Somatic genome variation in animals, plants, and microorganisms 9781118647028, 1118647025, 9781118647110, 1118647114, 9781118647127, 1118647122

Table of contents :
Content: Polyploidy in animal development and disease / Jennifer L. Bandura and Norman Zielke --
Large-scale programmed genome rearrangements in vertebrates / Jeramiah J. Smith --
Chromosome instability in stem cells / Paola Rebuzzini, Maurizio Zuccotti, Carlo Alberto Redi and Silvia Garagna --
Mechanisms of induced inheritable genome variation in flax / Christopher A Cullis --
Environmentally induced genome instability and its inheritance / Andrey Golubov --
The mitochondrial genome, genomic shifting and genomic conflict / Gregory G. Brown --
Plastid genome stability and repair / Eric Zampini, Sebastien Truche, Etienne Lepage, Samuel Tremblay-Belzile , and Normand Brisson --
RNA-mediated somatic genome rearrangement in ciliates / John R. Bracht --
Somatic genome instability in yeast and other fungi / Adrianna Skoneczna and Marek Skoneczny --
Genome variation in archaeans, bacteria, and asexually reproducing eukaryotes / Xiu-Qing Li --
RNA polyadenylation site regions : highly similar in base composition pattern but diverse in sequence : a combination ensuring similar function but avoiding repetitive-regions-related genomic instability / Xiu-Qing Li and Donglei Du --
Insulin signaling pathways in humans and plants / Xiu-Qing Li and Tim Xing --
Developmental variation in the nuclear genome primary sequence / Xiu-Qing Li --
Ploidy variation of the nuclear, chloroplast, and mitochondrial genomes in somatic cells / Xiu-Qing Li, Benoit Bizimungu, Guodong Zhang, and Huaijun Si --
Molecular mechanisms of somatic genome variation / Xiu-Qing Li --
Hypotheses for interpreting somatic genome variation / Xiu-Qing Li --
Impacts of somatic genome variation on genetic theories and breeding concepts / Xiu-Qing Li --
Somatic genome variation : what it is and what it means for agriculture and human health / Xiu-Qing Li.

Citation preview

Somatic Genome Variation in Animals, Plants, and Microorganisms

Somatic Genome Variation in Animals, Plants, and Microorganisms Edited by Xiu-Qing Li Agriculture and Agri-Food Canada, Fredericton, New Brunswick, Canada

This edition first published 2017 © 2017 John Wiley & Sons, Inc. Registered Offices John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA Editorial Office 111 River Street, Hoboken, NJ 07030, USA For details of our global editorial offices, customer services, and more information about Wiley products visit us at www.wiley.com. The right of Xiu-Qing Li to be identified as the author of the editorial material in this work has been asserted in accordance with law. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by law. Advice on how to obtain permision to reuse material from this title is available at http://www.wiley.com/go/permissions. Wiley also publishes its books in a variety of electronic formats and by print-on-demand. Some content that appears in standard print versions of this book may not be available in other formats. Limit of Liability/Disclaimer of Warranty The publisher and the authors make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of fitness for a particular purpose. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for every situation. In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of experimental reagents, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each chemical, piece of equipment, reagent, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. The fact that an organization or website is referred to in this work as a citation and/or potential source of further information does not mean that the author or the publisher endorses the information the organization or website may provide or recommendations it may make. Further, readers should be aware that websites listed in this work may have changed or disappeared between when this works was written and when it is read. No warranty may be created or extended by any promotional statements for this work. Neither the publisher nor the author shall be liable for any damages arising herefrom. Library of Congress Cataloging-in-Publication Data applied for

ISBN: 9781118647066 Cover Design: Wiley Cover Image: (Top) © Gio.tto/Shutterstock;(Bottom) © Fernan Federici/Gettyimages

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Contents List of Contributors  xv Preface and Introduction  xix Acknowledgments  xxi About the Editor  xxiii Part I 1

Somatic Genome Variation in Animals and Humans 

1

Polyploidy in Animal Development and Disease  3 Jennifer L. Bandura and Norman Zielke

1.1 Introduction  3 1.2 Mechanisms Inducing Somatic Polyploidy  4 1.2.1 Cell Fusion  4 1.2.2 Acytokinetic Mitosis  4 1.2.3 Endomitosis  5 1.2.4 Endoreplication  5 1.2.5 Gene Amplification  7 1.2.6 Ploidy Reversal  7 1.3 The Core Cell Cycle Machinery  8 1.4 Genomic Organization of Polyploid Cells  9 1.5 Endoreplication: An Effective Tool for Post-Mitotic Growth and Tissue Regeneration  10 1.6 Initiation of Endoreplication in Drosophila 11 1.6.1 Endocycle Entry in Ovarian Follicle Cells  11 1.6.2 Signaling Pathways Regulating Endocycle Entry in Follicle Cells  13 1.6.3 Endocycle Entry in Other Tissues  14 1.7 Mechanisms of Endocycle Oscillations in Drosophila 15 1.7.1 An Autonomous Oscillator Drives Endocycling in the Salivary Gland  15 1.7.2 Alternative Modes of Endoreplication  17 1.8 Gene Amplification in Drosophila Follicle Cells  17 1.8.1 Molecular Mechanism of Gene Amplification  17 1.8.2 The Endocycle-to-Amplification Switch  19 1.9 Endocycle Entry in the Trophoblast Lineage  19 1.10 Mechanisms of Endocycle Oscillations in Trophoblast Giant Cells  22 1.11 Cardiomyocytes  23

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1.11.1 Upstream Control of Cardiomyocyte Polyploidization  23 1.11.2 Mechanisms of Cardiomyocyte Polyploidization  24 1.11.3 Polyploidization as a Response to Tissue Damage  25 1.12 Hepatocytes  25 1.12.1 Mechanisms of Hepatocyte Polyploidization  25 1.12.2 The Ploidy Conveyor Model  26 1.12.3 Liver Regeneration  26 1.13 Megakaryocytes  28 1.13.1 Mechanisms of MKC Polyploidization  28 1.14 Concluding Remarks  30 Acknowledgments  31 References  31 2

Large-Scale Programmed Genome Rearrangements in Vertebrates  45 Jeramiah J. Smith

2.1 Introduction  45 2.1 Hagfish  46 2.2.1 Content of Eliminated DNA  47 2.2.2 Results and Mechanisms of Deletion  47 2.3 Sea Lamprey  48 2.3.1 Content of Eliminated DNA  48 2.3.2 Results and Mechanisms of Deletion  48 2.4 Zebra Finch  48 2.4.1 Mechanisms of Deletion  49 2.4.2 Content of Eliminated DNA  49 2.5 Emerging Themes and Directions  49 2.5.1 The Biological Function of PGR  49 2.5.2 Mechanisms of Deletion  50 2.5.3 Other Vertebrates?  51 References  51 3

Chromosome Instability in Stem Cells  55 Paola Rebuzzini, Maurizio Zuccotti, Carlo Alberto Redi and Silvia Garagna

3.1 Introduction  55 3.2 Pluripotent Stem Cells  56 3.2.1 Primate Embryonic Stem Cells  56 3.2.2 Mouse Embryonic Stem Cells  57 3.2.3 Parthenogenetic Embryonic Stem Cells  57 3.2.4 Induced Pluripotent Stem Cells  58 3.3 Somatic Stem Cells  58 3.3.1 Mesenchymal Stem Cells  58 3.3.2 Neural Stem Cells  59 3.4 Mechanisms of Chromosomal Instability  59 3.4.1 Dysfunction in the Spindle Assembly Checkpoints  60 3.4.2 Defects of Microtubule Attachment to the Kinetochore  60

Contents

3.4.3 Supernumerary Centrosomes  61 3.4.4 Sister Chromatids Cohesion  62 3.5 Mechanisms of Chromosomal Instability in Stem Cells  63 References  63 Part II Somatic Genome Variation in Plants  4

75

Mechanisms of Induced Inheritable Genome Variation in Flax  77 Christopher A. Cullis

4.1 Introduction  77 4.2 Restructuring the Flax Genome  79 4.3 Specific Genomic Changes  80 4.4 What Happens When Plastic Plants Respond to Environmental Stresses?  83 4.5 When Do the Genomic Changes Occur and Are they Adaptive?  83 4.6 Is this Genomic Response of Flax Unique?  84 4.7 Concluding Remarks  87 Acknowledgments  87 References  87 5

Environmentally Induced Genome Instability and its Inheritance  91 Andrey Golubov

5.1 Introduction  91 5.2 Stress and its Effects on Genomes  92 5.2.1 Genetic Changes  92 5.2.2 DNA Repair  92 5.2.3 Epigenetic Changes  93 5.2.3.1 DNA Methylation  93 5.2.3.2 Histone Modifications  95 5.2.4 The Link between Genetic and Epigenetic Changes  95 5.3 Transgenerational Inheritance  96 5.4 Concluding Remarks  97 Acknowledgments  97 References  97 6

The Mitochondrial Genome, Genomic Shifting, and Genomic Conflict  103 Gregory G. Brown

6.1 Introduction  103 6.2 Heteroplasmy and Sublimons  105 6.3 Cytoplasmic Male Sterility (CMS) in Plants  108 6.4 Mitochondrial Sublimons and CMS  109 6.5 Restorer Gene Evolution: Somatic Genetic Changes Drive Nuclear Gene Diversity?  111 6.6 Concluding Remarks  112 References  113

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7

Plastid Genome Stability and Repair  119 Éric Zampini, Sébastien Truche, Étienne Lepage, Samuel Tremblay‐Belzile and Normand Brisson

7.1 Introduction  120 7.2 Characteristics of the Plastid Genome  121 7.2.1 General Composition of the Plastid Genome  121 7.2.2 The Structure of the Plastid Genome  123 7.3 Replication of Plastid DNA  124 7.3.1 Plastid DNA Content during Development  124 7.3.2 Plastid DNA Replication Machinery  125 7.3.3 Replication Mechanisms  126 7.3.4 Origins of Replication  129 7.3.5 Nucleus and Plastid Coordination during DNA Replication  130 7.4 Transcription in the Plastid  130 7.5 The Influence of Replication and Transcription on Plastid Genome Stability  131 7.6 Plastid Genome Stability and DNA Repair  133 7.6.1 Oxidative Stress, Photo‐Adaptation, and ROS Detoxification  133 7.6.2 UV‐Induced DNA Damage  138 7.6.3 Recombination and DNA Double‐Strand Break Repair  141 7.7 Outcomes of DNA Rearrangements  145 7.8 Concluding Remarks  147 References  148 Part III Somatic Genome Variation in Microorganisms  8

165

RNA-Mediated Somatic Genome Rearrangement in Ciliates  167 John R. Bracht

8.1 Introduction  168 8.2 Ciliates: Ubiquitous Eukaryotic Microorganisms with a Long Scientific History  168 8.3 Two’s Company: Nuclear Dimorphism in Ciliates  170 8.4 Paramecium: Non-Mendelian Inheritance Comes to Light  171 8.5 Tetrahymena and the Origin of the scanRNA Model  173 8.6 Small RNAs in Stylonychia and Oxytricha 175 8.7 Long Noncoding RNA Templates in Genome Rearrangement  176 8.8 Long Noncoding RNA: An Interface for Short Noncoding RNA  177 8.9 Short RNA-Mediated Heterochromatin Formation and DNA Elimination  179 8.10 Transposable Elements and the Origins of Genome Rearrangements  182 8.11 Transposons, Phase Variation, and Programmed Genome Engineering in Bacteria  185 8.12 Transposases, Noncoding RNA, and Chromatin Modifications in VDJ Recombination of Vertebrates  186 8.13 Concluding Remarks: Ubiquitous Genome Variation, Transposons, and Noncoding RNA  187

Contents

Acknowledgments  187 References  187 9

Mitotic Genome Variations in Yeast and Other Fungi  199 Adrianna Skoneczna and Marek Skoneczny

9.1 Introduction  199 9.2 The Replication Process as a Possible Source of Genome Instability  200 9.2.1 DNA Polymerases as Guardians of Genome Maintenance  201 9.2.2 dNTP Cellular Level and their Pool Bias Contribute to Genome Stability  205 9.2.3 Mismatch Repair (MMR) and Ribonucleotide Excision Repair (RER) Are Used to Clean‐up after Replication  218 9.3 Post-Replicative Repair (PRR) or Homologous Recombination (HR) Are Responsible for Error-Free and Error-Prone Repair of Blocking Lesions and Replication Stall-Borne Problems  219 9.3.1 Sumoylated PCNA‐, Srs2‐, and Replicative Polymerase‐dependent DNA Synthesis on Damaged Template  221 9.3.2 Ubiquitinated PCNA‐ and Specialized Pol‐Dependent Translesion Synthesis  223 9.3.3 The Polyubiquitinated PCNA‐ and Rad5‐Dependent Damage Avoidance Pathway  225 9.3.4 The Alternative PCNA‐, RPA‐, and 5′‐Junction‐Dependent Pathway Involved in Gap Filling and Telomere Maintenance  226 9.3.5 Crosstalk between RFC Complexes Adapts Cellular Response to Different Stresses Arising from Genome Perturbations  226 9.3.6 Break‐Induced Replication (BIR) Is a Vastly Inaccurate Repair Pathway  227 9.4 Ploidy Maintenance and Chromosome Integrity Mechanisms  229 9.4.1 Processes that Affect Aneuploidy in Yeasts  230 9.4.2 Ploidy Changes in Yeasts  231 9.4.3 Possible Mechanism of Ploidy Change in Yeast  232 9.5 Concluding Remarks  234 References  235 Part IV General Genome Biology  10

251

Genome Variation in Archaeans, Bacteria, and Asexually Reproducing Eukaryotes  253 Xiu-Qing Li

10.1 Introduction  254 10.2 Chromosome Number in Prokaryote Species  254 10.3 Genome Size Variation in Archaeans and Bacteria  255 10.4 Archaeal and Bacterial Genome Size Distribution  256 10.5 Genomic GC Content in Archaeans, Bacteria, Fungi, Protists, Plants, and Animals  257 10.6 Correlation between GC Content and Genome or Chromosome Size  259

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10.7

Genome Size and GC-Content Variation in Primarily Asexually Reproducing Fungi  260 10.8 Variation of Gene Direction  263 10.9 Concluding Remarks  263 Acknowledgments  264 References  264 11

RNA Polyadenylation Site Regions: Highly Similar in Base Composition Pattern but Diverse in Sequence—A Combination Ensuring Similar Function but Avoiding Repetitive-Regions-Related Genomic Instability  267 Xiu-Qing Li and Donglei Du

11.1

General Introduction to Gene Number, Direction, and RNA Polyadenylation  268 11.2 Base Selection at the Poly(A) Tail Starting Position  269 11.3 Most Frequent Upstream Motifs in Microorganisms, Plants, and Animals  271 11.4 Motif Frequencies in the Whole Genome  273 11.5 The Top 20 Hexamer Motifs in the Poly(A) Site Region in Humans  273 11.6 Polyadenylation Signal Motif Distribution  273 11.7 Alternative Polyadenylation  275 11.8 Base Composition of 3′UTR in Plants and Animals  276 11.9 Base Composition Comparison between 3′UTR and Whole Genome  276 11.10 Base Composition of 3′COR in Plants and Animals  277 11.11 Base Composition Pattern of the Poly(A) Site Region in Protists  278 11.12 Base Composition Pattern of the Poly(A) Site Region in Plants  280 11.13 Base Composition Pattern of the Poly(A) Site Region in Animals  280 11.14 Comparison of Poly(A) Site Region Base Composition Patterns in Plants and Animals  280 11.15 Common U-A-U-A-U Base Abundance Pattern in the Poly(A) Site Region in Fungi, Plants, and Animals  284 11.16 Difference between the Most Frequent Motifs and Seqlogo-Showed Most Frequent Bases  284 11.17 RNA Structure of the Poly(A) Site Region  286 11.18 Low Conservation in the Overall Nucleotide Sequence of the Poly(A) Site Region  286 11.19 Poly(A) Site Region Stability and Somatic Genome Variation  286 11.20 Concluding Remarks  287 Acknowledgments  288 References  288 12

Insulin Signaling Pathways in Humans and Plants  291 Xiu‐Qing Li and Tim Xing

12.1 Introduction  291 12.2 Ranking of the Insulin Signaling Pathway and its Key Proteins  293 12.3 Diseases Caused by Somatic Mutations of the PI3K, PTEN, and AKT Proteins in the Insulin Signaling Pathway  293

Contents

12.4 Plant Insulin and Medical Use  295 12.5 Role of the Insulin Signaling Pathway in Regulating Plant Growth  295 12.6 Concluding Remarks  295 References  296 13

Developmental Variation in the Nuclear Genome Primary Sequence  299 Xiu-Qing Li

13.1 Introduction  299 13.2 Genetic Mutation, DNA Damage and Protection, and Gene Conversion in Somatic Cells  300 13.3 Programmed Large-Scale Variation in Primary DNA Sequences in Somatic Nuclear Genome  302 13.4 Generation of Antibody Genes in Animals through Somatic Genome Variation  303 13.5 Developmental Variation in Primary DNA Sequences in the Somatic Cells of Plants  303 13.6 Heritability and Stability of Developmentally Induced Variation in the Somatic Nuclear Genome in Plants  303 13.7 Concluding Remarks  304 References  305 14

Ploidy Variation of the Nuclear, Chloroplast, and Mitochondrial Genomes in Somatic Cells  309 Xiu‐Qing Li, Benoit Bizimungu, Guodong Zhang and Huaijun Si

14.1 Introduction  310 14.2 Nuclear Genome in Somatic Cells  311 14.2.1 Ploidy Variation of the Individual or Species in Plants and Animals  311 14.2.2 Effects of Species Ploidy Variation on the Growth of Animals and Plants  312 14.2.3 Ploidy of Bacteria  313 14.2.4 Endopolyploidy in Animal and Plant Somatic Cells  313 14.2.5 Somatic Cell Haploidization  315 14.2.6 Aneuploid Cells in Plant Somatic Tissues  315 14.2.7 Aneuploid Cells in Cancerous Masses  316 14.2.8 Nuclear B Chromosomes in Somatic Cells  316 14.3 Plastid Genome Variation in Somatic Cells  317 14.3.1 Types of Plastids  317 14.3.2 Plastid Genome and its Size in Somatic Cells  317 14.3.3 Recombination among Repeated Sequences in the Plastid Genome  318 14.3.4 Integrity of the Organelle Genome in Green Leaves under Light  318 14.3.5 Plastid Genome Ploidy or Copy Number Variation in Somatic Cells  319 14.4 Mitochondrial Genome in Somatic Cells  320 14.4.1 Mitochondrial Genome and its Size  320 14.4.2 Recombination among Repeated Sequences and Subgenomic Molecules in Mitochondria  321 14.4.3 Mitochondrial Subgenome Copy Number Variation in Somatic Cells  322

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14.4.4 Nuclear and Tissue‐Specific Regulation of Mitochondrial Gene Expression  322 14.4.5 Stoichiometric Variation and Effects on Mitochondrial Subgenomic Molecules  323 14.5 Organelle Genomes in Somatic Hybrids  324 14.6 Effects of Nuclear Genome Ploidy on Organelle Genomes  325 14.7 Concluding Remarks  326 Acknowledgments  326 References  326 15

Molecular Mechanisms of Somatic Genome Variation  337 Xiu-Qing Li

15.1 Introduction  338 15.2 Mutation of Genes Involved in the Cell Cycle, Cell Division, or Centromere Function  338 15.3 DNA Damage  338 15.4 Variation in Induction and Activity of Radical-Scavenging Enzymes  339 15.5 DNA Cytosine Deaminases  340 15.6 Variation in Protective Roles of Pigments against Oxidative Damage  340 15.7 RNA-Templated DNA Repair  341 15.8 Errors in DNA Repair  341 15.9 RNA-Mediated Somatic Genome Rearrangement  342 15.10 Repetitive DNA Instability  342 15.11 Extracellular DNA  343 15.12 DNA Transposition  343 15.13 Somatic Crossover and Gene Conversion  343 15.14 Molecular Heterosis  344 15.15 Genome Damage Induced by Endoplasmic Reticulum Stress  344 15.16 Telomere Degeneration  344 15.17 Concluding Remarks  344 References  345 16

Hypotheses for Interpreting Somatic Genome Variation  351 Xiu-Qing Li

16.1 Introduction  352 16.2 Cell-Specific Accumulation of Somatic Genome Variation in Somatic Cells  352 16.3 Developmental Age and Genomic Network of Reproductive Cells  353 16.4 Genome Generation Cycle of Species  353 16.5 Somatic Genome Variation and Tissue-Specific Requirements during Growth or Development  354 16.6 Costs and Benefits of Somatic Genome Variation  354 16.7 Hypothesis on the Existence of a Primitive Stage in both Animals and Plants  355 16.8 Sources of Genetic Variation from in Vitro Culture Propagation  357 16.9 Hypothesis that Heterosis Is Created by Somatic Genome Variation  357

Contents

16.10

Genome Stability through Structural Similarity and Sequence Dissimilarity  358 16.11 Hypothesis Interpreting the Maternal Transmission of Organelles  358 16.12 Ability of Humans to Deal with Somatic Genome Variation and Diseases  359 16.13 Concluding Remarks  360 References  360 17

Impacts of Somatic Genome Variation on Genetic Theories and Breeding Concepts, and the Distinction between Mendelian Genetic Variation, Somagenetic Variation, and Epigenetic Variation  363 Xiu‐Qing Li

17.1 Introduction  364 17.2 The Term ‘Somatic Genome’  365 17.3 Mendelian Genetic Variation, Epigenetic Variation, and Somagenetic Variation  365 17.4 What Is a Gene?  367 17.5 Breeding Criteria, Genome Cycle, Pure Lines, and Variety Stability  368 17.6 The Weismann Barrier Hypothesis and the Need for Revision  370 17.7 Implications for Species Evolution  370 17.8 Concluding Remarks  371 References  372 18

Somatic Genome Variation: What it is and What it Means for Agriculture and Human Health  377 Xiu-Qing Li

18.1 Introduction  378 18.2 Natural Attributes of Somatic Genome Variation  378 18.3 Implications of Somatic Genome Variation for Human and Animal Health  380 18.3.1 Cellular-Level Variation  380 18.3.2 Ploidy and Chromosome Number Variation of the Whole Organism  380 18.3.3 Endoploidy Variation  381 18.3.4 DNA Cytosine Deaminases, Somatic Mutation, Immunoglobulin Diversity, and Tumors  381 18.3.5 Mitochondrial Genome Sequence or DNA Amount Variation  384 18.3.6 Nuclear or Ooplasmic Transfer-Based Therapy  385 18.3.7 Differential Treatments of Beneficial and Harmful SGVs  385 18.4 Implications of Somatic Genome Variation for Agriculture  385 18.4.1 Cellular-Level Variation  385 18.4.2 Ploidy and Chromosome Number Variation of the Whole Organism  386 18.4.3 Endoploidy Variation  387 18.4.4 Intra- and Interchromosomal Variation  387 18.4.5 Dedifferentiation- and Redifferentiation-Induced Variation  388 18.4.6 DNA Damage, Epigenetics, Gene Mutation, and Bud Mutation  389

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18.4.7 Plastid Genome Sequence or DNA Amount Variation  389 18.4.8 Mitochondrial Genome Sequence or DNA Amount Variation  390 18.4.9 DNA Transfer, Organelle Transmission, and Organelle Genome Segregation  390 18.4.10 Intercompartmental Interaction and DNA Exchange  391 18.5 Concluding Remarks  391 Acknowledgments  392 References  392 Index  405

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List of Contributors Jennifer L. Bandura

Christopher A. Cullis

Biology Department Roanoke College Salem USA

Department of Biology Case Western Reserve University Cleveland USA

Benoit Bizimungu

Donglei Du

Fredericton Research and Development Centre Agriculture and Agri‐Food Canada Fredericton New Brunswick Canada

Quantitative Methods Research Group Faculty of Business Administration University of New Brunswick Fredericton Canada Silvia Garagna

Department of Biology American University Washington, DC USA

Laboratorio di Biologia dello Sviluppo Dipartimento di Biologia e Biotecnologie Università degli Studi di Pavia Pavia Italy

Normand Brisson

Andrey Golubov

Department of Biochemistry Université de Montréal Montréal Canada

Department of Biological Sciences University of Lethbridge Lethbridge Canada

Gregory G. Brown

Étienne Lepage

Department of Biology McGill University Montreal Canada

Department of Biochemistry Université de Montréal Montréal Canada

John R. Bracht

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List of Contributors

Xiu-Qing Li

Marek Skoneczny

Fredericton Research and Development Centre Agriculture and Agri‐Food Canada Fredericton New Brunswick Canada

Institute of Biochemistry and Biophysics Polish Academy of Sciences Department of Genetics Warsaw Poland Jeramiah J. Smith

Paola Rebuzzini

Laboratorio di Biologia dello Sviluppo Dipartimento di Biologia e Biotecnologie Università degli Studi di Pavia Pavia Italy Carlo Alberto Redi

Laboratorio di Biologia dello Sviluppo Dipartimento di Biologia e Biotecnologie Università degli Studi di Pavia Pavia Italy Huaijun Si

Gansu Provincial Key Laboratory of Aridland Crop Science Gansu Key Laboratory of Crop Genetic and Germplasm Enhancement College of Life Science and Technology Gansu Agricultural University Lanzhou People’s Republic of China

Department of Biology University of Kentucky Kentucky USA Samuel Tremblay-Belzile

Department of Biochemistry Université de Montréal Montréal Canada Sébastien Truche

Department of Biochemistry Université de Montréal Montréal Canada Tim Xing

Department of Biology Carleton University Ottawa Canada

Adrianna Skoneczna

Éric Zampini

Institute of Biochemistry and Biophysics Polish Academy of Sciences Laboratory of Mutagenesis and DNA Repair Warsaw Poland

Department of Biochemistry Université de Montréal Montréal Canada

List of Contributors

Guodong Zhang

Maurizio Zuccotti

Gansu Provincial Key Laboratory of Aridland Crop Science Gansu Key Laboratory of Crop Genetic and Germplasm Enhancement College of Life Science and Technology Gansu Agricultural University Lanzhou People’s Republic of China

Dipartimento di Scienze Biomediche, Biotecnologiche e Traslazionali (SBIBIT) Università degli Studi di Parma Parma Italy

Norman Zielke

Genome‐Scale Biology Research Program Institute of Biomedicine University of Helsinki Helsinki Finland

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Preface and Introduction The success of genetic analysis and breeding using Mendel’s laws, Johannsen’s concepts of genotype and phenotype, Weismann’s germ‐plasm continuity hypothesis, Morgan’s linear arrangement of genes on chromosomes, and Muller’s mutation theory leads to a belief—all genes are given by parents and stay the same except for having mutation occasionally caused by external mutagens, somatic cells have no contribution to inheritance, the gene transcript abundance is purely determined by the promoter activity and RNA stability, and clones are genetically identical. However, as shown in the present book, the somatic genome (the sum total of genetic materials in the cytoplasm and somatic nucleus) actually has environmental and developmental variations; for example: (1) many antibody genes are somatically produced; (2) some ciliate somatic genomes are generated using RNA templates and are therefore phenotypes of the germline genomes but are also the genotypes for many other traits; (3) gene transcription and some endogenous enzymes can induce mutation; (4) certain foods and drinks affect DNA stability and tumor growth; (5) the DNA fingerprint of an animal or plant has certain variations among somatic tissues; (6) various somagenetic and epigenetic variations are transgenerational, and some DNA is translocatable among cells; (7) some genes and repetitive DNA have copy number variation; (8) the chloroplast and mitochondrial genomes vary in ploidy and DNA amount; (9) some DNA sequences are functional through protein binding or DNA‐fragment release; (10) DNA damage is sometimes not repaired; (11) clones are genetically mosaic to a certain degree; and (12) the average ploidy level varies among tissues. This book attempts to assemble the evidence of somatic genome variation in microorganisms, plants, animals, and humans, update various basic concepts in genetics and breeding, determine the implications of somatic genome variation for human health and agriculture, and propose an updated synthesis of inheritance.

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Acknowledgments This book represents the cumulative work of 24 authors from 14 research institutes/ universities in six countries. I wish to express my gratitude to all authors who have contributed manuscripts to this book. I thank my Research Centre and my family for their continued support. I would like to extend my very special thanks to the ­commissioning editor, Mr Justin Jeffryes (the Editorial Director, Natural Sciences, The Americas, at Wiley), Rebecca Ralf, Managing Editor of Life Sciences Books, Ms Divya Narayanan, the primary contact, Ms Bhargavi Natarajan, the former primary contact, Kavitha Chandrasekar, the production editor, Mrs Julie Musk, the copyeditor, and the entire production team at Wiley for their support and high professional standards. Xiu‐Qing Li

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About the Editor

Dr Xiu‐Qing Li has been a Research Scientist of Molecular Genetics at Agriculture and Agri‐Food Canada, Government of Canada (the Fredericton Research and Development Centre, Fredericton, New Brunswick, Canada) since 1997, and an adjunct professor at the University of New Brunswick since 1998. Dr Li received his degree of Docteur d’Etat of France in natural science from Université de Paris‐Sud (1987), was an associate professor of biotechnology at Peking University (1987–1993), Invited Professor of Genetics (Université de Paris‐Sud, 1988), Research Scientist of Molecular Genetics (Chargée de recherche, level II) at the Centre National de la Recherche Scientifique (CNRS), Gif‐sur‐ Yvette, France (1987), Visiting Professor at Purdue University (1991–1992), and researcher at McGill University (1992–1997). Dr. Li is an Academic Editor of PloS ONE, co‐editor of the book Somatic Genome Manipulation (Li et al. eds, Springer, 2015), on the editorial boards of several other journals, and serves as the Communication Director of the Canadian Association of Plant Biotechnology. He is the organizer and chair for the annual Somatic Genome Workshops (San Diego, since 2010) and a co‐organizer of the Genome Features and Chromosome Functionality Workshop (since 2016) at the International Plant and Animal Genome Conferences. Dr. Li has extensive research experience and numerous publications on genetics, genomics, RNA polyadenylation, plant somatic genome, plant genetic engineering, plant cell biology, somaclonal variation, chloroplasts, mitochondria, plant male sterility, plant carbohydrate metabolism, potato genetic improvement, bioinformatic analysis, and genome evolution.

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Part I Somatic Genome Variation in Animals and Humans

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1 Polyploidy in Animal Development and Disease Jennifer L. Bandura1 and Norman Zielke 2* 1 2

Biology Department, Roanoke College, Salem, USA Genome-Scale Biology Research Program, Institute of Biomedicine, University of Helsinki, Helsinki, Finland

Abstract Somatic polyploidization is a developmentally controlled process that can be found in many animal species, including mammals. Polyploidy is utilized as a mechanism to amplify gene expression via an increase in DNA copy number, to drive cellular growth, and to initiate cellular differentiation. Moreover, polyploidy has been implicated in the response to injury or disease. This chapter discusses the processes by which polyploid cells arise and the underlying molecular mechanisms, with a focus on endoreplication in Drosophila and mice. In addition, it summarizes recent progress on the related subjects of endomitosis, ploidy reversal, and gene amplification. Keywords  polyploidy; endoreplication; endomitosis; gene amplification

1.1 ­Introduction A fundamental characteristic of the canonical eukaryotic cell cycle is that the genome is only replicated once per cycle, and becomes then equally distributed between two daughter cells during the following mitosis. This strict constraint on DNA replication ensures that each daughter cell receives exactly one diploid set of chromosomes. However, some cell types depart from this rule and differentiate into viable polyploid cells, in which the entire set of chromosomes has been multiplied. While the term polyploid refers to cells with virtually any chromosomal configuration, the term polytene refers to a subclass of polyploid cells in which sister chromatids remain closely associated in parallel arrays. The degree of polyploidy is indicated by the C-value, which represents the DNA content as a multiple of the normal haploid genome. Hence, tetraploid cells have a DNA content of 4 C, while octoploid cells are 8 C. Polyploidy is clearly

*Corresponding author: [email protected] Bandura J.L. and Zielke N. (2017) Polyploidy in animal development and disease. In: Li X.-Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley-Blackwell, Hoboken, NJ, Ch. 1, pp. 3–44. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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distinguishable from aneuploidy, which refers to cells containing an aberrant number of chromosomes that is not a multiple of the haploid genome. Polyploidy is often ­confused with the term re-replication, which describes genome duplications originating from unscheduled initiation of DNA replication during S phase. DNA re-replication is an aberrant event that produces cells characterized by a heterogeneous DNA content, incomplete chromosome duplications, stalled replication forks, and DNA damage. In  cells with a compromised DNA damage response, re-replication events can cause genomic instability and thereby promote cancer formation. Polyploidy occurs in two forms that can be distinguished by their mechanisms of origin: Germline polyploidy refers to genome duplications in the germline, which are often caused by chromosome mis-segregation during meiosis. In this case, all the cells in the resulting progeny will be polyploid and, because the multiplied chromosomes may be inherited by future generations, germline polyploidization is generally thought to promote speciation. In this chapter, we focus on the more common somatic polyploidy, which is often intrinsically connected to cellular differentiation and is therefore also referred to as developmentally programmed polyploidy. Somatic polyploidy occurs frequently in ferns, flowering plants, mollusks, arthropods, amphibians, and fish, but is also found in a few specialized cell types in mammals. Somatic polyploidy is utilized to increase tissue size, to initiate cell differentiation, and to multiply gene copy number to enhance tissue-specific functions. Somatic polyploidy has also been implicated in tissue regeneration and the suppression of tumorigenesis.

1.2 ­Mechanisms Inducing Somatic Polyploidy The formation of somatic polyploid cells relies on four distinct mechanisms: cell fusion, acytokinetic mitosis, endomitosis, and endoreplication (Figure 1.1; Table 1.1). 1.2.1  Cell Fusion

Cell fusion describes the fusion of two cells residing in G1 phase, which results in the formation of a multinucleated cell. A striking example is found during the development of nematodes such as Caenorhabditis elegans, whose hypodermis is a single syncytium that grows through successive cell fusion events (Flemming et al. 2000). Cell fusion also occurs during the fusion of skeletal muscle myoblasts into myotubes, when monocytes differentiate into osteoclasts, and during the formation of syncytiotrophoblasts in the human placenta (Cross 2005). 1.2.2  Acytokinetic Mitosis

Acytokinetic mitosis refers to an abortive cell cycle that includes mitosis but lacks cytokinesis and thereby leads to the formation of multinucleated cells. The stereotypical example of this mechanism happens during post-natal liver development, when the complex process of hepatocyte polyploidization is initiated by an acytokinetic division (Gentric et al. 2012). In addition, many cardiomyocytes undergo acytokinetic mitosis during early post-natal development to create binucleate polyploid cells (Soonpaa et al. 1996; Pandit et al. 2013).

1.2  Mechanisms Inducing Somatic Polyploidy

Figure 1.1  Mechanisms introducing somatic polyploidy. (A) Nuclear configurations in mitotic and polyploid cell types. C, C-value, multiples of the haploid DNA content arising from different mechanisms. (B) Overview of non-canonical cell cycle variants leading to somatic polyploidy. Hepatocytes (HPC) bypass cytokinesis, and as a consequence this cell cycle variant is called an acytokinetic mitosis. Megakaryocyte polyploidization occurs through endomitosis, which is another abortive mitosis lacking anaphase B, telophase, and cytokinesis. Mouse trophoblast giant cells (TGC) and Drosophila salivary glands (SG) both undergo endocycles, which are completely devoid of M phase. TGCs undergo full genome replications, while SGs exit S phase before completing late replication. (C) Schematic of an under-replicated region, which results from differential firing of replication origins and within the euchromatin usually encompasses 100–400 kb. (D) Genomic organization of amplified loci, which arise from reiterative origin firing, resulting in copy number gradients stretching approximately 100 kb.

1.2.3 Endomitosis

Endomitosis is a related to acytokinetic mitosis and also characterized by a truncated M phase, but in this case cells abort the cell cycle without completing anaphase or undergoing cytokinesis, resulting in the formation of a single polyploid nucleus. A well-known example of endomitosis takes place in the bone marrow of mice and humans when megakaryoblasts differentiate into polyploid non-proliferating megakaryocytes (Ravid et al. 2002; Bluteau et al. 2009). 1.2.4 Endoreplication

Endoreplication cycles, also referred to as endocycles, are the most extreme deviation from the canonical cell cycle, as endocycling cells execute multiple rounds of DNA replication without any intervening mitosis or cytokinesis (Edgar and Orr-Weaver 2001;

5

Skin/keratinocytes

Throughout development

Note: This is not a comprehensive list of all polyploid tissues in animals. a) There are 65 nuclei per hypodermis. b) Also undergo ploidy reversal. c) Additional unidentified mechanisms.

H. sapiens

Embryo (onset at E4.5)

Postnatal (onset at day 4)

Throughout development

Epidermis

Heart/cardiomyocytes

Adult

Midgut

Embryo (onset at E7.5)

Larva and pupa

Hindgut/rectal papillae

Megakaryocytes

Pupa

Mechanosensory bristles

M. musculus, H. sapiens

Adult

Malpighian tubules

Placenta/trophoblast giant cells

Adult

Fat body

M. musculus

Larva

Brain/subperineural glia

Postnatal (onset ~ 3 weeks)

Adult

Ovary/nurse cell

Liver/hepatocytes

Adult

Ovary/follicle cell

O. dioica

Larva

Larva

Salivary gland

Intestine

D. melanogaster

Larva and adult

Hypodermis

C. elegans

Developmental stage

Tissue/cell type

Organism

12 C

8C

128 C

512 C–1024 C

8 C–16 C

1300 C

64 C

16 C

8C

256 C

256 C

22 C

1500 C

16 C

1035 C

32 C

16 C a

Maximal ploidy

Table 1.1  Polyploidization achieved by varying mechanisms across animal species.

Endocycles, Acytokinetic mitosis

Acytokinetic mitosis c

Endomitosis, Acytokinetic mitosis

Endocycles c

(Zanet et al. 2010; Gandarillas 2012)

(Soonpaa et al. 1996; Pandit et al. 2013)

(Nagata et al. 1997; Ravid et al. 2002; Sher et al. 2013)

(Barlow and Sherman 1974; Varmuza et al. 1988; Sher et al. 2013)

(Ganot and Thompson 2002)

(Lamb 1982)

(Fox et al. 2010)

(Hartenstein and Posakony 1989; Audibert et al. 2005)

(Lamb 1982)

(Richards 1980)

(Unhavaithaya and Orr-Weaver 2012)

(Hammond and Laird 1985a)

(Hammond and Laird 1985a)

(Hammond and Laird 1985b)

(Hedgecock and White 1985)

(Flemming et al. 2000)

Reference

Acytokinetic mitosis b, c (Guidotti et al. 2003; Wirth et al. 2006; Margall-Ducos et al. 2007)

Endocycles

Endocycles

Endocycles b

Endocycles

Endocycles

Endocycles

Endocycles, Acytokinetic mitosis

Endocycles

Endocycles

Endocycles

Endocycles

Cell fusion, Endocycles

Mechanism of polyploidization

1.2  Mechanisms Inducing Somatic Polyploidy

Lilly and Duronio 2005; Lee et al. 2009; Ullah et al. 2009a; Ullah et al. 2009b; Fox and Duronio 2013; Zielke et al. 2013). Endocycles generally include Gap (G) phases between each S phase, which is the most obvious distinction from re-replication. By executing consecutive endocycles, cells can dramatically increase their nuclear DNA content and attain ploidies of up to 200,000 C. This mechanism of somatic polyploidy has been intensively studied in the fruit fly Drosophila melanogaster, where virtually all postmitotic growth is achieved by endoreplication (Edgar and Nijhout 2004). In mammals, endoreplication cycles occur in the extra-embryonic tissue of the placenta, where they are involved in the formation trophoblast giant cells (TGCs) (Hu and Cross 2010). Endoreplication has also been observed in epithelial keratinocytes (Gandarillas 2012) and hepatocytes regenerating from damage after injury or stress (Duncan 2013; Pandit et al. 2013). 1.2.5  Gene Amplification

Gene amplification is a developmentally controlled process that results in massive expansion of specific genomic loci (Calvi and Spradling 1999; Calvi and Spradling 2001; Tower 2004; Claycomb and Orr-Weaver 2005; Calvi 2006). Amplification is achieved by reiterative firing of specialized origins of replication while origins throughout the remainder of the genome are silenced. This mechanism does not result in polyploidy as it only affects certain regions of the genome, but it was included because it represents a striking example of somatic genome variation. The regulation of gene amplification has been most thoroughly analyzed in the follicular epithelium of the Drosophila ovary, in which the chorion genes and a few other loci become amplified to facilitate eggshell formation (Calvi et  al. 1998b; Claycomb et  al. 2004a; Bandura et  al. 2005; Kim et  al. 2011). Gene amplification also occurs in the ciliate Tetrahymena thermophila, in which portions of the rDNA locus become amplified about 5000 times (Kapler 1993), as well as the II/9A-1 locus of the fly Sciara coprophila, which undergoes a 17-fold amplification (Gerbi et al. 1993). 1.2.6  Ploidy Reversal

Somatic polyploidy is generally thought to be an irreversible process that is crucial for differentiation into specialized cell types. However, certain polyploid cell types are capable of executing reductive divisions, a process that is also referred to as ploidy reversal. The biological relevance of this process is only poorly understood, but recent work has suggested that ploidy reversal could serve as a mechanism to increase genomic variation of somatic cells and thereby enhance the robustness of tissues and organs. On the other hand, ploidy reversal might represent a mechanism by which tumor cells to gain or lose chromosomes and thereby acquire a competitive advantage favoring tumor growth. Polyploid cells undergoing reductive division are prevalent among plants, but only rarely observed in animals. The existence of polyploid mitosis in insects has been known for decades (Berger 1938; Grell 1946), but remained largely unstudied until recently, when it was reported that the cells of the Drosophila rectal papilla enter mitosis after executing two or more endocycles (Fox et al. 2010). Further examples of insect cells undergoing ploidy reversal are found in the hindgut of the mosquito Culex pipens (Berger 1938; Grell 1946), the larval epidermis of another mosquito, Aedes aegypti (Risler 1959), the epidermis of the tobacco hornworm Manduca sexta (Kato et al. 1987),

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and the fat body of the kissing bug, Rhodnius (Wigglesworth 1967). Arguably the most well-understood example of ploidy reversal in animals is found in the liver of mice and humans where tetra- or octoploid hepatocytes can revert to the diploid or tetraploid state (Duncan et al. 2010; Duncan et al. 2012b).

