnny Huard

10 Role of 14-3-3 Proteins in Eukaryotic Signaling and Development Dawn L. Darling, Jessica Yingling, and Anthony Wynshaw-Boris

Volume 69 1 Flipping Coins in the Fly Retina Tamara Mikeladze-Dvali, Claude Desplan, and Daniela Pistillo

2 Unraveling the Molecular Pathways That Regulate Early Telencephalon Development Jean M. He´bert

3 Glia–Neuron Interactions in Nervous System Function and Development Shai Shaham

4 The Novel Roles of Glial Cells Revisited: The Contribution of Radial Glia and Astrocytes to Neurogenesis Tetsuji Mori, Annalisa Buffo, and Magdalena Go¨tz

5 Classical Embryological Studies and Modern Genetic Analysis of Midbrain and Cerebellum Development Mark Zervas, Sandra Blaess, and Alexandra L. Joyner

6 Brain Development and Susceptibility to Damage; Ion Levels and Movements Maria Erecinska, Shobha Cherian, and Ian A. Silver

7 Thinking about Visual Behavior; Learning about Photoreceptor Function Kwang-Min Choe and Thomas R. Clandinin

324

Contents of Previous Volumes

8 Critical Period Mechanisms in Developing Visual Cortex Takao K. Hensch

9 Brawn for Brains: The Role of MEF2 Proteins in the Developing Nervous System Aryaman K. Shalizi and Azad Bonni

10 Mechanisms of Axon Guidance in the Developing Nervous System Ce´line Plachez and Linda J. Richards

Volume 70 1 Magnetic Resonance Imaging: Utility as a Molecular Imaging Modality James P. Basilion, Susan Yeon, and Rene´ Botnar

2 Magnetic Resonance Imaging Contrast Agents in the Study of Development Angelique Louie

3 1H/19F Magnetic Resonance Molecular Imaging with Perfluorocarbon Nanoparticles Gregory M. Lanza, Patrick M. Winter, Anne M. Neubauer, Shelton D. Caruthers, Franklin D. Hockett, and Samuel A. Wickline

4 Loss of Cell Ion Homeostasis and Cell Viability in the Brain: What Sodium MRI Can Tell Us Fernando E. Boada, George LaVerde, Charles Jungreis, Edwin Nemoto, Costin Tanase, and Ileana Hancu

5 Quantum Dot Surfaces for Use In Vivo and In Vitro Byron Ballou

6 In Vivo Cell Biology of Cancer Cells Visualized with Fluorescent Proteins Robert M. Hoffman

7 Modulation of Tracer Accumulation in Malignant Tumors: Gene Expression, Gene Transfer, and Phage Display Uwe Haberkorn

8 Amyloid Imaging: From Benchtop to Bedside Chungying Wu, Victor W. Pike, and Yanming Wang

9 In Vivo Imaging of Autoimmune Disease in Model Systems Eric T. Ahrens and Penelope A. Morel

Contents of Previous Volumes

325

Volume 71 1 The Choroid Plexus-Cerebrospinal Fluid System: From Development to Aging Zoran B. Redzic, Jane E. Preston, John A. Duncan, Adam Chodobski, and Joanna Szmydynger-Chodobska

2 Zebrafish Genetics and Formation of Embryonic Vasculature Tao P. Zhong

3 Leaf Senescence: Signals, Execution, and Regulation Yongfeng Guo and Susheng Gan

4 Muscle Stem Cells and Regenerative Myogenesis Iain W. McKinnell, Gianni Parise, and Michael A. Rudnicki

5 Gene Regulation in Spermatogenesis James A. MacLean II and Miles F. Wilkinson

6 Modeling Age-Related Diseases in Drosophila: Can this Fly? Kinga Michno, Diana van de Hoef, Hong Wu, and Gabrielle L. Boulianne

7 Cell Death and Organ Development in Plants Hilary J. Rogers

8 The Blood-Testis Barrier: Its Biology, Regulation, and Physiological Role in Spermatogenesis Ching-Hang Wong and C. Yan Cheng

9 Angiogenic Factors in the Pathogenesis of Preeclampsia Hai-Tao Yuan, David Haig, and S. Ananth Karumanchi

Volume 72 1 Defending the Zygote: Search for the Ancestral Animal Block to Polyspermy Julian L. Wong and Gary M. Wessel

2 Dishevelled: A Mobile Scaffold Catalyzing Development Craig C. Malbon and Hsien-yu Wang

3 Sensory Organs: Making and Breaking the Pre-Placodal Region Andrew P. Bailey and Andrea Streit

326

Contents of Previous Volumes

4 Regulation of Hepatocyte Cell Cycle Progression and Differentiation by Type I Collagen Structure Linda K. Hansen, Joshua Wilhelm, and John T. Fassett

5 Engineering Stem Cells into Organs: Topobiological Transformations Demonstrated by Beak, Feather, and Other Ectodermal Organ Morphogenesis Cheng-Ming Chuong, Ping Wu, Maksim Plikus, Ting-Xin Jiang, and Randall Bruce Widelitz