1.3 ­The Core Cell Cycle Machinery All mechanisms leading to somatic polyploidy, with the exception of cell fusion, are based on non-canonical cell cycles, which involve many of the known cell cycle regulators. Therefore, to ease the discussion of the mechanisms of these variant cell cycles it is imperative to include an overview of the core cell cycle machinery. However, the interested reader can find a more comprehensive review of this topic in Budirahardja and Gonczy (2009) and Nordman and Orr-Weaver (2012). Cell cycle transitions in mitotic cells are largely regulated by the sequential activation of cyclin-dependent kinase (CDK) complexes. Entry into mitosis relies on Cdk1, which forms complexes with the mitotic cyclins Cyclin A (CycA), Cyclin B (CycB), or Cyclin B3 (CycB3) and the Stg/Cdc25 phosphatase. Progression through mitosis requires proteasomal destruction of several cell cycle proteins, including the mitotic cyclins, which are targeted by the anaphase-promoting complex/cyclosome (APC/C). The APC/C is large multimeric E3 ubiquitin ligase, whose activity depends on the adapter proteins Fizzy (Fzy/Cdc20) and Fizzy-related (Fzr/Cdh1). M phase is divided into two periods: mitosis, in which the duplicated genome is subdivided into two daughter nuclei and therefore is also referred to as karyokinesis; and cytokinesis, which describes the division of the cytoplasm. Cytokinesis starts with the bundling of microtubules between the separating anaphase chromosomes (Carmena et al. 2012; Fededa and Gerlich 2012). The resulting structure is called the midzone and serves as a scaffold for the assembly of the chromosomal passenger complex, which includes Aurora B kinase, INCENP, and survivin, as well as the centralspindlin complex consisting of MKLP1 and MgcRacGAP. An actin–myosin ring mediates the ingression of the cleavage furrow. The formation and the contraction of the actin–myosin ring rely on the GTPase RhoA, whose activity is regulated by Aurora B and the GTPase-activating protein MgcRacGAP. RhoA promotes the movement of actin filaments by maintaining active myosin II, likely through its association with Anillin, a cytoskeletal scaffold protein which links RhoA, actin, and myosin. After remodeling of the cell cortex, the daughter cells split during the process of abscission, which involves disassembly of the actin–myosin ring and fusion of membranes. In Drosophila, Cdk2 triggers S phase in conjunction with Cyclin E (CycE), whereas in mammals, Cdk2 associates with both CycE and CycA. Cdk2 activity is restricted by Cyclin-dependent kinase inhibitors (CKIs) of the CIP/KIP family, which consists of only Dacapo (Dap) in flies and p21/Cip1, p27/Kip1, and p57/Kip2 in vertebrates. In addition, the E2F transcription factor serves as an important regulator of G1–S transition. Different E2F family members act as either transcriptional activators or repressors and they control the transcription of many cell cycle genes. S phase initiation in mitotic cycles is also controlled by the binding and activation of proteins on chromatin at origins of DNA replication. Prior to S phase, pre-replication complexes (pre-RCs) are assembled at DNA replication origins distributed throughout

1.4  Genomic Organization of Polyploid Cells

the genome. This process is referred to as ‘origin licensing’ and involves the sequential assembly of the origin recognition complex (ORC), Cdc6, and Cdt1, which together load the Mcm2-7 DNA helicase onto chromatin. Restricting genome duplication to once per cell cycle requires that cells do not reinitiate nuclear DNA replication within regions that have already been replicated until cell division is complete. Pre-RC assembly occurs during anaphase and G1 phase when CDK activity is suppressed. Activation of the pre-RC, by contrast, requires CDK and Dbf4-dependent Cdc7 kinase (DDK) activities, which are up-regulated during the G1 to S phase transition. To prevent premature initiation of DNA replication prior to cell division, both flies and mammals use CDKs and ubiquitin ligases and the inhibitory protein Geminin to inactivate one or more pre-RC proteins.

1.4 ­Genomic Organization of Polyploid Cells Early studies of Drosophila polyploid tissues led to the realization that the genome is not entirely replicated during each endocycle S phase, as the heterochromatic regions flanking the centromere were not visible during cytological analyses of polytene chromosomes (Gall et al. 1971) and measurement of DNA content resulted in C-values that were not exact multitudes of the diploid genome (Hammond and Laird 1985a; Hammond and Laird 1985b; Smith and Orr-Weaver 1991). The invention of arraybased comparative genomic hybridization (aCGH) allowed the systematic analysis of differential replication, and therefore led to the identification of under-replicated regions in a variety of Drosophila polyploid tissues, including larval salivary glands, fat body, and midgut (Nordman et al. 2011; Sher et al. 2012). Furthermore, these studies demonstrated that under-replication is not restricted only to heterochromatin, but also occurs in non-repetitive gene coding regions. The comparison of aCGH profiles from different Drosophila tissues revealed tissue-specific differences in the sites and the degree of under-replication, suggesting that differential replication may contribute to tissue differentiation. The tissue-specific differences in the extent of under-replication do not appear to correlate with the degree of polyploidization, however. For example, the larval fat body and midgut undergo the same number of endocycles, but differ signficantly in their aCGH profiles. A crucial question that has yet to be fully answered is how differential replication is achieved at the molecular level. ChIP-chip analysis has been used to detect an enrichment of the repressive H3K27me3 mark in under-replicated regions, while binding of ORC protein was significantly reduced. Furthermore, the under-replicated regions are associated with the protein Suppressor of under-replication (Su(UR)) (Pindyurin et al. 2007). Mutation of Su(UR) results in full replication of the vast majority of under-­ replicated regions in Drosophila (Belyaeva et al. 1998; Nordman et al. 2011; Sher et al. 2012). However, tethering of Su(UR) to specific sites was insufficient to induce underreplication (Sher et al. 2012). The reduced association of ORC protein with chromatin in under-replicated regions was not restored in Su(UR)-deficient salivary glands, whereas the H3K27me3 mark was absent. Analysis of gene amplification in ovarian follicle cells revealed that loss of Su(UR) accelerates replication fork progression, and thus Su(UR) may reduce the replication efficiency in specific chromosomal regions. Altogether, these data suggest a model in which reduced ORC binding combined with

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impeded progression of replication forks initiated from distant origins give rise to chromosomal regions that are not replicated and thus are under-represented in ­ Drosophila polyploid tissues. Surprisingly, a recent study revealed that the genomes of megakaryocytes and TGCs are fully replicated, including heterochromatic sequences (Sher et  al. 2013). These results demonstrate that the mechanisms of polyploidization differ between mice and flies, but at the moment we can only speculate about the reasons for this phenomenon. Gene expression profiling revealed that S phase genes are down-regulated in endoreplicating salivary glands, but were expressed at normal levels in TGCs (Maqbool et al. 2010; Sher et al. 2013). The reduced expression of replication factors may slow down S  phase progression, and thus possibly cause under-replication of late replicating regions in Drosophila salivary glands, but not in TGCs. However, polyploid MKCs also showed reduced expression of S phase genes, albeit significantly higher than in salivary glands, suggesting that this cannot be the sole explanation. Alternatively, the discrepany could result from differences in genomic organization, as most of the heterochromatin of Drosphila is organized in large blocks, whereas in mice it is distributed throughout the genome.

1.5 ­Endoreplication: An Effective Tool for Post-Mitotic Growth and Tissue Regeneration Endoreplication cycles are the best-understood mechanism for inducing somatic polyploidy and are often utilized to increase cell size under conditions prohibiting mitotic divisions (Edgar and Orr-Weaver 2001; Lilly and Duronio 2005; Lee et al. 2009; Zielke et  al. 2013). The larval and adult growth of many invertebrate species, including the nematode Caenorhabditis elegans, the tunicate Oikopleura dioica, and the fruit fly Drosophila melanogaster, is largely achieved via endoreplication, and in these organisms cell size is roughly protional to ploidy. For example, a survey among nematode species revealed a correlation between body size and hypodermal ploidy, and thus gave rise to the hypothesis that the degree of polyploidy is a major determinant for the evolution of body size. Inhibition of endoreplication in the hypodermis of C. elegans by treatment with hydroxyurea or mutation of the essential S phase regulators CycE or MCM4 results in dwarfish worms (Lozano et  al. 2006; Korzelius et  al. 2011), demonstrating that endoreplication is essential for hypodermal growth. Likewise, a striking correlation between cell size and endocycle progression was observed in the epithelium of Oikopleura, which is entirely comprised of polyploid cells but displays regional difference in the degree of polyploidization (Ganot and Thompson 2002). Many larval tissues of Drosophila, including the salivary glands, fat body, epidermis, midgut, trachea, and Malpighian tubules, are comprised of polyploid cells (Smith and Orr-Weaver 1991). Arguably, the best-studied example are the giant salivary gland cells, which undergo approximately 10 endocycles, resulting in a final ploidy approximating 1035 C (Hammond and Laird 1985b). Starvation experiments in the fat body revealed that endocycle progression in larval tissues is responsive to nutrient availability (Britton and Edgar 1998), underscoring the interdependency of polyploidization and cellular growth. Endoreplication is also crucial for female germline development in Drosophila, as the nurse cells, which provide the maturating oocyte with mRNA and proteins, attain a

1.6  Initiation of Endoreplication in Drosophila

final ploidy of approximately 1500 C (Hammond and Laird 1985a), and the somatic follicle cells that surround the egg become 16 C. Loss of the ability to undergo endoreplication in nurse cells results in sterility, indicating that endoreplication is essential for oogenesis (Lilly and Spradling 1996). Importantly, a recent study also highlights the physiological relevance of polyploidization for cell growth during organ development. The subperineural glia (SPG), which encapsulate the neurons of the Drosophila brain, constitute an important part of the blood–brain barrier (Unhavaithaya and Orr-Weaver 2012). During larval development, the SPGs increase in size by polyploidization, which allows the SPGs to accommodate the growing neuronal mass without undergoing mitotic divisions in order to constantly maintain a barrier around the brain. Inhibition of SPG polyploidization disrupted the blood–brain barrier, demonstrating that polyploidization allows growth without losing tissue integrity. SPG polyploidization occurs partly through endoreplication, but also a large fraction of multinucleated cells was observed, implying that other mechanisms contribute to this process. This strategy may also be employed in vertebrate tissues, for example the trophoblasts that provide a barrier between embryonic and maternal tissues (Brandner et al. 2006), and the polyploid keratinocytes, which seal the skin epithelia (Wang et al. 2004; Gandarillas 2012). As endoreplication is an effective strategy for post-mitotic growth it is often associated with cellular hypertrophy, which refers to volume increases of organs or tissues due to enlargement of its cells. Compensatory cellular hypertrophy has a pivotal role during liver regeneration (Pandit et al. 2013). After injury or damage the remaining hepatocytes re-enter the endocycle, and the accompanying increase in volume alleviates the loss of cell mass. Likewise, the polyploid cells of the Drosophila follicular epithelium compensate for the loss of defective, out-competed cells by increased endoreplication and cellular hypertrophy (Tamori and Deng 2013).

1.6 ­Initiation of Endoreplication in Drosophila 1.6.1  Endocycle Entry in Ovarian Follicle Cells

The mitotic-to-endocycle switch has been most extensively studied in ovarian follicle cells of Drosophila, where mitotic and endoreplicating cells are found alongside each other and can be easily distinguished by morphological criteria (Spradling 1993). A single fly ovary is composed of approximately 16 ovarioles, each of which is composed of several egg chambers of sequentially maturing developmental stages (Figure 1.2). Each egg chamber consists of an oocyte and 15 germline nurse cells surrounded by a layer of somatic follicle cells. These follicle cells undergo multiple mitotic divisions, and then undergo three endocycles after stage 6 of oogenesis. The transition from a mitotic cycle into an endocycle involves re-wiring of the cell cycle regulatory network, which is primarily achieved by eliminating the activity of mitotic cyclin/Cdk complexes by proteasomal degradation of the cyclins or termination of their transcription (Figure 1.2). In endoreplicating follicle cells the mRNAs encoding the mitotic cyclins are constantly expressed, but the cyclin proteins become immediately ubiquitinated by APC/C–Fzr and degraded by the proteasome (Schaeffer et  al. 2004). Follicle cells lacking the APC/C activator Fzr accumulated high levels of the mitotic cyclins A and B and failed to execute endocycles (Schaeffer et al. 2004). Although

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Figure 1.2  Signaling pathways controlling cell cycle transitions in Drosophila ovarian follicle cells. (Top) A schematic depicts the development of the Drosophila egg chamber and delineates the points where follicle cells undergo cell cycle transitions. Each egg chamber consists of 16 germline cells, the diploid oocyte (red), and 15 polyploid nurse cells (orange), surrounded by a layer of somatic follicle cells (green). The follicle cells proliferate mitotically until stage 6, and then undergo three endocycles. At stage 10B, all genome-wide DNA replication ceases and amplification initiates at six specific loci. (Bottom left) Activation of the Notch signaling pathway in follicle cells is the major stimulus for the mitotic-to-endocycle transition. Notch signaling blocks mitosis in these cells by activating the transcriptional repressor Hnt. Hnt then represses the expression of Stg, as well as Cut, which triggers the degradation of the mitotic cyclins A and B by the APC/C–Fzr complex. In addition, Notch downregulates the expression of Dap, preventing it from inhibiting S phase. CoREST is a transcriptional cofactor required for proper Notch signaling. (Bottom right) The endocycle-to-amplification transition requires the inactivation of Notch signaling to allow signaling through EcR. The crucial event downstream of both Notch silencing and EcR activation is increased expression of the transcriptional repressor Ttk69, which is required for the switch to amplification. miR-7 negatively regulates Ttk69 expression and can block amplification when overexpressed. Notch silencing also causes the down-regulation of Hnt expression, which allows Cut to be expressed. It is likely that additional signaling pathways are also involved in this cell cycle transition.

these cells lacking Fzr accumulated high levels of CycA and CycB, the phase of mitotic proliferation was not extended beyond stage 6 of oocyte development. However, extra mitoses were induced in fzr mutants by ectopic expression of the Cdk1 phosphatase Stg, demonstrating that initiation of endoreplication requires both up-regulation of APC/C– Fzr activity and down-regulation of Stg (Schaeffer et  al. 2004). In follicle cells, the repression of the CycE/Cdk2 inhibitor Dap also appears to be a component of the mitotic-to-endocycle switch, as ectopic expression of Dap prevented endocycle entry

1.6  Initiation of Endoreplication in Drosophila

(Shcherbata et al. 2004). Follicle cells lacking Dap function displayed no obvious phenotypes, suggesting that Dap is not required for the follicle cell endocycle (Shcherbata et al. 2004; Hong et al. 2007). 1.6.2  Signaling Pathways Regulating Endocycle Entry in Follicle Cells

A key upstream regulator of the mitotic-to-endocycle transition in Drosophila follicle cells is the Notch signaling pathway (Deng et al. 2001; Lopez-Schier and St Johnston 2001). The oocyte expresses the Notch ligand Delta, which activates the Notch receptor and downstream signaling in the surrounding follicle cells. Deletion of Notch in follicle cells, or germline-specific deletion of Delta, prevented the initiation of endoreplication (Deng et al. 2001; Lopez-Schier and St Johnston 2001). Conversely, experimental activation of Notch signaling resulted in repression of Stg and Dap as well as up-regulation of Fzr (Schaeffer et al. 2004; Shcherbata et al. 2004). Moreover, deletion of Delta from the germline resulted in accumulation of Dap, demonstrating that activation of Notch signaling represses Dap in endoreplicating follicle cells (Shcherbata et al. 2004). Notch mediates the mitotic-to-endocycle transition in follicle cells through a cascade of transcription factors (Figure 1.2). Based on multiple lines of evidence, Sun and Deng proposed that Notch signaling activates the transcriptional repressor Hindsight (Hnt), which subsequently suppresses the expression of Stg/Cdc25 as well as another transcription factor, Cut (Sun and Deng 2005; Sun and Deng 2007). The down-regulation of Cut at the mitotic-to-endocycle transition allows Fzr to accumulate, which in turn mediates the proteasomal degradation of CycA and CycB. It remains unclear, however, whether Fzr and Stg are direct targets of these transcription factors, and whether Hnt and Cut mediate the Notch-dependent down-regulation of Dap. In summary, these data suggest that endocycle entry in follicle cells occurs in two steps: First, the Hnt-mediated down-regulation of Stg arrests the cell at the G2/M transition, and then the down-regulation of Cut and subsequent de-repression of Fzr bypasses mitosis and pushes the cell directly into a G1-like state (Sun and Deng 2007). A recent study has also identified the transcriptional cofactor CoREST as a positive modulator of Notch signaling during the mitotic-to-endocycle switch in follicle cells (Domanitskaya and Schupbach 2012). Notch signaling is faulty in CoREST mutant cells, which fail to up-regulate Hnt and down-regulate Cut and do not switch from the mitotic cycle to the endocycle on schedule. The authors of this study found that CoREST regulates Notch signaling downstream of Notch receptor proteolytic cleavage but upstream of Hnt expression. Further, CoREST mutant follicle cells experience increased histone H3K27 tri-methylation and H4K16 acetylation at the stage when they should be beginning endoreplication, suggesting that CoREST may epigenetically modify Notch target gene loci and thereby affect the transcription of those genes. Intriguingly, the Notch pathway may also be important for the mitotic-to-endocycle transition in the ovarian follicle cells of another insect species, the beetle Tribolium castaneum (Baumer et  al. 2012). Tribolium Notch is expressed ubiquitously in both germline and somatic cells of the ovary, and Delta, similar to Drosophila, is expressed only in the germline. However, in Tribolium ovaries that have been treated with either Notch or Delta RNAi, it appears that mitotic cycles switch prematurely to endocycles, indicating that Notch signaling in this context has the opposite effect to that in Drosophila in that it maintains mitotic proliferation.

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Are other pathways besides Notch signaling involved in the mitotic-to-endocycle transition? Follicle cells mutant for patched (ptc), a negative regulator of the signaling molecule Hedgehog (Hh), fail to initiate endocycles, suggesting that the Hh pathway may antagonize the endocycle-promoting Notch signal (Zhang and Kalderon 2000). Sun and Deng (2007) have proposed a mechanism in which Hnt terminates the phase of mitotic proliferation by suppressing the transcription of the Hh-effector cubitus interruptus (Ci). Furthermore, genetic interaction experiments suggested that down-regulation of Ci by Hnt also requires the continuously expressed Zn-finger transcription factor Tramtrack69 (Tkk69) (Jordan et al. 2006; Sun and Deng 2007). Additionally, the Salvador–Warts–Hippo (SWH) tumor-suppressor pathway has been demonstrated to play an important role in the entry into endocycles specifically in the posterior follicle cells, which are important for body axis formation (Meignin et al. 2007; Polesello and Tapon 2007; Yu et al. 2008). The core of the SWH pathway consists of the serine/threonine kinases Hippo (Hpo) and Warts (Wts) and the scaffold proteins Salvador (Sav) and Mats, which function to inactivate the transcription factor Yorkie (Yki). In the posterior follicle cells, mutation of hpo, wts, or sav, or overexpression of yki, disrupts Notch signaling. As a result, these cells do not express Hnt or repress the expression of Cut after stage 6 and they continue to divide mitotically. 1.6.3  Endocycle Entry in Other Tissues

The mechanisms of endocycle initiation have also been studied in the Drosophila salivary gland, which is technically challenging as the transition to the endocycle occurs during embryogenesis and involves only a relatively small number of cells (Smith and Orr-Weaver 1991). As in follicle cells, endocycle entry in salivary glands requires APC/C–Fzr-mediated degradation of mitotic cyclins (Sigrist and Lehner 1997), but in this case it also involves the transcriptional down-regulation of mitotic regulators (Zielke et  al. 2008; Maqbool et al. 2010). Recent work has demonstrated that suppression of mitotic regulators in endoreplicating salivary glands involves E2F-mediated repression (Zielke et al. 2011). The relatively small fly genome contains only a single activator E2F, E2F1, and a single repressor, E2F2, which both associate with the same dimerization partner, namely dDP (van den Heuvel and Dyson 2008). Salivary glands derived from E2F2 mutants, or from animals overexpressing E2F1, displayed ectopic expression of Cdk1, CycA, and CycB3 mRNAs (Zielke et al. 2011). Cdk1 and CycA have been identified as E2F targets in mitotic cells (Ishida et al. 2001; Ren et al. 2002), suggesting that E2F2 may act as a selectivity factor, preventing E2F1 from activating mitotic targets during endoreplication. The decision to switch to endoreplication cycles also involves the Escargot (Esg) protein, which belongs to the family of Snail transcription factors. Esg is expressed in most imaginal tissues, but is absent in the polyploid tissues of the larvae (De Miglio et  al. 1999; Polesello and Tapon 2007). Abdominal histoblasts lacking Esg become polyploid, whereas ectopic expression of Esg abolishes endoreplication in larval salivary glands (Fuse et  al. 1994; De Miglio et  al. 1999). The esg phenotype resembles that of Cdk1 mutants, which also undergo ectopic endocycles, and further experiments revealed that Esg indeed genetically interacts with Cdk1 (Stern et al. 1993; Hayashi 1996; Weigmann et al. 1997). Tellingly, the abdominal histoblasts in hypomorphic esg mutants showed reduced levels of CycA protein (Hayashi 1996). These findings suggest that Esg is

1.7  Mechanisms of Endocycle Oscillations in Drosophila

required in diploid cells to sustain mitotic Cyclin/Cdk activity and that its function is dampened when cells switch to endoreplication cycles. Altogether, evidence from several Drosophila cell types indicates that the suppression of mitotic regulators upon entry into endocycles involves multiple mechanisms that act at both the transcriptional and post-transcriptional level.

1.7 ­Mechanisms of Endocycle Oscillations in Drosophila 1.7.1  An Autonomous Oscillator Drives Endocycling in the Salivary Gland

Endoreplicating cells utilize the same set of S phase genes as mitotic cells, but upon endocycle entry the underlying regulatory network becomes reorganized to adjust to the fact that mitosis is suppressed. The expression of many genes involved in DNA replication culminates right before S phase, and relies, as in all higher eukaryotes, on the activity of the transcriptional activator E2F1 (Duronio et al. 1995). Deletion of E2F1 impairs the polyploidization of larval tissues, whereas mild overexpression of E2F1 promotes endoreplication in the salivary gland, resulting in hyperpolyploidy (Royzman et al. 1997; Zielke et al. 2011). Like in mitotic cells, E2F1 protein accumulates during G phase, but is rapidly destroyed upon initiation of S phase (Asano et al. 1996; Weng et al. 2003; Reis and Edgar 2004). The S phase-specific degradation of E2F1 is mediated by the CRL4-Cdt2 E3 ligase and relies on the fact that activation of CRL4-Cdt2 requires chromatin-bound PCNA, which exists only at active replication forks (Shibutani et al. 2007; Shibutani et al. 2008; Havens and Walter 2009; Zielke et al. 2011). Substrates of CRL4-Cdt2 are characterized by the PCNA-interacting protein (PIP) motif, which is crucial for the association with PCNA (Arias and Walter 2006; Havens and Walter 2009). Consequently, mutation of the N-terminal PIP box in E2F1 inhibits its turnover (Shibutani et al. 2008). Tellingly, either depletion of CRL4-Cdt2 or stabilization of E2F1 by deletion of the PIP box blocks endocycle progression in salivary glands (Zielke et al. 2011). Altogether, these data demonstrated that endocycling in Drosophila salivary glands relies on an autoregulatory feedback loop comprised of E2F1 and CRL4-Cdt2, and thus provides the basis for the periodic accumulation of S phase genes (Figure 1.3). E2F1 is essential for both the expression and oscillation of CycE protein in Drosophila during both mitotic cell cycles and endocycles (Duronio and O’Farrell 1995; Duronio et al. 1996). In CycE-deficient embryos, DNA synthesis is abolished in cells undergoing either mitotic cell cycles or endocycles (Knoblich et al. 1994), and endocycles in salivary glands are eliminated by tissue-specific removal of Cdk2/cdc2c (Zielke et  al. 2011). CycE is an unstable protein (Lilly and Spradling 1996; Weng et al. 2003), whose proteasomal degradation relies on the E3 ubiquitin ligase CRL1-Ago and the specificity factor minus (mi) (Moberg et al. 2001; Szuplewski et al. 2009). So far, there is no evidence that CRL1-Ago activity oscillates in endoreplicating cells. Therefore its main function may be to render CycE constitutively unstable, rather than acting as part of the endocycle core oscillator. Salivary glands and follicle cells deficient for either ago or mi and salivary gland cells continuously overexpressing CycE fail to undergo DNA replication (Follette et al. 1998; Weiss et al. 1998; Shcherbata et al. 2004; Szuplewski et al. 2009; Zielke et al. 2011), implying that endocycle progression depends on the periodic accumulation and degradation of CycE/Cdk2 activity. The inhibitory effect of CycE is unique

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(b)

Figure 1.3  Endocycle oscillations. (A) Model of the autonomous oscillator that drives the salivary gland endocycles. The transcriptional activator E2F1 accumulates during the late G(ap) phase and stimulates CycE transcription, which triggers S phase in conjunction with its kinase partner Cdk2. Ongoing DNA replication activates the E3 ubiquitin ligase CRL4-Cdt2, which marks E2F1 for proteasomal degradation. The CRL1–Ago complex continuously targets CycE for degradation, and hence this negative feedback loop causes a drop in CycE levels, which is a prerequisite for origin licensing during the next G phase. CycE/Cdk2 also inhibits APC/C–Fzr activity, thereby allowing accumulation of the licensing inhibitor Geminin. (B) The regulatory network that controls the endocycle progression in trophoblast giant cells (TGC). S phase in TGCs relies on Cdk2, which associates with either CycE or CycA. During late G phase a peak of CycE/Cdk2 activity triggers S phase and concomitantly inhibits the ACP/C–Fzr complex, thereby promoting the accumulation of CycA and Geminin. CycA/Cdk2 phosphorylates CycE, which in turn will be recognized by E3 ligase CRL1-Fbw7 and thus marked for proteasomal degradation. The cessation of CycE/Cdk2 activity permits accumulation of CKI p57, which is no longer antagonized by the CRL1–Skp2 complex. The synergistic action of three factors, CRL1-Fbw7, CRL3, and p57, ensures low levels of CycE/Cdk2 activity during G phases, thereby allowing origin licensing to occur.

to endoreplicating tissues, as ectopic expression of CycE did not interfere with DNA synthesis in mitotically proliferating wing discs (Neufeld et  al. 1998) or follicle cells undergoing gene amplification (Calvi et al. 1998a). A critical CycE/Cdk2-target is the APC/C–Fzr complex, whose activity is essential for endocycle progression in salivary glands and follicle cells (Narbonne-Reveau et al. 2008; Zielke et al. 2008). High levels of CycE/Cdk2 activity trigger S phase and concomitantly inhibit APC/C–Fzr activity (Sigrist and Lehner 1997; Reber et al. 2006), thereby promoting the accumulation of Geminin and Orc1. Low CycE/Cdk2 activity during G phase, by contrast, releases the APC/C–Fzr complex, which subsequently degrades Geminin and thus allows assembly of pre-RCs. Consistently, it was found that depletion of APC/C–Fzr activity stabilizes Geminin and blocks endocycle progression (Zielke et al. 2008). Although Geminin is essential for cell proliferation in flies (Quinn et al. 2001), it appears to be dispensable for normal endoreplication in salivary glands, suggesting that CycE/Cdk2 prevents re-replication in endoreplicating salivary glands through multiple redundant mechanisms (Zielke et al. 2011).

1.8  Gene Amplification in Drosophila Follicle Cells

1.7.2  Alternative Modes of Endoreplication

Currently, the autoregulatory feedback loop based on E2F1 and CRL4-Cdt2 has only been demonstrated in the Drosophila salivary gland, but considering the diversity of species undergoing endocycles, it seems likely that a number of variant mechanisms have evolved. Interestingly, the mechanisms of endoreplication appear to vary even among Drosophila tissues. The endocycles of the ovarian nurse cells represent a notable exception, as these cycles involve the CKI Dap, which is dispensable in most other endoreplicating tissues including salivary glands, follicle cells, and socket and shaft cells of the bristle lineage (Shcherbata et al. 2004; Audibert et al. 2005; Zielke et al. 2011). In endoreplicating nurse cells, Dap accumulates only in cells expressing high levels of CycE/Cdk2, which leads to a subsequent decrease of CycE/Cdk2 activity (de Nooij et al. 2000). Late replicating heterochromatic sequences are under-represented in many polyploid Drosophila tissues, including the ovarian nurse cells (Nordman et al. 2011; Sher et al. 2012), but in Dap-deficient nurse cells where CycE/Cdk2 activity is extended, S phase is lengthened and these sequences are nearly fully replicated (Hong et  al. 2007). It has been hypothesized that the negative feedback loop between CycE/Cdk2 activity and Dap creates a window of low CDK activity during the next G phase, and thus allows pre-RC formation. However, Dap-mutant nurse cells can undergo fairly normal endocycles (Hong et al. 2003), suggesting that redundant mechanisms safeguard pre-RC formation during the Gap phase and that Dap is only required for ensuring proper replication timing. The endoreplicating cells of the mechanosensory bristle lineage employ another endocycle variant that involves oscillations of CycA protein (Salle et al. 2012). The presence of CycA in endoreplicating shaft and socket cells is surprising, as the expression of many mitotic regulators, including CycA, is dampened in salivary glands (Zielke et al. 2008; Maqbool et al. 2010). In shaft and socket cells, CycA protein specifically accumulates during late S phase, and both loss and gain of function results in decreased ploidy, delayed S phase, and dramatically affected association of Orc2 with heterochromatic regions (Salle et al. 2012), suggesting that CycA/Cdk1 regulates Orc2 localization and thereby modulates replication timing in endoreplicating bristle cells.

1.8 ­Gene Amplification in Drosophila Follicle Cells 1.8.1  Molecular Mechanism of Gene Amplification

During Drosophila oogenesis, six loci within the ovarian follicle cells undergo developmentally programmed DNA amplification by repeated firing of origins of replication. Amplification at these sites is required to meet the demand for high levels of expression of proteins needed for eggshell synthesis. Indeed, a reduction in amplification leads to thin eggshells and female sterility. These six loci are known as Drosophila amplicons in follicle cells (DAFCs) (Claycomb et al. 2004b; Kim et al. 2011), and they are named by their cytological positions in the genome. The amplicons DAFC-7F and DAFC-66D correspond to clusters of the chorion (eggshell) genes on the X and 3rd chromosomes, respectively (Spradling and Mahowald 1980; Claycomb et  al. 2004a). Several genes expressed in the follicle cells of late-stage egg chambers are encoded in the DAFC-30B and DAFC-62D amplified loci, including the gene yellow-g, which is likely required for

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proper vitelline membrane formation (Claycomb et  al. 2004a). The genomic regions amplified in DAFC-22B and DAFC-34B have been identified most recently, and DAFC34B contains the gene Vm34Ca that encodes a structural component of the vitelline membrane (Kim et al. 2011). DAFC-66D, the amplicon that achieves the highest increase in DNA copy number, begins to amplify periodically during the asynchronous S phases that occur during the three follicle cell endocycles, attaining an approximately four-fold increase in copy number by the end of endoreplication (Calvi et al. 1998a). All six amplified loci then synchronously enter into continuous amplification at stage 10B, immediately after endocycles have completed (Calvi et al. 1998a). The maximal increases in DNA copy number that have been observed by quantitative real-time PCR are 30-fold for DAFC-66D, 14-fold for DAFC-7F, and 4- to 6-fold for DAFC-30B, DAFC-62D, DAFC-22B, and DAFC-34B (Claycomb et al. 2002; Claycomb et al. 2004a). The amplified region at each locus is a gradient with the highest DNA copy number near the origins of replication and each spans in total 75–100 kb (Spradling and Mahowald 1980; Spradling 1981; Claycomb et al. 2004a; Kim et al. 2011). For DAFC-7F, DAFC-66D, and DAFC-30B, all DNA replication initiation events have completed by stage 10B or 11, and after that, further increases in DNA copy number only occur at greater distances from the origins by elongation by existing replication forks (Claycomb et  al. 2002; Claycomb et  al. 2004a). In contrast, DAFC-62D undergoes a late replication initiation event between stages 12 and 13, resulting in a relatively small maximally amplified region (Claycomb et al. 2004b). Amplification can be visualized by a punctate pattern of BrdU incorporation in follicle cells from stage 10B on (Calvi et al. 1998a). In addition, amplifying loci can be visualized by immunostaining for several proteins that bind DNA at origins of replication, such as ORC1, ORC2, ORC5, CDC45, DUP/Cdt1, PCNA, and the MCM2-7 complex, as the factors known to function at all origins of DNA replication also function at the DAFCs (Asano and Wharton 1999; Austin et al. 1999; Royzman et al. 1999; Loebel et al. 2000; Whittaker et al. 2000; Claycomb et al. 2002). As further evidence that DNA replication factors are required for amplification of the DAFCs, hypomorphic mutations in genes encoding necessary DNA replication proteins such as double-parked (dup/cdt1), origin recognition complex subunit 2 (orc2), chiffon (chif, dbf4-like), proliferating cell nuclear antigen (pcna, mus209), minichromosome maintenance factor 6 (mcm6), and humpty dumpty (hd, fs(3)272) result in reduced amplification, thin eggshells, and female sterility (Underwood et al. 1990; Landis et al. 1997; Landis and Tower 1999; Henderson et al. 2000; Whittaker et al. 2000; Schwed et al. 2002; Bandura et al. 2005). Moreover, amplification is under the control of the S phase regulators E2F and CycE/Cdk2. The levels of both E2F1 protein and CycE/Cdk2 activity oscillate in mitotic and endoreplicating follicle cells, but coincident with the onset of amplification both markers become evenly distributed specifically within the follicle cells undergoing amplification (Calvi et al. 1998a; Sun et al. 2008). Either inhibition of CycE/Cdk2 with Dap or mutation of E2f1, E2f2, Dp, or Rbf selectively disrupts amplification (Calvi et al. 1998b; Royzman et al. 1999; Bosco et al. 2001; Cayirlioglu et al. 2001). If the same factors required for DNA replication at all origins are also utilized at the chorion origins and other DAFCs during amplification, what instructs the amplifying origins to fire repeatedly while all other origins in the genome remain silent? The answer to this question has not been fully elucidated, but epigenetic regulation appears to be part of the explanation. For example, it has been demonstrated that histone acetylation

1.9  Endocycle Entry in the Trophoblast Lineage

at the DAFCs is crucial for activity of these origins during amplification. Hyperacetylation of histones H3 and H4 has been observed specifically at the DAFCs in amplificationstage follicle cells (Aggarwal and Calvi 2004; Hartl et al. 2007; Liu et al. 2012). Further, increasing the levels of acetylation throughout follicle cell nuclei by either mutating or inhibiting histone deacetylases (HDACs) resulted in extra genomic DNA replication during amplification stages (Aggarwal and Calvi 2004). Consistent with these data, tethering a histone acetyltransferase (HAT) to an amplification reporter containing the chorion origin at DAFC-66D increased amplification, whereas tethering an HDAC reduced it. A recent study has extended these results by identifying two HATs, Chameau/ HBO1 and CBP/Nejire, that bind to DNA at DAFC-66D and contribute substantially to the acetylation seen at all DAFCs (McConnell et al. 2012). In addition, the Myb-MuvB (MMB) or dREAM complex binds to the regulatory DNA sequences of at least one of the DAFCs and constrains genomic DNA replication during amplification (Beall et al. 2002; Beall et al. 2004; Beall et al. 2007). MMB/dREAM recruits chromatin modifiers and is believed to act as either an activator or repressor of replication, depending on developmental context and genomic location (Beall et al. 2002; Beall et al. 2004; Korenjak et al. 2004; Lewis et al. 2004; Beall et al. 2007; Georlette et al. 2007). This is consistent with earlier work that had demonstrated roles for both E2F2 and Rbf in restraining genomic DNA replication in amplification-stage follicle cells, as both E2F2 and Rbf are members of MMB/dREAM (Cayirlioglu et al. 2001; Cayirlioglu et al. 2003). 1.8.2  The Endocycle-to-Amplification Switch

It is currently only poorly understood how the switch from endocycles to amplification is regulated. Continuous Notch activation suppresses gene amplification in follicle cells, which instead execute an additional round of endoreplication, suggesting that downregulation of Notch signaling is a prerequisite for the endocycle-to-amplification switch (Sun et al. 2008). In addition, the endocycle-to-amplification switch coincides with the up-regulation of the insect hormone ecdysone, and ectopic expression of a dominantnegative ecdysone receptor (EcR) prevents the proper initiation of gene amplification (Sun et al. 2008). Thus both down-regulation of Notch and activation of EcR are essential for the endocycle-to-amplification switch (Figure 1.2). Increased expression of the transcriptional repressor Ttk69 at the endocycle-to-amplification transition appears to be the crucial event downstream of Notch down-regulation and EcR activation, and expression of Ttk69 at this transition is further modulated by the microRNA miR-7 (Sun et al. 2008; Huang et al. 2013). The combination of Notch down-regulation and activation of EcR, or direct up-regulation of Ttk69, were not sufficient to induce a premature transition to amplification, and so other signaling pathways are likely involved in this process (Sun et al. 2008). Further, the mechanism by which these signaling events may lead to chromatin modifications specifically at the DAFC origins has yet to be illuminated.

1.9 ­Endocycle Entry in the Trophoblast Lineage TGCs are the paradigm for endoreplication in mammals and contain, like the endoreplicating tissues of Drosophila, large polytene chromosomes with DNA contents up to 512 C (Barlow and Sherman 1974; Varmuza et  al. 1988; Sher et  al. 2013). TGCs are

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derived from the trophectoderm of blastocysts, the outer layer of epithelial cells that give rise exclusively to the various cells comprising the placenta. The large size of the TGCs facilitates their function as a barrier between the maternal blood supply and the embryo proper (Rossant and Cross 2001; Cross 2005; Hu and Cross 2010). TGCs originate from trophoblast stem cells (TSCs) and initiate endoreplication without undergoing mitotic divisions in the absence of fibroblast growth factor 4 (FGF4) (Tanaka et al. 1998). TSCs can be maintained in culture and differentiated into endoreplicating TGCs, but under these conditions their DNA content rarely exceeds 64 C (Tanaka et al. 1998). Although cultivated TGCs only execute a limited number of endocycles, they are currently the vertebrate cell type most suitable for in vitro studies on endoreplication. Similar to endocycle entry in Drosophila, the onset of endoreplication in TGCs requires the suppression of mitotic CDK activity, while the activity of S phase Cdks is not affected (Ullah et al. 2008). Endoreplication in TSCs is triggered by withdrawal of FGF4, which induces the expression of the CKIs p57 and p21 and thereby prevents the activation of Cdk1 and initiation of mitosis (Ullah et al. 2008). The switch to endoreplication also involves Checkpoint Kinase-1 (Chk1), a kinase that is part of the DNA damage checkpoint (Ullah et  al. 2011). In proliferating TSCs, Chk1 phosphorylates and thereby targets p57 and p21 for proteasomal degradation. The expression of Chk1 ceases when FGF4 is withdrawn from TSCs, thus allowing unphosphorylated p57 and p21 proteins to accumulate and the subsequent switch to endoreplication. However, only p57 is essential for the mitotic to endocycle transition, as FGF4 deprivation induces endoreplication in p21-deficient TSCs, whereas p57-depleted TSCs undergo acytokinetic mitosis instead (Ullah et  al. 2008). Treatment with the Cdk1 inhibitor RO3306 induces endoreplication in wild-type and p57-deficient TSCs, demonstrating that the main function of p57 is to constrain Cdk1 activity and that its activity is negligible after the onset of endoreplication (Hochegger et al. 2007; Ullah et al. 2008). Examination of p57 knockout mice revealed multiple developmental abnormalities, including placentomegaly and hyperplasia of labyrinthine trophoblasts and spongiotrophoblasts, but the DNA content of the TGCs was not significantly changed (Zhang et al. 1998; Takahashi et al. 2000; Kanayama et al. 2002), suggesting that trophoblasts can initiate endoreplication by alternative, p57-independent mechanisms. Knockout of the APC/C activator protein Cdh1 results in embryonic lethality due to defective endoreplication in extraembryonic tissues (Garci-Higuera et al. 2008; Li et al. 2008), and a recent study revealed that deletion of Cdh1 prevents the onset of endoreplication in cultivated TSCs, which instead continued proliferating (Naoe et al. 2013). The deletion of Cdh1 was accompanied by stabilization of CycA and CycB, indicating that APC/C-mediated suppression of mitotic cyclins is a general mechanism of endocycle entry that is conserved between flies and mice. In addition, the Cdh1-deficient cells exhibited reduced expression of TGC markers, implying interdependency between endoreplication and differentiation. In the trophoblast lineage the switch between mitotic proliferation and endoreplication also involves the Snail transcription factor mSma, which shows striking structural and functional similarities with Drosophila Escargot (Nakayama et al. 1998). Consistent with the fly model, the expression of mSma mRNA is restricted to the proliferating TGC precursors, but absent in endoreplicating TGCs. Moreover, overexpression of mSma in the rat choriocarcinoma-derived cell line Rcho-1 (Faria and Soares 1991) impaired differentiation into giant cells, while the number of differentiating Rcho-1 cells increased upon mSma depletion. The overexpression was accompanied with

1.9  Endocycle Entry in the Trophoblast Lineage

elevated levels of the mitotic Cyclins A and B, but did not affect endocycle progression. The latter observation stands in contrast to the inhibitory effect in Drosophila salivary glands, but can most likely be explained by the fact that CycA is normally present in endoreplicating Rcho-1 cells (MacAuley et al. 1998) and CycB does not affect S phase. Altogether, we conclude that the mSna and Escargot transcription factors either directly or indirectly control the expression of mitotic regulators, and thus antagonize differentiation into endoreplicating cell types. Down-regulation of Geminin also has been implicated in endocycle entry, because loss of Geminin in the mouse embryo or embryonal carcinoma and embryonic stem (ES) cells resulted in over-replication and expression of trophoblast-specific genes (Gonzalez et  al. 2006; Yang et  al. 2011). These results, however, are controversial as other studies did not describe increased DNA contents in Geminin-depleted mouse ES cells (Yellajoshyula et al. 2011), or report DNA re-replication and apoptosis at the eightcell stage of Geminin-deficient embryos (Hara et al. 2006). Considering that Geminin levels do not change during TGC differentiation (Ullah et al. 2008), and that Geminin oscillates during endocycles in Drosophila (Zielke et al. 2008), the most plausible explanation is that Geminin prevents re-replication during endocycles, ensuring that the genome is duplicated in discrete entities. Recent work has demonstrated that the Rb–E2F pathway regulates the onset of endoreplication in the trophoblast lineage. The mammalian E2F family includes nine members, of which three are transcriptional activators (E2F1–3a) and six are transcriptional repressors (E2F3b–8) (van den Heuvel and Dyson 2008; Chen et al. 2009). E2F1-6 forms heterodimers with members of the DP family (Dp1, Dp2, and Dp4), which is crucial for DNA binding. The atypical repressors E2F7 and E2F8, in contrast, form homo- or heterodimers and function independently of a distinct dimerization partner (Lammens et al. 2009). Deletion of E2F7/8 results in defects during placental development and reduced fetal viability (Ouseph et al. 2012), while placentation is unaffected by deletion of the canonical repressors E2F4 or E2F5 (Chen et al. 2012). TGC-specific deletion of E2F7/8 was accompanied by a significant increase in cells executing aberrant mitosis, resulting in the formation of abnormal binucleate cells (Chen et  al. 2012). Moreover, expression profiling of E2F7/8-deficient TGCs revealed increased levels of genes associated with mitosis, but no effect on p57 mRNA expression (Chen et al. 2012). Concomitant deletion of both isoforms of CycA rescued the reduced DNA contents in E2F7/8-deficient TGCs (Chen et al. 2012), strongly supporting the idea that the defects in E2F7/8-mutant placentas is due to unscheduled mitotic divisions. E2F-mediated suppression of mitotic genes appears to be a general feature of endoreplicating cell types, as ectopic accumulation of mitotic regulators was also observed in Drosophila salivary glands deficient for the repressor E2F2 (Zielke et al. 2011). Surprisingly, reduced ploidy in E2F7/8-deficient TGCs did not affect placental development or reduce fetal viability (Chen et al. 2012). Although most cells maintained residual polyploidy, these results suggest that high ploidy (more than 64 C) is dispensable for placenta function. An explanation could be that increased ploidy is only advantageous under stressed conditions requiring high metabolic activity. Furthermore, it remains to be elucidated whether impaired polyploidy affects age-related processes like cancer formation. In contrast to Drosophila, where endoreplication cannot occur in the absence of the activator E2F1 (Zielke et al. 2011), the deletion of all activator E2Fs (E2F1–3) in TGCs was accompanied by increased ploidy (Chen et  al. 2012), implying that mammalian

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activator E2Fs restrict endocycling in TGCs. Concomitant knockout of either E2F1 or E2F3a rescued the phenotypes observed in E2F7/8 mutants (Chen et al. 2012; Ouseph et al. 2012), underscoring the notion that the balance between activator and repressor E2Fs controls the switch between mitotic and endoreplication cycles. Deletion of Dp1 resulted in embryonic lethality, which was accompanied by severely compromised extra-embryonic tissues, reduced numbers of trophoblast precursors, and impaired endoreplication in TGCs (Kohn et  al. 2003), suggesting that Dp1 is involved in endocycle entry and progression. The pleiotropic nature of the Dp1 phenotype presumably results from the fact that loss of Dp1 affects both transcriptional activation and repression. Mammals possess three Retinoblastoma-like pocket proteins (Rb, p107, p130), which associate with different subsets of E2F proteins. RB acts as transcriptional repressor when bound to either E2F1, E2F2, or E2F3a, whereas complexes consisting of RB and E2F4, E2F5, or E2F3b allow recruitment of chromatin-modifying enzymes to E2F-regulated promoters and thereby silence these genes (van den Heuvel and Dyson 2008). Rb knockout mice displayed defects in erythropoiesis and placental architecture as well as pronounced hyperproliferation and apoptosis, whereas p107 and p130 knockout mice developed essentially normally (Lipinski and Jacks 1999). Rb functions specifically in TSCs and its loss resulted in ectopic proliferation of trophoblasts due to elevated expression of the activator E2F3a (Wenzel et  al. 2007). Consequently, this phenotype was rescued by simultaneous deletion of Rb and E2F3a (Chong et al. 2009).

1.10 ­Mechanisms of Endocycle Oscillations in Trophoblast Giant Cells The molecular mechanism of endoreplication in mammals is only poorly understood, but data derived from knockout mice and endoreplicating cell lines suggest that endoreplication in TGCs relies on a two-phased oscillator involving S phase-specific CDK complexes as well as multiple CKIs and E3 ligases providing negative feedback (Figure 1.3). Initial studies on Rcho-1 cells revealed that levels of CycE and CycA remain high in endocycling TGCs, while the expression of CycB ceases upon onset of endoreplication (MacAuley et al. 1998). Deletion of both CycE (CcnE) alleles in mice abolishes endocycle progression in TGCs (Geng et al. 2003; Parisi et al. 2003), demonstrating that CycE-dependent kinase activity is essential for the execution of endoreplication cycles in both flies and mammals. The level of p57 protein oscillates during the endocycles of TGCs as well as Rcho-1 cells, while p57 mRNA is constantly expressed (Hattori et al. 2000; Ullah et al. 2008). The oscillation of p57 relies on CRL1-Skp2, which ubiquitinates CDK-phosphorylated p57 and thereby marks it for proteasomal degradation (Kamura et  al. 2003). Ectopic expression of a stabilized p57 variant blocks DNA synthesis in Rcho-1 cells (Hattori et  al. 2000), indicating that the oscillation of p57 is crucial for endocycle progression. p57 accumulates only during the G phase of endocycles, but is rapidly degraded upon initiation of S phase. The periodic accumulation of p57 antagonizes CycE/Cdk2 activity during S phase, and thereby releases the APC/C–Cdh1 complex. The subsequent degradation of CycA creates a window of low Cdk2 activity that allows pre-RC formation in the next G phase.

1.11 Cardiomyocytes

However, endoreplication occurred essentially normally in p57-depleted TGCs that were differentiated by drug treatment (Ullah et al. 2008). Moreover, TGCs were found in p57-deficient mice (Zhang et al. 1998), suggesting that p57 either is not a part of the core oscillator that drives the TGC endocycle or has redundant functions. At least a partial explanation could be that the accumulation of CycE protein is also negatively regulated by two different cullin-RING E3 ligases. The first pathway involves Cullin 1 and the F-box protein Fbw7 (the mammalian ortholog of Drosophila ago), whereas the second pathway(s) involves at least one Cullin 3-based E3 ligase. Placentas derived from Cullin 1 or Fbw7-deficient mice displayed high levels of CycE, but in either case the TGCs continued endoreplicating (Wang et  al. 1999; Tetzlaff et  al. 2004). Deletion of Cullin 3 resulted in early embryonic lethality and an abnormally developed trophoectoderm with fewer and smaller TGCs (Singer et al. 1999). The Cul3-depleted TGCs accumulated high levels of CycE protein, but failed to incorporate BrdU.

1.11 ­Cardiomyocytes The heart is one of the first organs to form in the embryo, and it must grow both prenatally and post-natally while still beating and pumping blood (Lacroix and Maddox 2012). Before birth, growth of the muscle cells of the heart (cardiomyocytes) is accomplished mainly due to mitotic divisions (hyperplasia). The myofibrils, the contractile filaments within each cardiomyocyte, are briefly disassembled during each cell division in order for cytokinesis to occur (Ahuja et al. 2004). In contrast, beginning shortly after birth the heart grows by enlargement of individual cardiomyocytes without any accompanying cell division (hypertrophy) (Leu et  al. 2001; Lacroix and Maddox 2012). In concert with the onset of cardiomyocyte hypertrophy, polyploidization is initiated in these cells at post-natal day 4 in rodents (Li et al. 1996; Pandit et al. 2013). By adulthood, up to 70% of human and 85% of rodent cardiomyocytes are polyploid (Soonpaa et al. 1996; Mollova et al. 2013). Cardiomyocytes generally become polyploid by means of an acytokinetic mitosis. Shortly after birth, these cells continue to undergo DNA synthesis and nuclear division, but lose almost all ability to undergo cytokinesis (Lacroix and Maddox 2012). After mitosis, these cardiomyocytes build an actin–myosin contractile ring but are unable to successfully complete cytokinesis, giving rise to polyploid cells containing two diploid nuclei (Li et al. 1997). Although acytokinetic mitoses are the most common mechanism by which polyploidy is achieved in cardiomyocytes, some cells with single 4 C or 8 C nuclei have been observed, suggesting that endoreplication likely also contributes to polyploidy in this cell type (Mollova et al. 2013). 1.11.1  Upstream Control of Cardiomyocyte Polyploidization

The mechanisms responsible for cardiomyocyte polyploidization have not been completely worked out, but several factors potentially implicated in this process have recently been identified. One possible mechanism involves the interactions between cardiomyocytes and the extracellular matrix surrounding them. It has been demonstrated that fibronectin, a component of the extracellular matrix that is particularly abundant during embryogenesis, acts through integrin β1 receptors present on the

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surface of cardiomyocytes to induce proliferation (Ieda et  al. 2009). Focal adhesion kinase (FAK), which plays an important role in integrin-mediated signal transduction, appears to be a critical mediator of the pro-proliferative effect of fibronectin, as inactivating FAK in embryonic cardiomyocytes results in reduced cell proliferation (Peng et al. 2008). FAK activity is antagonized by expression of an endogenous inhibitor FAKrelated non-kinase (FRNK), and while the expression of FAK is comparatively steady throughout heart development, FRNK expression in cardiomyocytes is dynamic and reaches a maximum just at the onset of polyploidization (DiMichele et al. 2009). It has been recently reported that cardiomyocytes in post-natal FRNK null mice undergo mitotic divisions characterized by ectopic DNA synthesis, nuclear division, and cytokinesis, which produced larger hearts containing increased numbers of cardiomyocytes (O’Neill et al. 2012). This suggests that FRNK and the dampening of FAK activity in post-natal cardiomyocytes comprises part of the regulatory mechanism triggering cell cycle exit and polyploidization in these cells, although the fact that FRNK null cardiomyocytes eventually stop dividing and achieve a similar level of multinucleation as controls indicates that additional mechanisms must also control polyploidization in cardiomyocytes. Similar effects have been observed with inhibition of p38 MAP kinase. p38 has been implicated in the differentiation of cardiomyocytes, and its activity has been demonstrated to be inversely correlated with cardiomyocyte proliferation (Eriksson and Leppa 2002; Engel et al. 2005). Endogenous p38 activity increases and remains continuously high in neonatal cardiomyocytes at approximately the time when these cells become polyploid. In combination with stimulation with growth factors such as FGF1, p38 inhibition triggered DNA synthesis, karyokinesis, and cytokinesis in neonatal and adult cardiomyocytes (Engel et al. 2005). Although inhibition of p38 alone had no proliferative effect, microarray analysis indicates that p38 inhibition resulted in up-regulation of genes involved in cytokinesis (Engel et al. 2006). The current model to explain these results is that growth factor stimulation is required for DNA synthesis in maturing cardiomyocytes, and this effect is potentiated by p38 inhibition. p38 inhibition then also induces the G2/M transition and cytokinesis by blocking the down-regulation of mitosis and cytokinesis genes usually caused by p38 activity (Engel et al. 2005). 1.11.2  Mechanisms of Cardiomyocyte Polyploidization

Normal post-natal cardiomyocytes can undergo DNA synthesis and mitosis, but fail to execute cytokinesis and thus become multinucleate. Therefore, mechanisms ­specifically preventing cytokinesis in post-natal cardiomyocytes are likely the most relevant for polyploidization. It has been suggested that terminally differentiated post-natal cardiomyocytes are incapable of cytokinesis because at this point in development the contractile apparatus has fully matured and cannot be completely disassembled, and the presence of myofibrils physically impedes cytokinesis (Ahuja et al. 2004; Lacroix and Maddox 2012). However, it is unlikely that this explains the lack of cytokinesis in postnatal cardiomyocytes, as more recent studies have demonstrated that neonatal and adult cardiomyocytes that are induced to divide can transiently disassemble their myofibrils, and this occurs even in cells that do not undergo cytokinesis and become binucleate (Engel et al. 2005; Engel et al. 2006; Bersell et al. 2009). An alternative hypothesis proposed by Engel and colleagues is based on their observation that the cytoskeletal

1.12 Hepatocytes

protein Anillin is mislocalized in cardiomyocytes stimulated to become binucleate in vitro (Engel et al. 2006). Anillin is nuclear during G1, S, and G2, is redistributed to the cell cortex at the beginning of mitosis, and then specifically localizes to the cleavage furrow at the onset of cytokinesis (Field and Alberts 1995; Oegema et al. 2000). It is required for cytokinesis to occur, as the cleavage furrow in cells lacking Anillin function cannot complete ingression (Oegema et al. 2000; Somma et al. 2002; Echard et al. 2004). Compared to cardiomyocytes that were induced to divide, in those that were induced to become binucleate, Anillin localization was more diffuse and failed to localize to the midbody (Engel et al. 2006). Furrow ingression was defective in these cells, suggesting that Anillin is required for proper actin–myosin ring formation and contraction and that it may be an important element of the mechanism preventing cytokinesis in terminally differentiated cardiomyocytes. 1.11.3  Polyploidization as a Response to Tissue Damage

In addition to the polyploidization and multinucleation that occur during early postnatal development, these processes also occur in response to myocardial injury. For example, an increase in cardiomyocytes with a DNA content of more than 8 C has been observed in human hearts after myocardial infarction (Herget et al. 1997). Furthermore, the ploidy of cardiomyocytes immediately adjacent to myocardial injury has been found to be significantly higher than at more distant sites (Meckert et al. 2005; Senyo et al. 2013). Detailed examination of the morphology of proliferating cardiomyocytes next to the injury revealed that endomitosis may be occurring in this situation (Meckert et al. 2005). Although it has also been demonstrated that terminally differentiated cardiomyocytes can undergo limited cell divisions after tissue damage, it is clear that polyploidization and hypertrophy also comprise a significant part of the adaptive response to myocardial injury (Senyo et al. 2013).

1.12 ­Hepatocytes 1.12.1  Mechanisms of Hepatocyte Polyploidization

The liver is comprised of a heterogeneous mix of hepatocytes that contain either one or two nuclei with DNA contents ranging from 2 to 16 C, although 16 C hepatocytes are only rarely found (Guidotti et al. 2003; Wirth et al. 2006; Margall-Ducos et al. 2007). The proportion of polyploid cells appears to vary among species, as more than 70% of the hepatocytes in rodents are polyploid, whereas human livers only contain 30–50% polyploid cells (Toyoda et  al. 2005; Duncan et  al. 2012b; Gentric et  al. 2012). Liver polyploidization is a developmentally controlled process that is initiated during post-natal development at the stage of weaning by glucose-independent Insulin/Akt signaling (Celton-Morizur et al. 2009). The ploidy of hepatocytes increases gradually by progression through a complex cell cycle program (Guidotti et al. 2003). After birth, rat hepatocytes are exclusively diploid, but during the third week of post-natal development a subpopulation of hepatocytes undergoes a cell cycle with incomplete cytokinesis, leading to the formation of binucleated, tetraploid cells (2×2 C) (Guidotti et al. 2003). These hepatocytes execute mitosis normally until anaphase, but fail to assemble a functional actin–myosin ring during telophase. Hepatocytes undergoing incomplete cytokinesis

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are incapable of accumulating actin along the cleavage site and cannot properly recruit myosin II to the cortex, due to diminished Rho Kinase activity (Margall-Ducos et al. 2007). Cytokinesis, including F-actin and RhoA localization along the cleavage site, is reconstituted by inhibition of Akt, but not by Rapamycin-mediated inhibition of mTORC1, suggesting that Akt regulates the cytokinesis program directly or indirectly through mTORC2 (Celton-Morizur et al. 2009). Remarkably, the binucleated hepatocytes are capable of entering the cell cycle (Guidotti et al. 2003). The binucleated hepatocytes contain two parental centrosomes, which are duplicated in S phase. During mitosis, two centrosomes cluster at opposite poles of the cell, thereby allowing the formation of a bipolar spindle. The subsequent division results in mononucleate cells with a 4 C DNA content (Guidotti et al. 2003; Margall-Ducos et al. 2007). A subpopulation of the mononucleated tetraploid hepatocytes undergoes another round of acytokinetic mitosis, thereby giving rise to binucleate octoploid cells (2×4 C), which may execute bipolar mitosis to produce mononucleate, octoploid cells (1×8 C) (Duncan 2013). 1.12.2  The Ploidy Conveyor Model

Hepatocyte ploidy subsequently increases during development, but recent studies on murine and human livers have demonstrated that subpopulations of tetra- or octoploid hepatocytes undergo reductive division, thereby producing diploid or tetraploid cells (Duncan et al. 2010; Duncan et al. 2012b). This observation is consistent with earlier studies reporting that ploidy transiently increases after partial hepatectomy, but returned to basal ploidy levels within 1 month (Sigal et al. 1999). The interplay of polyploidization and ploidy reversal led to the postulation of the ploidy conveyor model, which explains how constant ratios of diploid and polyploid hepatocytes are maintained (Duncan et al. 2010). The ploidy reversal in hepatocytes relies on multipolar mitosis, a process that is prone to chromosome mis-segregation (Ganem et  al. 2009), and thus gives rise to a heterogeneous population of aneuploid cells. This observation generated the notion that this mechanism generates a pool of genetically divergent daughter cells, and thus extends the genetic variety of hepatocytes, which might contribute to the remarkable robustness of liver cells. In support of this idea, Duncan et  al. recently reported that a specific chromosome carrying a mutation that suppresses tyrosinemia was enriched in liver repopulation assays (Duncan et al. 2012a). This remarkable finding suggests that polyploidy-induced aneuploidy plays an important role in the development and maintenance of somatic tissues. 1.12.3  Liver Regeneration

Hepatocytes exhibit a remarkable capacity for regeneration after injury (Duncan 2013; Pandit et al. 2013). Adult hepatocytes are metabolically active, but are quiescent under normal conditions. In response to liver damage, the dormant hepatocytes enter the cell cycle and regenerate the liver (Guidotti et al. 2003; Wirth et al. 2006; Margall-Ducos et al. 2007). In principle, the tissue loss upon damage can be restored by two possible mechanisms: The first option is that the remaining cells could re-enter the cell cycle to replace the dead cells. Alternatively, the tissue lost could be compensated by cellular hypertrophy, an increase in the volume of an organ or tissue due to the enlargement of its cells. In rats, liver regeneration is accompanied by higher percentages of polyploid cells. Livers regenerating from radiation-induced oxidative damage or after partial

1.12 Hepatocytes

hepatectomy exhibited increased numbers of tetra- and octaploid hepatocytes with a single nucleus, while the fraction of binucleated cells decreases to less than 5% (Sigal et al. 1999; Gorla et al. 2001). Early studies revealed that most of the hepatocytes (84%) in the rat liver enter S phase during regeneration (Grisham 1962; Bucher and Swaffield 1964). Genetic tracing experiments in mice recently revealed that only half of the regenerating hepatocytes undergo cell division (Miyaoka et  al. 2012), implying that a subpopulation of hepatocytes compensates for cell loss with hypertrophy, which may involve endoreplication cycles. Another study, however, which used in-situ hybridization with a probe specific to the y chromosome, reported that upon partial hepatectomy the majority of wild-type cells in the liver are diploid (70%) and only minor subpopulations are tetraploid, octoploid, or higher (Nevzorova et al. 2009). In any case, we can conclude that cellular hypertrophy and polyploidization are part of the normal response to liver damage. Liver-specific deletion of Cdk1 or Separase, which are crucial for the initiation or progression through mitosis, respectively, did not affect the regenerative capacity of damaged livers (Wirth et al. 2006; Diril et al. 2012). Both studies reported an increase in ploidy after partial hepatectomy, suggesting that these cells compensated for their inability to execute mitosis by undergoing endoreplication cycles. Remarkably, single-cell DNA measurements demonstrated that hepatocytes derived from Separase-deficient livers attained DNA contents exceeding 32 C, whereas wild-type hepatocytes only reached 16 C, indicating that the rate of endocycling is accelerated under conditions prohibiting cell division (Wirth et al. 2006). Thus cellular hypertrophy through endoreplication may represent an effective strategy for liver regeneration under conditions that do not permit mitotic divisions, for example upon DNA damage. This hypothesis is underscored by the finding that hepatocytes deficient for the essential telomere protein TRF2 exhibited normal liver regeneration after partial hepatectomy (Lazzerini Denchi et  al. 2006). Mutation of TRF2 in proliferating cells results in telomere dysfunction, which leads to p53 activation and p21-mediated G1 arrest, followed by apoptosis or senescence (van Steensel et  al. 1998; Smogorzewska and de Lange 2002; Celli et  al. 2006). The TRF2-deficient hepatocytes showed severe chromosomal abnormalities and a strong DNA damage response (gamma-H2AX staining), and failed to complete mitosis. During recovery from partial hepatectomy, the TRF2-deficient livers exhibited an essentially normal pattern of S phase, but an increased number of polyploid cells, indicating that liver regeneration was mediated by endoreplication. Conversely, it has been observed that liver regeneration was unaffected by hepatocyte-specific knockout of the atypical E2Fs, E2F7 and E2F8, which are crucial for hepatocyte polyploidization (Chen et  al. 2012; Pandit et  al. 2012). Recovery from partial hepatectomy or treatment with the xenobiotic drug DDC was accompanied by an increased number of diploid cells, indicating that the inability to undergo endocycles was compensated by increased mitotic proliferation, thus allowing normal liver regeneration. Altogether, we conclude that liver regeneration relies on both mitotic proliferation of hepatocytes and cellular hypertrophy due to increased hepatocyte polyploidization and that the balance between the two pathways strongly depends on the type of damage. Increased mitotic proliferation may represent the stereotypical response to liver injury, but after DNA damage hepatocytes may preferentially undergo endoreplication to prevent aberrant cell divisions. Currently, it is unclear how the decision between these two options is made, but it may involve the ATR/ATM–Chk1/Chk2

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pathways. Notably, it has recently been demonstrated that p53/Rb-deficient cells with irreparably damaged DNA become polyploid (Davoli et al. 2010). The damaged cells initially arrest at the G2/M transition due to ATR/ATM–Chk1/Chk2-mediated downregulation of the mitosis-promoting phosphatase Cdc25A, but later escape this arrest and initiate endoreplication cycles.

1.13 ­Megakaryocytes Megakaryocytes (MKCs) are specialized blood cells that give rise to platelets (thrombocytes), which are crucial for blood clotting and angiogenesis. The cellular process resulting in the formation of platelets is referred to as megakaryocytopoiesis. MKCs are characterized by multilobulated nuclei and reach ploidies of up to 128 C, although 16 C appears to be the most prevalent state (Ravid et al. 2002). A single 32 C MKC yields approximately 3000 platelets (Winkelmann et al. 1987). Although platelets are enucleated, it has been proposed that the large size of the polyploid MKCs indirectly facilitates platelet formation by providing an enormous resource for membranes and other cellular components (Lacroix and Maddox 2012). MKCs differentiate from hematopoietic stem cells in response to activation of the receptor c-MPL upon binding of the cytokine thrombopoietin (Tpo) (Kaushansky et al. 1994; Wendling et al. 1994). 1.13.1  Mechanisms of MKC Polyploidization

Polyploidization of murine and human MKCs relies on endomitosis (Figure 1.4), a noncanonical cell cycle without karyokinesis and cytokinesis (Nagata et al. 1997; Vitrat et al. 1998). During each round of endomitosis the entire genome is replicated (Sher et al. 2013), and thus MKCs always have DNA contents that are exact multiples of the haploid genome (4 C, 8 C, 16 C, and 32 C, etc.). Chromosome condensation and centrosome duplication occur during prophase, as in mitotic cells. Prometaphase in MKCs is characterized by breakdown of the nuclear envelope and assembly of the mitotic spindle; in cells that are already polyploid, the presence of multiple centrosomes results in the formation of multipolar spindles. During metaphase, the chromosomes align at right angles to the spindle microtubules. In anaphase, sister chromatids separate and are pulled towards the spindle poles, but in MKCs the replicated copies of the chromosome are incorporated into the same nucleus during reformation of the nuclear envelope. The multipolar spindle allows chromosomes to attach in multiple configurations, resulting in asymmetrical distribution of chromosomes among nuclear lobes (Roy et al. 2001). Live microscopy has revealed that MKCs undergo two distinct phases of endomitosis (Geddis and Kaushansky 2006; Geddis et al. 2007; Lordier et al. 2008; Papadantonakis et al. 2008; Gao et al. 2012). In the majority of MKCs, the first endomitosis resulting in the formation of tetraploid (4 C) cells involves cleavage furrow ingression and subsequent regression, while in high-ploidy MKCs (more than 4 C), endomitosis usually occurs without any cleavage furrow formation. MKCs show correct localization of Aurora B, MKLP2, and MagRacGAP, implying that midzone formation occurs normally during endomitosis (Geddis and Kaushansky 2004; Geddis and Kaushansky 2006). In high-ploidy MKCs, the GTPase RhoA is absent from the midzone, thus explaining the lack of a contractile actin–myosin ring. During

1.13 Megakaryocytes

Figure 1.4  Abortive division cycles. (A) Schematic of normal mitotic cell division. During prophase, chromatin condenses into distinctive chromosomes and the mitotic spindle begins to assemble between duplicated centrosomes. Breakdown of the nuclear envelope marks the onset of prometaphase, thereby allowing spindle microtubules to connect with chromosomes. During metaphase, chromosomes align along the equator of the spindle and both of their kinetochores are attached to microtubules. At anaphase A, sister chromatids separate and are then pulled towards spindle poles by shortening of kinetochore microtubules. Anaphase B is characterized by lengthening of polar microtubules, thereby pushing spindle poles apart. In telophase (not shown) chromosomes decondense and the nuclear envelope starts to assemble. Finally, a contractile actin–myosin ring forms during cytokinesis and splits cytoplasm into two daughter cells. (B) During acytokinetic mitosis the mitotic program is executed normally, but cells fail to establish a contractile actin–myosin ring and cannot cleave, which leads to formation of binucleate cells with two centrosomes. (C) Megakaryocytes undergoing endomitosis normally progress through prophase, prometaphase, metaphase, and anaphase A, but fail to execute anaphase B, telophase, or cytokinesis. As a consequence, the reassembling nuclear envelope encloses the sister chromatids in a single nucleus, thereby giving rise to a characteristic lobular structure. Higher levels of ploidy are accomplished by consecutive rounds of endomitosis, which involve centrosome duplication and formation of multipolar spindles. (See plate section for color representation of this figure.)

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the first endomitosis, by contrast, RhoA is readily detectable at the cleavage furrow, but the majority of low-ploidy MKCs fail to assemble a contractile actin–myosin ring (Lordier et al. 2008). This obstacle was explained by a recent study that utilized a FRETbased biosensor and thereby demonstrated that RhoA is correctly localized, but inactive in low-ploidy MKCs (Gao et  al. 2012). Moreover, the activation of GTPases like RhoA requires switching from the GDP-bound to the GTP-bound form and is mediated by guanine nucleotide exchange factors (GEFs). During cytokinesis in mitotic cells, the activity and localization of RhoA relies on the GEFs ECT2 and GEF-H1 (Birkenfeld et al. 2007). Depletion of ECT2 abolishes cleavages furrow formation and cytokinesis, while knockdown of GEF-H1 only affects later stages of cytokinesis. Tellingly, the expression of both GEFs is down-regulated upon induction of polyploidization with TPO, and involves the transcriptional co-factor MKL1 (Gao 2012). The expression of GEF-H1 is absent during the first endomitosis but recovers during subsequent cycles, whereas ECT2 persists until higher-ploidy states. Hence, the presence of ECT2 in lowploidy MKCs supports recruitment of RhoA to equatorial region MKCs, but due to the lack of GEF-H1 most of the RhoA is inactive and cytokinesis cannot be completed. In high-ploidy MKCs the reduction of ECT2 prevents RhoA recruitment, and thus explains the absence of a distinct cleavage furrow. Consistent with this model, ectopic expression of GEF-H1 increases the number of diploid MKCs, but a subpopulation of MKCs still undergoes polyploidization. Conversely, overexpression of ECT2 prevents tetraploid MKCs from attaining high-ploidy states, but does not influence the proliferation diploid cells. Concomitant overexpression of both GEFs completely abolishes MKC polyploidization. The exact purpose of the differential regulation of the initial endomitosis is currently unclear, but it was observed that a fraction of diploid MKCs (13%) completes cytokinesis, suggesting that the first endomitotic cycle acts as a restriction point that balances differentiation and precursor proliferation (Shivdasani 2012). As MKC polyploidization results from a modified M phase, it is intuitive that the rest of the endomitotic cell cycle essentially relies on the same mechanisms as mitotic cells. MKCs display normal expression levels of Cdk1 and CycB, which are the major regulators for the execution of mitosis (Nagata et al. 1997). Consistent with the observation that CycB degradation occurs with normal timing during metaphase, it was found that MKCs express both APC/C activators Cdc20 and Cdh1 (Roy et  al. 2001). Moreover, polyploid MKCs treated with the microtubule-destabilizing agent nocodazole undergo a non-permanent arrest, indicating that the spindle assembly checkpoint is functional during endomitosis (Roy et al. 2001). Interestingly, it was found that concomitant deletion of CycE1/E2 blocks MKC polyploidization (Geng 2003), thus implying that S phase entry in MKCs exclusively relies on CycE-dependent kinase activity and CycA cannot fulfill this function.

1.14 ­Concluding Remarks In this chapter, we have summarized how somatic polyploidy arises in multiple tissues in diverse organisms and which functions it fulfills during normal development and under adverse circumstances. These functions include determining developmental fate, providing cells with the ability to produce large quantities of proteins or membrane to promote their function, and facilitating cellular growth under conditions

  References

where cell divisions would compromise tissue integrity. Furthermore, we have highlighted the role polyploidization plays in providing tissues with flexibility during recovery from injury and its proposed function as a mechanism to acquire genetic diversity. However, additional studies are needed to completely understand the molecular mechanisms leading to somatic polyploidy and how polyploidization enforces the physiology of organisms.

­Acknowledgments We are grateful to our postdoc advisor Bruce A. Edgar who encouraged us to pursue this project independently. The research in the Edgar laboratory was supported by ERC Advanced Grant 268515 and the DKFZ.

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2 Large-Scale Programmed Genome Rearrangements in Vertebrates Jeramiah J. Smith* Department of Biology, University of Kentucky, Kentucky, USA

Abstract In some species, large-scale rearrangements occur in a tightly regulated manner, as a normal part of development. This phenomenon, broadly termed programmed genome rearrangement (PGR), is known have evolved independently in several eukaryotic lineages, including ciliated protozoans, some species of roundworms, copepod crustaceans, sciarid flies, and midges. The repeated evolution of PGR suggests recurrent selection for regulated dynamism of genome primary structure. Keywords  vertebrates; somatic cells; DNA elimination; programmed genomic changes

2.1 ­Introduction Interestingly, a few vertebrate species have been observed to undergo reproducible large‐scale rearrangements during the normal course of development that eliminate segments of chromosomes, or entire chromosomes. The first examples of such recurrent and large‐scale change were identified in several species of bandicoots and a few other marsupials (Hayman et al. 1969; McKay et al. 1984; Watson et al. 1998). In bandicoots (Isoodon) these changes result in the elimination of sex chromosomes from some rapidly dividing cell lineages (epithelial and hematopoetic cells) and likely occurs during post‐natal development in the pouch. The elimination of sex chromosomes seems to be correlated with the generally delayed timing of DNA synthesis for the sex chromosomes in rapidly dividing cell lineages and varies within and among cell lineages. Other vertebrates have been observed to undergo arguably more programmatic rearrangement events, which result in broadscale genomic differentiation of germline versus somatic cells (Figure 2.1). The first vertebrate identified to undergo such changes was the hagfish (Eptatretus burgeri) (Kohno et al. 1986), many species of which were subsequently found to eliminate entire chromosomes, or chromosomal segments, *Corresponding author: [email protected] Smith J.J. (2017) Large-scale programmed genome rearrangements in vertebrates. In: Li X.-Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley-Blackwell, Hoboken, NJ, Ch. 2, pp. 45–54. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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Figure 2.1  Distribution of germline‐specific DNA and patterns of DNA elimination across the vertebrate phylogeny. Germline‐specific sequences have been observed in lamprey, hagfish, and zebra finch, and studies have begun to reveal the causes and consequences of programmed genome rearrangement in these groups. It is not known if hagfish and lamprey retain programmed genome rearrangement as an aspect of their biology inherited from their common ancestor or if similar genome biologies evolved more than once in the two lineages. The germline‐specific chromosome of zebra finch appears to have arisen relatively recently and likely represents an independent evolutionary acquisition of germline‐specific DNA. Most vertebrate lineages have not been rigorously tested for the presence or absence of germline‐specific DNA. MYA, Million years ago.

from most somatic cell lineages. Subsequently, similar phenomena were observed in zebra finch (Taeniopygia guttata) (Pigozzi and Solari 1998) and the sea lamprey (Petromyzon marinus) (Smith et al. 2009). The discovery of these species and dissection of the molecular details of DNA loss and genetic differentiation of the germline and soma provides a novel perspective on the causes and consequences of change in the somatic genome. This chapter reviews our current state of knowledge regarding the molecular details of these novel vertebrate systems and attempts to identify emerging themes and directions of inquiry that build on these studies.

2.1 ­Hagfish Hagfish are living representatives of an ancient chordate lineage that diverged from extant vertebrate lineages in the late Precambrian or early Cambrian, prior to the

2.1 Hagfish

evolution of jaws and paired appendages. Patterns of DNA elimination have been described for several species of hagfish and have been detected in all species examined, though details regarding the developmental progression of PGR are lacking, largely due to the rarity of hagfish embryos. Prior to the recent identification of pharyngula‐stage hagfish embryos in a laboratory setting, it had been more than a century since any hagfish embryos had been collected (Janvier 2007; Ota et al. 2007; Ota et al. 2011). Evidence for chromosome elimination in hagfish has therefore necessarily relied on examination of DNA and chromosomes prepared from adult cells. Differences between germline and somatic cells have been reported for the genera Myxine (two species), Eptatretus (four species), and Paramyxine (two species), with the proportion of germline DNA that is eliminated from somatic cells varying extensively across species, ranging from approximately 0.21 on E. burgeri to more than 0.70 in P. sheni (Nakai et al. 1991; Nakai et al. 1995; Goto et al. 1998; Nabeyama et al. 2000; Kojima et al. 2010). 2.2.1  Content of Eliminated DNA

Karyological studies indicate that much of the eliminated DNA is packaged as heterochromatin in the adult germ cells within which this DNA is retained, although some studies also support the idea that some of the eliminated DNA is euchromatic in primary spermatocytes (Kohno et al. 1986; Nakai et al. 1991; Kubota et al. 1992; Kubota et al. 1993; Kubota et al. 1994; Nakai et al. 1995; Kubota et al. 1997; Goto et al. 1998; Nabeyama et  al. 2000; Kubota et  al. 2001; Kojima et  al. 2010). Restriction digestion/ Southern blotting experiments have identified several sequences that are present in the germline, but absent from the soma (or highly reduced) (Kubota et al. 1993; Goto et al. 1998; Nabeyama et al. 2000; Kubota et al. 2001; Kojima et al. 2010). All such sequences identified to date correspond to high‐abundance tandem repeats. Interestingly, one of these sequences (known as EEEo2) has been identified in all species of Eptatretus and Paramyxine examined to date (Nabeyama et al. 2000; Kojima et al. 2010). The presence of this sequence in the eliminated DNA of several species provides an indication that at least some fractions of eliminated sequence in these hagfish species are homologous and consistently deleted across taxa. 2.2.2  Results and Mechanisms of Deletion

The initial discovery of DNA elimination in hagfish was based on the observation of reproducible differences between somatic and germline karyotypes. In all species of hagfish examined thus far, the number of chromosomes in somatic tissues is smaller than the number of chromosomes in germline cells. Although intrachromosomal fusions or rearrangements could also result in reduction in chromosome number, observations strongly suggest the chromosome loss is a major driver of DNA loss in all hagfish species. For three other species (P. sheni, E. stoutii, and E. okinoseanus—subtype B) observations support the elimination of portions of chromosomal arms by DNA fragmentation, followed by re‐establishment of the telomeres (Kubota et  al. 1992; Kubota et al. 1994; Kojima et al. 2010). Thus far, examinations indicate the existence of at least two linked mechanisms of DNA elimination in hagfish and appear to neither support nor refute the possibility of smaller deletion events.

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2.3 ­Sea Lamprey The lampreys are members of a second ancient lineage that also diverged from other extant vertebrate lineages in the late Precambrian/early Cambrian. Programmed elimination of DNA has been found to occur in one species (the sea lamprey: Petromyzon marinus) and thus far has not been assayed for other lamprey species (Smith et al. 2009). 2.3.1  Content of Eliminated DNA

As in hagfish, the eliminated fraction of the genome contains a substantial complement of repetitive DNA. In fact, the discovery and characterization of programmed genome rearrangement in lamprey employed several common repetitive elements, in addition to a small handful of low copy sequences (Smith et al. 2009). Subsequent sequencing studies further revealed that likely thousands of germline‐specific genes are reproducibly eliminated from somatic tissues (Smith et  al. 2012). Eliminated genes appear to function in the development and maintenance of germline DNA, but also hold the potential to contribute to cancer or other deleterious outcomes if misexpressed in somatic cells. 2.3.2  Results and Mechanisms of Deletion

Programmed genome rearrangement results in the elimination of approximately 20% of germline DNA from somatic cells, and is initiated at or near the beginning of gastrulation (at approximately 2.5 days post fertilization) (Smith et  al. 2012). Comparison of somatic and spermatogonial karyotypes reveals differences in karyotype number, with germline possessing approximately 99 chromosomes and somatic cells possessing 84 (Smith et  al. 2010b), though it is again important to note that multiple mechanisms could account for observed differences in chromosome number. Bacterial artificial chromosome (BAC) sequencing studies reveal that large fragments of DNA (in excess of 100 kb) are eliminated in their entirety, indicating that individual deletion events frequently involve large portions of chromosomes or entire chromosomes (Smith et al. 2010b). Intriguingly, survey sequencing of sperm DNA identified several sequences that were consistent with deletion of somatic sequence via recombination, although three experimentally validated recombination sites showed little evidence of a specific consensus sequence that could target site‐specific recombination (Smith et al. 2012).

2.4 ­Zebra Finch Among the jawed vertebrates, the zebra finch (Taeniopygia guttata) and its close relative the Bengalese finch (Lonchura domestica) are the only known species to undergo large‐scale programmed elimination of DNA that results in the reproducible differentiation of germline and somatic cell lineages (Pigozzi and Solari 1998; del Priore and Pigozzi 2014). The initial discovery of these elimination events resulted from the identification of a single chromosome (the ‘germline‐restricted chromosome,’ GRC) that is present in the germ cells of both sexes, but absent from all somatic cells. In the germline, the GRC is the largest chromosome, being visibly larger than finch chromosome 1.

2.5  Emerging Themes and Directions

2.4.1  Mechanisms of Deletion

Given that programmed rearrangement of the zebra finch genome apparently involves the elimination of a single chromosome from apparently all somatic cell types, it seems reasonable to speculate that the chromosome is lost through modifications of the segregational machinery during the establishment of somatic cell lineages in the early embryo (Pigozzi and Solari 1998). Elimination of the zebra finch GRC has not yet been described in the finch embryo, but studies have examined its segregation in the adult germline (Pigozzi and Solari 1998; Pigozzi and Solari 2005; Itoh et al. 2009; Goday and Pigozzi 2010; Schoenmakers et  al. 2010). The GRC is maintained in oocytes, largely packaged as euchromatin, and undergoes recombination during female meiosis, but is eliminated from spermatocytes during meiosis. The elimination of the GRC involves repressive modifications to histones (e.g., methylated H3 lysine 9) and packaging into heterochromatin followed by fragmentation, packaging into a micronuclear exclusion body, and expulsion (Pigozzi and Solari 1998; Pigozzi and Solari 2005; Goday and Pigozzi 2010; Schoenmakers et al. 2010). 2.4.2  Content of Eliminated DNA

A single testes‐restricted sequence was isolated by screening a panel of arbitrary primers for testes‐specific amplification (Itoh et al. 2009). This sequence (TRAP5) could be amplified from testes and ovary DNA, but not from blood, liver, or brain of either sex. The TRAP5 element corresponds to a tandemly repetitive element that is broadly distributed across the GRC, and is apparently not homologous to any known protein coding sequence. Intriguingly, a homolog of the GRC is present on an autosome in zebra finch, specifically chromosome 3. This observation seems to suggest that the GRC may have originated (at least in part) from this autosome and that the GRC might contain homologs of some genes present of zebra finch chromosome 3, though genic sequences have yet to be isolated from the GRC.

2.5 ­Emerging Themes and Directions 2.5.1  The Biological Function of PGR

It has been observed that the germline‐specific fraction of the lamprey genome contains hundreds and likely thousands of genes. Although studies on eliminated DNA from finch and hagfish have yet to confirm the existence of germline‐specific genic sequences (eliminated from soma), it seems reasonable to argue that several genic sequences are deleted, given the sheer amount of somatically eliminated DNA that is maintained in the germline of these species. The patterns of observed deletions in lamprey seem to suggest that programmed rearrangements act to satisfy the differential genetic requirements of germline and soma in a way that contributes positively to organismal fitness. At some level, it can be argued that germline and somatic cells possess conflicting genetic requirements, though in most species these two fundamental cell types share exactly the same complement of genes. The germline must undergo meiotic recombination between homologous chromosomes, and retain, or develop, the ability to give rise to all cell types that develop

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during an organism’s life (totipotency). These unique functional requirements of the germline set it at odds with the requirements of other cell lineages and raise the possibility that some of the genes that contribute to germline functions could have deleterious effects on somatic tissues (somatic antagonistic pleiotropy). Here the term antagonistic pleiotropy (Williams 1957) is used in its broadest sense. This includes not only cases wherein a gene contributes to several phenotypes, with mutations having differential effects on resulting traits, but also cases wherein mutations in a gene cause ectopic expression, with untoward effects. Such mutations are not necessarily transmitted to subsequent generations, but may have profound consequences with respect to an organism’s survival and reproduction. An overt example of this type of antagonism is oncogenesis, wherein misexpression of genes may impart functions that contribute to the development of cancer (e.g., limitless cellular replication, genomic reprogramming, self‐sufficiency in growth signals, tissue invasion) (Hanahan and Weinberg 2000). Notably, genes that are targeted for somatic elimination by PGR include a lamprey homolog of the human gene that encodes cancer/testis antigen 68 (CT68). The cancer testes antigens are an evolutionary diverse class of genes that share a common feature in that they are normally expressed germline and not expressed in somatic cells, except in the context of cancer (Simpson et al. 2005; Hofmann et al. 2008). However, the expression of such pleiotropic effects is not necessarily limited to cancer and could presumably involve any disease that is related to pluripotency or somatic recombination (e.g., Fanconi anemia, aging). Thus, the simple presence of such a gene in the genome may be considered a potential risk factor for disease, and somatic pleiotropic mutations need not be transmitted to subsequent generations in order to affect an individual’s fitness. As future sequencing studies begin to characterize the gene content of eliminated DNA from hagfish and zebra finch, it should be particularly interesting to understand whether eliminated genes generally fall into functional categories similar to those observed for lamprey or if data from these divergent lineages reveal relationships between PGR and other conserved aspects of genome or developmental biology. 2.5.2  Mechanisms of Deletion

Data from hagfish, lamprey, and zebra finch suggest several common themes with respect to the likely mechanism of DNA elimination, though it should be recognized that the deep divergence times between these three groups and the lack of evidence for PGR in chicken and other jawed vertebrates presents substantial potential for independent evolution. Shared features include genomic differentiation at the fundamental level of germline versus soma in all three groups and preferential elimination of heterochromatin in hagfish and finch. The observation that programmed rearrangements result in genomic differentiation at the fundamental level of germline versus soma indicated that these rearrangements take place early in development. Germ cell induction is not described in hagfish, but in birds it is known that the germline is established during early gastrulation. The fact that programmed rearrangement takes place in lamprey in early gastrulation suggests that PGR (and perhaps germ cell fate) is induced during gastrulation. Given that embryos possess thousands of cells by this stage in development, the elimination of DNA must

  References

be coordinated across a relatively large number of cells in order to yield the broad‐scale patterns observed in adults. In both hagfish and zebra finch, it is observed that eliminated regions are packaged into transcriptionally repressive chromatin in the developing spermatocytes. However, it should be noted that these may not reflect the chromatin state of germline or somatic cells earlier in development. This packaging may reflect the fact that many eliminated genes are not expressed during male meiosis, rather being expressed at other stages of germ cell development. In this regard it is worth noting that the finch GRC is euchromatic in meiotic oocytes and that elimination of euchromatic regions occurs in some hagfish species (Pigozzi and Solari 1998; Pigozzi and Solari 2005; Kojima et al. 2010), indicating that some fraction of eliminated genes may be expressed in meiotic cells. Indeed, several genes eliminated from the lamprey germline are expressed in meiotic germline, whereas others are expressed earlier in development (Smith et al. 2009; Smith et al. 2012). Defining the functional relationship between transcriptionally repressive chromatin and DNA elimination (absolute repression) will necessarily require studies of DNA packaging and loss during early embryonic development. If it is ultimately found that the mechanisms of programmed genome rearrangement in hagfish and lamprey are derived from a common ancestral mechanism, this would have profound implications on our own understanding of human genome biology. A shared mechanism would imply that the ancestor of all vertebrates (or a very close relative) possessed programmed rearrangement as a fundamental aspect of its developmental biology (Smith et al. 2010a), although the existence of some shared components might also be explained by convergent evolution. 2.5.3  Other Vertebrates?

Known examples of programmed rearrangement are the result of fortuitous discoveries that were facilitated by careful observations and the overt nature of resulting changes. The discovery of PGR in diverse taxa over the last century begs the question as to how widespread the phenomenon truly is. For example, are patterns of somagenetic variation observed in some plants (including flax, ciliates, tobacco, potato, and maize) also due to programmatic changes (see Chapters 4, 8, and 13, this study)?. Even for the vast majority of species with ‘sequenced’ genomes, we simply haven’t looked. Modern sequencing approaches are opening the door to ever more powerful means of accessing genomic information for a diversity of species and assays for detecting differences between genomes. It is anticipated that future studies will leverage these technologies to identify additional examples of programmed genome rearrangement in the vertebrate lineage and shed light on the causes and consequences of PGR in all of these systems.

­References del Priore L. and Pigozzi M.I. (2014) Histone modifications related to chromosome silencing and elimination during male meiosis in Bengalese finch. Chromosoma 123:293–302.