6 Fur Seal Adaptations to Lactation: Insights into Mammary Gland Function Julie A. Sharp, Kylie N. Cane, Christophe Lefevre, John P. Y. Arnould, and Kevin R. Nicholas

Volume 73 1 The Molecular Origins of Species-Specific Facial Pattern Samantha A. Brugmann, Minal D. Tapadia, and Jill A. Helms

2 Molecular Bases of the Regulation of Bone Remodeling by the Canonical Wnt Signaling Pathway Donald A. Glass II and Gerard Karsenty

3 Calcium Sensing Receptors and Calcium Oscillations: Calcium as a First Messenger Gerda E. Breitwieser

4 Signal Relay During the Life Cycle of Dictyostelium Dana C. Mahadeo and Carole A. Parent

5 Biological Principles for Ex Vivo Adult Stem Cell Expansion Jean-Franc¸ois Pare´ and James L. Sherley

6 Histone Deacetylation as a Target for Radiosensitization David Cerna, Kevin Camphausen, and Philip J. Tofilon

7 Chaperone-Mediated Autophagy in Aging and Disease Ashish C. Massey, Cong Zhang, and Ana Maria Cuervo

8 Extracellular Matrix Macroassembly Dynamics in Early Vertebrate Embryos Andras Czirok, Evan A. Zamir, Michael B. Filla, Charles D. Little, and Brenda J. Rongish

Contents of Previous Volumes

327

Volume 74 1 Membrane Origin for Autophagy Fulvio Reggiori

2 Chromatin Assembly with H3 Histones: Full Throttle Down Multiple Pathways Brian E. Schwartz and Kami Ahmad

3 Protein–Protein Interactions of the Developing Enamel Matrix John D. Bartlett, Bernhard Ganss, Michel Goldberg, Janet Moradian-Oldak, Michael L. Paine, Malcolm L. Snead, Xin Wen, Shane N. White, and Yan L. Zhou

4 Stem and Progenitor Cells in the Formation of the Pulmonary Vasculature Kimberly A. Fisher and Ross S. Summer

5 Mechanisms of Disordered Granulopoiesis in Congenital Neutropenia David S. Grenda and Daniel C. Link

6 Social Dominance and Serotonin Receptor Genes in Crayfish Donald H. Edwards and Nadja Spitzer

7 Transplantation of Undifferentiated, Bone Marrow-Derived Stem Cells Karen Ann Pauwelyn and Catherine M. Verfaillie

8 The Development and Evolution of Division of Labor and Foraging Specialization in a Social Insect (Apis mellifera L.) Robert E. Page Jr., Ricarda Scheiner, Joachim Erber, and Gro V. Amdam

Volume 75 1 Dynamics of Assembly and Reorganization of Extracellular Matrix Proteins Sarah L. Dallas, Qian Chen, and Pitchumani Sivakumar

2 Selective Neuronal Degeneration in Huntington’s Disease Catherine M. Cowan and Lynn A. Raymond

3 RNAi Therapy for Neurodegenerative Diseases Ryan L. Boudreau and Beverly L. Davidson

4 Fibrillins: From Biogenesis of Microfibrils to Signaling Functions Dirk Hubmacher, Kerstin Tiedemann, and Dieter P. Reinhardt

328

Contents of Previous Volumes

5 Proteasomes from Structure to Function: Perspectives from Archaea Julie A. Maupin-Furlow, Matthew A. Humbard, P. Aaron Kirkland, Wei Li, Christopher J. Reuter, Amy J. Wright, and G. Zhou

6 The Cytomatrix as a Cooperative System of Macromolecular and Water Networks V. A. Shepherd

7 Intracellular Targeting of Phosphodiesterase-4 Underpins Compartmentalized cAMP Signaling Martin J. Lynch, Elaine V. Hill, and Miles D. Houslay

Volume 76 1 BMP Signaling in the Cartilage Growth Plate Robert Pogue and Karen Lyons

2 The CLIP-170 Orthologue Bik1p and Positioning the Mitotic Spindle in Yeast Rita K. Miller, Sonia D’Silva, Jeffrey K. Moore, and Holly V. Goodson

3 Aggregate-Prone Proteins Are Cleared from the Cytosol by Autophagy: Therapeutic Implications Andrea Williams, Luca Jahreiss, Sovan Sarkar, Shinji Saiki, Fiona M. Menzies, Brinda Ravikumar, and David C. Rubinsztein

4 Wnt Signaling: A Key Regulator of Bone Mass Roland Baron, Georges Rawadi, and Sergio Roman-Roman

5 Eukaryotic DNA Replication in a Chromatin Context Angel P. Tabancay, Jr. and Susan L. Forsburg

6 The Regulatory Network Controlling the Proliferation–Meiotic Entry Decision in the Caenorhabditis elegans Germ Line Dave Hansen and Tim Schedl

7 Regulation of Angiogenesis by Hypoxia and Hypoxia-Inducible Factors Michele M. Hickey and M. Celeste Simon

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