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Goday C. and Pigozzi M.I. (2010) Heterochromatin and histone modifications in the germline‐restricted chromosome of the zebra finch undergoing elimination during spermatogenesis. Chromosoma 119:325–336. Goto Y., Kubota S. and Kohno S. (1998) Highly repetitive DNA sequences that are restricted to the germ line in the hagfish Eptatretus cirrhatus: a mosaic of eliminated elements. Chromosoma 107:17–32. Hanahan D. and Weinberg R.A. (2000) The hallmarks of cancer. Cell 100:57–70. Hayman D.L., Martin P.G. and Waller P.F. (1969) Parallel mosaicism of supernumerary chromosomes and sex chromosomes in Echymipera kalabu (Marsupialia). Chromosoma 27:371–380. Hofmann O., Caballero O.L., Stevenson B.J., Chen Y.T., Cohen T., Chua R., Maher C.A., Panji S., Schaefer U., Kruger A., Lehvaslaiho M., Carninci P., Hayashizaki Y., Jongeneel C.V., Simpson A.J.G., Old L.J., and Hide W. (2008) Genome‐wide analysis of cancer/ testis gene expression. Proc Natl Acad Sci USA 105:20422–20427. Itoh Y., Kampf K., Pigozzi M.I. and Arnold A.P. (2009) Molecular cloning and characterization of the germline‐restricted chromosome sequence in the zebra finch. Chromosoma 118:527–536. Janvier P. (2007) Evolutionary biology: born‐again hagfishes. Nature 446:622–623. Kohno S., Nakai Y., Satoh S., Yoshida M. and Kobayashi H. (1986) Chromosome elimination in the Japanese hagfish, Eptatretus burgeri (Agnatha, Cyclostomata). Cytogenet Cell Genet 41:209–214. Kojima N.F., Kojima K.K., Kobayakawa S., Higashide N., Hamanaka C., Nitta A., Koeda I., Yamaguchi T., Shichiri M., Kohno S., and Kubota S. (2010) Whole chromosome elimination and chromosome terminus elimination both contribute to somatic differentiation in Taiwanese hagfish Paramyxine sheni. Chromosome Res 18:383–400. Kubota S., Nakai Y., Kuro‐o M. and Kohno S. (1992) Germ line‐restricted supernumerary (B) chromosomes in Eptatretus okinoseanus. Cytogenet Cell Genet 60:224–228. Kubota S., Kuro‐o M., Mizuno S. and Kohno S. (1993) Germ line‐restricted, highly repeated DNA sequences and their chromosomal localization in a Japanese hagfish (Eptatretus okinoseanus). Chromosoma 102:163–173. Kubota S., Nakai Y., Sato N., Kuroo M. and Kohno S. (1994) Chromosome elimination in northeast Pacific hagfish, Eptatretus stoutii (Cyclostomata, Agnatha). J Hered 85:413–415. Kubota S., Ishibashi T. and Kohno S. (1997) A germline restricted, highly repetitive DNA sequence in Paramyxine atami: an interspecifically conserved, but somatically eliminated, element. Mol Gen Genet 256:252–256. Kubota S., Takano J., Tsuneishi R., Kobayakawa S., Fujikawa N., Nabeyama M. and Kohno S. (2001) Highly repetitive DNA families restricted to germ cells in a Japanese hagfish (Eptatretus burgeri): a hierarchical and mosaic structure in eliminated chromosomes. Genetica 111:319–328. McKay G.M., McQuade L.R., Murray J.D. and von Sturmer S.R. (1984) Sex‐chromosome mosaicism in the lemur‐like possum Hemibelideus lemuroides (Marsupialia: Petauridae). Austr J Biol Sci 37:131–135. Nabeyama M., Kubota S. and Kohno S. (2000) Concerted evolution of a highly repetitive DNA family in eptatretidae (Cyclostomata agnatha) implies specifically differential homogenization and amplification events in their germ cells. J Mol Evol 50:154–169.

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Nakai Y., Kubota S. and Kohno S. (1991) Chromatin diminution and chromosome elimination in four Japanese hagfish species. Cytogenet Cell Genet 56:196–198. Nakai Y., Kubota S., Goto Y., Ishibashi T., Davison W. and Kohno S. (1995) Chromosome elimination in three Baltic, south Pacific and north‐east Pacific hagfish species. Chromosome Res 3:321–330. Ota K.G., Kuraku S. and Kuratani S. (2007) Hagfish embryology with reference to the evolution of the neural crest. Nature 446:672–675. Ota K.G., Fujimoto S., Oisi Y. and Kuratani S. (2011) Identification of vertebra‐like elements and their possible differentiation from sclerotomes in the hagfish. Nature Comms 2:373. Pigozzi M.I. and Solari A.J. (1998) Germ cell restriction and regular transmission of an accessory chromosome that mimics a sex body in the zebra finch, Taeniopygia guttata. Chromosome Res 6:105–113. Pigozzi M.I. and Solari A.J. (2005) The germ‐line‐restricted chromosome in the zebra finch: recombination in females and elimination in males. Chromosoma 114:403–409. Schoenmakers S., Wassenaar E., Laven J.S., Grootegoed J.A. and Baarends W.M. (2010) Meiotic silencing and fragmentation of the male germline restricted chromosome in zebra finch. Chromosoma 119:311–324. Simpson A.J., Caballero O.L., Jungbluth A., Chen Y.T. and Old L.J. (2005) Cancer/testis antigens, gametogenesis and cancer. Nat Rev Cancer 5:615–625. Smith J.J., Antonacci F., Eichler E.E. and Amemiya C.T. (2009) Programmed loss of millions of base pairs from a vertebrate genome. Proc Natl Acad Sci USA 106:11212–11217. Smith J.J., Saha N.R. and Amemiya C.T. (2010a) Genome biology of the cyclostomes and insights into the evolutionary biology of vertebrate genomes. Integr Comp Biol 50:130–137. Smith J.J., Stuart A.B., Sauka‐Spengler T., Clifton S.W. and Amemiya C.T. (2010b) Development and analysis of a germline BAC resource for the sea lamprey, a vertebrate that undergoes substantial chromatin diminution. Chromosoma 119:381–389. Smith J.J., Baker C., Eichler E.E. and Amemiya C.T. (2012) Genetic consequences of programmed genome rearrangement. Curr Biol 22:1524–1529. Watson C.M., Margan S.H. and Johnston P.G. (1998) Sex‐chromosome elimination in the bandicoot Isoodon macrourus using Y‐linked markers. Cytogene Cell Genetics 81:54–59. Williams G.C. (1957) Pleiotropy, natural‐selection, and the evolution of senescence. Evolution 11:398–411.

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3 Chromosome Instability in Stem Cells Paola Rebuzzini1, Maurizio Zuccotti2, Carlo Alberto Redi1 and Silvia Garagna1* 1 2

Laboratorio di Biologia dello Sviluppo, Dipartimento di Biologia e Biotecnologie, Università degli Studi di Pavia, Pavia, Italy Dipartimento di Scienze Biomediche, Biotecnologiche e Traslazionali (SBIBIT), Università degli Studi di Parma, Parma, Italy

Abstract Stem cells (SCs) are able to differentiate into many different cell types of the body during early life and growth. They are unspecialized cells capable of self-renewal and, when challenged, they can be induced to differentiate into specific and mature cell types. SCs can be classified into two different types: somatic stem cells (SSCs) and embryonic stem cells (ESCs). One major problem concerning in vitro SC culture is the difficulty to maintain their genetic and karyotype integrity during long-term culture. This chapter describes the different types of abnormalities accumulated by ESCs, derived from different species, and SSCs, of different somatic origin, during long-term culture in vitro. It outlines the main processes that are implicated in the onset of chromosome abnormalities in cells during culture. Among these processes, only one has been so far described in SCs. Keywords  stem cells; aneuploidy; chromosome instability

3.1 ­Introduction Stem cells (SCs) are unspecialized cells capable of self‐renewal and, when challenged, they differentiate into specific cell types with distinct and specific functions. They can be classified into two different types: somatic stem cells (SSCs) and embryonic stem cells (ESCs). In the 1960s, the haematopoietic SCs of the bone marrow were the first SSCs to be described. They are able to differentiate into all types of blood cells and into the stromal (or mesenchymal) SCs, the latter precursors of the variety of SCs from which the connective tissues (i.e., bone, cartilage, fat, blood cells, and the proper connective tissue) originate (Till and McCullough 1961). Up to now, SSCs have been identified in

*Corresponding author: [email protected] Rebuzzini P., Zuccotti M., Alberto Redi C., and Garagna S. (2017) Chromosome instability in stem cells. In: Li X.-Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley-Blackwell, Hoboken, NJ, Ch. 3, pp. 55–74. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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most organs where they maintain tissue homeostasis and participate in repairing processes. Within the body, SSCs are able to differentiate either in one, few, or multiple cell lineages and thus are classified as unipotent—for example, spermatogonia SCs, oligopotent (e.g., neural stem cells (NSCs)) or multipotent (e.g., mesenchymal stem cells (MSCs)). In vitro, SSCs may show a greater differentiation potential than in vivo. For example, MSCs are able to differentiate into mesodermic cells such as osteoblasts, adipocytes, chondrocytes, or myocytes, as well as ectodermic cells like neurons (Jiang et al. 2002; Franco Lambert et al. 2009), indicating their great plasticity. ESCs were derived for the first time in 1981 from pre‐implantation mouse embryos at the blastocyst stage (mESCs) (Evans and Kaufman 1981; Martin 1981); about 20 years later, the first human embryonic stem cell (hESC) line was generated from human blastocysts (Thomson et  al. 1998). In vitro, they have the potential to differentiate into almost any fetal or adult cell type and thus they are defined as pluripotent SCs. When transplanted into a blastocyst, they participate in the formation of both the somatic and germlines of the new individual (Rossant 2001; Smith 2001; Cockburn and Rossant 2010). Although the identification of both SSCs and ESCs represents a promise in regenerative medicine (Grompe 2012) and for toxicological studies (Seiler and Spielmann 2011), one major hurdle with their culture for prolonged periods of maintenance and differentiation is the difficulty to keep their genetic and chromosomal integrity (Zapata et al. 2012). Surprisingly, while SC derivatives for therapeutic purposes must pass strict quality controls (expression of specific markers, phenotype, mycoplasma contamination, viability and bacteriological testing, etc.; Durnaoglu et al. 2011; Vaes et al. 2012), at present, the evaluation of chromosomal and genomic stability is not included in the protocol of compulsory pre‐transplantation controls (Wu et al. 2011). The aim of this chapter is to give a detailed overview of the chromosome abnormalities described in both pluripotent (ESCs; induced pluripotent stem cells (iPSCs) and pathenogenetic embryonic stem cells (pESCs)) and SSCs and describe the main mechanisms involved in the generation of numerical abnormalities.

3.2 ­Pluripotent Stem Cells 3.2.1  Primate Embryonic Stem Cells

The variations in the chromosome complement occur non‐randomly in cultured human embryonic stem cells (hESCs). The chromosome changes observed affect more frequently chromosomes 12, 17, 20, and X. Many independent laboratories described a whole or partial gain of chromosomes 12 and 17, aneuploidy of chromosome X, or duplication of the 20q11.21 region (Brimble et al. 2004; Draper et al. 2004; Inzunza et al. 2004; Mitalipova et al. 2005; Imreh et al. 2006; Ludwig et al. 2006; Baker et al. 2007; Lefort et al. 2008; Spits et al. 2008; Lefort et al. 2009), suggesting that these specific types of mutations may confer a selective and/or proliferative advantage (Clark et al. 2004; Rosler et al. 2004; Caldas and Brenton 2005; Baker et al. 2007; Maitra et al. 2005; Navarro et al. 2008; Lefort et  al. 2008; Spits et  al. 2008; Blum and Benvenisty 2009; Werbowetski‐Ogilvie et al. 2009; Amps et al. 2011; Oliveira et al. 2014; Rebuzzini et al. 2015, 2016). The analysis of a remarkable number of hESC lines showed an additional recurrent genomic

3.2  Pluripotent Stem Cells

aberration, involving the amplification of a segment of chromosome 1. The duplication of 1q11–1q32 has been demonstrated to be associated with advantages in proliferation and metastasis formation; particularly, alterations in the 1q region may have a significant role in disease evolution by providing a growth and/or survival advantage (Ghose et al. 1990; Hanamura et al. 2006; Shaughnessy et al. 2007). A number of studies showed the variation of the chromosome complement of non‐ human primate embryonic stem cell lines (nhpESC) in Rhesus monkey (Macaca mulatta), common marmoset (Callithrix jacchus), and Cynomolgus monkey (Macaca fascicularis) (Thomson et al. 1995; Thomson et al. 1996; Nakatsuji and Suemori 2002). In contrast to hESC lines, early studies showed no clonal or recurrent abnormalities in all the nhpESC lines analyzed (Thomson et al. 1995; Thomson et al. 1996; Nakatsuji and Suemori 2002). Using a serum‐free medium in combination with trypsin detachment, the majority of Cynomolgus and Rhesus monkey ESC lines are able to maintain a normal chromosome complement, even after over a year of continuous culture (Nakatsuji and Suemori 2002). More recently, three unstable nhpESC lines (ORMES‐1, ORMES‐2, and ORMES‐5 from Rhesus monkey) have been characterized, showing, even at low passages, important structural abnormalities (translocations or inversions) (Mitalipov et al. 2006). 3.2.2  Mouse Embryonic Stem Cells

The first significant report that has highlighted the alterations in the chromosome make‐up of mESC lines was published by Longo et al. (1997). These authors made the important observation that when mESCs were injected into a mouse blastocyst, the absolute number of generated chimaeras and the ratio of chimaeras to total pups born was strictly correlated with the frequency of euploid metaphases in the ESC population (Longo et al. 1997). During prolonged culture, it is a general feature of mESCs to undergo chromosome number variations, as also documented in a very extensive cytogenetic study on 540 mESC lines (Sugawara et al. 2006). Depending on the cell line, the modal chromosomal number was found to be 41, 42, and 39, with recurrent trisomies of chromosomes 8 and 11 (Sugawara et al. 2006). Guo et al. (2005) reported both numerical (41, 43, 44, sub‐ or tetraploid chromosome complement) and structural (trisomy of chromosomes 8, 12, 14, and 15, deletion of chromosome 6q, and other aberrations with low frequency) alterations in four different lines. Similar results were obtained in our laboratory, when the chromosome complement of the R1 mESC line was analyzed at passage 30–32 (Rebuzzini et  al. 2011; Rebuzzini et  al. 2012). Importantly, in other mESC lines, we found a very heterogeneous spectrum of abnormalities that varied during a 3‐month culture period, suggesting their continuous arising and lack of selective pressure in favor or against a specific chromosome complement (Rebuzzini et al. 2008a; Rebuzzini et al. 2008b). 3.2.3  Parthenogenetic Embryonic Stem Cells

Parthenogenetic embryonic stem cell (pESC) lines have been derived from primate parthenogenetic embryos. Because the generation of pESC lines does not require the use of a human embryo, they do not raise the ethical problems of hESCs. Non‐human primate (Cibelli et  al. 2002; Vrana et  al. 2003) and more recently human (Kim et  al. 2007;

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Revazova et al. 2007; Brevini et al. 2009; Brevini et al. 2012) pESCs were generated and maintained in culture for many passages. Non‐human primate pESC lines remained karyotypically normal, with a diploid set of 42 chromosomes, even after in vitro culture for more than 2 years (80–120 passages) (Wei et al. 2011). Human pESC lines are more variable. Some cell lines (chHES‐32, hPES‐1, and hPES‐2) are stable and have normal karyotype (46, XX) even after more than 50–100 passages in culture (Lin et al. 2007; Mai et al. 2007). Following long‐term culture, other pESC lines lose one chromosome X, and the fraction of cells with this abnormality gradually increases in the population with culture passage (Liu et al. 2011). Chromosome translocations or deletions have also been described for hPES‐2 cell line, cultivated for more than 100 passages (Mai et al. 2007). Although mouse parthenogenetic embryonic stem cell (mpESC) lines were established in the 1980s (Robertson et al. 1983), few cytogenetic data are available. mpESCs are reported to be stable in culture, maintaining a normal karyotype (Shao et al. 2007; Ju et al. 2008; Shan et al. 2012). 3.2.4  Induced Pluripotent Stem Cells

The possibility to obtain in vitro pluripotent cells by the reprogramming of adult somatic cells represents an important advancement in SC research, as it may allow researchers to avoid the use of embryos, and, by the reprogramming of a patient’s own cells, is likely to overcome immunogenic responses. Since Takahashi and Yamanaka (2006) first reported the reprogramming of mouse fibroblast into pluripotent SCs (iPSCs), several methods have been used to reprogram different types of cells of different species (Jaenisch and Young 2008; Jaenisch 2009; Zhou and Ding 2010), but little attention has been given to the karyotype of the generated iPSC lines. iPSC lines seem to maintain a normal karyotype during the early culture passages (Takahashi et  al. 2007; Lowry et  al. 2008), but disparate chromosomal abnormalities, translocations, megabase‐scale duplications or deletions, and point mutations arise when cells are subjected to continuous passaging (about 60 passages) (Aasen et al. 2008; Mayshar et al. 2010; Gore et al. 2011; Martins‐ Taylor and Xu 2012; Rebuzzini et al. 2015, 2016). Chromosomes 8, 12, 17, and/or 20 are more frequently involved in aberrations, the same alterations found in hESCs (Aasen et al. 2008; Mayshar et al. 2010; Boulting et al. 2011; Peterson and Loring 2014). As suggested for hESCs, the presence of clonal aberrations in iPSC lines may facilitate the adaptation process during reprogramming (Mayshar et al. 2010). It has been demonstrated that karyotypically abnormal iPSC lines differentiate, for example in motor neurons, with an efficiency similar to that of karyotypically normal iPSC lines (Boulting et al. 2011).

3.3 ­Somatic Stem Cells 3.3.1­  Mesenchymal Stem Cells

Human adult mesenchymal stem cells (MSCs) are rare elements present in various organs. They have been initially found in the bone marrow; however, MSCs can also be isolated from other tissues including cord blood, peripheral blood, fallopian tube, skeletal muscle, and fetal liver and lung (for a review see Pittenger and Martin 2004; Da Silva et  al. 2006). MSCs have the capability to differentiate into various cell types

3.4  Mechanisms of Chromosomal Instability

(e.g., chondrocytes, adipocytes, and osteoblasts) and they are already used in therapy, that is, for Crohn’s disease and for tendon injury repair (Kraus and Kirker‐Head 2006; Granero‐Molto et al. 2008; Chao and Feng 2009; Ahmad et al. 2012; de la Portilla et al. 2012). The karyotypic stability of MSC cultures intended for transplantation was intensively studied and from this point of view MSCs are well characterized. The majority of cytogenetics data have been collected for MSCs derived from human bone marrow (hBM), with contradictory results (Pittenger and Martin 2004; Soukup et al. 2006; Bernardo et al. 2007; Grigorian et al. 2010; Nikitina et al. 2011; Tamaki et al. 2012; Zhao et al. 2012). Some authors showed that hBM‐derived MSCs remain chromosomally stable throughout long‐term culture (Pittenger and Martin 2004; Soukup et al. 2006, Bernardo et al. 2007; Tamaki et al. 2012; Zhao et al. 2012). In contrast, other laboratories demonstrated the onset of spontaneous numerical and structural chromosome aberrations, together with a spontaneous malignant transformation in long‐term cultures. hBM‐derived MSCs showed aneuploidy of autosomes 6, 8, and 11 and of the sex chromosomes, in particular trisomy of chromosome 8 and loss of chromosome Y (Grigorian et al. 2010; Nikitina et al. 2011; Rebuzzini et al. 2015). MSCs derived from other tissues appear to maintain a more stable karyotype. Human adipose‐derived (hAD) MSCs have a normal chromosome complement until passage 12, the last passage described (Ra et al. 2011; Yang et al. 2011), as well as chorionic villi (CV)‐ and amniotic fluid (AF)‐derived MSCs (Poloni et al. 2011; Angelo et al. 2012). MSCs have been derived also from human dental pulp (hDP) (Gronthos et al. 2000). These cells showed a normal karyotype until the Haflick limit (passage 65) (Suchánek et al. 2007; Tamaki et al. 2012). Beyond this limit, some chromosome abnormality has been described (2n = 44, XX, t (13, 14), –22; Suchánek et al. 2007). More recently, polyploid and aberrant metaphases (loss of chromosomes 2, 3, 5, 6, 7, 8, 9, and 16) were found in four hDP‐derived MSCs lines, in association with the presence of structural abnormalities, such as ring chromosomes (Duailibi et al. 2012). 3.3.2  Neural Stem Cells

Neural progenitor stem cells (NSCs) can be established either from the sub‐ventricular zone or from the ventral telencephalon of the brain. They show a limited capacity of mitotic expansion in vitro, and for this reason few papers have tackled the analysis of the chromosome complement. The few studies available report stability of their karyotype. Neither numerical nor structural chromosomal abnormalities were detected up to 70 passages (Bai et al. 2004; Jiang et al. 2008; Zhang et al. 2008a). In a paper by Wu et al. (2011), it has been shown that human neural SCs (human fetal striatum neural SCs) cultured until passage 17, undergo spontaneous malignant transformation, associated with different numerical (from 60 to 70 chromosomes per metaphase) and structural chromosomal aberrations.

3.4 ­Mechanisms of Chromosomal Instability A drawback that needs to be addressed when using SCs in regenerative medicine is the surprising complete lack of knowledge on the mechanisms implicated in the onset of aneuploidy. To date, a meagre number of papers only describe the involvement of

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supernumerary centrosomes in determining chromosome number variability in SCs (Zhang et al. 2007; Holubcova et al. 2011; Brevini et al. 2012). Clearly, other mechanisms must be causal to chromosome instability (CIN), as extensively demonstrated on terminally differentiated cells both in vivo and in vitro (Thompson et al. 2010; Holland and Cleveland 2012a). In the following sections, the main mechanisms and their role are detailed. 3.4.1  Dysfunction in the Spindle Assembly Checkpoints

During evolution, eukaryotic cells have developed a fine mechanism, known as the spindle assembly checkpoint (SAC) (for a review see Lew and Burke 2003), with the role of controlling erroneous attachment of chromosomes to the mitotic spindle. It represents a feedback control mechanism whose activity ensures accurate chromosome segregation (Musacchio 2011) and, when abnormalities may not be fixed, the cell enters apoptosis (Rieder and Maiato 2004; Musacchio and Salmon 2007). The SAC is controlled by the Bub–Mad pathway (Mad1/Mad1L1, Mad2/Mad2L1, BubR1/Bub1B, Bub3/Bub3; yeast or human/mouse protein name) and by ubiquitin (Ub)‐mediated degradation of regulators of sister chromatid cohesion by anaphase‐promoting complex/Ciclosome (APC/C). In addition, a group of modulator proteins, named Chfr, Rae1, Nup98, Cenp‐E, and Apc, cooperates in the activity of SAC (Li and Murray 1991; Weiss and Winey 1996; Amon 1999). The activation of SAC induces a delay of the cell cycle progression by negatively regulating CDC20, an essential regulator of cell division, in order to prevent the activation of the polyubiquitylation activities of APC and give the cells enough time to mend potential defects. Defects or dysfunction in proteins involved in SAC cause genomic instability or genetic material loss in the daughter cells and thus aberrant cells are generated (Musacchio and Salmon 2007; Rao et al. 2009). Knockout mice for Mad2 and Bub3 display a reduction of embryonic viability and high level of chromosome mis‐segregation (Dobles 2000; Kalitsis 2000). Moreover, mutations in Bub1, BubR1, Bub3, Mad1, and Mad2 genes have been detected in various types of cancers, in association with the onset of CIN (Cahill et al. 1998; Nomoto et al. 1999; Reis 2001). 3.4.2  Defects of Microtubule Attachment to the Kinetochore

Recently, live cell imaging has shown that the most common cause of CIN is due to wrong attachment of spindle microtubules to the kinetochore (Thompson and Compton 2008; Bakhoum et al. 2009). Erroneous attachments may result in the mis‐segregation of chromosomes during anaphase, leading to aneuploid cell progeny (Torres 2008; Tanaka and Hirota 2009). The correct kinetochore–spindle microtubule binding is the amphitelic (or biorientation) attachment, where the two sister chromatids’ kinetochores are exclusively connected to microtubules originating from the opposite spindle poles (Gerton 2012). The tension produced by the amphitelic attachment is likely on the basis of the high stability of this configuration, thus preventing chromosome mis‐segregation (Crighton et al. 2006). Several errors can occur in the microtubule–kinetochore coupling, leading to aneuploidy (Guerrero et  al. 2010). In monotelic attachment, only one of the two sister

3.4  Mechanisms of Chromosomal Instability

kinetochores attaches to the microtubules originated from one pole of the spindle, while the other does not attach to any microtubule of the opposite pole. Monotelic attachment is a frequent event in the early stages of chromosome orientation and can cause a mitotic delay (Silkworth et al. 2012). Two other types of erroneous kinetochore–microtubule binding can occur during spindle assembly, particularly in the first phase of the mitotic process: the syntelic or the merotelic attachments (Ault and Rieder 1992; Cimini et al. 2003; Lampson et al. 2004). Both types can cause CIN through the induction of chromosome mis‐segregation. In syntelic attachment, both sister kinetochores interact with microtubules that arise from a single spindle pole. In the merotelic attachment, a single kinetochore is connected to microtubules emanating from both poles of the spindle. Merotelic kinetochore attachment is considered the most frequent mechanism that gives rise to aneuploidy in mitotic cells (Gregan 2011) and it is the primary described mechanism of CIN in tumor cells (Cimini et al. 2001). The chromosomal passenger complex (CPC) is responsible for the correction of these anchoring errors. The complex consists of Aurora B kinase, that is the enzymatic core, its regulatory subunits Inner Centromere Protein (INCENP), borealin/Dasra‐B, and survivin (for a review see Adams et al. 2001; Ruchaud et al. 2007). Aurora B is activated and guided to its specific locations in the mitotic cell by INCENP, borealin, and survivin (Klein et al. 2006; Vader et al. 2006). The abolishment of the CPC in dominant negative mutants, by RNAi or by drugs, causes an accumulation of errors in chromosome anchoring, with the consequent onset of abnormalities (Adams et al. 2001; Kallio et al. 2002; Guerrero et al. 2010). In particular, it has been shown that Aurora B inhibition leads to an increase in the number of syntelic attachments (Hauf et  al. 2003), leading to the formation of polyploid cells (Gürtler et  al. 2010). In contrast, abnormally elevated levels of Aurora B give rise to irregular chromosomal separation during mitosis, resulting in the formation of cells with abnormal numbers of chromosomes (Katayama et  al. 2003; Chen et  al. 2009). Moreover, borealin reduction produces incorrect localization of chromosomes at the equatorial plane of the mitotic spindle and, consequently, two or more nuclei per cell are frequently formed (Yamanaka et al. 2008; Zhang et al. 2009). 3.4.3  Supernumerary Centrosomes

The centrosome is the principal microtubule‐organizing centre of eukaryotic cells, monitoring the number, organization, and polarity of microtubules (Figure 3.1A). During mitosis, aberrant centrosome duplication may adversely affect the maintenance of cell polarity. The presence of an abnormal number of centrosomes causes multipolar spindles and multipolar mitoses, resulting in chromosomal segregation abnormalities and aneuploid daughter cells (Salisbury et al. 1999; Krämer et al. 2002) (Figure 3.1B). Supernumerary centrosomes have been described in many tumor types and have long been considered to contribute to CIN and tumorigenesis (Krämer et  al. 2002; Nigg 2002). Nevertheless, some authors showed that centrosome amplification does not necessarily lead to spindle multipolarity (Quintyne et al. 2005), because extra centrosomes can cluster and the cells may divide with a ‘normal’ bipolar spindle (Acilan et al. 2008) (Figure 3.1C).

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(A)

(B)

(C)

Figure 3.1  Mitotic spindle organization: (A) normal bipolar spindle with two centrosomes; (B) multipolar spindle with three centrosomes; (C) bipolar spindle with clustered supernumerary centrosomes. In SCs only multipolar spindles with three or more centrosomes were described. Light gray bars represent mitotic chromosomes.

Several cell cycle kinases act on the centriole duplication (Strnad and Gönczy 2008; Loncarek and Khodjakov 2009; Nigg and Stearns 2011), and, in particular among these, the Polo‐like kinase 4 (Plk4) has a key role in centriole biogenesis. This kinase is in fact directly involved in numerical control of procentriole formation (Holland et  al. 2010). It has been clearly demonstrated that the decrease of Plk4 expression inhibits centriole assembly, and Plk4 overexpression induces formation of multiple procentrioles (centrosome amplification) around the mother centriole (Bettencourt‐Dias et al. 2005; Kleylein‐Sohn et al. 2007; Holland et al. 2010). In addition, it has been demonstrated that both Aurora A and CDK2 have an important role in centrosomal maturation, because their overexpression leads to centrosome amplification and chromosomal instability (Loncarek and Khodjakov 2009; Lukasiewicz and Lingle 2009). 3.4.4  Sister Chromatids Cohesion

During mitosis, the sister chromatids have to be correctly distributed into two daughter cells. Sister chromatid cohesion is the process by which sister chromatids are paired and held together for alignment on the metaphase spindle. The molecular basis of sister chromatid cohesion is a chromosomal protein complex, called cohesin (Michaelis et al. 1997). The cohesin complex is composed of four subunits, which form together a ring structure (Gruber et  al. 2003), rounded by two ABC‐type ATPases called structural maintenance chromosome 1 and 3 (Smc1 and Smc3) (Weitzer et al. 2003). In addition, two further essential cohesin subunits, Scc1 and Scc3, associate with the Smc heads (Haering et al. 2004). Sister chromatid cohesion is destroyed by separase to trigger the final segregation into daughter cells (Nasmyth 2001; Barbero 2011). Errors in the control of sister chromatid cohesion frequently lead to cell death or aneuploidy (Holland and Cleveland 2012b). Somatic mutations in genes involved in the maintenance and regulation of sister chromatid cohesion (i.e., cohesin) were identified in human tumors, resulting in CIN (Barber et al. 2008). The mechanism by which CIN is induced in cells carrying cohesin‐mutated genes is still unknown, but it seems that the lack of cohesins induces premature sister separation or failed chromosome disjunction at the beginning of anaphase (Zhang et al. 2008b; Iwaizumi et al. 2009).

  References

3.5 ­Mechanisms of Chromosomal Instability in Stem Cells To date, only one mechanism has been studied and shown to be involved in the appearance of chromosome abnormalities in SCs (Holubcova et  al. 2011; Brevini et  al. 2012): hESCs suffer centrosome amplification (Holubcova et  al. 2011). In 12 independent hESC lines, the presence of an inadequate number of centrosomes (more than two centrosomes per cell) (Figure 3.1B) was detected in both interphasic and mitotic cells. Although the causes of this incorrect number of centrosomes are still unclear, it has been suggested that the higher expression of Aurora A and CDK2 in growing hESCs compared to human somatic cells could be responsible for the deregulation of centrosome number (Holubcova et al. 2011). In hpESCs, the occurrence of centrosome amplification has been correlated to an altered expression level of mitotic spindle‐related molecules such as BUB1, CENPE, and MAD2 (Brevini et  al. 2012). In SSCs, supernumerary centrosomes have been observed, but, up to now, the molecular mechanism that induces the alteration of the number of centrosomes is still unknown (Zhang et al. 2007). Nevertheless, it has been shown that supernumerary centrosomes interfere and often disrupt the asymmetric division, leading to an improper expansion of the SC population with subsequent increase of CIN and malignant transformation (Wodarz and Gonzalez 2006; Zhang et al. 2007; Basto et al. 2008; Zhang et al. 2008b). In this chapter, we aimed to bring to light an aspect of SC biology that has been so far underestimated; that is, the need for systematic control of the maintenance of a correct karyotype complement when cultured in vitro. As a consequence, we highlighted the urgency for studies on the mechanisms involved in determining chromosome instability, exploiting the knowledge acquired earlier on other cell types.

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Part II Somatic Genome Variation in Plants

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4 Mechanisms of Induced Inheritable Genome Variation in Flax Christopher A. Cullis* Department of Biology, Case Western Reserve University, Cleveland, USA

Abstract The characteristics of environmentally induced genomic changes in flax are reviewed in this chapter. The changes have been identified through characterization of specific regions of the genome as well as through complete genome sequencing. The characteristics of the genomic changes that have been observed in the progeny of lines after growth for a single generation under stress environments are compared with those observed while responsive flax varieties are growing under the various environments. The characteristics of the flax phenomenon are compared with seemingly analogous observations from a wide range of organisms. This enables these observations to be looked at in the wider context of other established instances of non-random frequent genomic alterations. Keywords  flax; genotrophs; genome restructuring; stress adaptations

4.1 ­Introduction Particular varieties of flax are capable of responding to environmental stress with the appearance of heritable phenotypic variations. The synopsis of these observations is that the growth of plants for a single generation in a particular environment resulted in heritable differences in phenotype that have persisted for more than 40 generations, during which the variant phenotypes (termed genotrophs) have been grown in a common environment. If the observations of heritable phenotypic modifications in flax in response to the growth environment (outlined in Figure 4.1) (Durrant 1962; Cullis 1977; Cullis 1979) were described in the current literature they would be interpreted as a typical example of epigenetic variation. Since the traits that differed included plant height and branching patterns, the pathways controlling these phenotypic characters would be examined for the presence of epigenetic marks that would explain the altered growth patterns. A suggestion that *Corresponding author: [email protected] Cullis C.A. (2017) Mechanisms of induced inheritable genome variation in flax. In: Li X.-Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley-Blackwell, Hoboken, NJ, Ch. 4, pp. 77–90. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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Pl (inducible line) Non-inducing environment

Low nutrient environment

Pl

S

DNA content

100 units

96 units

rDNA

2000 copies

1000 copies

LIS-1

Absent

Present

Inducible

Seeds breed true

Figure 4.1  Induction of heritable changes in flax. The growth of the original line Stormont Cirrus (Pl) under different environmental conditions can result in new stable phenotypes with associated genomic variation. Shown is the appearance of the small genotroph following growth for a single generation under inducing conditions, and individuals after 40 additional generations of growth under standard non‐inducing conditions where the original phenotypic and genotypic differences have persisted.

there would be any DNA sequence variation in the generation of these responses would be deemed redundant since the genome is essentially conserved through generations. However, the observations were made at a time when the information concerning the possibility of such epigenetic control was minimal and the accepted view was that such heritable phenotypes would be conditioned in changes in the genome. Therefore the genomes of these stable phenotypes have been compared and the accumulated data have demonstrated unequivocally that the flax genome can undergo restructuring in response to the growth environment. What still remains to be demonstrated is a direct link between any of the genomic changes and the resultant phenotypes, leaving open

4.2  Restructuring the Flax Genome

the possibility that the heritable variation is a true epigenetic phenomenon and the genomic restructuring is an inconsequential aberration. However, the nature of the variants and their appearance only under certain growth conditions makes it more likely that they are responsible for the phenotypic variation. The flax varieties that undergo restructuring of their genomes, with concomitant phenotypic variation in the following generation, in response to the growth environment have been termed ‘plastic,’ with the most studied example being the variety Stormont Cirrus. Particular nutrient regimes, such as an imbalance of nitrogen or starvation, consistently lead to significant DNA changes while the plant is growing (Evans et al. 1966; Cullis and Charlton 1981; Cullis 2005). These variations are inherited in the following generation, resulting in either stable lines (termed genotrophs) or unstable lines still capable of responding to environmental cues (Evans et al. 1966; Cullis 1981). Of particular note is that the immediate progeny of the genotrophs resulting from a single generation’s growth in the extreme stress environments bred true; that is, all the changes that had occurred had become homozygous within that single generation.

4.2 ­Restructuring the Flax Genome Given the historical perspective of when the original observations were made (1954), since these stable lines had been altered through a single generation growth in specific environments and the variation was heritable, the obvious place to start investigating the cause of the variation was in the nuclear DNA. The only available technique was to measure the total DNA content by Feulgen cytophotometry. These measurements demonstrated a substantial difference between the lines and a correlation existed between these nucleotypic and phenotypic changes (Evans et  al. 1966; Evans 1968). These original observations have been amplified using the developing DNA analysis tools since the early 1970s through renaturation kinetics (Cullis 1973), copy number variations of cloned repetitive sequences (Cullis 1979; Cullis and Cleary 1986a; Cullis and Cleary 1986b; Schneeberger and Cullis 1991), identification of single copy variants (Chen et  al. 2005; Cullis 2005; Chen et  al. 2009), and finally the complete genome sequencing of the original line and three genotrophs. The differences between the total nuclear genomes were initially confirmed using renaturation kinetics (Cullis 1973). Subsequently, sub‐regions of the genome that were altered in the genotrophs when compared to the original parent line were identified. These regions included the ribosomal RNA (rRNA) genes and a series of tandemly arrayed repetitive families (Cullis 1976; Cullis 1979; Cullis 1983; Cullis and Cleary 1986a; Cullis and Cleary 1986b; Cullis 1987). Of particular interest was that specific subsets within these repetitive families were differentially targeted for variation (Cullis 1973; Schneeberger and Cullis 1992), which was consistent with the hypothesis that regions of the genome were particularly susceptible to this restructuring, while the majority of the genome remained stable (Cullis 1977). In spite of the total nuclear DNA content and specific subsets of the genome being altered between the original line and the genotrophs, these were difficult characteristics to measure during the growth of the Pl plants under inducing conditions. However, both the nuclear DNA (Evans et al. 1966; Evans 1968) and the rRNA gene number (Cullis and Charlton 1981) were shown to vary within the plant as it was growing under various nutrient regimes; that is, the genomic

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changes were occurring in somatic cells and not restricted to the germline. The genomic changes could be transmitted to subsequent generations, confirming that the variation had to occur in the meristematic cells prior to flower formation. This was the first observation that the environment in which the plant was growing could affect the actual genome and that the variation could occur during this vegetative growth phase and still be passed to subsequent generations.

4.3 ­Specific Genomic Changes The genomic changes are spread over all of the 15 chromosomes, and include variation in tandemly arrayed highly repetitive sequences as well as low copy‐number sequences (Cullis et al. 1999; Oh and Cullis 2003; Chen et al. 2005). In the flax genome, both the overall copy number and specific subsets of many of the tandemly arrayed highly repetitive sequence families, including the genes that code for the rRNAs, can be modulated during growth under various nutrient regimes and these altered numbers can be inherited in subsequent generations (Cullis 1983; Cullis and Cleary 1986a). A single copy sequence that can also alter in response to the nutritional environment is a novel type of insertion termed Linum Insertion Sequence 1 (LIS‐1) (Chen et  al. 2005). This single‐ copy 5.7‐kb DNA fragment generally appeared in response to growth under poor environments but not under optimal growth conditions, and is a precise insertion into a single copy site. However, the intact LIS‐1 cannot be detected in the line in which it arises prior to its appearance. Not only does LIS‐1 appear reproducibly at a unique site in a single generation after an environmental stress from a source that cannot be identified as an intact donor element, but also it becomes homozygous almost immediately. The identification of LIS‐1 was the first indication that a novel mechanism was operating in this pathway of genome restructuring (Chen et al. 2005; Cullis 2005; Chen et al. 2009). The novel observation was that this element was not present intact in the progenitor genome prior to its appearance (Chen et al. 2005; Chen et al. 2009). The absence was initially demonstrated through hybridization to genomic Southern blots and by PCR amplifications using primer pairs distributed across the element. The absence of an intact element in the progenitor Pl line has been confirmed by whole genome shotgun sequencing. However, LIS‐1 has been shown to be present in some fiber flax varieties as well as some of the accessions of Linum bienne, the wild progenitor of cultivated flax, confirming that this element is a recognizable, if unstable element of the Linum genome. That the appearance of LIS‐1 may not be a unique event has been verified through the complete next‐generation sequencing of Pl, and two small genotrophs (S and C3). These genome sequences have been aligned to the assembled genome of the oilseed reference variety Bethune (Wang et al. 2012). These alignments have been interrogated to identify differences between Pl and the genotrophs, but since they are all aligned to the Bethune genome, any differences in regions that are limited to the genomes of fiber varieties are missing from the data. Many differences have, however, been identified between Pl and two small genotrophs (S and C3) that can be generally classified into three groups. The first is where Pl and Bethune are identical but the genotrophs have a deletion in this region (Figure 4.2). The deletion of a region can occur by a number of different mechanisms. In some of the characterized regions the deleted fragment is flanked by a short direct repeat, which is consistent with the involvement of a transposase in the rearrangement.

4.3  Specific Genomic Changes

Figure 4.2  Alignment of next‐generation sequencing reads (approximately 100× sequencing) from Pl, and the two small genotrophs S and C3, to the Bethune genome. Both S and C3 have a deletion in a region where Pl has the same sequence as Bethune. The deletion has been confirmed by PCR across the region, where the lane MI is Bioline molecular marker I.

The second is where Pl differed from the genotrophs, with the region not present in Pl but present in the genotrophs. The sequence present in the genotrophs was identical to the sequence present in Bethune (Figure 4.3). This category is similar to the observations with LIS‐1, where a sequence is present in the genotrophs that is missing from the progenitor line. The origin of such a sequence is more difficult to explain. The restructuring of genomes with transposons, be they retro‐elements, DNA transposons, or heliotrons, depends on an element that is already present in the genome being mobilized. The data

Figure 4.3  Alignment of next‐generation sequencing reads (approximately 100× sequencing) from Pl, and the two small genotrophs S and C3, to the Bethune genome. Pl differs from the genotrophs with a region not present in Pl but present in the genotrophs, and that region in the genotrophs is identical to the sequence in Bethune. The deletion has been confirmed by PCR across the region, where the lane MI is Bioline molecular marker I.

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in flax are the first that are a result of novel genomic sequence arising de novo. The identification of some of these sequences as similar or identical to those already identified in Bethune indicates a programmed process by which the genomic regions arise. LIS‐1 is, to date, the only example that has been identified that is not present in Bethune, but this may simply be an artifact of the analysis thus far. The comparisons of the genomes of Pl and the genotrophs have all been done by alignment with the published Bethune sequence. Although LIS‐1 is present in some flax and Linum bienne lines, it is not present in Bethune so cannot be identified in the whole genome comparisons with the alignments to the Bethune genome. Therefore any regions (such as LIS‐1) that are not present in Bethune will not yet have been identified. As the assembly of the Pl and genotroph genomes proceeds, regions that are not present in Bethune will be identified and more LIS‐1‐type of events are likely to be uncovered. That the examples of both these deletions and insertions were present in the genomes and not an artifact of the assembly and alignment processes has been confirmed by PCR (Figures 4.2 and 4.3). The third category is where there are copy number differences between Pl and the genotrophs, which confirms some of the quantitative polymorphisms previously identified (Figure 4.4). These quantitative polymorphisms account for most of the nuclear DNA differences observed in the earlier nuclear DNA amount measurements. As seen in Figure 4.4, the repetitive sequence is reduced in S and C3. Other examples with an amplification of the repetitive sequence in the genotrophs have also been observed. As hypothesized previously (Cullis 1977), the total nuclear differences between Pl and the genotrophs was the result of a balance between amplifications and deletions, a larger genome having more amplifications than deletions, while a smaller genome had the inverse relationship. The regions that are modified between Pl and the genotrophs will be characterized to determine if there are conserved motifs that are involved in the molecular mechanisms by which the genome is restructured. In addition the chromatin structure of the regions that are highly modified can be determined in order to understand if this is the basis of the recognition of the variable compartment of the genome.

Figure 4.4  Alignment of next‐generation sequencing reads (approximately 100× sequencing) from Pl, and the two small genotrophs S and C3, to the Bethune genome. There are copy number differences between Pl and the genotrophs. The number of reads from the Pl sample is five to six times greater than that from either S or C3. S and C3 appear slightly different from each other, which has previously been observed in the genotrophs where the 5S and large rRNA genes vary but were not exactly identical in all lines.

4.5  When Do the Genomic Changes Occur and Are they Adaptive?

4.4 ­What Happens When Plastic Plants Respond to Environmental Stresses? Most of the salient features of the process of the induction of heritable genomic changes in flax are demonstrated by considering the features of three sets of sequences, namely the genes for the 18S and 25S rRNAs , the genes for the 5S rRNA, and the sequence LIS‐1. All three sets of sequences have been shown to be affected during growth under inducing conditions (Cullis 1976; Cullis and Charlton 1981; Cullis 1983; Cullis and Cleary 1986a; Cullis 2005). In each case, changes occurred during the growth of the plants under inducing conditions and these changes were inherited in the next generation (and in the genotrophs, or stable lines, for a large number of subsequent generations). A particular subset of the 5S rDNA (the gene encoding for the 5S ribosomal RNA) was preferentially deleted (Cullis and Cleary 1986a; Cullis and Cleary 1986b) and specific polymorphisms were present after the genomic changes occurred (Schneeberger and Cullis 1991), indicating a non‐random targeting of this gene family. In every case, the progeny of the induced generation were homozygous for any inherited induced genomic changes. In other words, when selfed, the genotrophs showed no segregation for copy number for any of the rDNA or 5S rDNA, or for the presence of LIS‐1. However the behavior of the rDNA and LIS‐1 in crosses between large and small genotrophs was different. The rDNA amount in the F1 individuals in a cross between two genotrophs with differing numbers of these genes did not fall into the expected range but had values spread through the range of the two parents (Cullis 1979). The F2 individuals from selected selfed F1s did not show segregation for the number of this gene family but had a value correlated with the number of genes in the F1 plant from which that F2 family was derived. Therefore there appeared to be a post‐fertilization equalization of the rDNA on the parental chromosomes in these crosses that was then stably inherited. This interaction is reminiscent of the loss of heterozygosity in some crosses in rice (Wang et al. 2001). In contrast, however, the inheritance of LIS‐1 in the F2 was, as expected, in a normal Mendelian fashion (Chen et al. 2005). Thus both these regions of the genome are affected during growth under inducing conditions, but LIS‐1 is stable in crosses between genotrophs, while the rDNA is unstable in the same crosses, again underscoring the selectivity of which regions of the genome are labile under particular circumstances. Data arising from the complete genome comparisons indicate that some of the variable regions show non‐Mendelian inheritance in the F2 from a cross between Pl and the oilseed variety Bethune.

4.5 ­When Do the Genomic Changes Occur and Are they Adaptive? It has been shown that the genomic changes occur during growth under inducing conditions prior to flowering (Evans et al. 1966; Cullis and Charlton 1981; Cullis 2005). There are subsets of the 5S genes, in particular the long and short forms of the sequence present (Goldsbrough and Cullis 1981). These two length variants are differentially modulated during growth under various nutrient regimes and are also differentially represented in the genotrophs. However, in individuals growing in the same environment no consistent

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end point for the relative numbers of the two variants appears to be achieved at flowering (Figure 4.5). A contrasting situation occurs with LIS‐1. Under some growth conditions (especially those with an imbalance of nutrients or low nutrients that reduce growth) LIS‐1 always arises, becomes homozygous, and is inherited by all the progeny of these induced plants, while under other growth conditions (good growth conditions) it is never inherited, although it can arise during growth of plants under these good growth conditions (Figure 4.5) (Cullis 2005). Under conditions where the induced changes are not stabilized after the first generation under inducing conditions but where LIS‐1 has been inserted, then LIS‐1 can be lost in the next generation. This can happen if the growth environment is changed to one where LIS‐1 is not observed to be induced, that is, from poor growth conditions in the first generation to good growth conditions in the second generation. This appearance and movement to homozygosity of LIS‐1 in 100% of the plants under some conditions, its complete absence under others, as well as its loss on removal of the specific inducing conditions is consistent with this locus being under positive selection. One possibility is that the presence of the insertion has a phenotypic effect within the meristem that is beneficial (adaptive) under those treatments where it arises, but is either deleterious or neutral in other growth environments. Therefore the cells in the meristem containing LIS‐1 would be selected for in one growth environment but selected against in others. The selection probably takes place in the meristem, since if the selection occurred in other organs then there should be no, or at least a variable, transmission to the next generation. Since this is not observed, it is likely that the origin of LIS‐1 is in the meristem, and that this is also the site of selection. The loss of LIS‐1 when not stabilized again supports the notion that this region is under some direct selection in various growth environments. Selection at the meristem level is also supported by the appearance of ‘adapted’ branches growing out under extreme environments (Cullis 1986). The lack of consistency relating to changes within the rDNA and the 5S rDNA suggests that all the changes within these gene families are not specifically adaptive, although it is possible that a subset within these families is adaptive but cannot yet be distinguished because of the ‘noise’ from all the other changes in these gene families. The identification of the large number of genomic loci that differ between Pl and the genotrophs can be used to follow the progression of genomic changes while Pl is growing under inducing conditions.

4.6 ­Is this Genomic Response of Flax Unique? Phenomena that may have a similar underlying mechanism include somaclonal variation, transposon restructuring of the genome, and loss of heterozygosity in certain crosses. Somaclonal variation is the genetic variation that is observed when plants are regenerated from cultured somatic cells. Various types of mutations have been described in somaclonal variants, including point mutations, gene duplication, chromosomal rearrangements, and chromosome number changes. The phenotypes that frequently arise in somaclonal variation in many species include plants of different stature (e.g., dwarf plants in banana) and flower mutations (e.g., the mantled mutation in oil palms). At

4.6  Is this Genomic Response of Flax Unique?

Figure 4.5  Changes in LIS‐1 and 5SrDNA following growth under various nutrient regimes. PCR amplifications from DNA extracted from the upper leaves of Stormont Cirrus plants grown under three different nutrient regimes, with primers (a) spanning the left junction of LIS1, (b) the uninserted site, (c) the right junction of LIS1, and (d) primers specific for the spacer region of 5SrDNA represented by the clone pBG13 (Goldsbrough and Cullis 1981). The two bands in (d) represent the two spacer length variants found in pBG13 and differ by 21 base pairs. Lanes 1 and 2 show two plants grown with an imbalance of nitrogen; lanes 3 and 4, two plants grown in soil without any added nutrients; lanes 5 and 6, two plants grown with balanced nutrients. M, Molecular weight marker VI (Roche). All the other plants grown under each of the set of nutrient conditions had the same LIS‐1 status as those shown in the figure. However, there was no consistent pattern of relative amounts of the two spacer‐length variants for the 5SrDNA among these same plants. This complete reproducible conversion of all the plants to homozygous LIS‐1 containing under two of the nutrient regimes and never under the third regime is consistent with LIS‐1 being selected for under some nutrient regimes and either neutral or deleterious under others.

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the genome level, a series of hot‐spots in the genome that are physically modulated in somaclonal variants has been identified (Cullis and Kunert 2000; Linacero et al. 2000). Therefore, as in the flax example, a limited suite of changes occurs in somaclonal variants at both the phenotypic and genomic levels. Of note is that many of the genomic regions that are modulated by growth conditions are also modulated in regenerated flax plants (Cullis and Cleary 1986b). Many of the somaclonal variants have been termed epigenetic, as they can revert (be unstable) during vegetative growth. However, as can be seen with LIS‐1, such revertants could also be accompanied by a physical genome modification at the sequence level, making the differentiation between genetic and epigenetic variation more difficult to confirm. Vegetatively propagating species are a probable source of active genome restructuring due to the lack of reassortment possibilities offered by sexual reproduction. The plantain cultivar Bise Egome 2 is the epitome of a hypervariable banana, where each cycle generates additional new off‐ types (Vuylsteke et al. 1991). A comparison of the genomes of various phenotypes of this plantain may shed more light on the nature of the hypervariable regions in this plant genome. The use of high‐throughput next‐generation sequencing has started to uncover unexpected genomic variation. For example, complete genome sequencing and analysis of different tissues within a single plant as well as clones of black cottonwood trees (Populus trichocarpa) has identified differences arising somatically (Yong 2012). The variation observed indicated that the differences within an individual could be greater than the differences between equivalent tissues within cloned relatives. This means that the genomic differences between the leaves and roots of an individual might be greater than the differences between the leaves of cloned relatives. As the data from sequencing of different tissues within an individual and across generations are reported, the fluidity of the genome and the response to the growth environment will become more apparent. The selectivity of LIS‐1 and the limitation of changes to restricted regions of the genome have some parallels with the restructuring of the grass genomes by transposons. A variety of ‘stresses’ are known to activate transposable elements, and these elements vary widely in their target site specificity (Lönnig and Saedler 2002). The hot‐spots for transposable element integration may define pre‐established rather than accidental chromosome rearrangements for non‐homologous recombination of host DNA mediated by these elements (Lönnig and Saedler 2002). In many cases there are short direct repeats at the margins of the regions that differ between Pl and the genotrophs, as with the insertion of LIS‐1 that is accompanied by a 3‐bp duplication at the insertion site, indicating the possible involvement of transposases in the mechanisms. A possible analogous mechanism exists in ciliates with the development of the macronucleus, where extensive DNA rearrangements result in the remodeling of the new somatic genome. These developmentally programmed DNA rearrangements precisely eliminate tens of thousands of short germline‐limited segments and variably excise transposon sequences (Chalker 2005). The process can be affected by PIWI‐interacting small RNAs (piRNAs) from the maternal nucleus and transposases that can specify genomic regions for retention or elimination in this process (Fang et al. 2012). Thousands of specific DNA elements (termed the internal eliminated sequences (IESs)) are deleted, with varying degrees of specificity, resulting in a genome‐wide scale of DNA rearrangements. As the details of the macronuclear rearrangement processes are identified they

  References

can be used as models to determine whether or not other organisms utilize the same mechanisms to restructure their genomes. The progress to homozygosity in all the induced changes in flax may have parallels in the loss of heterozygosity in certain crosses of rice (Wang et al. 2001). In hybrids that included a particular parent and some of the subsequent selfed generations, somatic variations have been identified in molecular genotypes and/or morphological phenotypes in vegetative parts of the same plant. Random amplified polymorphic DNA (RAPD) markers were found to be uniformly present or absent in all plants within some or all F2 panicle rows derived from F1 hybrids involving the specific parent, although the expected Mendelian segregation for these dominant markers in panicle rows derived from control hybrids was observed. Some of the regions of the genome which appear to be highly variable in response to the growth environment also are inherited in a non‐Mendelian fashion. In some cases there is a preponderance of alleles from one of the parents in the F2, while other loci segregate in the expected fashion. Since the F1 parents show the expected heterozygosity, the interaction post fertilization is restricted to some of the genome. Until more is known about the genes controlling the response in these systems the parallels between the two sets of observations will have to remain at a phenomenological level.

4.7 ­Concluding Remarks The first reports of the environmental induction of heritable changes in flax (Durrant 1958) set the system apart with no comparable observations. However, the ability to dissect genomes has both increased the number of organisms in which potential adaptive variation has been demonstrated and identified the methods by which these organisms achieve their ends. Currently, the flax system still shows the largest range and frequency of identified genomic variants. An understanding of how these variants are generated, the genetic control of the response to environmental stress, and the way the labile regions of the genome are differentiated will be useful in understanding other potentially analogous systems. LIS‐1 and the sets of loci that have similar characteristics have intrinsic interest in terms of how they arise, as well as potential use as a marker to map the genes controlling plasticity in flax.

­Acknowledgments Thanks go to T. Yael Moss, Kyle Logue, and David Serre for assistance with the whole genome sequencing and analysis. Some of the work described here was supported through the Oglebay Fund.

­References Chalker, D.L. 2005. Genome rearrangements: mother knows best! Curr Biol 15(20):R827–R829. Chen, Y., R.G. Schneeberger, and C.A. Cullis. 2005. A site‐specific insertion sequence in flax genotrophs induced by environment. New Phytol 167:171–180.

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Chen, Y., R. Lowenfeld, and C.A. Cullis. 2009. An environmentally induced adaptive (?) insertion event in flax. J Genetics Mol Biol 1:38–47. Cullis, C.A. 1973. DNA differences between flax genotypes. Nature 243:515–516. Cullis, C.A. 1976. Environmentally induced changes in ribosomal RNA cistron number in flax. Heredity 36:73–80. Cullis, C.A. 1977. Molecular aspects of the environmental induction of heritable changes in flax. Heredity 38:129–154. Cullis, C.A. 1979. Quantitative variation in the ribosomal RNA genes in flax genotrophs. Heredity 42:237–246. Cullis, C.A. 1981. Environmental induction of heritable changes in flax: defined environments inducing changes in rDNA and isozyme band pattern. Heredity 47:87–94. Cullis, C.A. 1983. Environmentally induced DNA Changes in plants. CRC Crit Rev Plant Sci 1:117–129. Cullis, C.A. 1986. Phenotypic consequences of environmentally induced changes in plant DNA. Trends Genetics 2:307–310. Cullis, C.A. 1987. The generation of somatic and heritable variation in response to stress. Am Nat (Suppl)103:562–573. Cullis, C.A. 2005. Mechanisms and control of rapid genomic changes in flax. Annal Bot 95:201–206. Cullis, C.A. and L.M. Charlton. 1981. The induction of ribosomal DNA changes in flax. Plant Sci Lett 20:213–217. Cullis, C.A. and W. Cleary. 1986a. Rapidly varying DNA sequences in flax. Can J Genet Cytol 28:252–259. Cullis, C.A. and W. Cleary. 1986b. DNA variation in flax tissue culture. Can J Genet Cytol 28:247–251. Cullis, C.A. and K. Kunert. 2000. Isolation of tissue culture‐induced polymorphisms in bananas by representational difference analysis. Acta Hortic 530:421–428. Cullis, C.A., Y. Song, and S. Swami. 1999. RAPD polymorphisms in flax genotrophs. Plant Mol Biol 41:795–800. Durrant, A. 1958. Environmental conditioning of flax. Nature 181:928–929. Durrant, A. 1962. The environmental induction of heritable change in Linum. Heredity 17:27–61. Evans, G.M. 1968. Nuclear changes in flax. Heredity 23:25–38. Evans, G.M., A. Durrant and H. Rees. 1966. Associated nuclear changes in the induction of flax genotrophs. Nature (Lond) 212:697–699. Fang, W., X. Wang, J.R. Bracht, M. Nowacki, and L.F. Landweber. 2012. Piwi‐interacting RNAs protect DNA against loss during Oxytricha genome rearrangement. Cell 151:1243–1255. Goldsbrough, P.B. and C.A. Cullis. 1981. Characterization of the genes for ribosomal RNA in flax. Nucleic Acid Res 9:1301–1309. Linacero, R., E.F. Alves, and A.M Vazquez. 2000. Hotspots of DNA instability revealed through the study of somaclonal variation in rye. Theoret Appl Genet 100:506–511. Lönnig, W.E. and H. Saedler. 2002. Chromosome rearrangements and transposable elements. Annu Rev Genet 36:389–410. Oh, T.J. and C.A. Cullis. 2003. Labile DNA sequences in flax identified by combined sample representational difference analysis (csRDA). Plant Mol Biol 52:527–536.

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Schneeberger, R.S. and C.A. Cullis. 1991. Specific DNA alterations associated and the environmental induction of heritable changes in flax. Genetics 128:619–630. Vuylsteke, D., R. Swennen, and E. De Langhe. 1991. Somaclonal variation in plantains (Musa spp. AAB group) derived from shoot‐tip culture. Fruits 46:429–439. Wang, R.R.‐C., X. Li, and J.N. Chatterton. 2001. A proposed mechanism for loss of heterozygosity in rice hybrids. Euphytica 118:119–126. Wang, Z., N. Hobson, L. Galindo, S. Zhu, D. Shi, J. McDill, L. Yang, S. Hawkins, G. Neutelings, R. Datla, G. Lambert, D.W. Galbraith, C.J. Grassa, A. Geraldes, Q.C. Cronk, C. Cullis, P.K. Dash, P.A. Kumar, S. Cloutier, A. Sharpe, G.K.‐S. Wong, J. Wang, and M.K. Deyholos. 2012. The genome of flax (Linum usitatissimum) assembled de novo from short shotgun sequence reads. Plant J 72(3):461–473. Yong, E. 2012. Tree’s leaves genetically different from its roots. Nature August 10, doi:10.1038/nature.2012.11156.

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5 Environmentally Induced Genome Instability and its Inheritance Andrey Golubov* Department of Biological Sciences, University of Lethbridge, Lethbridge, Canada

Abstract The necessity of research on environmentally induced genome instability and its inheritance is dictated by our curiosity and attempts to improve crops that are important for our lives. A considerable body of data, despite sometimes being controversial, is available for uncovering the molecular mechanisms of plant stress tolerance, and the adaptation and passage of the induced genomic changes through generations. This chapter reviews publications on this subject, characterizes stress-induced genome rearrangements and plant stress responses at the molecular level, and investigates how alterations in gene expression after stress exposure can be inherited. Keywords  environmental exposure; genome instability; epigenetic modifications; genetics changes; DNA methylation; histone modifications; transgenerational inheritance

5.1 ­Introduction Plants cannot escape harsh environmental conditions. There is a very high chance that plantlets will experience biotic and abiotic stresses in a similar way to their predecessors. It has been shown that stress in plants does not necessarily damage DNA directly; rather it leads to epigenetic modifications that can result in changes in the genome stability. Plants need to sense environmental stimuli and react accordingly; they also need to adjust their metabolic pathways in order to maintain the stability of their genomes (referred to as genome plasticity). Stressful conditions usually cannot persist for many generations, which means that there should be a mechanism of reversible inheritable changes that would allow some degree of genome plasticity, yet maintaining genome stability. Plantlets pre‐programed for a certain type of stress will have evolutionary advantages over plantlets from non‐stressed parental plants. *Corresponding author: [email protected] Golubov A. (2017) Environmentally induced genome instability and its inheritance. In: Li X.-Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley-Blackwell, Hoboken, NJ, Ch. 5, pp. 91–102. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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Recent genetic studies have linked changes in DNA methylation patterns to the plant immune response, suggesting that some genes involved in the immune response may be directly modulated by cytosine methylation (Alvarez et al. 2010). In this chapter, we describe plant responses at the molecular level to various types of stresses in terms of genome instability and also discuss the mechanisms of transgenerational inheritance of stress‐induced genomic changes.

5.2 ­Stress and its Effects on Genomes Plants often grow under various types of stresses: biotic stresses caused by interactions with other plants, viruses, bacteria, fungi, insects, and abiotic stresses due to strong winds, extreme temperatures, drought, flood, UV light, salinity, nutrient deficiency, and high or low soil pH. All these types of stresses may induce heritable genomic changes. 5.2.1  Genetic Changes

Biotic or abiotic stresses may sometimes induce various types of DNA damage. For example, UV‐A light can generate DNA‐damaging free radicals, UV‐B light can induce pyrimidine dimmers, ionizing radiation may cause DNA double‐strand breaks, and high temperatures can increase the rate of hydrolytic depurination, deamination, and backbone scission (Hutchinson 1985; Lindahl 1993). We have found that abiotic factors, such as cold, salt, UV‐C, ionizing radiation, heavy metals, temperature, drought, and flood, can increase the frequency of point mutations, homologous recombination, and microsatellite instability in somatic cells in Arabidopsis plants (Boyko et al. 2005; Boyko et al. 2006a; Boyko et al. 2006b; Alvarez et al. 2010; Boyko et al. 2010b). Endogenous toxic molecules are another type of DNA‐damaging agents. They are represented by reactive oxygen species (ROS), such as superoxide anion, hydrogen peroxide, and hydroxyl radicals. It has been shown that up to 2% of oxygen consumption leads to superoxide formation in plant tissues (Puntarulo et al. 1988). ROS can heavily damage DNA, resulting in double‐ and single‐strand breaks, base modifications (oxidation, alkylation, and hydrolysis), and the formation of DNA–protein crosslinks. A good example of genetic changes caused by biotic stress in plants is large genomic rearrangements, point mutations, and an increased microsatellite instability in Arabidopsis plants after infection with oilseed rape mosaic virus (Yao et al. 2011). Therefore stressful factors, either biotic, abiotic, endogenous, or exogenous, can cause significant adverse effects on plant metabolic and cellular processes. Stressed plants can produce a variety of metabolites that can work as messengers to their neighbors—unstressed plants—and help them activate their systemic defense in advance. Good examples of such metabolites are methyl jasmonate (MeJA), methyl salicylate (MeSA), and salicylic acid (SA) (Shulaev et al. 1995; Ryan and Moura 2002; Kessler et al. 2006; Zavala and Baldwin 2006; Heil and Bueno 2007). 5.2.2  DNA Repair

Biotic and abiotic stresses are largely unavoidable for plants. Thus plants, during their evolution, have developed elaborate mechanisms to adapt to stressful conditions.

5.2  Stress and its Effects on Genomes

These mechanisms promote the activation of checkpoint pathways and result in the initiation of one or several DNA repair mechanisms: base excision repair (BER), nucleotide excision repair (NER), mismatch repair (MMR), homologous recombination (HR), and non‐homologous end joining (NHEJ). Changes in the frequencies of HR and NHEJ are probably the most crucial for the maintenance of genome stability due to the involvement of these two pathways in the repair of single‐ and double‐ strand breaks (SSBs and DSBs). The main difference between NHEJ and HR is the presence of proteins in the HR pathway that interact with a homologous DNA template. On the other side, the NHEJ pathway utilizes regions of microhomology to repair DSBs. Such rejoining of the broken ends may be accompanied by nucleotide deletions and/or insertions of various sizes. This, in turn, can result in chromosomal alterations, including translocations, copy‐number variations, deletions, and so on. Furthermore, if the DNA repair pathways fail to maintain DNA sequence integrity, it can lead to mutations and changes in cell fate, including apoptosis/necrosis, or, preferably, higher resilience to stress. 5.2.3  Epigenetic Changes

Epigenetic changes are enzyme‐mediated reversible changes in the status of epigenetic marks, including DNA methylation and histone post‐translational modifications/ replacements that occur without the alteration of nucleotide sequence. Changes in the levels of expression of the small non‐coding RNA (sncRNA) also can be considered epigenetic in nature (Huettel et al. 2007; Matzke et al. 2007). It has been shown that sncRNAs are responsible for the formation of locus‐specific heterochromatin (Chan 2008). The RNA‐directed DNA methylation (RdDM) pathway is a key mechanism of de novo methylation that silences transposons and therefore can drastically reshape the genome structure and its transcriptome profile. 5.2.3.1  DNA Methylation

DNA methylation in plant genomes occurs in cytosines at both symmetric (CpG and CpNpG) and asymmetric (CpNpN) sites where N can be either A, T, or C. Cytosine methylation is reversible by special enzymes (Table 5.1). For a more comprehensive description of enzymes involved in the DNA methylation/demethylation machinery, see a review by He et al. (2011). RNA‐directed DNA methylation is a de novo DNA methylation mechanism in plants described by Wassenegger et al. (1994). The authors demonstrated de novo DNA methylation of genes driven by a sequence‐specific interaction with siRNAs generated from their own mRNAs. To date, many components of the RdDM pathway have been discovered. In transposons and other DNA repeat regions, these components are: the DNA‐ dependent RNA polymerase IV (Pol IV), the chromatin remodeling protein CLSY, the RNA‐dependent RNA polymerase RDR2, the Dicer‐like protein DCL3, and the ARGONAUTE proteins AGO4, AGO6, or AGO9. In the intergenic non‐coding (IGN) regions, these components are DNA‐dependent RNA polymerase V (Pol V), RDM4/ DMS4, DRD1, DMS3, RDM1, Pol II, KTF1, IDN2, and the de novo DNA methyltransferase DRM2 (He et al. 2011). The components and the proposed mechanism of action of RdDM have been summarized in an excellent review and will not be covered here (Mahfouz 2010). The RdDM pathway is considered as a general transcriptional

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Table 5.1  Enzymes involved in DNA methylation. Context

Enzyme

Function

Reference

CG

MET1 (DNA Methyltransferase 1)

Maintenance of cytosine methylation

(Ronemus et al. 1996)

CNG

DNA methyltransferase CMT3 (Chromomethylase 3), histone methyltransferase KYP (Kryptonite/ SUVH4)

Maintenance of cytosine methylation

(Bartee et al. 2001; Lindroth et al. 2001; Jackson et al. 2002; Malagnac et al. 2002)

CNN

DRM2 (Domains Rearranged Methyltransferase 2)

Maintenance and de novo cytosine methylation

(Cao et al. 2000; Cao and Jacobsen 2002; Johnson et al. 2002)

CG, CNG, CNN

DNA glycosylases of the DEMETER (DME) family, which have both DNA glycosylase and apurinic/apyrimidinic (AP) lyase activities, ROS1 (Repressor Of Silencing 1), DML2, and DML3 (Demeter‐Like 2 and 3)

Cytosine demethylation

(Gong et al. 2002; Gehring et al. 2006; Penterman et al. 2007)

silencing mechanism in plants involved in many epigenetic effects, including transposon suppression, gene imprinting, and transgene silencing (Zaratiegui et al. 2007; Xie and Qi 2008; Law and Jacobsen 2010). Biotic and abiotic stresses modulate global genomic DNA methylation and sequence‐ specific methylation patterns, thus altering the expression of stress‐responsive genes. For example, it has been shown that exposure of Arabidopsis plants to various abiotic stresses (salt, UVC, cold, heat, and flood) results in an increase in global genome methylation and subsequently in the elevated rates of homologous recombination, and genome instability (Boyko et al. 2010a). Also, other studies have revealed DNA hypermethylation upon heavy metal ion stress in various plants, changes in methylation patterns in response to cadmium in oilseed rape and radish, and changes upon exposure to chromium in oilseed rape. Besides, these changes have been correlated with the intensity of stress (Labra et al. 2004; Filek et al. 2008). Interestingly, just like abiotic stresses, biotic stresses also have a profound effect on DNA methylation profiles. Boyko et al. (2007) analyzed genome stability and methylation in Nicotiana tabacum plants infected by tobacco mosaic virus (TMV). They found global genome hypermethylation, hypomethylation in several leucine‐rich repeat (LRR)‐containing loci, substantial hypermethylation of an actin locus, and no changes in methylation in the loci of repetitive elements (5.8S rRNA). The authors hypothesized that such differences between global DNA methylation and local methylation patterns might indicate an involvement of different parts of a protective mechanism against stress. Global genome hypermethylation might be a general protection mechanism against stress; local hypomethylation might be associated with a higher recombination rate and thus could be a sign of an adaptive response.

5.2  Stress and its Effects on Genomes

5.2.3.2  Histone Modifications

There are four nucleosome core histones (H2A, H2B, H3, and H4) that usually undergo various covalent chemical modifications (e.g., phosphorylation, acetylation, methylation, ubiquitylation, and sumoylation) which are flexible in response to stress. These modifications can lead to changes in chromatin structure and thus play an important role in epigenetic events and genome stability. A variety of histone modifications allow a highly specific regulation of gene expression and complex signaling mechanisms. It has been shown that a specific stress can lead to specific histone modifications that might be associated with particular transcriptional effects. For example, gene activation can be associated with H3 methylation at lysine 4 (H3K4) and 36 (H3K36), whereas gene silencing is associated with H3 deacetylation, H3K9 methylation, and H3K27 methylation (Lang‐Mladek et al. 2010; Liu et al. 2010; van Dijk et al. 2010) (Table 5.2). For a comprehensive review on histone modifications, see Luo et al. (2012). 5.2.4  The Link between Genetic and Epigenetic Changes

A considerable body of research shows that biotic and abiotic stresses can destabilize genomes through epigenetic changes (DNA methylation, histone modifications, and sncRNA expression), followed by the variation in the frequency of somatic homologous recombination, point mutation frequency, and microsatellite instability (Lebel et  al. 1993; Kovalchuk et al. 2003; Boyko et al. 2006b; Molinier et al. 2006; Pecinka et al. 2009; Schuermann et  al. 2009; Boyko et  al. 2010b; Kathiria et  al. 2010; Yao and Kovalchuk 2011; Yao et al. 2011). Interestingly, a recent work by Melamed‐Bessudo and Levy (2012) on the effect of DNA methylation on chromosomal recombination demonstrated the repressive role of methylation in meiotic recombination in euchromatic regions. The authors also suggested that some additional factors might have a role in controlling the suppression of recombination in heterochromatin. Epigenetic modifications are also responsible for changes in the mobility of repetitive and transposable elements known as powerful recombinogenic substrates whose activities can have a great impact on rearrangements of genomic structure, the creation of new gene networks, and thus genome stability (Mc 1950; Hedges and Deininger 2007; Zhang et al. 2009; Matsunaga et al. 2012; Staton et al. 2012; Tian et al. 2012). For additional reading on this subject, see an excellent review describing the link between Table 5.2  Stress and histone modifications. Stress

Histone modifications

Reference

Hypoxia

Acetylation of histone H3, conversion of di‐methyl H3K4 to tri‐methyl H3K4

(Tsuji et al. 2006)

High salinity and cold

H3 phosphoacetylation and H4 acetylation

(Sokol et al. 2007)

Drought

Acetylation at H3K23 and H3K27

(Kim et al. 2008)

Abscisic acid and salt stress

H3K9K14 acetylation, H3K4 trimethylation, decreased H3K9 dimethylation

(Chen et al. 2010; Chen and Wu 2010)

95

96

5  Environmentally Induced Genome Instability and its Inheritance Environmental stress (biotic or abiotic) Epigenetic changes (DNA methylation, histone modifications, sncRNA)

Genetic changes (point mutations, insertions, deletions, etc.)

Soft inheritance

Hard or Mendelian inheritance Adaptation and/or speciation

Figure 5.1  A hypothetical scheme connecting environmental stress and epigenetic/genetic changes that promote soft and hard (Mendelian) inheritance.

epigenetics, mobile and repetitive elements, and their impact on genome stability and evolution (Rebollo et al. 2010). Therefore we can draw a hypothetical line that connects an environmental stress, epigenetic changes, gene transcription, and genetic changes (insertions, deletions, translocations, etc.) with a selective force that promotes soft and hard (Mendelian) inheritance (Figure 5.1). The definitions of soft and hard inheritance are given in the following section.

5.3 ­Transgenerational Inheritance As already discussed, epigenetic changes can lead to the increased frequencies of genetic recombination, which in turn can give rise to new gene combinations that might improve plant tolerance to environmental stresses. Interestingly, it has been shown that a stress‐induced increase in the frequency of spontaneous somatic homologous recombination is heritable and can be traced even in the fourth consecutive unstressed generation of Arabidopsis plants (Molinier et al. 2006). The most obvious inheritance was observed in the first unstressed generation of plants, with a steep diminution in the following generations without additional treatments (Chan 2008; Boyko and Kovalchuk 2010; Boyko et al. 2010a; Boyko et al. 2010b; van Dijk et al. 2010; Rahavi et al. 2011; Herman et al. 2012). Several other studies also found that changes in DNA methylation patterns of stressed plants were inheritable, with a tendency to diminish in the consecutive generations, which was similar to the frequency of homologous recombination (Verhoeven et al. 2010; Kou et al. 2011; Ou et al. 2012). All these facts have confirmed an idea of ‘soft inheritance’ developed by Mayr (1980). Nowadays, soft inheritance is often referred to as a reversible form of inheritance of the changed epigenetic traits (DNA methylation and histone modifications) without changes in the DNA sequence (Boyko and Kovalchuk 2011). For those interested in transgenerational epigenetic inheritance, there is a comprehensive review on that subject (Hauser et al. 2011). In contrast to soft inheritance, hard or Mendelian inheritance is based on changes in the DNA sequence (mutations, insertions, deletions, translocations, etc.). Hard inheritance is stable over many generations.

  References

5.4 ­Concluding Remarks Nowadays, this is a matter of fact that environmental stimuli have a tremendous influence on epigenetics, genetics, and plant evolution. There is a considerable body of research that either directly or indirectly links stressful factors, genome instability, and plant tolerance to reversible changes in the epigenome and irreversible changes in the genome. The reversible changes in the epigenome that persist only for one to four (even five) generations might reflect a quick attempt of plants to adapt to harsh environmental conditions. If those conditions persist and keep stimulating epigenetic plant responses, then epigenetic changes may lead to irreversible genetic changes through homologous recombination and point mutations. Obviously, both two proposed types of inheritance, soft and hard, form the distinctive mechanisms of plant tolerance, adaptation, and probably evolution. There are still controversies that need to be resolved in order to improve our understanding of plant epigenetics, genome rearrangements, and inheritance. However, many parts of the inheritance machinery of plants are still unknown: it is still unclear what the physiological mechanisms of transgenerational inheritance are, what the role of sncRNAs across stressed and unstressed generations is, and whether changes in the levels of sncRNAs can be maintained in the non‐stressed progeny. Also, it would be very interesting to reveal how we could apply our knowledge in plant epigenetics and genetics for improving stress tolerance of cultivated plants.

­Acknowledgments I would like to thank Dr Xiu‐Qing Li for providing an exciting opportunity to write this chapter and Valentina Titova for proofreading the text. I am really grateful to Dr Igor Kovalchuk and Dr Andrii Bilichak for critical reading of the manuscript.

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6 The Mitochondrial Genome, Genomic Shifting, and Genomic Conflict Gregory G. Brown* Department of Biology, McGill University, Montreal, Canada

Abstract Mitochondrial DNA (mtDNA) contributes to the genetic differences between somatic cells in a number of ways. In animals, the mtDNA high mutation rate generates heteroplasmy, differences among the mtDNAs within an individual, and these differences may vary among different organs and cell types. In plants, the rate at which point mutations occur in mtDNA is, in general, much lower than in animals, but plant mitochondrial genomes undergo very active recombination and are subject to frequent rearrangement. Rearranged plant mtDNA forms may be maintained at very low levels over many generations and then become the predominant form via a process termed substoichiometric shifting. Mitochondrial genomes are cytoplasmically inherited, and in angiosperm plants and in animals this is reflected in their maternal inheritance, that is, inheritance primarily or exclusively from the female parent. In plants, the potential conflict between two genomes with different modes of inheritance functioning within the same organism is manifested in the related phenomena of cytoplasmic male sterility (CMS) and nuclear fertility restoration. In this genetic arms race, the spread of mitochondrial male sterility genes is countered by the evolution of nuclear genes that suppress the male sterility. The role of substoichiometric shifting in this process is discussed in this chapter. Keywords  mtDNA; heteroplasmy; genome rearrangement; maternal inheritance; genomic conflict; substoichiometric shifting; cytoplasmic male sterility; nuclear fertility restoration

6.1 ­Introduction In 1967, Lynn Margulis (then Lynn Sagan) proposed that the mitochondrion, the double membrane bound organelle that is the seat of cellular energy production, is the descendant of a free‐living prokaryotic ancestor (Sagan 1967). In the years since, molecular and cell biological findings have validated and refined this view, the weight of evidence now *Corresponding author: [email protected] Brown G.G. (2017) The mitochondrial genome, genomic shifting, and genomic conflict. In: Li X.‐Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley‐Blackwell, Hoboken, NJ, Ch. 6, pp. 103–118. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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supporting a ‘symbiogenesis’ scenario in which the origin of mitochondria is ­inextricably linked to the origin of the eukaryotic cell (Koonin 2010; Gray 2012). In this view, a non‐respiring archaeal (archaebacterial) cell formed a close symbiotic relationship with a respiring eubacterium most similar to a present‐day alpha‐proteobacteria, eventually engulfing it as an intracellular symbiont to form a proto‐eukaryotic organism. In the eons that followed, the host evolved a complex intracellular membrane system, possibly as a consequence of the energetic advantage afforded by the symbiont (Lane and Martin 2010), including a nucleus that served as a repository for the increasing load of genetic information associated with this increasing complexity. As the size of the host genome increased, its DNA became organized as multiple linear chromosomes requiring a mitotic apparatus to ensure equal partitioning of genetic information into daughter cells during cell division. Meanwhile, much of the genetic information contained within the eubacterial endosymbiont was lost or transferred to the nucleus (endosymbiont gene transfer (EGT)), and mechanisms for the targeting and translocation of nuclear encoded eubacterial proteins to the symbiont and other sites within the emerging eukaryotic cell evolved. The endosymbiont retained its role of providing respiratory metabolism to the host, and evolved into the mitochondrion of the last eukaryotic common ancestor. As a consequence of EGT, the size of the endosymbiont genome became greatly reduced; in some eukaryotic lineages it was even lost altogether, along with the capacity for respiratory metabolism (Hjort et al. 2010). Nevertheless, the vast majority of eukaryotes have retained respiration and, along with it, a genome derived from the eubacterial genome in the form of modern mitochondrial DNA (mtDNA). This genome specifies RNAs of mitochondrial‐specific ribosomes as well as mitochondrial proteins and tRNAs, the number and types of which vary among different eukaryotic lines of descent (Gray 2012). Importantly, mitochondrial genomes are not subject to the restrictions on ploidy associated with nuclear genes, and their copy number can vary greatly between different cell and tissue types within an individual. For example, there are roughly 7 × 103 mtDNAs per diploid genome in human cardiac muscle but only about half that number in skeletal muscle (Miller et al. 2003). In addition, the mode of inheritance of mtDNA is markedly different. Mitochondrial genes are not allocated into gametes via meiosis, and consequently their copy number, relative to nuclear genes, is higher in larger, female gametes than in male gametes. As a result, mitochondrial genes are usually maternally inherited, that is, inherited primarily or exclusively from the female, in anisogametic organisms such as animals and land plants. Even in isogametic organisms such as yeast, mitochondrial inheritance is generally uniparental, since only one of the two parental mt genotypes is found in a given diploid progeny individual due to the stochastic sorting process of vegetative, or mitotic, segregation that occurs during the divisions following zygote formation (Birky 2001). The mitochondrial genetic systems found in most modern eukaryotes bear little similarity to those of either prokaryotes or the eukaryote nucleus‐cytoplasm. Even among mitochondrial genomes, there are enormous differences in gene organization and expression among different groups of eukaryotes. For example, animal mitochondrial genomes, with only a few exceptions, exist as physical circles of less than 20 kb with the same set of 2 rRNA, 22 tRNA, and 13 protein coding genes arranged in a highly compact format (Boore 1999). These genomes experience a high rate of nucleotide substitution as a result of a high mutation rate (Brown and Simpson 1982; Larsson 2010) and do

6.2  Heteroplasmy and Sublimons

not undergo appreciable recombination. In contrast, the mitochondrial genomes of flowering plants are large, ranging in size from 200 to 2900 kb, and encode 3 rRNAs, 15–21 tRNAs, and roughly 35 proteins, the number and types of which vary, principally due to variation in ribosomal protein gene content (Kubo and Newton 2008). Plant mtDNAs, though they map as circles, appear to exist physically primarily as linear and branched molecules (Bendich 1996; Oldenburg and Bendich 1996), experience a low rate of nucleotide substitutions (with a few notable exceptions, e.g. Palmer et al. 2000; Mower et  al. 2007), and contain introns, repeat sequences which undergo frequent recombination, very significant amounts of non‐coding DNA, and large numbers of non‐conserved open reading frames, some of which are associated with the phenomenon of cytoplasmic male sterility (Kubo and Newton 2008). In contrast to animal mitochondrial genomes where gene order is relatively conserved across groups, plant mitochondrial genomes evolve primarily through rearrangements, and gene order shows little conservation, even among closely related species (Palmer and Herbon 1988; Kubo and Newton 2008). Most mitochondrial gene products function in macromolecular assemblages, either in mitochondrial ribosomes or in the respiratory complexes of the inner membrane, where they interact tightly and specifically with partner subunits encoded in the nucleus. The function of these assemblages is therefore dependent on interactions between partner components encoded by two genomes that undergo different modes of inheritance and may be subject to different rates of mutation and recombination. This mandates that mechanisms exist to ensure the coordinate regulation of components produced in the two compartments and has implications for their modes of evolution, since nuclear gene variation can drive mitochondrial gene variation, and vice versa. Moreover, the different modes of inheritance can pose an evolutionary conflict of interest between the nucleus/host and mitochondrial/symbiont. In the sections that follow, certain aspects of mitochondrial gene variation, as it pertains to variation in the genetic makeup of somatic cells, are discussed. The latter part of this chapter deals with a novel aspect of mitochondrial gene variation in plants related to the evolutionary conflict between nucleus and mitochondrion, the phenomenon of cytoplasmic male sterility and nuclear fertility restoration.

6.2 ­Heteroplasmy and Sublimons Variation in the restriction fragment profiles provided the first evidence for extensive mtDNA polymorphism in animal species (Potter et  al. 1975; Francisco and Simpson 1977; Upholt and Dawid 1977). Importantly, these studies also indicated that only a single mtDNA form was observed within an individual animal, a situation designated as homoplasmy. The occurrence of widespread variation between individuals but not within individuals was presumed to result from the combination of maternal inheritance, with little or no ‘leakage’ of paternal mtDNA into progeny individuals (Francisco et al. 1979; Gyllensten et al. 1985), and the stochastic process of mitotic (or vegetative) segregation, through which mutant mtDNAs would be expected to sort out into pure clones within a relatively small number of cellular generations (Birky 2001). In 1985, Harrison and co‐workers described a clear example of animal mtDNA heteroplasmy, that is, variation within individuals, in this case, in the length of a portion of

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cricket mtDNA (Harrison et al. 1985). Subsequent studies documented relatively widespread heteroplasmy in animal species, primarily in the form of length variation in the D‐loop or control region. Importantly, Holt et al. (1988) described human mitochondrial heteroplasmy involving deletions of tRNA, rRNA, and protein coding regions that occurred in muscle, but not blood, cells of individuals expressing symptoms of mitochondrial myopathy. Numerous examples of point and indel mitochondrial heteroplasmy associated with variable expression of many maternally inherited diseases of humans have since been reported (Wallace 2011; Shapira 2012). Cumulatively, these studies have indicated that: (1) human mtDNA mutations occur frequently—perhaps as high as 103 times that of nuclear genes (Sosa et  al. 2012); (2) through the combined processes of replication and mitotic segregation, these mutant mtDNAs can, on occasion, predominate in certain tissues (Jenuth et al. 1996; Jenuth et al. 1997); (3) females can pass on mutant mtDNAs, usually in a heteroplasmic state, to their progeny – such mitochondrial disorders show a characteristic pattern of maternal inheritance; (4) more severe mitochondrial mutations, large deletions for example, can generate pathological states or phenotypes in tissues when present at a lower level than milder mutations (Wallace 2011). More recently, it has been reported that low‐level heteroplasmy appears to be universal in human mitochondrial DNA (Payne et al. 2013). Thus the mitochondrial genome plays an important part in determining the genetic differences among different somatic cell tissues, and in this respect it is a key element of the somatic genome of animals. Widespread mitochondrial heteroplasmy has also been detected in plant tissues, though in plants this phenomenon is more difficult to define due to the complex mode of plant mtDNA organization, which results in a lack of uniformity in restriction fragment stoichiometry (Palmer and Shields 1984). Plant mitochondrial genomes are conventionally represented as a ‘master‐circular’ form containing a variable number of long (more than 1‐kb) repeated sequences capable of undergoing frequent recombination (recombination repeats); recombination between direct repeats on the master‐circle results in subgenomic circles which can undergo intermolecular recombination to regenerate the master‐ circle (Lonsdale et al. 1984; Palmer and Shields 1984). This circular genome model, based on physical mapping data, can explain differences in stoichiometry observed among fragments generated by restriction endonuclease digestion of plant mtDNA preparations. It is at odds, however, with visual and electrophoretic analysis of plant mitochondrial genomes, which indicate that linear and branched forms predominate (Backert et al. 1996; Bendich 1996; Oldenburg and Bendich 1996). This conflict can be reconciled with a model in which plant mtDNAs exist principally as circularly permuted linear molecules, as reported by Oldenburg and Bendich (2001) for the hornwort Marchantia. Newton and colleagues provided clear evidence for plant mitochondrial heteroplasmy by showing that the striping phenotype of maize non‐chromosomal stripe (NCS) mutants resulted from the mitotic segregation of normal and mutant mtDNAs inherited from the female (Newton and Coe 1986). Small et al. (1987) first suggested that a novel type of heteroplasmy might exist in plant mitochondria through an analysis of sequence arrangements surrounding the mitochondrial atpA gene in different maize cytoplasms. It was found that the predominant arrangement for one cytoplasm could frequently be detected in cytoplasms with structurally distinct mtDNAs, albeit at much

6.2  Heteroplasmy and Sublimons

lower, substoichiometric levels. They termed these substoichiometric arrangements ‘sublimons.’ It was suggested that selective amplification of such sublimons might provide an explanation for several observed dramatic shifts in mitochondrial genome organization, as, for example, the distinct recombinant mtDNAs generated in plant cytoplasmic hybrids (Belliard et  al. 1979). Small et  al. (1989) subsequently explained how the N cytoplasm maize mitotype could have arisen from a progenitor form via stoichiometric shifts resulting from infrequent recombination events across short repeat sequences, followed by intermolecular recombination between substoichiometric forms across longer repeats. The notion that stoichiometric shifting (or substoichiometric shifting) could explain mitochondrial rearrangements observed in plants regenerated from tissue culture received support from Shrizadegan et al. (1989). who showed that segments of rearranged mtDNAs in tissue cultures of Brassica rapa (formerly designated campestris) could be detected in the parental plant lines of the explant source. They suggested that the tissue culture‐induced rearrangements resulted from the amplification of pre‐­existing low‐ abundance forms present in the parental line. Similar results were reported by Vitart et al. (1992) and Hartmann et al. (1994), raising the possibility that other tissue‐culture‐induced rearrangements, such as those that accompany the reversion of cytoplasmic male sterility in maize (Gengenbach et al. 1981), might also be a reflection of stoichiometric shifting. In recent years it has become evident that mtDNA of most plant species encompasses multiple forms of varied stoichiometry and that these forms not only may differ in gene arrangement but also may have minor DNA sequence content differences (Woloszynska 2010). Moreover, the presence of substoichiometric mtDNA forms characteristic of deleted mtDNAs associated with pathological states has been detected in unaffected human tissues (Kajander et al. 2000), suggesting that this type of mitochondrial heteroplasmy may be widespread in animals as well. Thus mitochondrial heteroplasmy due to low‐abundance DNA types that may be maintained over successive sexual generations and that can occasionally become the predominant type in a cell may be regarded as a general aspect of the somatic genome. It is unclear how substoichiometric plant mtDNA forms can be replicated and maintained over multiple generations and not be eliminated or segregated into pure clones through mitotic segregation. One possibility is that some mechanism exists to ensure the transmission of the complete complement of mitochondrial genetic information to daughter cells during mitosis, with sublimon forms drifting to lower abundance levels in the non‐dividing, expanding cells that constitute the bulk of plant tissues. Consistent with this possibility, Arrieta‐Montiel et al. (2001) noted that a bean sublimon associated with cytoplasmic male sterility (discussed in greater detail in the next section) was notably more abundant in root sections that included the meristematic zone than in vegetative sections. Sugui‐Simarro et al. have, in fact, shown that during cell division in the shoot apical meristem, mitochondria fuse to form a cage‐like structure surrounding the nucleus that contains 80% of the total mtDNA, thus providing a structural mechanism that might allow for transmission of low‐abundance mtDNA forms to daughter cells (Segui‐Simarro et al. 2008). Alternatively, low‐abundance forms could be recurrently generated through low‐frequency recombination events (Bellaoui et al. 1998), although this mechanism apparently cannot account for most examples of sublimon maintenance (Woloszynska 2010).

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6.3 ­Cytoplasmic Male Sterility (CMS) in Plants Because of the essential contribution of mitochondrially encoded gene products to respiratory function, in both animals and plants most loss‐of‐function mtDNA mutations have severe phenotypes or are lethal in the homoplasmic state. In flowering plants, however, mtDNA variation is often associated with the phenotype of male sterility—the inability of the plant to produce functional pollen. This common trait occurs in at least 147 different plant species (Laser and Lersten 1972), and is so named because its cytoplasmic, or maternal, mode of inheritance distinguishes it from other forms of male sterility arising from mutations in nuclear genes. Numerous investigations have shown that CMS results from the expression of novel open reading frames (ORFs) in the mitochondrial genome (Hanson and Bentolila 2004). These novel ORFs are unique to each type of CMS, although multiple types, and hence associated ORFs, can exist in a single species. CMS is often phenotypically latent and revealed only through interspecific or wide intraspecific crosses. This latency results from the presence of nuclear genes termed restorers‐of‐fertility (Rf genes), which can specifically down‐regulate expression of the corresponding sterility‐associated mitochondrial ORFs and thereby suppress the CMS phenotype (Hanson and Bentolila 2004; Chase 2007). CMS is of interest to plant breeders because it provides a facile means of producing higher yielding hybrid seed. Since CMS plants are unable to self‐fertilize, when a CMS line is planted alongside a male fertile line, all of the seed that forms on the CMS plants will have a hybrid nuclear genotype and the CMS cytoplasm. If the pollen donor line carries a dominant nuclear restorer gene, the hybrid will also be male fertile, a necessity for crops such as wheat, maize, and canola, where seed is the harvested product (Braun et al. 1992). CMS is also of evolutionary interest, since it is unclear why this apparently disadvantageous trait would be so widespread across the angiosperm division of the plant kingdom. Some insight into this question has come from studies of gynodioecy, the term for naturally occurring plant populations consisting of a mixture of hermaphroditic and female (i.e., male sterile) individuals. Genetic studies generally indicate that gynodioecy reflects a polymorphism of male sterile cytoplasmic genes and nuclear restorer genes (Frank 1989), and theoretical studies indicate that a maternally inherited male sterile mutation can spread in a population of hermaphroditic plants if it offers females a slight selective advantage, such as increased fecundity (Delph et al. 2007). The frequency of females can then rise until pollen becomes limiting, thereby creating the selective pressure for the evolution of a nuclear restorer gene that could specifically suppress the CMS mutation. This could result in a maintained state of gynodioecy, the loss of the CMS mutation, or the fixation of both the CMS mutation and the nuclear restorer gene, as illustrated in Figure 6.1. This last alternative would effectively mask the presence of the CMS mutation and explain why CMS can often be revealed only through wide crosses that allow mating between genotypes with and without the relevant restorer gene. The ongoing recurrence of CMS‐causing mitochondrial mutations followed by the evolutionary appearance of corresponding nuclear restorer genes has been likened to the ‘gene‐for‐gene’ model for plant–pathogen interactions. New virulent forms of plant pathogens arise through the mutation of avirulence (Avr) genes, and the spread of such forms can be countered by the appearance of a new allele at a plant disease resistance

6.4  Mitochondrial Sublimons and CMS 1. Appearance of new CMS mutation

Increase in frequency of females 2.

Gynodioecious populations 4.

3.

Appearance of new Rf gene

Fixation of new CMS/Rf combination

Figure 6.1  Genetic events implicated in the occurrence of gynodioecy in plant populations. In a population of hermaphroditic plants (open symbols), the appearance of a new mitochondrial CMS mutation (1, filled, female symbols) can spread due to its maternal mode of inheritance, resulting in a rise in the proportion of female, male sterile plants (2). This eventually places a limitation on pollen production that creates the selective force to drive a response by the nuclear genome, the evolution of a new fertility restorer gene (3). This suppresses the male sterile trait, and leads to the appearance of hermaphroditic plants with the mitochondrial male sterility gene (filled hermaphroditic symbols). In the case shown here, the new CMS–restorer combination eventually spreads to fixation (4), masking the presence of the CMS mitochondrial genome.

(R) locus that blocks the action of the Avr gene (Flor 1971; Ma et al. 2006). Both paradigms can be viewed as molecular ‘arms races,’ in one case between host and pathogen and in the other between host and the mitochondrial symbiont (Touzet and Budar 2004). What then are the molecular events that mediate this arms race by controlling the appearance of CMS mutations and nuclear restorer genes?

6.4 ­Mitochondrial Sublimons and CMS As mentioned above, plant mitochondrial genomes evolve primarily through sequence rearrangements and experience a relatively low rate of nucleotide substitutions. These rearrangements are thought to result from relatively rare recombination events across short repeated sequences that are dispersed throughout the genome (Palmer and Herbon 1988; Kubo and Newton 2008). In contrast to the very active recombination across longer repeats, recombinations across such short repeats can be viewed as aberrant since they can lead to sectoring mutations and their frequency becomes elevated when null mutations are introduced into any of a number of genes that participate in mtDNA metabolism, including homologs of bacterial recA, mutS, and single strand DNA binding proteins (SSBs) (Abdelnoor et al. 2003; Zaegel et al. 2006; Shedge et al. 2007). An important consequence of elevated levels of aberrant recombination events is mitochondrial genome rearrangement through substoichiometric shifting (SSS).

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One intriguing feature of plant mitochondrial genomes is the presence of a large number of ORFs of unknown function. For example, the Brassica rapa genome contains 44 such ORFs (Chang et  al. 2011). These ORFs are not conserved in more distantly related genomes, lack any known function, and in general do not appear to be expressed. A small subset of these ORFs is expressed, however, and such ORFs have been linked to CMS. Many of these CMS‐associated ORFs are chimeric genes encoding proteins in which a portion of a functional gene is joined in frame to a sequence without a known function. For example, two such proteins associated with CMS systems of Brassica napus consist of the 58 N‐terminal amino acids of the atp8 gene fused to sequences displaying about 50% nucleotide identity with an ORF of unknown function in Arabidopsis mtDNA (Brown 1999; Brown, G.G., unpublished observations). Such sequences clearly arise through low‐frequency recombination events. The frequency with which such events could give rise de novo to expressed, CMS‐associated ORFs is expected to be low, however, perhaps too low to account for the recurrent appearance of CMS and fertility restoration postulated for gynodioecious populations. It is possible that in many cases, the recurrent appearance of CMS mutations results not from the de novo generation of the associated mitochondrial ORF, but from amplification of a sublimon containing the CMS‐associated ORF by SSS. Studies on CMS in the common bean, Phaseolus vulgaris, provided the first evidence for such a possibility. Phaseolus possesses an unusual restorer gene, Fr, that, rather than suppressing CMS by down‐regulating expression of an associated ORF, conditions reversion to fertility by altering mtDNA organization (Mackenzie et al. 1988). The mitochondrial genome of the CMS line maps as three circular forms capable of undergoing inter‐ and intramolecular recombination. In revertant lines, clones containing the CMS‐associated ORF, pvs, were not detected (Janska and Mackenzie 1993). The CMS configuration was found at low, substoichiometric levels in progenitor lines, while revertant lines possess substoichiometric levels of both the progenitor and CMS arrangements, suggesting that the generation of both the CMS and revertant forms took place by SSS (Janska et al. 1998). Since the CMS arrangement is found in a wide variety of wild and domesticated strains of Phaseolus vulgaris, usually at low but occasionally at high copy number, this genomic shifting occurs in natural populations and has taken place since the species originated (Arrieta‐Montiel et al. 2001). It is important to emphasize that SSS is essentially a somatic genetic phenomenon, since it can be induced through tissue culture, in the same manner as somaclonal variation. Protoplast culture‐regenerated plants in Nicotiana sylvestris provided the first evidence that CMS can be generated through mitochondrial substoichiometric mutations from a fertile cytoplasm in plants (Li et al. 1988). It is not clear to what extent SSS can contribute to the appearance or reversion of CMS in plant genera other than Phaseolus, but several observations suggest that this is a widespread phenomenon. High levels of heteroplasmy have been found in plant genera in which gynodioecy is common, such as Silene, where multiple CMS forms may be found in single populations (Stadler and Delph 2002; McCauley et al. 2005; Houliston and Olson 2006). Mutations in the Arabidopsis MSH1 gene, formerly CHM (Martinez‐ Zapater et al. 1992), which encodes a protein with a mutS domain characteristic of a bacterial gene involved in mis‐match repair and suppression of ectopic recombination, lead to mitochondrial genome rearrangements that likely reflect SSS (Abdelnoor et al. 2003). RNAi‐mediated down‐regulation of MSH1 in tomato and tobacco induces

6.5  Restorer Gene Evolution: Somatic Genetic Changes Drive Nuclear Gene Diversity?

mitochondrial rearrangements associated with male sterility, with multiple events leading to the same alterations, again suggesting SSS (Sandhu et al. 2007). Finally, reversion of CMS in sorghum is associated with amplification of a pre‐existing mtDNA sublimon (Feng et al. 2009). In none of the above examples could the appearance or reversion of CMS be associated with SSS of a known CMS‐associated mitochondrial gene arrangement. Recent analysis of Brassica napus mitochondrial genomes has provided evidence, however, for the substoichiometric persistence of naturally occurring CMS‐associated genes. Brassica napus is a good model for molecular analysis of evolutionary events associated with CMS and fertility restoration. Two forms of CMS native to the species, nap and pol, are associated with closely related chimeric genes and the restorers for the two forms map to the same nuclear locus, observations suggesting a relatively recent evolutionary origin (Li et al. 1998; Brown 1999). The orf222 gene associated with nap CMS has been found to be present at substoichiometric levels in plants with the related pol cytoplasm (Chen et al. 2011), and observations in the author’s laboratory indicate that both orf222 and the pol CMS‐associated orf224 gene are present at substoichiometric levels in plants possessing the fertile cam cytoplasm (Gallant and Brown, unpublished observations). Thus sublimons containing CMS genomes are apparently present in all three Brassica napus mitochondrial genomes. Not only do these observations provide strong support for the view that SSS can drive the appearance of CMS in natural populations, but also they suggest that SSS may be able to drive the recurrent appearance of the same CMS multiple times.

6.5 ­Restorer Gene Evolution: Somatic Genetic Changes Drive Nuclear Gene Diversity? As shown in Figure 6.1, the appearance of CMS in a plant population, possibly via SSS, is predicted to provide the evolutionary drive for the appearance of a corresponding nuclear restorer gene. Although the molecular mechanisms through which this process takes place are not known, some possibilities are suggested by the molecular characteristics of known restorer gene loci and the more thoroughly examined analogous phenomena governing the appearance of new resistance genes in response to new pathogen forms. The first restorer gene to be cloned, Rf2 for cms‐T maize, encodes an aldehyde dehydrogenase (Cui et  al. 1996). This gene is an atypical restorer, however, in that it does not affect expression of the associated mitochondrial T‐urf13 gene and also affects the male fertility of plants with normal N cytoplasm (Liu et  al. 2001). More typical restorer genes from petunia (Bentolila et al. 2002), radish (Brown et al. 2003; Desloire et al. 2003; Koizuka et al. 2003), rice (Kazama and Toriyama 2003; Akagi et al. 2004; Komori et al. 2004), and sorghum (Klein et al. 2005), which down‐regulate expression of CMS‐associated genes, have all been found to encode proteins with tandem repeats of a degenerate 35 amino acid motif, the pentatricopeptide repeat (PPR) (Small and Peeters 2000). In all but the sorghum example, the Rf genes are found within clusters of highly related genes with two or more PPR protein‐encoding genes arranged in tandem repeats. The PPR gene family has been greatly expanded in land plants to over 400 members and the proteins all appear to modulate gene expression post‐transcriptionally by binding to specific RNA molecules.

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According to the current view of the gene for gene model for host–pathogen ­interactions, avr genes encode proteins that engage with host cells. In resistant hosts, these Avr proteins encounter products of disease resistance or R genes that specifically bind to them and trigger a cascade that results in death of the infected host cell, restricting the capacity of the pathogen to replicate and invade neighboring cells (Ellis et al. 2000). Like Rf genes, R genes also contain repeat motifs and are often situated within tandem repeats of related genes. Most characterized R genes encode proteins with tandem leucine‐rich repeats (LRRs). These repeats are involved in specific protein binding: the LRR folds into consecutive alpha‐helical, beta‐turn, and beta‐sheet domains, with the beta‐turn/beta‐sheet domains exposed to the solvent where they can associate with partner proteins. Analysis of closely related resistance gene sequences has shown that the rate at which non‐synonymous nucleotide substitutions accumulate (Ka) is higher than the rate of synonymous substitutions (Ks), a hallmark of genes under positive or diversifying selective pressure. This result would be expected if recurrent selection were operating to modulate the binding properties of their products in response to altered pathogen Avr proteins in a genetic arms race. The occurrence of these genes in clusters of related sequences may be a reflection of a birth and death process governing the evolution of multi‐gene families under positive selection (Michelmore and Meyers 1998). The radish Rfo locus is a cluster of three highly related sequences potentially capable of encoding PPR proteins, with the central protein functioning as the restorer gene Rfo (Brown et al. 2003). These three sequences all share more similarity with one another than with any other known sequence, suggesting they evolved from a common ancestor via relatively recent gene duplications. A comparison of the Rfo locus gene sequences revealed a higher degree of divergence at coding than at non‐coding nucleotides, suggesting, as with R genes, that they were subject to positive evolutionary selection (Geddy and Brown 2007) and functioning as participants in a genetic arms race. More recently, Fujii et al. (2011), through a systematic analysis of sequenced plant genomes, showed that restorer genes represent a distinct clade of PPR‐encoding genes, Rf‐PPR‐like or RFL genes, that display the non‐synonymous > synonymous substitution rates characteristic of positive selection. Moreover, it was further shown that positive selection in these genes was most evident at codons encoding residues in the PPR motif that have since been shown to be involved in RNA recognition (Barkan et al. 2012). These observations indicate that not only genes at known Rf loci, but also an entire subfamily of plant PPR genes are undergoing positive selection pressure that most likely reflects participation in an intracellular arms race. In short, somatic genetic changes resulting from mitochondrial genomic shifting can introduce a new CMS form in a population, and drive nuclear gene evolution.

6.6 ­Concluding Remarks Due to copy number differences, mtDNA contributes significantly to the genetic differences among different cell and organ types in multicellular eukaryotes and thus represents a substantial component of the somatic genome. Due to the high rate at which animal mtDNA sustains mutations, there are sequence differences among the mtDNA molecules within an individual (heteroplasmy). As a result of mitotic segregation, some

  References

variants will be more abundant in some cells and tissues than in others, a further contribution to somatic genetic differences. In individuals heteroplasmic for mutations that lead to mitochondrial dysfunction, this can result in dramatic phenotypes, such as the symptoms of mitochondrial neurodegenerative and myopathic diseases. In plants, the novel phenomenon of mitochondrial substoichiometric shifting underlies the appearance of the cytoplasmic male sterile phenotype in plant populations. This then produces an evolutionary selective force that drives nuclear genetic change, resulting in the ‘birth’ of a new restorer gene encoding a PPR protein with an RNA binding specificity that allows repression of the CMS‐associated mitochondrial gene. This intracellular genetic arms race reflects the dynamic tension between the nuclear genome and mitochondrial symbiont and is partially responsible for the expansion of one of the largest gene families in flowering plants.

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Komori T., Ohta S., Murai N., Takakura Y., Kuraya Y., Suzuki S., Hiei Y., Imaseki H., and Nitta N. (2004) Map‐based cloning of a fertility restorer gene, Rf‐1, in rice (Oryza sativa L.). Plant J 37:315–325. Koonin E.V. (2010) The origin and evolution of eukaryotes in the light of phylogenomics. Genome Biol 11:209. Kubo T. and Newton K. (2008) Angiosperm mitochondrial genomes and mutations. Mitochondrion 8:5–14. Lane N. and Martin W. (2010) The energetic of genome complexity. Nature 467:929–934. Larsson N.‐G. (2010) Somatic mitochondrial mutations in mammalian aging. Annu Rev Biochem 79:683–706. Laser K.D. and Lersten N.R. (1972) Anatomy and cytology of microsporogenesis in cytoplasmic male sterile angiosperms. Bot Rev 38:425–454. Li X.‐Q., Chetrit P., Mathieu C., Vedel F., De Paepe R., Remy R., and Ambard‐Bretteville F. (1988) Regeneration of cytoplasmic male sterile protoclones of Nicotiana sylvestris with mitochondrial variations. Curr Genet 13:261–266. Li X.‐Q., Jean M., Landry B.S., and Brown G.G. (1998) Restorer genes for different forms of Brassica cytoplasmic male sterility map to a single nuclear locus that modifies transcripts of several mitochondrial genes. Proc Natl Acad Sci USA 95:10032–10037. Liu F., Cui X., Horner H.T., Weiner H., and Schnable P. (2001) Mitochondrial aldehyde dehydrogenase activity is required for male fertility in maize. Plant Cell 13:1063–1078. Lonsdale D.M., Hodge T.P., and Fauron C.M.‐R. (1984) The physical map and organisation of the mitochondrial genome from the fertile cytoplasm of maize. Nucleic Acids Res 12:9249–9261. Ma W., Dong F.T.D., Stavrinides J., and Guttman D.S. (2006) Type III effector diversification via both pathoadaptation and horizontal transfer in response to a coevolutionary arms race. PLoS Genet 2:e209. Mackenzie S.A., Pring D.R., Basset M.J., and Chase C.D. (1988) Mitochondrial DNA rearrangement associated with fertility restoration and cytoplasmic reversion to fertility in cytoplasmic male sterile Phaseolus vulgaris L. Proc Natl Acad Sci USA 85:2714–2717. Martinez‐Zapater J.M., Gil P., Capel J., and Somerville C.R. (1992) Mutations at the Arabidopsis CHM locus promote rearrangements of the mitochondrial genome. Plant Cell 4:889–899. McCauley D.E., Bailey M.F., Sherman N.A., and Darnell M.Z. (2005) Evidence for paternal transmission and heteroplasmy in the mitochondrial genome of Silene vulgaris, a gynodioecious plant. Heredity 95:50–58. Michelmore R.W. and Meyers B.C. (1998) Clusters of resistance genes in plants evolve by divergent selection and a birth‐and‐death process. Genome Res 8:1113–1130. Miller F.J., Franklin L., Rosenfeldt F.L., Zhang C., Linnane A.W., and Nagley P. (2003) Precise determination of mitochondrial DNA copy number in human skeletal and cardiac muscle by a PCR‐based assay: lack of change of copy number with age. Nucleic Acids Res 31:e61. Mower J.P., Touzet P., Gummow J.S., Delph L.F., and Palmer J.D. (2007) Extensive variation in synonymous substitution rates in mitochondrial genes of seed plants. BMC Evol Biol 7:135. Newton K.J. and Coe E.H. (1986) Mitochondrial DNA changes in abnormal growth (nonchromosomal stripe) mutants of maize. Proc Natl Acad Sci USA 83:7363–7366.

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7 Plastid Genome Stability and Repair Éric Zampini, Sébastien Truche, Étienne Lepage, Samuel Tremblay‐Belzile and Normand Brisson* Department of Biochemistry, Université de Montréal, Montréal, Canada

Abstract Plastids, like mitochondria, result from an ancient endosymbiosis event and contain a distinct genome. Though many plastid genes have since been transferred to the nuclear genome, the small plastid genome still encodes between 90 and 100 genes, which are notably involved in translation, transcription, and energy metabolism in the plastid. The many roles of this organelle, the most familiar being photosynthesis in chloroplasts, make it essential for the development of higher plants. As such, the ability of the plastid to maintain the stability of its genome represents a crucial element of plant life. The physical organization of the genome itself can have an influence on DNA metabolism, with its large inverted repeats acting as templates for recombination. Furthermore, the localization of chloroplast DNA near elements of the electron transport chain increases the importance of DNA repair mechanisms, as reactive oxygen species (ROS) appear as by‐products of photosynthesis. These ROS, along with UV radiation and DNA double‐strand breaks, create a genotoxic stress through their respective ability to oxidize nucleotides, link DNA bases, or rearrange the structure of the genome. To minimize the deleterious effects of these events, different mechanisms present in the nucleus such as homologous recombination exist in plastids. Some less conservative mechanisms based on sequences of microhomology are also found, and sometimes lead to copy‐number variation in certain areas of the plastid genome. While some of these changes can remain silent, others can be linked to phenotypes such as variegation. Keywords  plastid genome; replication; transcription; DNA repair; homologous recombination; photo‐adaptation; reactive oxygen species; UV light; DNA double‐strand breaks; variegation

*Corresponding author: [email protected] Zampini É., Truche S., Lepage É., Tremblay‐Belzile S., and Brisson N. (2017) Plastid genome stability and repair. In: Li X.‐Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley‐Blackwell, Hoboken, NJ, Ch. 7, pp. 119–164. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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7.1 ­Introduction When thinking about plant genomes, one must keep in mind that plants own three distinct genomes sorted in three compartments: the nucleus, mitochondria, and plastids. Plastids are a family of organelles that contain many different entities: from the proplastid, precursor of all types of plastids, to the chloroplast, responsible for photosynthesis in leaves. Less well known, however, are the etioplast, precursor of the chloroplast in the dark, and the amyloplast, for the storage of starch (reviewed in Wise 2006). Plastids are present in all parts of the plant. Proplastids usually reside in meristematic cells, while chromoplasts are found in fruits, flowers, or roots. Chloroplasts are present in leaves, and amyloplasts in roots, stem, leaves, and seeds (reviewed in Wise 2006). As their variety suggests, plastids have multiple roles in plant life, the most familiar being photosynthesis, on which the phototrophic lifestyle of plants depends. However, plastids are not limited to energy production, and they have been shown to be essential for the development of higher plants. Indeed, a bioinformatics study revealed that of 323 embryo‐defective mutants resulting in an arrest in seed development, one‐third of the mutated genes encoded plastid‐targeted proteins (Hsu et al. 2010), revealing a major role in embryogenesis. Moreover, plastids play roles in leaf and root development, gravitropism, and temperature acclimation (reviewed in Inaba and Ito‐Inaba 2010). Mesophyll cells of Arabidopsis thaliana leaves contain around 100 chloroplasts, with some variation between ecotypes (Koniger et  al. 2008). This number is believed to increase with cell size, as shown in Arabidopsis (Pyke and Leech 1992) and spinach leaves (Possingham and Saurer 1969). Plastids arose from the endosymbiosis of ancient photosynthetic bacteria that occurred approximately 1.5 billion years ago. The first to propose this hypothesis was Mereschkowsky in 1905 (English translation in Martin and Kowallik 1999). After the endosymbiosis event, a genome reduction step occurred, leading to the plastid organelle. This genome reduction consists of a massive gene transfer to the host’s nucleus that occurred during the evolution of the plastid. In fact, present plastids of higher plants have a rather small genome carrying few genes with some variation between species, as will be discussed in more detail in this chapter. There exist many theories as to why this transfer occurred. Some suggest that the reactive oxygen species (ROS) produced by the electron transport chain could act as a source of pressure for the displacement of genes to the nucleus, or that a sexual mode of inheritance for the genome favored the transfer (Martin and Herrmann 1998). For now, the question remains unanswered, but the transfer of plastid genes to the nucleus has been shown to occur at a high frequency in selection experiments (Stegemann et al. 2003) and large regions of organellar DNA can be transferred (Martin 2003). This process is thought to occur in three steps: (1) endosymbiont DNA escapes into the host cell by lysis or by a replication/division event and is transferred into the nucleus; (2) the organellar and nuclear copies of genes then coexist, during which time the nuclear copies are subject to random recombination and modification to acquire promoter and targeting sequences; (3) finally, when the nuclear copies become functional with satisfactory expression and targeting to the organelle, the organellar copies can be eliminated (Martin and Herrmann 1998; Dyall et al. 2004). The coexistence step has been observed in some angiosperms (Millen et al. 2001). As a consequence of this transfer, the plastid genome encodes only 5% or less of total plastid proteins (Martin and

7.2  Characteristics of the Plastid Genome

Herrmann 1998), while the rest is nucleus‐encoded and transported into the plastid through the TIC/TOC transmembrane import machinery (reviewed in Soll and Schleiff 2004). Like mitochondria, plastids are semi‐autonomous organelles. Indeed, they can divide independently of the cell by cytokinesis, as bacteria do. Moreover, the plastid genome is maternally inherited in most higher plants (Reboud and Zeyl 1994). Consequently, their genome is considered to propagate asexually. Due to this lack of sexuality and recombination between parental copies, the plastid genome should be subject to Muller’s ratchet and accumulate deleterious mutations over time (Muller 1964). However, since every plastid carries more than one copy of its genome, any deleterious mutation occurring on one gene is masked by the multiple wild‐type copies that coexist in the plastid. The effect of Muller’s ratchet would therefore be amplified by the plastid’s polyploidy (Day and Madesis 2008). Yet, the plastid genome was shown to be subject to lower mutation rates than the nuclear genome of plants (Wolfe et al. 1987). Thus plastids have developed mechanisms to avoid Muller’s ratchet, which implies the existence of plastid repair pathways such as gene conversion between different copies (Khakhlova and Bock 2006). Because plastids harbor their own DNA, they are subject to genome variation, as is the case for the nucleus. In this chapter, we review the mechanisms that occur in plastids and that are currently known to impact genome variation pathways such as repair, recombination, replication, and transcription.

7.2 ­Characteristics of the Plastid Genome 7.2.1  General Composition of the Plastid Genome

When compared to other genomes, the plastid genome is rather unique in its size and structure. In fact, its only close relative in terms of size is the mitochondrial genome, which also originated from an endosymbiotic event. The plastid genome maintains a somewhat constant size within the plant kingdom and is usually represented as two large inverted repeats (IR) that separate two single copy regions, designated as large and small. For higher plants, the total size ranges from 120 kb to 180 kb (reviewed in Green 2011) and the difference is mainly attributed to the number of genes present in the IR (Palmer 1990). With a broader look at the phylogenetic tree, the plastid genome can range from 35 kb in parasites such as Plasmodium falciparum to 521 kb in Chlorophyta (the family of Chlamydomonas reinhardtii). It is interesting to note that this difference in size can be linked to significant divergence between organisms. For instance, the plastid of parasite species has lost the genes involved in photosynthesis (Blanchard and Hicks 1999), while the plastid genome in Chlorophyta is inflated by abundant DNA repeats scattered throughout (Maul et al. 2002). In most organisms, the plastid genome contains between 90 and 100 genes needed for the plastid translation apparatus, the multiple‐subunit RNA polymerase, and most proteins of the electron transport chain (reviewed in Green 2011). Interestingly, gene positions and orientations on this dense genome show a high level of similarity between species. Notably, the majority of ribosomal RNA genes are located in the IR (reviewed in Green 2011). Others, such as the RPO genes encoding one of the RNA polymerases

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of the plastid, are clustered on the genome and are transcribed as polycistronic mRNA (Drechsel and Bock 2011). Unlike the nuclear genome, the plastid genome can be present in very high copy numbers. Not only are there many plastid genome copies within a single plastid, but also this high level of ploidy is exacerbated by the abundance of plastids within a single cell. The copy number per cell varies from several hundred in Arabidopsis (Leutwiler et al. 1984) to more than 10,000 copies in wheat (Triticum aestivum) and barley (Hordeum vulgare). This polyploidy can have various effects on the evolution of the genome. Indeed, it is possible to observe distinct populations of plastid genomes within individual plants, a phenomenon called heteroplasmy. This can lead to a slow drift from one form of the genome to another, depending on the nature of the mutation. If the mutation leads to increased fitness, the mutated copy can be selected and preferentially transmitted. If the mutation is deleterious, it can be discarded through gene conversion, a process through which a genome copy is transformed from one particular sequence to another (Khakhlova and Bock 2006). Therefore gene conversion can serve as a DNA repair mechanism by removing mutated copies of the genome. In diploid cells, this generally leads to a loss of heterozygosity by transforming the sequence of one sister chromatid to that of the other. In polyploid cells, however, this mechanism usually leads to the conservation of the original sequence, due to the overabundance of one form of the genome (Khakhlova and Bock 2006). Furthermore, in Arabidopsis, there exists a bias toward GC to AT genome conversion rather than AT to GC, which could explain the higher abundance of AT base pairs in the plastid genome (Khakhlova and Bock 2006). Conversely, more variation is observed between the plastid genomes of different organisms at the sequence level. The GC content is generally low and can vary from about 30% to 55% in plants (Shimada and Sugiura 1991; Tsuji et al. 2007; Smith 2009), but can be as small as 19.5% in parasites. The sequence divergence varies a lot from gene to gene and in non‐coding sequences. The genes involved in photosynthesis show an extremely low rate of non‐synonymous/synonymous mutations, suggesting a strong selective pressure on these genes, while other plastid genes show a higher, more variable rate (Magee et al. 2010). The most divergent region is clustered around the RbcL and Ycf4 genes. It is speculated that this region might be a fragile site, leading to a higher frequency of DNA breakage and repair (Ogihara et al. 1992; Magee et al. 2010). These successive rounds of repair would lead to increased substitutions, deletions, and polymorphisms. DNA repeats constitute an important proportion of the plastid genome, mostly because of the two long, perfectly homologous IR sequences. Despite the intensive characterization of these perfect repeats, little is known about their biological relevance. In rare cases, plastid genomes do not include such IR, suggesting that they are not essential. Nevertheless, at the sequence level, the IR are more stable than the single‐ copy regions (Yamane et al. 2006), probably due to more efficient repair in the repeated sections (Wolfe et  al. 1987). Also, many short repeats up to 60 base pairs exist, and could be involved in rapid evolution of the plastid genome (Kawata et  al. 1997). As mentioned earlier, the abundance of repeats varies from species to species and is particularly high in Chlorophyta. Interestingly, there is evidence that these repeats might be involved in the repair of DNA double‐strand breaks (DSBs) in severe stress conditions, such as treatments with mutagenic agents (Cappadocia et  al. 2010; Kwon et al. 2010).

7.2  Characteristics of the Plastid Genome

The study of plastid genomes also provides information that has been widely used to trace the phylogenetic tree for the plant kingdom (Martin et al. 2005). The number of genes and their orientation within the genome are the most useful hints to relate the different species. More recently, results have shown the necessity of having larger and more complex datasets to confirm the phylogeny of species (Wodniok et al. 2011). In that situation, plastid genomes should be linked to information from the nuclear genome to get a more robust look at the evolution of different species. As mentioned earlier, most plastid proteins are encoded in the nucleus, and an important proportion of the plastid genome, which can reach up to 99%, is duplicated in the nuclear genome (Timmis et al. 2004; Kumar and Bendich 2011). The fact that plastid genome sequences can be found in the nucleus further supports the importance of genetic transfer between the two cellular compartments. The function of these duplicated DNA fragments and of the proteins involved in this transfer is still largely unknown, but recent evidence suggests that it would be an ongoing process (Wang et al. 2012). 7.2.2  The Structure of the Plastid Genome

For many years, it has been considered common knowledge that the plastid genome is a circular molecule, similar to what has been observed in prokaryotes and mitochondria. However, this hypothesis has now been re‐evaluated, with some electronic microscopy and pulse field gel electrophoresis results showing that most plastid DNA molecules are linear (reviewed in Bendich 2004). This has also been observed in some prokaryotes (Cui et al. 2007), in yeast mitochondria (Valach et al. 2011), and plant mitochondria (reviewed in Bendich 2004). It is now hypothesized that plastid genomes would be present as a mixture of circular, linear, and branched molecules. Nevertheless, the fraction of each form and their contribution in DNA replication and transcription remain elusive. A study showed that ciprofloxacin, a gyrase inhibitor that specifically affects the organelle genomes of plants, has an important effect on plastid genome structure (Rowan et  al. 2010). Indeed, this group reported that the complex branched structure is less abundant in the stressed plants. A similar reduction of the branched structure was also observed in cpreca mutant plants (Rowan et al. 2010). Taken together, these results link the structure of the plastid genome with the DNA metabolism activity of the plastid, insofar as recombinases and gyrases are concerned. However, some questions remain to be clarified to support a linear plastid genome model: What mechanism is used for DNA replication? Where are the extremities of these molecules? How are the extremities replicated? To ensure optimal efficiency of DNA metabolism, living organisms organize their genome in a highly structured nucleoproteic complex, hence the compaction of nucleus DNA in nucleosomes composed mainly of histones and DNA (Olins and Olins 1974). Although prokaryotic genomes are less structured, they are nevertheless compacted by different DNA‐binding proteins (Luijsterburg et al. 2006). In plastids, such a DNA/protein complex, named nucleoid, exists and has been widely characterized. In 1976, Hallick et  al. were the first to describe what they called the plastid transcriptionally active chromosome (pTAC) (Hallick et al. 1976). This complex was proposed to be composed of DNA, RNA, and proteins involved in transcription—similar to what is observed

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in prokaryotes—and the proteins identified by this procedure were called the pTAC proteins (numbered from 1 to 18). More recently, experiments have led to the purification of new proteins composing the pTAC. So far, 35 proteins of A. thaliana and Sinapis alba have been identified from this complex (Pfalz et al. 2006). Among them are subunits of DNA and RNA polymerases, gyrases, and superoxide dismutases (SODs), as well as the original pTAC proteins. Interestingly, a different group identified 11 proteins that were all part of the pTAC by immunoprecipitation of the plastid‐encoded RNA polymerase (PEP) active protein complex (Suzuki et  al. 2004). This finding supports the hypothesis that the pTAC contains all the components of the PEP in addition to other factors involved in transcription. Improvements in mass spectrometry and protein quantification allowed the identification of proteins in nucleoid‐enriched fractions (Olinares et al. 2010; Schroter et al. 2010; Majeran et  al. 2012). These studies strongly suggest that transcription, mRNA processing, mRNA editing, and ribosome assembly take place in the nucleoid. More specifically, Majeran et al. identified 33 proteins involved in DNA replication, repair, or organization of the nucleoid. These proteins were among the most enriched within the nucleoid, which suggests their high importance in this nucleoprotein complex. Notably, DNA gyrases A and B, the main DNA polymerase POLIA, some endonucleases, single‐ stranded DNA‐binding proteins (OSB and WHY1), MUTS, and RECA homologs were found. In addition, 131 proteins of RNA metabolism and 52 proteins without a known function in DNA/RNA metabolism were enriched in the nucleoid. Of these uncharacterized proteins, many had a pentatricopeptide repeat, a tetratricopeptide repeat, a DnaJ motif, or a mitochondrial terminal elongation factor domain. More recently, the SWIB family of proteins was also proposed to be involved in plastid genome compaction (Melonek et al. 2012). Taken together, these results suggest that the nucleoid is a highly active super‐complex, similar to the prokaryote chromosome. Within the chloroplast, the nucleoid is usually bound to the thylakoid in the stromal section, but its location nevertheless varies between species and during development. Indeed, during the transition from proplastid to chloroplast, the nucleoid has been demonstrated to relocalize from the envelope to the thylakoid in some species such as Nicotiana tabacum and Spinacia oleracea (Sellden and Leech 1981) but not in others (e.g., Triticum aestivum (Sellden and Leech 1981; Miyamura et al. 1986)). Despite the fact that the proteins that anchor the nucleoid to the thylakoid have not yet been clearly identified, the subcellular localization of MFP1 and TCP34 in the plastid would argue for such a role (Jeong et al. 2003; Weber et al. 2006). Being located near the thylakoid puts the plastid genome in close proximity with the electron transport chain, which means it is more exposed to ROS. This could justify the presence of SODs at the vicinity of the genome to scavenge these mutagenic molecules and allow genome stability. This is discussed in further detail in Section 7.6 on plastid genome stability and DNA repair.

7.3 ­Replication of Plastid DNA 7.3.1  Plastid DNA Content during Development

As the plant is growing, its plastid DNA is replicated and the quantity present in the plant cell is subject to change. The plastid genome copy number varies within a wide

7.3  Replication of Plastid DNA

range, and estimations differ even within a single species. In Arabidopsis, according to separate studies, each cell has been shown to contain between 1000 and 1500 plastid DNA copies (Zoschke et  al. 2007) or between 400 and 1200 copies (Leutwiler et  al. 1984), allowing for a consensus value of approximately 1000 (Zoschke et al. 2007). This represents about 25% by mass of the total cellular DNA (Draper and Hays 2000). However, this number varies in other species. For instance, almost 10,000 plastid DNA copies per cell have been reported in pea (Lamppa and Bendich 1979) and in barley (Baumgartner et  al. 1989). This number seems to be tissue dependent, since it decreases to almost 500 plastid DNA copies per cell in non‐green tissue (Lamppa and Bendich 1979). Presently, there is a discrepancy among studies regarding the cellular plastid DNA content during leaf development. Indeed, several studies have shown that the plastid genome copy number remains constant as the plant grows, whereas others reported a decline in chloroplast DNA content very early in the development. For instance, Rowan et al. stated a two‐ to seven‐fold reduction in plastid DNA content from immature to mature leaves using DAPI fluorescence on isolated chloroplasts and qPCR analysis (Rowan et al. 2004; Rowan et al. 2009). In some chloroplasts, plastid DNA was not even detectable (Rowan and Bendich 2009). According to the authors, the plastid DNA degradation occurring in mature leaves undergoing photosynthesis would be the cellular response to the increase in ROS‐mediated plastid DNA damage (Kumar and Bendich 2011). These results remain controversial, as another study showed, through qPCR, that the plastid genome copy number per cell remains remarkably constant during leaf expansion and early senescence (Zoschke et al. 2007). The authors of the latter study state that technical artifacts could explain the controversial results of Bendich et  al. (Rowan and Bendich 2009). Indeed, several other studies, using DAPI fluorescence (Rauwolf et al. 2010), plastid DNA gel blot analysis (Li et al. 2006), or qPCR analysis (Draper and Hays 2000) do not support the finding of plastid DNA degradation during plant development. A more recent study reveals that in maize, even the apical leaf tissue retains roughly 1000 copies of the plastid genome per nuclear genome complement (Udy et al. 2012). 7.3.2  Plastid DNA Replication Machinery

While many studies attempt to elucidate the replication mechanisms and to identify replication origins of the plastid nucleoid, little attention has been brought to the machinery involved. Two DNA polymerases involved in plastid DNA replication were identified, purified, and characterized: polymerase I‐like A (POLIA) and B (POLIB) (Uchiyama et al. 2002; Ono et al. 2007). The two polymerases appear to be targeted to both plastids and mitochondria in Arabidopsis and Nicotiana (Mori et al. 2005; Ono et al. 2007; Parent et al. 2011). In Arabidopsis, the PolIA and PolIB genes respectively encode 117‐ and 115‐kDa proteins that share a similar conformation with the bacterial DNA polymerase I (Mori et al. 2005). In vitro experiments suggest that these proteins are sufficiently processive to allow the replication of the entire plastid genome. Furthermore, in vivo characterizations suggest a redundant role for the two POLI‐like proteins in replicating the plastid genome, as both single knock‐out plants have the same decrease in plastid DNA content per cell compared to the wild‐type. Furthermore, a double knock‐out plant does not appear to be viable, supporting their essential role in

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DNA replication (Parent et al. 2011). Despite this shared function, POLIB seems to have evolved a specific role in the repair of plastid DSBs in Arabidopsis (Parent et al. 2011). The authors state that POLIB could be recruited to DSBs in order to mediate its repair through homologous recombination (HR), a process that includes a DNA polymerization step (Parent et al. 2011). In addition, the maize co‐ortholog of POLIA and POLIB, WHITE2, has recently been implicated in the replication of the maize chloroplast genome (Udy et al. 2012). Several other plastid‐targeted proteins have been linked to plastid DNA replication in higher plants. First, a DNA primase has been identified in pea, and in vitro analyses link this protein to the initiation of replication (Nielsen et al. 1991). A DNA helicase involved in the unwinding of double‐stranded DNA during replication has also been identified in pea (Tuteja and Phan 1998). Additionally, the PEND protein (Plastid ENvelope DNA‐ binding) in pea would allow the recruitment of the plastid nucleoid to the plastid envelope during the first stage of chloroplast development in order to facilitate the replication process (Sato et al. 1998). A 43‐kDa protein interacting with the pea POLI‐like protein has also been identified and shown to stimulate DNA polymerase activity (Chen et al. 1996b; Gaikwad et  al. 1999). More recently, a 28‐kDa protein similar to the single‐ stranded DNA‐binding protein (SSB) of Escherichia coli has been identified in soybean as a plastid DNA replication origin binding protein (Lassen et al. 2011). Finally, the two plastid DNA gyrases, GYRA and GYRB (similar to the nucleus topoisomerase II), appear to play diverse roles that contribute to the processivity of the replication machinery. Not only would they relax the plastid DNA supercoils ahead, but also they would resolve the partitioning of the two newly created plastid DNA genomes after replication, just before the onset of plastid division (Cho et al. 2004) (Figure 7.1G). Further studies will be needed to investigate the binding capacity of the two Arabidopsis POLI‐ like proteins to plastid DNA replication origins and their putative interaction with the different previously mentioned proteins. This would in turn contribute to the elucidation of the structure and function of the plastid DNA replication machinery. 7.3.3  Replication Mechanisms

Presently, three plastid DNA replication models have been proposed: (1) the D‐loop model, (2) the rolling circle model, and (3) the recombination‐mediated replication model. The first two were proposed by Kolodner and Tewari in 1975 (Kolodner and Tewari 1975a). Using electron microscopy, they observed the topology of the plastid DNA during replication and detected the presence of D‐loops (displacement loops) and extended D‐loops (Kolodner and Tewari 1975a). They proposed that these loops were replication initiation sites where replication begins unidirectionally, replicating only one of the two parental strands. They found two D‐loop regions on the two strands, expanding in opposite directions from each other (Kolodner and Tewari 1975a) (Figure  7.1A and B). This replication model goes through a Cairn‐type intermediate during which the two expanding strands pass each other, leading to a bidirectional replication bubble (θ structure) (Figure 7.1C and D). The replication continues on the leading strand and starts discontinuously on the lagging strand as the other origin of replication is passed (Figure 7.1D and E). These two leading strands encounter each other at almost 180° of the origin of the D‐loop (Figure 7.1F). At this point, the two daughter genomes are separated and the replication of each one is terminated by filling

7.3  Replication of Plastid DNA (a)

(b)

(c)

(d)

(e)

(f)

(g)

Figure 7.1  The D‐loop plastid DNA replication model. In (a) to (f ), black lines represent the two parental strands. Red and blue lines represent the two daughter leading strands and lagging strands, respectively. In (a) to (g), arrows represent direction of DNA polymerization (3′ end). (See plate section for color representation of this figure.)

the remaining gap (Kolodner and Tewari 1975a) (Figure 7.1G). Some evidences favor the hypothesis of a unique replication origin as it has been observed in the unicellular Euglena gracilis (Ravel‐Chapuis et al. 1984; Hedrick et al. 1993). Contrary to the D‐loop model, replication arising from this unique origin would expand bidirectionally. Rolling circle‐mediated replication was proposed later by the same authors as the D‐loop model as a DNA replication mechanism that derives from D‐loop‐mediated replication (Kolodner and Tewari 1975b). After one round of replication, just after the separation of the two daughter genomes (Figure 7.1G), the 3′ end of the newly synthesized strand encounters its 5′ end (lagging strand), creating a gap (Figure 7.2A). The 5′ end is then displaced by the replication fork (Figure 7.2B and C), leading to continuous polymerization on the circular genome and discontinuous polymerization on the tail of the rolling circle (Kolodner and Tewari 1975b) (Figure 7.2D). This mode of replication allows the creation of head‐to‐tail tandem repeats of the plastid genome in one round of replication (Kolodner and Tewari 1975b). Rolling circles have also been observed in dinoflagellates as the mode of replication for minicircles (Leung and Wong 2009). Since

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(b)

(c)

(d)

Figure 7.2  Rolling circle plastid DNA replication. Plain lines represent parental strands. Dotted and gray lines represent leading and lagging strands, respectively. Arrows represent direction of DNA polymerization (3′ end).

its proposal, this model seems to have been largely accepted (Hedrick et al. 1993; Day and Madesis 2008) and is also supported by a bioinformatics study (Krishnan and Rao 2009). Even though Kolodner and Tewari envisioned this mechanism as consecutive to a round of D‐loop‐mediated replication (Figures 7.1G and 7.2A), evidence showing that the two could be independent has been brought by Williamson et  al. (2002). In this study, the authors showed that novobiocin (a type‐II topoisomerase inhibitor) is able to decrease D‐loop‐mediated replication of the plastid genome of Plasmodium falciparum. Rolling circle‐mediated replication was also affected but to a lesser extent, supporting the idea that the latter could arise independently of D‐loop‐mediated replication (Williamson et al. 2002). These first two models were challenged by the discovery of multiple topologies of the plastid genome (Oldenburg and Bendich 2004) and the targeted inactivation of replication origins (Scharff and Koop 2007). Indeed, Scharff and Koop found that the inactivation of three of the four putative origins of replication in the tobacco plastid genome do not disable its capacity for replication (Scharff and Koop 2007). This result is incompatible with both D‐loop and rolling circle models, suggesting that another mode of replication could coexist (Scharff and Koop 2007). Oldenburg and Bendich have also estimated that only approximately 3–4% of the maize plastid genome is found in a circular form, while 50% exists in branched and complex forms. They found that the ends of these linear molecules mapped near the previously suggested origins of replication. They therefore suggested that these ends would invade a double‐stranded template and create a D‐loop, initiating the recombination‐mediated replication of the plastid genome (Oldenburg and Bendich 2004). Oldenburg and Bendich suggest two different DNA replication models: first, a rolling circle model in which a template switching event occurs, leading to a head‐to‐tail concatemer linear molecule where the long single

7.3  Replication of Plastid DNA

copy region would be repeated in opposite orientations. They also propose a model in which replication is initiated through the invasion of a circular or linear template by the extremity of a molecule, leading to linear multimers with the long single copy region repeated in the same or opposite orientations (Oldenburg and Bendich 2004). These recombination events involve a Holliday junction that needs to be resolved in order to separate the two daughter genomes and re‐establish a replication fork. If this resolution step does not occur correctly, the two linear molecules would remain in a branched‐ form. Consequently, as suggested by Day and Madesis, the complexity of the genome would increase, as observed in the maize plastid genome (Oldenburg and Bendich 2004; Day and Madesis 2008). This mechanism has also been proposed in the chloroplast of C. reinhardtii as a way to bypass the inhibition of the D‐loop‐mediated replication by the gyrase inhibitor novobiocin (Woelfle et al. 1993) and as a model for the replication of minicircles in the plastid of Adenoides eludens (Nelson and Green 2005). Finally, the ‘double rolling circle’ observed in the circular 2‐µm yeast plasmid model could also occur in plastids. If a recombination event occurs between the two IR in the θ structure of D‐loop‐mediated replication, it gives rise to a large circular multimer. A second recombination event would then allow the resolution of this structure (Futcher 1986; Kornberg and Baker 1992). Nevertheless, this mode of replication has never been observed for the plastid genome. 7.3.4  Origins of Replication

The mapping of origins of replication has been widely investigated, but many discrepancies exist regarding their location and their requirement. The location of origins of replication in the plastid genome varies from one organism to another, but also within an organism in a tissue‐dependent manner. In pea, two plastid DNA replication origins have been mapped by Meeker et al. in 1988 (Meeker et al. 1988). They found that the first origin, named OriA, was located between the 16S and 23S rRNA genes, and that the second, named OriB, was found after the 23S rRNA gene (Meeker et  al. 1988; Nielsen et al. 1993). The presence of these two origins was later confirmed in tobacco by the Nielsen group. They mapped OriA between the 16S and 23S rRNA genes in one of the two IR (Lu et al. 1996; Kunnimalaiyaan and Nielsen 1997) and OriB upstream of the 23S rRNA gene in the same IR (Lu et al. 1996; Kunnimalaiyaan and Nielsen 1997; Kunnimalaiyaan et al. 1997). Because of their presence in the IR, it suggests that these origins of replication are each present twice in the tobacco plastid genome. Whereas these studies argue in favor of the conservation of replication origins in the plastid genome of higher plants, others provide seemingly contradictory conclusions. Indeed, in proplastids from tobacco‐cultured cells, Takeda et al. found only one replication origin situated very close to the 23S rRNA gene in the IR (Takeda et al. 1992). Also, in Oenothera hookeri, the two origins of replication were found flanking the 16S rRNA gene, suggesting a variation among plant species (Chiu and Sears 1992; Sears et  al. 1996). In Oryza sativa, it has been shown that replication origins differ in location and in number depending on the tissue (Wang et al. 2003). Despite the fact that the deletion of OriA does not seem to affect the replication of the plastid genome, whether or not in vivo replication is possible through another mechanism remains to be determined (Muhlbauer et al. 2002). Indeed, linear molecules with defined ends near OriA have been found in tobacco (Scharff and Koop 2006). The

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possibility that these molecules could be responsible for recombination‐mediated replication even after the targeted inactivation of the OriA sequences must be investigated. As in rice, an organism can use different modes of replication or different origins of replication depending on the tissue (Wang et al. 2003), increasing the difficulty of determining whether a replication origin has a relevant function in vivo. Whereas no strong conclusion can be drawn regarding the in vivo role of OriA, it has been suggested that the presence of one of the two copies of OriB is indispensable for the replication of plastid DNA (Muhlbauer et al. 2002). However, Scharff and Koop suggested that this might not be the case, since the plastid genome can still be replicated when one OriB is deleted and the other mutated (Scharff and Koop 2007). They have also shown that if three of the four origins were deleted, the genome still replicates, suggesting that the two D‐loops model of replication is not sufficient to explain the replication of the plastid DNA (Scharff and Koop 2007). Finally, origins of replication have also been mapped outside of the IR in Glycine max (Saski et al. 2005), Zea mays (Carrillo and Bogorad 1988), and N. tabacum (Ohtani et al. 1984). The discrepancies regarding the location, numbers, and usage of plastid genome origins of replication might indicate a species‐ or tissue‐dependent use of these origins. Also, different modes of replication might coexist in the same organism, as shown in rice (Wang et al. 2003), increasing the difficulty of mapping origins of replication and testing whether or not they are necessary in vivo. 7.3.5  Nucleus and Plastid Coordination during DNA Replication

For many years, nuclear DNA replication (NDR) and plastid DNA replication, or more widely organellar DNA replication (ODR), were thought to be independent processes. Kobayashi et al. however, demonstrated that NDR was dependent on the onset of ODR in the unicellular algae Chlamydomonas merolae. Using ODR‐ or NDR‐specific inhibitors such as nalidixic acid or aphidicolin, they showed that NDR is unable to take place if ODR does not (Kobayashi et al. 2009). They demonstrated that this coordination is dependent on tetrapyrrole Mg‐ProtoIX, a precursor of heme and chlorophyll also important in plastid‐to‐nucleus signaling (Kobayashi et  al. 2009). More recently, the same laboratory observed NDR activation after the completion of ODR, demonstrating the implication of the ubiquitin E3 ligase subunit FBX3. This protein frees CYCLIN1, preventing its degradation and leading to NDR activation (Kobayashi et al. 2011). The mechanism leading to Mg‐ProtoIX signaling after ODR completion and the question of whether a similar NDR and ODR coordination exists in higher plants remain to be determined.

7.4 ­Transcription in the Plastid Transcription in plastids is a process that involves two types of plastid RNA polymerases: the nuclear‐encoded polymerase (NEP) and the plastid‐encoded polymerase (PEP). Two NEPs are found in Arabidopsis plastids: RPOTP and RPOTPM. RPOTP is targeted only to plastids, whereas RPOTPM is targeted to both plastids and mitochondria. The PEP, composed of four subunits, is encoded partly in an operon‐like sequence that includes RPOB, RPOC1, and RPOC2, and partly in another gene encoding RPOA.

7.5  The Influence of Replication and Transcription on Plastid Genome Stability

The PEP is found in association with six different sigma factors (named SIG1–6 or SIGA–F), which share some similarity with the σ70 factor found in bacteria (Hakimi et  al. 2000). Each of these plastid sigma factors directs the PEP to specific genes to activate their transcription. Among these factors, SIG2 and SIG6 seem to mediate a plastid retrograde signal coordinating nuclear and plastid gene expression since a single knock‐out mutant of these genes leads to nuclear gene expression reprogramming (Woodson et al. 2012). Finally, SIG5 has been implicated in the regulation of the circadian rhythm of plastid gene transcription (Noordally et al. 2013). Because there are three different RNA polymerases found in plastids, transcription must be tightly regulated and, accordingly, each RNA polymerase is thought to use different promoters to initiate transcription. Chloroplast genes are therefore divided into three classes: class I genes, transcribed exclusively by the PEP; class II genes, transcribed by the PEP and the NEP; class III genes, transcribed by the NEP (Schweer et al. 2006). Despite this classification, it has been shown that almost all the chloroplast genes have NEP‐ and PEP‐type promoters (Zhelyazkova et al. 2012). Indeed, a PEP‐deficient barley chloroplast is able to transcribe all its genes (Zhelyazkova et al. 2012) and an Arabidopsis sigma mutant, lacking PEP function for some genes, can rescue the transcription by using upstream SOS promoters for the NEP (Schweer et  al. 2006). Furthermore, a switch in RNA polymerase during plant development is thought to occur. As the plastids develop, the NEP would be the first RNA polymerase present, transcribing the chloroplast transcriptional and translational machinery, including the PEP. Later in the development, the PEP would play its main role of transcribing the photosynthesis‐ related genes (Hanaoka et al. 2005). The pTAC is also thought to play a role in the PEP‐dependent plastid transcription (Pfalz et al. 2006). Among the 35 proteins found in this complex in Arabidopsis, many were shown to play a role in replication, ROS detoxification, transcription, translation, or even repair pathways. Many of the mutants for these proteins display an albino or pale‐green phenotype, suggesting a role in plastid DNA transcription (Pfalz et al. 2006; Gao et al. 2011). Further investigation will be needed to elucidate the role of these proteins in the plastid transcription process and to study their interaction with the PEP‐ type RNA polymerase.

7.5 ­The Influence of Replication and Transcription on Plastid Genome Stability Some evidences show that replication and transcription are not independent from genome stability. Replication can have a major impact on genomic variation as it has the deleterious potential to introduce mutations in the daughter genome. Replication can also be responsible for DNA structure instability, since secondary structures, proteins, or processes involving DNA represent obstacles for the movement of the replication fork through the genome. Transcription, DNA‐binding proteins, and non‐conventional DNA structures such as triplex DNA molecules and DNA–RNA hybrids have all been  shown to cause stalling of the replication fork during replication (Burhans and Weinberger 2007; Mirkin and Mirkin 2007). A possible fate of this stalled replication fork is fork reversal, during which an intermediate named ‘chickenfoot’ is created (Mirkin and Mirkin 2007; Kim et  al. 2012b). This structure can be resolved by

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recombination‐mediated replication fork re‐establishment (Mirkin and Mirkin 2007) or by Holliday‐junction‐type resolving followed by a restart of DNA replication (Mirkin and Mirkin 2007; Rudolph et  al. 2007). If this replication fork stalling is caused by a lesion in the DNA template such as a nucleotide dimer, a translesion‐type polymerase can be recruited in order to restart DNA replication (Mirkin and Mirkin 2007). Finally, the stalled replication fork can collapse, resulting in DNA DSB and recombination‐ mediated replication fork re‐establishment (Burhans and Weinberger 2007). Among those processes, the DNA DSB, and the recombination‐mediated re‐establishment of the replication fork are prone to introduce genomic variation, such as DNA rearrangements, if the processing of DSBs uses semi‐conservative repair pathways or if the template used in recombination is not perfectly homologous during the re‐establishment of DNA replication. Aside from this deleterious effect on genome stability, replication is also implicated in the maintenance of genome stability. Conservative repair by break‐induced replication consists in the establishment of a replication fork through homologous recombination in order to repair a DSB (Malkova et al. 1996). Therefore depletion in a component of the replication machinery would have a deleterious impact on genomic stability. Indeed, an Arabidopsis knock‐out mutant for the plastid DNA polymerase POLIB has been shown to accumulate unprocessed DSBs, suggesting a direct role for this polymerase in plastid DNA repair (Parent et al. 2011). It is therefore hypothesized that this DNA polymerase plays a role in break‐induced replication, its absence leading to semi‐­conservative repair of DSBs. Transcription can also be responsible for genomic instability. During transcription, the newly synthesized RNA is dissociated from the DNA template, leading to a short transcription bubble. In some cases, however, this RNA re‐anneals with the DNA, leading to an extended transcription bubble, called an R‐loop (Li and Manley 2006; Kim and Jinks‐Robertson 2012). One of the major threats to genome stability posed by R‐loops is the concomitant formation of a long region of single‐stranded DNA (ssDNA). This ssDNA can be more susceptible to nucleotide modification by ROS, strand‐breaks, or as a template for non‐homologous recombination (Li and Manley 2006; Kim and Jinks‐ Robertson 2012). Indeed, yeast mutants showing more numerous R‐loops than wild‐ type have been shown to accumulate transcription‐associated recombination and loss of chromosomes (Gonzalez‐Aguilera et al. 2008). In plastids, because of the coupling of transcription and translation, ribosomes binding on nascent mRNA can be a way to prevent the appearance of R‐loops, as seen in bacteria (Li and Manley 2006). Finally, a conflict between transcription and replication has been shown to occur in bacteria and yeast. Indeed, RNA polymerase and DNA polymerase can encounter each other while interacting with DNA. If the RNA polymerase is transcribing the leading strand of the replication fork, they are co‐directional, and if it is the lagging strand that is transcribed by the RNA polymerase, they are convergent (Bermejo et al. 2012). In these two cases, there is a risk of collision between the two structures as the replication machinery has been shown to process faster than the transcription machinery in prokaryotes (Mirkin and Mirkin 2007). In the first case, the collision is known as co‐ directional collision, and in the second, as a head‐on collision. The second type of collision seems to have a more damageable impact on genome stability as it favors a pause of the replication fork, ultimately leading to a stalled replication fork (Kim and Jinks‐ Robertson 2012). Also, in the case of a head‐on collision between a replication fork and

7.6  Plastid Genome Stability and DNA Repair

RNA polymerase, positive supercoils accumulate behind the two structures, leading to replication fork reversal and possible changes to the genome (Bermejo et al. 2012; Kim and Jinks‐Robertson 2012). Whether these processes are relevant in plastid genome instability remains to be investigated, but their relevant roles in the genomic stability of many species suggest that they could play a major role in plastids as well. As frequent processes involving the plastid genome, chloroplast DNA replication and transcription are major factors that are susceptible to having an effect on plastid genome stability. Furthermore, these processes are not as well understood in the plastid as they are in the nucleus, and it is therefore possible that their impact on chloroplast genome variation is largely underestimated. Indeed, plastid DNA damage or repair processes are usually thought to be independent of these two plastid activities. Nevertheless, as demonstrated by the implication of one of the two POLI‐like proteins in the repair of plastid DNA (Parent et al. 2011) and by the replication slippage occurring in Oenothera chloroplast DNA (Stoike and Sears 1998), these processes have to be kept in mind when one thinks about plastid genome variation and stability. The challenge for the next few years will be to investigate the real role of replication and transcription on plastid genome stability/instability in higher plants and to integrate these to a plastid DNA maintenance model.

7.6 ­Plastid Genome Stability and DNA Repair Genomic stability is essential for all living organisms to ensure proper cell functions and transmission of their genetic material. This is also true for the chloroplast genome which is essential for photosynthesis and, consequently, for plant energy metabolism. Chloroplast genome integrity is, however, constantly compromised by numerous environmental and endogenous agents that can spontaneously damage DNA. Chloroplast genome stability is therefore maintained through the action of specialized proteins involved in the replication, recombination, and repair of DNA. The following section thus focuses on the main threats the chloroplast genome encounters, the associated repair pathways, and the principal actors involved. 7.6.1  Oxidative Stress, Photo‐Adaptation, and ROS Detoxification

Photosynthesis is the major energy source for autotrophic organisms. However, the photosynthetic electron transport (PET) chain is also one of the main sources of photo‐ oxidative stress that constantly threatens the photosynthesis apparatus and genome integrity (reviewed in Asada 2006). Indeed, ROS readily damage all components of the cell including nucleic acids, proteins, and lipids. Numerous environmental conditions such as high light intensities and herbicides stimulate ROS production. It was notably shown that singlet oxygen is the main type of ROS produced under high illumination in Arabidopsis cell suspension cultures (Gonzalez‐Perez et al. 2011) and that it accounts for the majority of photo‐oxidative damage in plants (Triantaphylides et  al. 2008). Therefore a complex set of mechanisms finely regulate the photosynthetic capacity and avoid an excessive generation of ROS. Chloroplasts act as sensors of fluctuating environmental conditions. To decrease ROS production, they exert both PET state transitions in the short term and transcriptome

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modulations in the long term. Two main chloroplast‐to‐nucleus signaling pathways are currently known to mediate this adaptation: the PET redox state and the accumulation of metabolites. The redox state of the plastoquinone pool, the electron acceptor of photosystem II (PSII), has long been proposed as the main site to perceive changes in the PET redox state (Escoubas et  al. 1995). Breakthrough studies identified the redox‐­ sensitive thylakoid protein kinases STT7 and STN7, respectively, in Chlamydomonas and Arabidopsis as such sensors (Depege et al. 2003; Bellafiore et al. 2005). Bellafiore et  al. also showed that STN7 is essential for light‐harvesting complex II (LHCII) phosphorylation‐­dependent state transition between photosystem I (PSI) and PSII, a process necessary for the correct light absorption capacity balance between both photosystems (reviewed in Wollman 2001). However, the plastoquinone redox state seems to be also involved in modifications of the thylakoid protein composition, leading to photosystems stoichiometry adjustment. Indeed, STN7 initiates a retrograde signaling cascade triggering the establishment of this response (Bonardi et al. 2005) by phosphorylation of a still‐unknown thylakoid protein, leading to modifications of the nuclear and chloroplast transcriptome patterns (Pesaresi et  al. 2009). However, transcriptomic studies showed that only a limited number of genes seem to be directly regulated by the plastoquinone redox state (Fey et  al. 2005), with PSI redox state being the most important (Piippo et al. 2006). In addition to the PET redox state signaling pathways that coordinate the photosynthetic activity with the changing environmental conditions, the accumulation of some specific metabolites in the chloroplast triggers retrograde signaling cascades. Of these metabolites, the ROS themselves are able to initiate retrograde signaling and likely constitute a stress response. Many ROS such as singlet oxygen (1O2), hydrogen peroxide (H2O2), and superoxide anion (O2–) are produced as by‐products of photosynthesis at many locations of the PET (Figure 7.3), making it difficult to differentiate between the signaling of each type of ROS. Singlet oxygen

1O

Energy 2

3O

e– 2

O2–

Superoxide anion

SODs + O2–, 2 H+ O2 +

Fe rea nto cti n Xs on AP Hydrogen peroxide H2O2

H2O

•OH +

–OH

Figure 7.3  ROS generation and scavenging. Energy or electron transfer to molecular oxygen generates, respectively, two types of ROS: singlet oxygen and superoxide anions. Superoxide dismutases and ascorbate peroxidases constitute the main scavenging process to eliminate superoxide and avoid production of highly reactive hydroxyl radical. 1O2, Singlet oxygen; 3O2, dioxygen; e–, electron; O2–, superoxide anion; SODs, superoxide dismutases; H2O2, hydrogen peroxide; APXs, ascorbate peroxidases; •OH, hydroxyl radical; –OH, hydroxide anion.

7.6  Plastid Genome Stability and DNA Repair

An effective tool to distinguish singlet oxygen from superoxide and hydrogen peroxide came from the flu mutant of Arabidopsis, which accumulates singlet oxygen upon a dark‐to‐light shift and undergoes, in mature plants, growth arrest and necrosis as a consequence (Meskauskiene et al. 2001; op den Camp et al. 2003). This mutant notably facilitated the identification of EXECUTER 1 and 2, two thylakoid‐localized proteins involved in singlet oxygen signaling. Indeed, an executer1/flu double mutant abrogates this phenotype and partially abolishes the nuclear response, while an executer1/­ executer2/flu triple mutant completely eliminates it (Wagner et al. 2004; Lee et al. 2007). This suggests that growth inhibition and necrosis are not the consequence of singlet oxygen toxicity but rather a genetically controlled response (op den Camp et al. 2003; Wagner et al. 2004; Lee et al. 2007). Although they do not seem to act directly as second messengers in this specific response, it was suggested that the products of singlet oxygen‐­mediated lipid peroxidation, which are synthesized enzymatically and are precursors of jasmonic acid, could be part of a plastid‐to‐nucleus signaling cascade (Przybyla et al. 2008). ROS retrograde signaling is thus not fully understood and further work will be needed to fully characterize each of these pathways. Nevertheless, an interesting study demonstrated that the hydrogen peroxide‐ and singlet oxygen‐mediated signaling pathways are not independent but rather have antagonistic effects that could account for a fine‐tuning of the response (Laloi et al. 2007). Furthermore, it was suggested that singlet oxygen accumulation is followed by loss of chloroplast integrity and subsequently by the collapse of the entire cell (Kim et al. 2012a). Interestingly, mutations of the EXECUTER proteins prevented chloroplast rupture, suggesting a role for these proteins in the signaling of these responses (Kim et al. 2012a). Adjustments of the plastid and nuclear genomes expression are thus complex, but allow adaptation to fluctuating environmental conditions. Indeed, it has been known for a long time that, in response to high light, LHC genes could be repressed (Escoubas et al. 1995; Ballottari et al. 2007) and antioxidant enzymes could accumulate to higher levels (Grace and Logan 1996). ROS are, however, not the only metabolites known to initiate retrograde signaling from the chloroplast. Other signaling molecules include MEcPP (methylerythritol cyclodiphosphate) and PAP (3′phosphoadenosine 5′phosphate), two metabolites involved in isoprenoid and sulfate compound synthesis, respectively (Estavillo et  al. 2011; Xiao et al. 2012). It has also been shown that β‐cyclocitral, a product of the oxidation of β‐carotene by singlet oxygen, constitutes a signal that initiates retrograde signaling and nuclear genetic reprogramming (Ramel et al. 2012). Interestingly, it has been shown that the genetic reprogramming associated with β‐cyclocitral is extremely similar to the one observed in the flu mutant (Ramel et al. 2012). Although some of these compounds might diffuse to the nucleus (Estavillo et al. 2012), how these signals travel through the cytosol is mostly unknown. One exception is the chloroplast outer‐­ membrane‐bound transcription factor PTM (PHD‐type transcription factor with transmembrane domains), which has been shown to be digested in response to retrograde signaling cascades. Once cleaved, the PTM N‐terminal domain diffuses to the nucleus and initiates genetic remodeling (Sun et al. 2011). Photo‐adaptation by genetic reprogramming is, however, a long term and complex process. Chloroplasts thus necessitate a particular scavenging machinery specialized in ROS detoxification. This is especially true for the superoxide anion that is formed by the PSI‐dependent photo‐reduction of molecular oxygen (O2) (Mehler 1951). Superoxide itself is only moderately reactive, but it poses a threat through its potential to be

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transformed, by the Haber–Weiss cycle, in the highly reactive hydroxyl radical (•OH), for which no enzymatic scavenging system is known (Halliwell and Gutteridge 1984; Halliwell 2006). Hydroxyl radicals notably jeopardize DNA integrity (Balasubramanian et al. 1998). SODs are thus central actors in these detoxification processes since they catalyze the conversion of the superoxide anion to hydrogen peroxide (reviewed in Pilon et al. 2011). Higher plants contain an assortment of iron, copper/zinc, and manganese SODs, but only the first two categories are found in the chloroplast (reviewed in Pilon et  al. 2011). The two chloroplast FeSODs, FSD2 and FSD3, were shown to be essential to protect the plastid nucleoid against oxidative stress (Myouga et al. 2008). Indeed, chloroplast development was compromised in fsd2 and fsd3 single mutants, which display a pale‐green phenotype. These mutations also render the plant extremely sensitive to a high light treatment, whereas a low light treatment could partially reverse this phenotype. There are many reports of the presence of FSD1 in chloroplasts (Kliebenstein et  al. 1998; Zybailov et  al. 2008), but its subcellular localization is still under debate. Apart from FeSODs, the only chloroplast member of the copper/zinc SOD family, CSD2 (Kliebenstein et al. 1998), was shown to be an important superoxide detoxifying enzyme (Sunkar et al. 2006). Interestingly, CSD2 is mainly found on the stromal side of the thylakoid membrane (Ogawa et  al. 1995) and, as a consequence, in proximity to chloroplast DNA. Myouga et al. demonstrated that, similarly to CSD2, FSD3 was localized specifically to plastid nucleoids, whereas FSD2 was found more uniformly all over the chloroplast. It was also shown that FSD2 and FSD3 interact with each other, suggesting that FSD3 can recruit FSD2 to the plastid nucleoid and thus promote ROS scavenging at that specific location (Myouga et al. 2008). This function for FeSODs seems to be conserved from dicotyledons to monocotyledons, since FSD2 and FSD3 are also found in maize plastid nucleoids (Majeran et al. 2012). These results are in accordance with several studies that found FSD2 and FSD3 as part of the plastid transcriptional machinery. Indeed, Pfalz et al. identified many proteins found in Arabidopsis and mustard pTAC, including FSD2 and FSD3 (Pfalz et al. 2006). More recently, using a mass spectrometry approach, Steiner et al. identified FSD2 and FSD3 as essential subunits of the PEP and classified these enzymes as PEP‐associated proteins (PAP) 4 and 9, respectively (Steiner et al. 2011). SODs thus appear to be of major importance for chloroplast genome stability and expression. Hydrogen peroxide constitutes the substrate for the final step of the Haber–Weiss cycle, called Fenton’s reaction, which generates hydroxyl radicals (Halliwell 2006). It is thus of extreme importance to avoid the release of free hydrogen peroxide in the stroma. SODs are therefore in close proximity to hydrogen peroxide scavenging enzymes (reviewed in Pilon et  al. 2011), particularly thylakoid‐bound ascorbate peroxidase (tAPX) (Miyake and Asada 1992). A soluble stromal ascorbate peroxidase (sAPX) is also found in the chloroplast. APXs catalyze the conversion of hydrogen peroxide to water (reviewed in Pilon et al. 2011) using ascorbate as the electron donor (Foyer and Halliwell 1976). Interestingly, although both APXs are important for hydrogen peroxide scavenging, loss of tAPX seems to have a greater impact on photo‐oxidative stress than sAPX (Maruta et al. 2010). This might point towards the importance of SODs–tAPX coupling at the thylakoid membrane. Additionally to APXs, chloroplasts contain glutathione peroxidases (Rodriguez Milla et al. 2003) and numerous other anti‐oxidant enzymes that prevent oxidative damage (reviewed in Gill and Tuteja 2010).

7.6  Plastid Genome Stability and DNA Repair O N

N

O H N

NH

oxidation

N

NH2

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8-oxoguanine

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oxidation O

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Thymine

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Thymine glycol

Figure 7.4  Oxidation of guanine and thymine mainly leads to generation of 8‐oxoguanine and thymine glycol, respectively.

Although photo‐adaptation and ROS scavenging machineries minimize the number of free oxygen radicals, DNA bases are frequently oxidized, generating products such as 8‐oxoguanine and thymine glycol (Figure 7.4). Such oxidized bases can mis‐pair in the DNA duplex and consequently yield mutations in the DNA sequence following few rounds of replication (Shibutani et al. 1991). This can be prevented by the multistep process base excision repair, which removes and replaces damaged nucleotides. Base excision repair initiates with the action of DNA glycosylases that recognize and excise the damaged base by hydrolysis of the N‐glycosidic bond. The sugar‐phosphate backbone is then cleaved on the 5′ side of the apurinic/apyrimidinic (AP) site by an endonuclease, the gap filled by the insertion of one or more nucleotides, the residual base pairs removed, and the backbone sealed again (reviewed in Zharkov 2008). Despite the fact that many enzymes involved in base excision repair are known in plants, the precise mechanism is still poorly understood in chloroplasts. Nevertheless, some studies identified enzymes that might be part of a chloroplast‐specific base excision repair machinery. Indeed. Babiychuk et al. found in Arabidopsis the ARP protein that functions as an AP endonuclease (Babiychuk et  al. 1994). Later, AtNTH1, an Arabidopsis homolog of the E. coli endonuclease III (encoded by the Nth gene), was identified and shown to exhibit both a DNA glycosylase activity against damaged pyrimidines (such as thymine glycol) and an AP lyase activity (Roldan‐Arjona et al. 2000). Such glycosylases that also exert an AP lyase activity are qualified as bifunctional and are able to cut on the 3′ side of the AP site. However, it was only recently that Gutman et al. showed that ARP, AtNTH1, and AtNTH2, another homolog of the E. coli endonuclease III, are localized at the chloroplast nucleoid (Gutman and Niyogi 2009). Apart from oxidized pyrimidines, ROS can also modify purines, notably generating 8‐­oxoguanine, which necessitates particular glycosylases in order to be excised. Although such glycosylases have not been identified in the chloroplast, plants are the only eukaryotes to encode a homolog of the prokaryotic MUTM protein, a bacterial‐type DNA glycosylase that targets oxidized purines (Ohtsubo et al. 1998). This suggests that this MUTM homolog

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has been acquired at the moment of plastid endosymbiosis, and could potentially still be targeted to the chloroplast even though its subcellular localization has not been confirmed (Ohtsubo et al. 1998). Finally, ROS can also induce the oxidative deamination of cytosines, converting them into uracil residues (Duncan and Miller 1980). Interestingly, a uracil‐DNA glycosylase activity has been observed in maize chloroplasts (Bensen and Warner 1987). 7.6.2  UV‐Induced DNA Damage

In addition to ROS generation, light harvesting has the inconvenience of exposing tissues to the harmful effects of UV‐B radiations. UV‐B rays (280–320 nm) represent a threat to DNA stability as they induce two main lesions, cyclobutane pyrimidine dimers (CPDs) and pyrimidine‐pyrimidone (6‐4) photoproducts, which are both formed through the covalent linkage of two adjacent pyrimidines (Figure 7.5). These types of DNA damage were shown to inhibit progression of both RNA and DNA polymerases and thus constitute an important risk to accurate gene expression and replication (Villani et al. 1978; Mitchell et al. 1989). Although translesion DNA synthesis allows a mutagenic bypass of the lesions during DNA replication (reviewed in Waters et  al. 2009), it remains essential to keep the O

H C

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Cyclobutane pyrimidine dimer (CPD)

UV-B

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OH

CH3

R

6-4 photoproduct Figure 7.5  UV‐B‐induced linkage of adjacent pyrimidines mainly leads to formation of cyclobutane pyrimidine dimers (CPDs) and 6‐4 photoproducts.

7.6  Plastid Genome Stability and DNA Repair

concentration of such dimers as low as possible to minimize the accumulation of mutations. Accordingly, specialized DNA repair mechanisms remove CPDs and 6‐4 products in the DNA helix. These mechanisms can be sorted into two classes, whether or not they involve a light‐dependent removal of the dimers, respectively called photoreactivation and dark repair. The latter is associated with the nucleotide‐excision repair pathway and, although it was shown to remove 6‐4 products in the nucleus of Arabidopsis (Britt et  al. 1993), no evidence suggests that it also takes place in the organelles. In contrast to the nucleotide‐excision repair pathway that necessitates DNA synthesis, photoreactivation constitutes the enzymatic separation of the dimer into two pyrimidine monomers. This process is mediated by a class of enzymes called photolyases that use blue light as their catalytic energy source. Photorepair of pyrimidine dimers has been extensively studied in the plant nucleus. Indeed, it was shown some time ago that Arabidopsis can repair CPDs in both the absence or presence of visible light, but that in the latter condition, removal of CPDs was faster (Pang and Hays 1991). In addition, a distinct pathway for the light‐dependent removal of 6‐4 products was shown to occur in Arabidopsis (Chen et al. 1994). These results were further confirmed by the identification of UV‐B‐sensitive mutants defective in each of these repair pathways. The uvr1, uvr2, and uvr3 mutations were, respectively, shown to compromise specifically dark repair, CPDs photorepair, and 6‐4 products photorepair (Britt et al. 1993; Jiang et al. 1997; Landry et al. 1997). Interestingly, cloning of a  gene for an Arabidopsis CPD photolyase, designated AtPhr1 (photoreactivating enzyme 1), and for a 6‐4 photolyase gene showed that they mapped to the same genes as the uvr2 and uvr3 mutations, respectively (Ahmad et al. 1997; Nakajima et al. 1998). Although these UV‐B DNA lesions are very deleterious in plant chloroplasts, the existence of photorepair in this compartment is rather controversial. In fact, the situation appears to vary between species, as some appear to exhibit photorepair in their chloroplasts while others do not. The green alga C. reinhardtii is one of the organisms for which the chloroplast photorepair of UV‐B‐induced DNA damage is fairly well established. Small and Greimann showed that CPD photorepair is active in both Chlamydomonas nucleus and chloroplast (Small and Greimann 1977). Accordingly, a gene encoding a CPD photolyase with a putative chloroplast‐targeting peptide was cloned and shown to complement an E. coli photolyase‐deficient mutant (Petersen et al. 1999). Interestingly, the protein encoded by this gene, PHR2, was shown to be dually targeted to the nucleus and chloroplast, and to necessitate the product of a second gene, Phr1 (which is not a homolog of AtPhr1), to carry out its function in both compartments (Petersen and Small 2001). Because a phr1 mutation leads to a photoreactivation deficiency in both the nucleus and chloroplast, it was hypothesized that this gene would also encode a photolyase (Petersen and Small 2001). Surprisingly, Chlamydomonas Phr1 was instead shown to encode an FO (7,8‐didemethyl‐8‐hydroxy-5-­deazariboflavin) synthase (Petersen and Ronan 2010). In addition to flavin adenine dinucleotide (FAD), FO often serves as a second chromophore for photolyase blue light harvesting and thus seems essential for proper PHR2 activity (Petersen and Ronan 2010). Chlamydomonas is, however, not the only organism to exhibit photorepair in these two DNA‐containing compartments. Maize and soybean were also shown to use photoreactivation for the repair of CPDs in both compartments (Cannon et al. 1995; Stapleton et al. 1997). This is also the case for rice, which encodes a single CPD photolyase gene that is targeted to all DNA‐containing compartments (Takahashi et al. 2011).

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Conversely, other plants apparently lack photorepair in their chloroplasts. This is the case for spinach, in which CPD photoreactivation could be detected in whole‐leaf extracts but not in purified chloroplast extracts (Hada et  al. 2000). The existence of chloroplast photorepair in the model plant Arabidopsis is also rather controversial. Chen et al. reported that, in both the presence and absence of visible light, CPDs were poorly repaired in organelle genomes of 5‐day‐old Arabidopsis seedlings (Chen et al. 1996a). In addition, overexpression of an Arabidopsis PHR1–GFP fusion in protoplasts resulted in an exclusive nuclear import (Kaiser et al. 2009). This is in agreement with the observation of an apparent lack of transit peptide in Arabidopsis PHR1 (Ahmad et al. 1997). It is, however, worth mentioning that plants such as maize require high light conditions for growth, thus probably making pyrimidine dimer repair more essential. On the other hand, this result contrasts with the observation that older Arabidopsis seedlings (14 days) efficiently remove CPDs and 6‐4 products from both their nuclear and chloroplast DNA in the presence of blue light (Draper and Hays 2000). Still, this study provided no proof that this chloroplast blue light‐dependent photoproducts removal was mediated by a photolyase. Interestingly, in a genetic screen for complementation of an E. coli UV‐sensitive mutant, a cDNA encoding Arabidopsis Drt101 (DNA‐damage Repair/Toleration) was isolated and predicted to possess an N‐terminal chloroplast targeting sequence (Pang et al. 1993b). An intriguing observation came from the identification of the first Arabidopsis blue light photoreceptor. Surprisingly, this protein was shown to share extensive sequence similarity with bacterial photolyases (Ahmad and Cashmore 1993). It was later shown that the blue light photoreceptors from several organisms, called cryptochromes, share similarity with DNA photolyases not only in amino acid sequence but also in chromophore composition and three‐dimensional structure (Cashmore et al. 1999; Brautigam et al. 2004). Indeed, crystal structure of Arabidopsis CRY1 revealed a folding very similar to several photolyases (Brautigam et  al. 2004). However, despite this apparent homology, no DNA photolyase activity could be detected for CRY1 (Lin et al. 1995). CRY1 and 2, the two initially characterized Arabidopsis cryptochromes, were instead shown to exert control over diverse processes such as de‐etiolation, photoperiodism, gene expression, and flowering (reviewed in Lin 2002). In addition to these two photoreceptors, a third Arabidopsis blue light photoreceptor, CRY3, has been identified and shown to belong to a newly characterized class of cryptochromes called DASH (to underline the relationship of the Arabidopsis and Synechocystis cryptochromes with their Drosophila and Homo sapiens counterparts) (Brudler et  al. 2003; Kleine et  al. 2003). Interestingly, it was reported that AtCRY3/DASH is targeted to both chloroplasts and mitochondria, and although no photolyase activity could be detected, it was shown to bind non‐specifically to DNA (Kleine et  al. 2003). Unexpectedly, breakthrough studies revealed that the DASH family of cryptochromes, including AtCRY3, has a specific photolyase activity towards CPDs in ssDNA only (Huang et  al. 2006; Selby and Sancar 2006). Accordingly, it was later shown that AtCRY3, and probably the other members of the DASH family, lost the capacity of flipping the CPD dimer out of the DNA duplex but retained its photolyase activity (Pokorny et  al. 2008). Pokorny et  al. suggested that the disrupted DNA helix during replication and transcription could be a relevant substrate for this specific class of CPD photolyases (Pokorny et al. 2008). It is therefore tempting to speculate that CRY3 is involved in Arabidopsis organelle genome stability.

7.6  Plastid Genome Stability and DNA Repair

7.6.3  Recombination and DNA Double‐Strand Break Repair

Compared to ROS and UV‐light‐induced DNA damages, a DSB is certainly the most threatening genotoxic stress, since it can lead to severe chromosomal rearrangements. DSBs are induced by the stalling and collapse of replication forks (Burhans and Weinberger 2007), by chemical agents that spontaneously damage the DNA (Heisig 2009), and during programmed genomic rearrangements such as meiotic and V(D)J recombination (Neale and Keeney 2006; Schatz and Ji 2011). Many specialized cellular processes have thus evolved to allow DSB surveillance and to promote the appropriate repair mechanisms. DSB repair mechanisms can be roughly divided into two classes, whether they involve the joining of two DNA ends or recombination. Homologous recombination (HR) is the most privileged and prevailing mechanism for the repair of DSBs since it permits a conservative repair of the break, and although the proteins involved differ, it is conserved from prokaryotes to eukaryotes. Briefly, HR consists of the 5′ end resection of the DNA ends by an ssDNA exonuclease, followed by the coating of the resulting ssDNA by an ssDNA‐binding protein (SSB/RPA). This ssDNA‐binding protein is then replaced by a recombinase (RECA/RAD51) that catalyzes the invasion of a homologous DNA duplex by the free 3′ end. A DNA polymerase then synthesizes the missing sequence using the homologous DNA as the matrix, and a resolvase catalyzes the resolution of the Holliday junction (reviewed in Helleday et al. 2007). Many HR pathways such as double‐strand break repair (DSBR), which often results in crossover, single‐ strand annealing (SSA) and synthesis‐dependent strand annealing (SDSA), have been shown to repair DSBs (Figure 7.6). In contrast to HR, the main end‐joining mechanism for the repair of DSBs, called non‐homologous end joining (NHEJ), acts by the ligation of two free DNA ends regardless of the homology (reviewed in Weterings and van Gent 2004). Although the study of HR has mainly focused on nuclear and prokaryotic genomes, the existence of recombination in the chloroplasts was recognized in the 1980s. Jeffrey D. Palmer showed in 1983 that the chloroplast genome of the common bean (Phaseolus vulgaris) consists of equimolar populations differing only by the relative orientation of their single copy DNA regions (Palmer 1983). It could thus be assumed that interconversion of the single‐copy regions was mediated by recombination between the IR. It was further shown that DSBs in the chloroplast genome of the green alga C. reinhardtii could be repaired by recombination between the IR. The IR of this organism contain the 23S ribosomal RNA gene, which, in turn, contains a group I intron with an open reading frame encoding the DNA endonuclease I‐CreI (Durrenberger and Rochaix 1991). Notably, it was shown that the I‐CreI cleavage site was located at the exon–exon junction, and that an intronless 23S cDNA transformed in the Chlamydomonas chloroplast systematically integrates the intron (Durrenberger and Rochaix 1991). Interestingly, intron‐homing only occurred if the intronless cDNA was in an inverted orientation relative to its homologous genomic DNA. In contrast, a direct repeat orientation led to a deletion between the exogenous and endogenous sequences, reminiscent of the SSA pathway (Durrenberger et  al. 1996). This is consistent with an active HR repair of DSBs in the chloroplasts, in which a homologous sequence elsewhere in the genome is used as the template. In fact, all three HR pathways were found to operate in Chlamydomonas chloroplasts (Odom et  al.

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DNA DSB

5ʹ-end resection Annealing of homologous sequences

5ʹ-end invasion of homologous duplex

Ligation

SSA

DNA synthesis

Separation/resolution and ligation

With Crossover

SDSA

Without Crossover

DSBR

Figure 7.6  Repair of a DSB by homologous recombination. Dotted lines represent newly synthesized DNA and purple lines represent stretches of homologous sequence. SSA, Single‐strand annealing; SDSA, synthesis‐dependent strand annealing; DSBR, double‐strand break repair. Source: Adapted from Hastings et al. 2009b. (See plate section for color representation of this figure.)

2008). Indeed, although Dürrenberger et al. were not able to identify crossover products (Durrenberger et  al. 1996), Odom et  al. showed that both DSBR and SDSA work to repair DSBs, but that the latter must prevail (Odom et al. 2008). Furthermore, when extensive homology for repair was lacking, repeats as short as 15 base pairs were used, consistent with the SSA pathway (Odom et al. 2008). In contrast, no evidence for NHEJ in Chlamydomonas chloroplasts was obtained. It is, however, worth mentioning that Chlamydomonas is extremely repeat rich. Indeed, more than 20% of its chloroplast genome represents short repeated DNA sequences (Maul et al. 2002). In comparison, Arabidopsis is repeat poor, and, hence, the preferred mechanism for the repair of DSBs could differ. A recent study assessed this question by generating conditional, chloroplast‐targeted I‐CreII Arabidopsis transgenic lines (Kwon et al. 2010). Induction of the endonuclease resulted in repair products that had undergone deletions on both sides of the DSB. Furthermore, in most of the products obtained, micro‐repeats were observed at the repair junctions (Kwon et al. 2010). This is reminiscent of the mammalian microhomology‐mediated end joining pathway (MMEJ), also called alternative‐NHEJ (alt‐NHEJ), that constitutes a distinct DSB repair mechanism (Bennardo et al. 2008). Additionally, in contrast to Chlamydomonas, some other repair junctions contained no micro‐ homologies, which therefore suggests the presence of NHEJ in the chloroplasts of

7.6  Plastid Genome Stability and DNA Repair

higher plants (Kwon et  al. 2010). Apart from the repair of DSBs, recombination in N. tabacum plastids has also been shown to eliminate deleterious mutations by gene conversion (Khakhlova and Bock 2006). Even though the chloroplast DSB repair pathways are fairly well characterized, they remain poorly understood from an enzymatic point of view. Nevertheless, certain proteins that catalyze some of the key steps of HR have been identified. This is especially true for the homologs of the bacterial RECA recombinase that have been found in the chloroplasts of higher plants (Cerutti et  al. 1992), of the green alga C. reinhardtii (Nakazato et al. 2003), and of the moss Physcomitrella patens (Inouye et al. 2008). In prokaryotes, this enzyme catalyzes the strand exchange between a single‐stranded 3′ end and a homologous duplex DNA. The function of the chloroplast RECA (cpRECA) was extensively studied in the 1990s by Cerutti et al. Indeed, an Arabidopsis stromal protein cross‐reacting with an E. coli RECA antibody was identified and shown to share more than 50% sequence identity with its bacterial counterparts (Cerutti et al. 1992). Its involvement in recombination was further shown in pea (Pisum sativum), in which the RECA homolog exhibits a strand exchange activity dependent on Mg2+, ATP, and a homologous duplex DNA (Cerutti and Jagendorf 1993). Additional evidences for its function in DNA repair came from the use of Chlamydomonas‐dominant negative RecA mutants that were hypersensitive to DNA‐damaging agents (Cerutti et al. 1995). Interestingly, the RecA expression level was also shown to increase following treatment of pea protoplasts with DNA‐ damaging agents (Cerutti et al. 1993). In a report, Rowan et al. further confirmed these results by the use of cpRecA T‐DNA insertion mutants that showed an altered structure of their chloroplast DNA (Rowan et al. 2010). This mutant showed an increase in its ssDNA content and a decrease in its total chloroplast DNA. It was also shown that the cpreca mutant is sensitive to ciprofloxacin, a DNA gyrase inhibitor producing DSBs in plant organelles (Rowan et al. 2010). All of these evidences thus support an important role for cpRECA in the maintenance of chloroplast genome integrity. Although not as much documented, other proteins have also been suggested to participate in the homology‐directed repair of the chloroplast DNA. It was shown that RECA2, another RECA homolog, is dually targeted to Arabidopsis chloroplasts and mitochondria (Shedge et al. 2007). Moreover, the cDNA of the putative chloroplast protein DRT100 was shown to complement the sensitivity to DNA‐damaging agents of a RECA‐deficient E. coli strain (Pang et al. 1992). However, this protein shares little in common with bacterial RECAs. Another study allowed the identification of DRT111 and DRT112, two other putative chloroplast‐targeted proteins, which complement the E. coli Holliday junction resolution‐deficient ruvC/recG double mutants (Pang et  al. 1993a). Interestingly, both proteins show no significant similarity to known prokaryotic Holliday junction resolvases, and DRT112 is highly homologous to plastocyanin (Pang et al. 1993a). POLIB, one of the two Arabidopsis chloroplast replicative DNA polymerases, is also thought to be specialized in DNA repair (Parent et al. 2011). It was actually shown that polIb mutants are sensitive to ciprofloxacin and accumulate more unrepaired DSBs in the DNA of their organelles than wild‐type plants upon treatment (Parent et al. 2011). Curiously, RAD52, which catalyzes RAD51 loading on ssDNA, is found in all Arabidopsis DNA‐containing organelles (Samach et al. 2011). This finding was unexpected, as the chloroplast genome maintenance apparatus mainly consists of prokaryotic‐like proteins. Although it was shown to be involved in mitochondrial

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homologous recombination, there is no evidence for its involvement in chloroplast DNA repair (Janicka et al. 2012). Despite its capacity to repair DSBs, recombination must be finely regulated in order to avoid drastic DNA rearrangements. Indeed, most processes of DNA metabolism require ssDNA intermediates with considerable recombinogenic properties. ssDNA thus has to be stabilized to avoid pairing of complementary sequences and, consequently, the occurrence of illegitimate recombination. For this purpose, plant organelles contain proteins specialized in preventing non‐homologous recombination. Of these proteins, the nuclear‐encoded protein MSH1 participates in illegitimate recombination surveillance in both the mitochondria and the chloroplast (Shedge et  al. 2007; Xu et al. 2011). MSH1 was formerly called ‘chloroplast mutator 1’ (CHM1), as homozygous mutations in this gene lead to distorted and variegated leaves (Rédei and Plurad 1973). It was later renamed MutS homolog 1 (MSH1) as the encoded protein showed homology to the E. coli mismatch‐repair protein MUTS (Abdelnoor et  al. 2003). Although no functions in mismatch‐repair were reported for this protein, MSH1 mutants accumulate organellar genome rearrangements in both the plastid and the mitochondria, suggesting an important role in recombination surveillance for this protein (Shedge et al. 2007; Xu et al. 2011). More recently, it has been shown that the loss of MSH1 is associated with methylome reprogramming (Virdi et  al. 2015). This reprogramming produces epigenetic changes, such as enhanced growth or dwarfism, which can be isolated through crosses (Xu et al. 2012; Virdi et al. 2015; Yang et al. 2015). In addition to MSH1, the Whirly proteins are involved in the prevention of organellar illegitimate recombination. They constitute a small family of ssDNA‐binding proteins almost exclusively found in the plant kingdom. Whirlies were first characterized as transcriptional activators of pathogenesis‐related genes in the nucleus (Desveaux et al. 2000; Desveaux et al. 2004), but it was later shown that these nuclear‐encoded proteins perform important functions in the organelles as well (Maréchal et  al. 2009). In Arabidopsis, AtWHY1 and AtWHY3 are targeted to the plastids, while AtWHY2 is targeted to the mitochondria (Krause et al. 2005). Upon tetramerization, these proteins bind ssDNA with limited sequence specificity and stabilize it (Cappadocia et al. 2010). It was further shown that tetramers, when bound to long ssDNA molecules, exert cooperative binding and assemble into higher complexity mega‐structures (Cappadocia et al. 2012). To elucidate the function of Whirly proteins in the plastids, Maréchal et al. obtained why1why3 double mutants and showed that these plants accumulate large amounts of  genomic rearrangements (Maréchal et  al. 2009). The identified rearrangements mainly consist of duplications and deletions of DNA fragments bordered by very short repeated sequences, suggestive of a particular type of illegitimate recombination called microhomology‐mediated recombination (MHMR) (Maréchal et al. 2009). This specific type of illegitimate recombination, also called microhomology‐mediated break‐ induced replication (MMBIR), was associated with copy number variation (CNV) observed in the human and yeast genomes (reviewed in Hastings et al. 2009a). It was thus suggested that Whirly proteins act as anti‐recombination proteins that stabilize ssDNA until it is correctly repaired. This hypothesis was further supported by the finding that why1why3 plants are sensitive to ciprofloxacin and accumulate even more DNA rearrangements after this treatment (Cappadocia et al. 2010). Therefore it seems

7.7  Outcomes of DNA Rearrangements

probable that following the occurrence of a DSB in the chloroplast DNA duplex, the ssDNA produced by the 5′ end resection is stabilized by the Whirly proteins until it is taken over by the homo­logous recombination machinery. Crystallographic characterization of the ssDNA‐­binding mode of Whirly supports this hypothesis as, in the DNA– protein complex, the nucleobases are unavailable for pairing with complementary sequences (Cappadocia et al. 2010). Interestingly, a cross between the polIb DNA polymerase mutant and the why1why3 plants yielded plants with severe growth defects and many more DNA rearrangements than the why1why3 plants, indicating a synergistic interaction between these proteins (Parent et al. 2011). As a consequence, Whirlies and POLIB most probably function in complementary pathways for the conservative repair of DSBs. Similarly to POLIB, the combination of the cpreca (reca1) and why1why3 mutations in Arabidopsis leads to a synergistic interaction, as the triple mutant why1why3reca1 exhibits severe growth retardation and a white variegation phenotype (Zampini et al. 2015). This synergy is also reflected at the level of chloroplast genome instability, since the triple mutant accumulates high levels of DNA rearrangements. In addition to the  generalized instability observed in why1why3polIb, why1why3reca1 accumulates approximately 60 times more short‐range inversions than wild‐type plants (Zampini et  al. 2015). As the occurrence of this type of genomic instability correlates with an apparent replication stress, it has been speculated that this type of genomic instability arises from a replication‐dependent mechanism. It was suggested that, when the progression of replication forks is hampered, short inverted microhomologies promote the annealing of the 3′ extremity of the leading strand to the opposite template strand, similarly to what has been observed in other systems (Mizuno et al. 2009; Paek et al. 2009; Mizuno et al. 2013). Replication re‐initiation on this wrong strand would then lead to the formation of a short‐range inversion, also called U‐turn‐like inversion (Zampini et  al. 2015). As the frequency of these U‐turn‐like inversions is highly increased in why1why3reca1 compared to wild‐type plants, it indicates that cpRECA (RECA1) and Whirly proteins both stabilize replication forks and preserve plastid genomic integrity (Zampini et al. 2015). Surprisingly, relatively high levels of U‐turn‐like inversions are also found in wild‐type Arabidopsis chloroplasts and mitochondria, suggesting that this  type of inversion constitutes a physiological characteristic of organelle DNA. Interestingly, U‐turn‐like inversions also seem to be a hallmark of human mitochondria, in which more than half of the genomic rearrangements seem to arise from this replication‐dependent process (Zampini et al. 2015).

7.7 ­Outcomes of DNA Rearrangements In this chapter, the mechanisms involved in maintaining plastid genome stability have been described. Nevertheless, in the field of DNA replication and repair, it is well known that error is an unavoidable compromise for processivity. Mutations and rearrangements therefore occur in the plastid genome and shape its evolution. Rearrangements in the plastid genome can have different impacts on plant development. One interesting phenotype that can arise from plastid genome instability is variegation. This phenotype is defined by the presence of differently colored zones on a tissue of a plant. Variegated mutants have been observed in many different plant

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species. In Oenothera, plastome mutator has been extensively described and characterized (Stoike and Sears 1998). The plastid genome of this mutant contains deletions at sites of small tandem repeats, particularly in the 16S‐TrnI gene. These deletions are thought to be produced by template slippage at the site of secondary structure formed by the repeat. These deletions could be linked to the appearance of variegated sectors on the plant. Moreover, the variegated phenotype is maternally inherited, and so directly linked to plastid genome transmission. Despite the extensive characterization of this plant, the nuclear gene that causes this chloroplast genome instability is still unidentified. In Arabidopsis, three variegated mutants are dependent on plastid genome instability: Whirly mutants (Maréchal et al. 2009), RecA mutants (Rowan et al. 2010), and Msh1 mutants (Xu et al. 2011). The latest was originally characterized for its variegated phenotype, and DNA rearrangements have only recently been identified in its plastid genome (Xu et al. 2011). In all of these mutants, the rearranged DNA molecules are present at a low, substoichiometric level (Small et al. 1987). This heteroplasmy could be the cause of the variegated phenotype, suggesting that rearranged plastid genomes could lead to the apparition of white sectors. The phenotype is also maternally inherited in Whirly mutants and msh1. Variegation has been observed in transgenic lines that conditionally generate chloroplast DSBs upon ß‐estradiol treatment, supporting the hypothesis that DNA rearrangements in the chloroplast can lead to this phenotype (Kwon et al. 2010). Other variegated mutants have been characterized in Arabidopsis, notably Var1, Var2, and Immutans, but were shown to be linked to ROS imbalance or chloroplast differentiation (Wetzel et al. 1994; Sakamoto et  al. 2002; Sakamoto et  al. 2004; Kato et  al. 2007; Rosso et  al. 2009) and independent of DNA rearrangements. More recently, the Arabidopsis why1why3polIb triple mutant was shown to have high levels of DNA rearrangements in its chloroplast genome (Parent et al. 2011). This high level of DNA rearrangements is associated with severe dwarfism, yellow variegation, photosynthetic defects, and increased chloroplast ROS production (Parent et al. 2011; Lepage et al. 2013). As the plastid genome encodes many components of the photosynthetic machinery, extensive plastid genome modifications could account for the appearance of these phenotypes. Another outcome of plastid genomic instability is the rare appearance of herbicide resistance. One case that has been widely characterized is that of atrazine resistance. This mutation was characterized in Solanum nigrum and linked to a single codon change in the PsbA gene (Goloubinoff et al. 1984). The same pattern of point mutations was shown in many other species such as A. thaliana (El‐Lithy et al. 2005), N. tabacum (Sato et  al. 1988), H. vulgare (Rios et  al. 2003), Senecio vulgaris (Park and Mallory‐ Smith 2006), and Amarantus blitoides (Sibony and Rubin 2003). The latter two are of particular concern because of their deleterious potential on many different crop cultures. Some chloroplast point mutations located in the ribosomal 16S and 23S genes have also been related to antibiotic resistance in C. reinhardtii (Harris et  al. 1989). Finally, in tobacco, a point mutation in the 23S ribosomal protein leads to lincomycin resistance (Cseplo et al. 1988), but the mechanism leading to these mutations is still a matter of debate. Some hypermutable regions have been identified in the plastid genome of legumes (Jansen et al. 2007; Magee et al. 2010). These regions appear to diverge more rapidly between species and could be linked to a higher frequency of DNA breakage, or fork

7.8  Concluding Remarks

collapse. It is proposed that most of these regions could be associated with a high abundance of tandem repeats of short DNA sequences. In fact, plants containing more DNA repeats in their chloroplasts tend to have even more unstable chloroplast genomes (Haberle et al. 2008). One of these hypermutable regions is located in the Ycf4 region. This 1.5‐kb DNA fragment has been demonstrated to evolve more rapidly than the rest of the genome. Furthermore, Ycf4 has been lost in some plants species, and the plastid genome of these plants tends to be more stable. Interestingly, this part of the genome is prone to rearrangements in both whirly and msh1 mutants, in A. thaliana and Zea maize (Maréchal et al. 2009; Xu et al. 2011). The genes surrounding this region, Ycf4‐ PsaI‐AccD‐Rps16, are subject to extensive rearrangements as well. The adaptation of many species to their local environment could be attributed to changes in their organelle genomes (reviewed in Budar and Roux 2011). Indeed, reciprocal crosses between species that spontaneously hybridize in nature showed that their cytoplasm plays an important role in the survival of plants in a given environment (Sambatti et al. 2008). One must remember that organelle genomes are inherited maternally, and so are linked to the transmission of cytoplasm. Furthermore, a divergence in plastid genome evolution has been observed within the same species, such as Chamaecrista fasciculata, in which it gives rise to a subtle phenotype that confers an advantage in Maryland plant populations (Galloway and Fenster 2001). A difference has also been observed in Arabidopsis lyrata between European and American populations (Leinonen et al. 2011). Finally, it is important to note that many mutations in the plastid genome can have deleterious effects. Most of the chloroplast genes are essential for plant viability. Mutations in photosystem proteins (Scharff and Koop 2007), ribosomal proteins (Tiller et al. 2012), and lipid biogenesis (Kuroda and Maliga 2003) were characterized to be lethal. Some point mutations were shown to have milder effects on plant development, such as photosynthetic defects (Schottler et al. 2007; Krech et al. 2012).

7.8 ­Concluding Remarks To fully understand the mechanisms ensuring the stability of the plastid genome, it will be important to pay attention to the regular processes taking place within plastids. As discussed in this chapter, pressure on the genome comes not only indirectly from the role of the chloroplast in energy metabolism, but also directly from the replication and transcription machineries. The response to these many sources of instability is itself directed by characteristics proper to the plastid genome, some of which are better understood than others. While the abundance of repeated sequences and the high level of ploidy are well associated to recombination, the roles of branched and linear forms of the genome in DNA metabolism remain largely unknown. It is also important to note that many mechanisms of DNA repair found in plastids were previously known from their existence in nuclear and prokaryotic genomes. As such, differences in the machinery and enzymatic mechanisms involved have not been as well characterized. Of particular interest, for instance, is the plastid‐to‐nucleus signaling that occurs in relation to the maintenance of organellar genomes. The dependence of the plastid on nucleus‐ encoded genes adds a level of complexity that is only beginning to emerge in the study of genome stability.

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Part III Somatic Genome Variation in Microorganisms

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8 RNA-Mediated Somatic Genome Rearrangement in Ciliates John R. Bracht* Department of Biology, American University, Washington, DC, USA

Abstract Ciliates are powerful model systems for the study of genome rearrangements, leading to important insights into the epigenetic mechanisms involved in restructuring their somatic genomes. A recurring motif is the use of noncoding RNA to sculpt the genetic information into its final functional form, which is transmitted to future generations without altering the germline genome. Therefore the somatic genome constitutes an epigenomic template, containing functional information not encoded in the germline and yet essential to the construction of proteincoding genomic sequences. This chapter describes the current understanding of the mechanisms of this programmed genome rearrangement in various ciliate species, with emphasis on the roles for noncoding RNAs in guiding the rearrange­ ment process. In addition to previously described somatic template RNAs, data are pre­ sented demonstrating that Oxytricha produces noncoding RNA from at least one class of noncoding germline sequences during genome rearrangements, consistent with observa­ tions in other ciliates. Transposons (and their associated transposase proteins) play key roles in ciliate genome rearrangements. The chapter discusses recent data from Tetrahymena and Oxytricha that support a single origin of genome rearrangement processes as a Tc1/mariner transposon invasion event in an ancient common ancestor. It concludes by describing common themes in the programmed genome rearrangements of ciliates, vertebrates, and some species of bacteria. Keywords  Oxytricha; Stylonychia; genome rearrangement; DNA elimination; noncoding template RNAs; small RNAs; short RNAs; nuclear dimorphism; non-Mendelian inheritance; transposable elements; programmed genome engineering; chromatin modifications; ubiquitous genome variation

*Corresponding author: [email protected] Bracht J.R. (2017) RNA‐mediated somatic genome rearrangement in ciliates. In: Li X.‐Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley‐Blackwell, Hoboken, NJ, Ch. 8, pp. 167–198. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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8.1 ­Introduction Because of the detrimental nature of genome instability and rearrangement in human cancers (reviewed in Mardis and Wilson (2009) and Hanahan and Weinberg (2011)) and developmental disease (reviewed in Weise et al. (2012)), it is frequently assumed that DNA sequences are fundamentally stable, at least in healthy tissues and organisms. It is further thought that cells within the same organism will contain generally the same genomic structure (with some exceptions, such as VDJ recombination in jawed verte­ brates, which is discussed in some detail in Section 8.12). Therefore reports of develop­ mentally programmed genome rearrangement in the soma of various organisms, starting in 1887 when Boveri described large‐scale chromatin elimination in the nema­ tode Parascaris univalens (Boveri 1887), have generally been dismissed as biological oddities. However, the accumulating examples are now sufficient to challenge the notion that genomes are normatively stable, at least in some groups. Indeed, large‐scale genome DNA elimination has been documented for at least ten species of parasitic nematode (reviewed in Tobler and Müller (2001)), for 18 of 30 examined copepod spe­ cies (Wyngaard and Rasch 2000), and in both living clades of jawless fish: hagfish (Kohno 1998) and lamprey, a vertebrate (Smith et al. 2009; Smith et al. 2012). The com­ plete elimination of a chromosome in somatic tissues has been reported in the bandi­ coot, a marsupial mammal (Hayman and Martin 1965), and the zebra finch (Pigozzi and Solari 1998). Pioneering studies of tissue‐ and cell‐specific somatic genome variation in humans and mouse (Abyzov et al. 2012; O’Huallachain et al. 2012; Shibata et al. 2012) highlight the plasticity of genomes within our own lineage. Ciliates have perfected the art of genome rearrangement as they keep separate ger­ mline and somatic nuclei within each cell, separating distinctive genetic architectures within each organelle. Due to their ease of culture, large cell size, and experimentally tractable biology, ciliates have emerged as important model systems to study the mech­ anisms controlling genome rearrangement. One of the most striking insights to emerge from these studies is the remarkable use of RNA to sculpt the genetic material. Two predominant RNA‐based pathways are known: longer noncoding template RNAs (Nowacki et al. 2008), as well as short RNAs that are known either as piRNAs (PIWI‐ interacting RNAs, analogous to those observed in mammalian germlines) (Aravin et al. 2006; Aravin et al. 2008; Fang et al. 2012; Zahler et al. 2012) or as scanRNAs (scnRNAs) (Mochizuki et al. 2002; Lepere et al. 2009).

8.2 ­Ciliates: Ubiquitous Eukaryotic Microorganisms with a Long Scientific History Ciliates are a group of widely distributed microorganisms that comprise a major com­ ponent of ocean plankton and play an important role in global marine food webs as consumers of bacteria and algae (Caron et al. 2012). They exist in both fresh and salt water, and though the vast majority may be still undescribed (Foissner et al. 2008), the estimated number of known species is currently around 4500 (Finlay 1998; Foissner et al. 2008). The genetic diversity within this ancient group of organisms is staggering, approximating the diversity that separates animals and plants (Prescott 1994; Parfrey et al. 2011). Even though they are composed of single cells, ciliates can become quite

8.2  Ciliates: Ubiquitous Eukaryotic Microorganisms with a Long Scientific History

large, in some cases exceeding 2 mm (Tartar 1961; Finlay 1998). They can exist as para­ sites (Coyne et al. 2011), although most feed on algae and bacteria, or other ciliates, and some harbor algal or bacterial symbionts (Preer 1969; Finlay 1998; Fujishima and Kodama 2012). They can exhibit extreme adaptations, including to anaerobic environ­ ments such as the intestines of cockroach (Ricard et al. 2008) and frog (Wichterman 1937). The remarkably varied cellular morphologies of ciliates (Finlay 1998) provide striking examples of the ‘endless forms most beautiful’ (Darwin 1859) that have long captured the imagination and ignited the curiosity of scientists. The origin of ciliate biology is properly traced to Antony van Leeuwenhoek (1632– 1723), a Dutch tradesman who created custom magnifying devices for his fabric busi­ ness, discovering a new world when he examined rainwater instead of cloth. He vividly described the ‘animalcules’ he discovered there (Dobell 1932; Corliss 1978; Fokin 2004) in reports to the Royal Academy. Van Leeuwenhoek’s observations ignited an entire field of research that reads as a scientific hall of fame. Among the early contributors to protozoology, in addition to van Leeuwenhoek himself, were Louis Pasteur, Ernst Haeckel, Camillo Golgi, and Otto Friderich Müller (Corliss 1978; Corliss 1979). For many years, the study of ciliates focused on morphology, inheritance, cell immortality, and the ways in which cytoplasmic determinants and maternal effects control mating type (Sonneborn 1977). The modern era of ciliate genetics originated in 1939 with Tracy Sonneborn’s seminal work elucidating the mating types of Paramecium aurelia (Sonneborn 1939)—a project that occupied several decades, culminated in the naming of 14 sibling species nearly 40 years later (Sonneborn 1975) (later expanded to the current total of 15 sibling species (Aufderheide et al. 1983)). While this work was progressing, another landmark develop­ ment occurred in 1952 at the Eel Pond at Woods Hole, Massachusetts, USA, with the isolation of conjugation‐competent Tetrahymena thermophila (originally T. pyriformis) as described by Nanney and Simon (2000)), which rapidly became one of the most use­ ful and studied ciliate models. In addition to two types of nuclei in each cell, ciliates perform extremely complex genomic rearrangements. Early ciliate researchers began to chronicle in detail the elaborate process by which one kind of genome (the germline) develops into another type (the soma) in Stylonychia and Euplotes (Kloetzel 1970; Ammermann 1971). Genetic comparisons of DNA from the two different nuclei, performed initially by David Prescott and co‐workers, uncovered a remarkable difference in sequence content, implying genomic diminution—in some cases, over 95% of the germline genome sequence is completely eliminated during the conversion to somatic genome (Bostock and Prescott 1972; Prescott et al. 1973; Lauth et al. 1976). Similar analyses in Tetrahymena (Yao and Gorovsky 1974) suggested that such genome rearrangements are universal among cili­ ates, differing in degree but not kind. However, at least two clades of ciliates, the Stichotrich and Phyllopharyngea classes, have added an extra layer of complexity: their germline genomes are often scrambled (Greslin et  al. 1989; Prescott 1994; Katz and Kovner 2010), a condition not observed to date with Tetrahymena or Paramecium (members of the class Oligohymenophorea). With the advent of next‐generation sequencing, these early studies are ripe for updat­ ing. The first complete ciliate germline genome sequence was recently obtained from Oxytricha trifallax (Chen et al. 2014) and compared to the somatic sequence from the same organism (Swart et al. 2013). These data provide the first comprehensive view of

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genome unscrambling, showing that, in addition to a 90% reduction in sequence com­ plexity, over 225,000 retained DNA segments are stitched together to make the somatic genome, and that the DNA segments can be interwoven in bafflingly complex patterns in the germline (Chen et al. 2014). Another study identified 45,000 sequences that are germline‐limited in Paramecium (Arnaiz et al. 2012) and whose removal occurs with­ out reordering DNA segments (i.e., unscrambling) as observed in Oxytricha (Chen et al. 2014).

8.3 ­Two’s Company: Nuclear Dimorphism in Ciliates Ciliates as a group are denoted by their hair‐like structures anchored in the cell cortex and by the fact that each vegetative cell has two distinct kinds of nuclei: they embody nuclear dimorphism. In practice, the number of nuclei in a cell can be considerably greater than two, as each type of nucleus can have multiple copies, and some species, such as Uroleptus, can even have hundreds of distinct nuclei, though only of two differ­ ent types (Inaba and Suganuma 1966; Prescott 1994). Regardless of the total number of nuclei, the underlying principle of nuclear dimorphism is retained: two types of nuclei are responsible for different aspects of cell function. The physically smaller micronu­ cleus (or MIC) functions as a germline within the single‐celled organism, and comprises the only DNA passed from parents to offspring in sexual reproduction. The physically larger macronucleus (or MAC), on the other hand, constitutes the soma, specialized for the production of proteins needed for cell growth and viability. During vegetative (or asexual) growth the MIC is thought to be transcriptionally silent and the MAC provides the RNA transcripts necessary for growth and reproduc­ tion. This conclusion is supported by the existence of viable amicronucleate ciliate cell lines (for examples see Dawson (1919) and Ammermann et al. (1989)) that, while able to reproduce asexually (Figure 8.1A), cannot undergo the sexual portion of the lifecycle (Figure 8.1B–G) since they have no germline. When (MIC‐containing) ciliates are gen­ tly starved of food they become reactive, allowing cells of compatible mating type to transiently cohere and enter into the sexual cycle (Figure 8.1B–G). Importantly, the sexual cycle does not result in reproduction—two cells enter and two cells exit—but it accomplishes genetic recombination and the replacement of the old MAC with a new (zygotic) one made from a genetic contribution from both cells (Figure 8.1B–G). The formation of the new (zygotic) MAC can be an exercise in extreme genome reduction, with some species eliminating 98% of their MIC genome sequences (Prescott 1994). This deleted sequence consists of repetitive elements, intergenic sequence, and spacer DNA elements termed internal eliminated sequences (IESs), which cripple most open reading frames in the MIC. The DNA segments retained in the new macronucleus are known as macronuclear‐destined sequences (MDSs). In the Stichotrich class of ciliates, which includes both Oxytricha and Stylonychia, the process of IES excision can fragment genes into 50 or more MDSs (Hoffman and Prescott 1996; Chang et al. 2005) and in Oxytricha a record‐holding 245 MDS gene has been reported (Chen et al. 2014). Scrambling patterns can be remarkably complex in Oxytricha, with examples of interwoven and reused MDSs among the total of 225,000 DNA segments that constitute the mature somatic nucleus (Chen et al. 2014). All these genic segments must be precisely reassembled to form functional genes in the new

8.4  Paramecium: Non-Mendelian Inheritance Comes to Light A

Reproductive asexual cycle

Non-reproductive sexual cycle

Mitosis of MIC

B

C

D

Amitosis of MAC n

io

at rv

d Fe

a St

Conjugation of compatible mating types

Meiosis of MIC

Reciprocal exchange of haploid MICs

MIC MAC

G

F

New MIC

zygotic (new) MAC

New diploid

E zygotic nuclei

H

old MAC (degrading) Late-stage “donut” cells

Division of diploid zygotic nuclei and differentiation of new MAC

Fertilization: formation of diploid zygotic nuclei

Figure 8.1  Simplified Oxytricha lifecycles. A: Reproductive asexual cycle. Cells divide by mitosis of the diploid germline micronucleus (MIC, indicated by a circle) and amitosis of the polyploid somatic macronucleus (MAC, indicated by an oval). B: In the laboratory, starvation induces conjugation between compatible mating types and initiation of a non‐reproductive sexual cycle. C: Meiosis of the MIC produces haploid gametic nuclei. D: Exchange of haploid micronuclei occurs. E: Fertilization produces genetically distinct, diploid zygotic nuclei. F: Zygotic nuclei divide mitotically and one differentiates into a new MAC, while one remains as a new MIC. G: Old MACs shrink as their genomic content is degraded, and these late‐stage cells are known as donut cells due to the very large, relatively transparent, developing zygotic MAC. H: At completion of the genome rearrangement process, the newly formed reproductive cell proliferates asexually (A) or can conjugate again (after a latency period), depending on nutrient availability.

macronucleus, producing a predominantly protein‐coding genome encoding approxi­ mately 18,000 proteins and comprised of at least 16,000 distinct chromosomes (Swart et  al. 2013). The resulting MAC nano‐chromosomes are tiny, usually contain only a single gene, have no centromeres, and are capped by short telomeres at their termini. These MAC chromosomes can be present at high ploidy, often thousands of copies (Heyse et  al. 2010; Nowacki et  al. 2010), probably to sustain the substantial protein synthesis required to support the very large sizes these cells can attain: Stylonychia can reach several hundred microns in length (Prescott 1994). The process of genome rearrangement in the oligohymenophorean ciliates Paramecium and Tetrahymena is less dramatic, resulting in loss of 10–30% of sequence complexity (Eisen et  al. 2006; Coyne et  al. 2008; Arnaiz et  al. 2012); see also Broad Institute, www.broadinstitute.org), but these cells still excise over 40,000 or 6000 IESs, respectively (Fass et al. 2011; Arnaiz et al. 2012).

8.4  ­Paramecium: Non-Mendelian Inheritance Comes to Light The Paramecium model system (in actuality a species complex consisting of 15 sibling species, of which P. tetaurelia is most studied (Sonneborn 1975; Catania et al. 2009)) provided early scientists with fertile grounds for analyzing inheritance. Tracy Sonneborn

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observed that mating type inheritance did not follow standard patterns of classical Mendelian genetics (Sonneborn 1937). Specifically, he found that there are two mating types, E and O, defined by an inability to mate with cells of the same type (Sonneborn 1937). Classical Mendelian inheritance would suggest that the initial (F1) progeny (from a mating of type E and type O parents) should be a mixture of the two mating types (with one being dominant and the other recessive), and the original mating types should re‐emerge in the next generation (F2). Instead, in P. tetaurelia, the progeny cell always retains the mating type of the parental cell. There appears to be no traditional ‘domi­ nant’ or ‘recessive’ categories or mixing of mating types (Nanney 1953). Instead, the macronucleus of the parental cell determines the mating type of the progeny cell and does not reflect the mating type of the conjugating partner cell. The new macronucleus is produced de novo by development from a fertilized zygotic nucleus during the sexual cycle (Figure 8.1, compare B and G), so the mating type of the old parental MAC is transmitted to the developing MAC genome through the cytoplasm. Parental‐determined inheritance in Paramecium was also observed in instances beyond mating type. One early example was the inclusion of an antigen gene in the MAC genome (Epstein and Forney 1984), but the effect extended to other genes as well (Scott et al. 1994). Experimentally manipulating the parental macronucleus by inserting genic DNA sequences which were originally absent could cause the retention of these sequences in the next generation’s MAC genome (Koizumi and Kobayashi 1989; You et al. 1991; Scott et al. 1994; Duharcourt et al. 1995; Duharcourt et al. 1998). Some IESs behave in the same way upon experimental introduction into the MAC (where they are not ordinarily found)—once present in the MAC they can template their own retention in offspring. IESs capable of inducing their own retention are ‘maternally controlled IESs’ (mcIESs), and approximately 1/3 of tested IESs have this property (Duharcourt et al. 1995; Duharcourt et al. 1998). A breakthrough discovery linked mating type in P. tetaurelia with a maternally con­ trolled IES‐like element (Singh et al. 2014). The IES‐like element encompasses both the transcriptional start‐site and ATG of the mating type A (mtA) gene, so it is required for gene expression—which leads to mating type E—while the absence of the IES‐like ele­ ment prevents gene expression and leads to mating type O (Singh et  al. 2014). Like mcIESs and exogenously introduced DNA sequence, the pattern of retention or exci­ sion in the parent is recapitulated in the progeny, making mating type a stable but parentally inherited phenomenon, as originally reported (Sonneborn 1937). It is important to note that in all cases, the parental MAC genome itself is not passed on to the progeny (it gets degraded), yet its structure—a genomic echo—controls the structure of the new, zygotic MAC. The echo must take the form of a signal from (old) somatic MAC to (new) somatic MAC, bypassing the germline genome, which is rele­ gated to supplying raw DNA but not its final configuration. (Curiously, mating type is determined completely differently in Tetrahymena thermophila: of seven different mating types, one is chosen at random in each progeny cell and is not linked to parental mating type (Cervantes et al. 2013). While noncoding RNA may be important in this process, this has not yet been experimentally demonstrated.) While the inheritance of mating type, IESs, and transgenes in P. tetaurelia are often parentally (or maternally) controlled (i.e., a sequence present in the MAC programs its retention in the progeny MAC), there are some curious exceptions that have revealed key mechanistic insights. In several cases, high‐copy transformation of the somatic

8.5  Tetrahymena and the Origin of the scanRNA Model

nucleus with a transgene caused the deletion of the corresponding endogenous genomic sequence in the next generation (Meyer 1992; Meyer et al. 1997). This apparent contra­ diction (relative to cases of parental control) was resolved by work showing that the eliminative effects were caused by the production of small RNA (or lack thereof ): if small RNAs were produced from the transgene, deletion of endogenous sequences occurred, while if no small RNAs were produced, the transgene caused retention and parental control (Garnier et  al. 2004). The proof of this model came in the form of experiments showing that simply introducing small RNAs, through feeding the Paramecium with Escherichia coli expressing double‐stranded RNA (Garnier et  al. 2004) or by microinjection (Lepere et al. 2008), was sufficient to induce deletion of cor­ responding sequences. Likewise, feeding Paramecium E. coli‐expressing small RNAs targeting the IES‐like element in the mtA gene was sufficient to induce its deletion in the next generation (Singh et al. 2014). The result was a conversion of a type E cell into a type O progeny that was thereafter stably maintained as type O in subsequent genera­ tions (Singh et al. 2014). Unified, these studies support a simple model (Figure 8.2A–C): The MIC produces ubiquitous endogenous small RNAs that guide deletions, and the MAC‐introduced DNA sequences can, through transcription into long noncoding RNA, absorb and block these endogenous small RNAs (thereby mediating retention and parental control) or, at high copy‐number, can generate their own small RNAs which mediate elimination of the corresponding sequence elements in the next generation.

8.5  ­Tetrahymena and the Origin of the scanRNA Model Meanwhile, several studies in the Tetrahymena model system provided additional sup­ port for the Paramecium studies described above, and these observations eventually led to the ‘scan RNA’ model (Mochizuki et al. 2002). The development of this model fol­ lowed a string of experimental results and observations mirroring those in Paramecium. Microinjection of DNA encoding an IES into the MAC of a parental Tetrahymena cell was sufficient to inhibit elimination of the homologous sequence in the developing MAC of the next generation, thus establishing maternal control of IES excision in this system (Chalker and Yao 1996). As in Paramecium, these epigenetic effects were stably inherited by future generations (Chalker and Yao 1996). Seminal work by Mochizuki et al. (2002) described a novel class of endogenous, approximately 28 nt small RNAs and showed that they associate with a small RNA‐binding protein: a member of the Argonaute family, from the PIWI clade (Hock and Meister 2008), known as TWI1. Knockdown of TWI1 led to the loss of the small RNAs as well as blocking the genome rearrangement process completely (Mochizuki et al. 2002). Furthermore, hybridization experiments showed that these small RNAs preferentially target eliminated MIC sequences (Mochizuki et al. 2002). These experiments led to the proposal of a scanning mechanism in which small RNAs originate from the entire germline genome, but over time those targeting somatic (i.e., macronuclear‐destined) sequences are selectively degraded (Figure 8.2A–C) (Mochizuki et al. 2002; Mochizuki and Gorovsky 2004; Chalker et al. 2005). Because they undergo this scanning and selection process, the small RNAs were termed scanRNAs or scnRNAs. Deep sequencing of small RNA pools during genome development has

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Figure 8.2  Summary of noncoding RNA (ncRNA) in ciliate genome rearrangements. Panels A–C represent nuclei simultaneously present in one Paramecium or Tetrahymena cell, while panels D–F represent nuclei present in one Oxytricha cell. A–C: ScanRNA model in Paramecium and Tetrahymena. ScanRNA biogenesis initiates in the micronucleus (MIC) (A), where all sequences are transcribed into long noncoding ncRNAs that are subsequently processed by Dicer‐like proteins into scanRNAs. B: Small RNAs are scanned against the parental macronuclear (MAC) genome via RNA–RNA interaction with long ncRNAs. Those that match are eliminated (indicated by scissors symbol) and only nonmatching scan RNAs are retained for use in the developing zygotic MAC (C). C: Elimination of IES sequence is accomplished by hybridization of scanRNAs to nascent ncRNAs which target chromatin modifying enzymes to eliminated sequences. D–F: piRNA and template model developed in Oxytricha. D: The MIC genomic sequence remains quiescent. E: Transcription of all MAC chromosomes produces templates covering all genomic sequence (labeled ‘templates’). PiRNAs are also produced, either from these sequences (as shown) or from independent precursor molecules (not shown). Both classes of noncoding RNA are imported into the zygotic MAC (F), where they play different functional roles. F: PiRNAs pair with nascent ncRNAs (produced by transcription of precursor DNA molecules) and direct retention of genomic sequences. Template RNA molecules guide segment reordering and ligation of cleaved DNA. In all panels, orange represents RNA, while DNA is indicated as a black double helix. Blue indicates the sequence to be eliminated in the new MAC, and can be encoded in DNA or RNA depending on species and developmental stage.

confirmed a selective loss of MDS‐targeting scanRNAs during Tetrahymena develop­ ment, although a biogenesis bias favoring eliminated sequences was observed (Schoeberl et al. 2012). Central to the scanning process is the involvement of long noncoding RNAs, produced from the parental MAC (Chalker and Yao 2001; Garnier et al. 2004; Aronica et  al. 2008) (Figure 8.2B), which act as molecular sponges for MDS‐targeting small RNAs and lead to their degradation. This natural genomic subtraction (Duharcourt

8.6  Small RNAs in Stylonychia and Oxytricha

et al. 2009) results in a small RNA pool that is enriched in molecules (scanRNAs) that target eliminated regions including IESs (Figure 8.2B and C). Confirming the central role for scanRNA in shaping the genome, Yao et al. (2003), found that microinjection of double‐stranded RNA into Tetrahymena cells during genome rearrangements was sufficient to induce the deletion of the corresponding DNA sequences. The microinjected double‐stranded RNAs are likely processed into small RNAs once introduced into the ciliate, where they may act in two ways to induce deletion: they may act as scanRNAs on their own, or they may feed into the RNA‐­ interference (RNAi) pathway (Holoch and Moazed 2015) to degrade the MAC long non­ coding RNA ‘sponges’ used in the scanning process (Figure 8.2B). The loss of the noncoding RNA ‘sponges’ would then prevent the destruction of scanRNAs targeting sequences for deletion. The net result of this ‘inhibition of the inhibitor’ is the release of endogenous scanRNAs targeting sequences for elimination (Figure 8.2B and C).

8.6 ­Small RNAs in Stylonychia and Oxytricha The development of the scanRNA model in oligohymenophorean ciliates led natu­ rally to the examination of other ciliates, particularly of the divergent Stichotrich class. The first study of a member of this class, Stylonychia lemnae, reported a broad agreement with the work in Paramecium and Tetrahymena (Juranek et al. 2005). A conjugation‐specific class of small RNAs was identified in these ciliates, which hybridize strongly to germline genomic DNA, and a conjugation‐specific PIWI‐­family protein, known as Macronuclear Development Protein 1 (MDP1) was identified as a key player in genome rearrangements (Fetzer et al. 2002; Paschka et al. 2003; Juranek et al. 2005). As in Tetrahymena (Mochizuki et al. 2002) and Paramecium (Bouhouche et al. 2011), inactivation of this protein leads to a lethal developmental arrest at an early stage (Paschka et al. 2003). However, in contrast to Tetrahymena and Paramecium, scanRNAs still are produced in the absence of the Stylonychia MDP1 protein (Juranek et al. 2005). The study of small RNAs in another stichotrichous ciliate, Oxytricha, reached broadly divergent conclusions (Fang et al. 2012; Zahler et al. 2012). This work initiated with the deep sequencing of a highly abundant class of conjugation‐specific 27‐nt small RNAs which exclusively target the MAC genome sequence, and thus appear to mediate reten­ tion and not elimination (Fang et al. 2012; Zahler et al. 2012) (Figure 8.2D–F). These startling observations were mechanistically verified by microinjection of artificial small RNAs designed to target IESs, leading to their retention in progeny cells (Fang et al. 2012). As was observed with Paramecium and Tetrahymena, these epigenetic IES‐ retention events were parentally inherited in the progeny of injected cells (Fang et al. 2012). As in the other ciliates described, a PIWI class protein, Otiwi1, was found to be essential for the genome rearrangement process, and its inactivation induced a deple­ tion of small RNAs coincident with developmental arrest and eventual cell death (Fang et al. 2012). Therefore these authors proposed a model in which small RNAs coat the retained regions of the genome (most likely by binding to noncoding RNA produced by transcription of the entire genome; see Figure 8.2E) and either block cleavage of DNA by a transposase protein (Nowacki et al. 2009) or mark the DNA for retention, possibly by chromatin marks of some type (Figure 8.2F).

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This striking ‘sign change’ of the small RNAs in Oxytricha relative to oligohymeno­ phorean ciliates makes economic sense. Oxytricha eliminates 90% of MIC sequences in the process of genome rearrangement (Lauth et  al. 1976; Prescott 1994; Chen et  al. 2014). Therefore small RNAs appear to target the minority category, whether elimi­ nated (10–30% for Tetrahymena and Paramecium) or retained (less than 10% for Oxytricha and Stylonychia).

8.7 ­Long Noncoding RNA Templates in Genome Rearrangement The precise excision of IESs in Oxytricha and Stylonychia raises a theoretical difficulty, as there appears to be insufficient information in the germline to specify exactly the pattern of rearrangements. Similar to Paramecium, Oxytricha and Stylonychia IESs are often very short (20–100 bp), and must be removed with basepair‐level accuracy because they interrupt protein coding regions (Greslin et  al. 1989; Eder et  al. 1993; Hoffman and Prescott 1996; Prescott and DuBois 1996; Gratias and Betermier 2003; Chang et al. 2005; Arnaiz et al. 2012). Paramecium has solved the informational prob­ lem by using specific motifs that clearly define IES boundaries (Klobutcher and Herrick 1995; Arnaiz et al. 2012) and are thought to guide the precise cleavage by transposase (Baudry et  al. 2009) and religation by the non‐homologous end‐joining pathway (Kapusta et al. 2011). Tetrahymena has taken a different approach to solving the problem: its IESs are nearly always in noncoding regions and their excision is generally imprecise, ranging over sev­ eral hundred basepairs (Austerberry et al. 1984; Austerberry and Yao 1988). DNA exci­ sion in Tetrahymena is controlled primarily by repressive histone modifications (Madireddi et al. 1996; Taverna et al. 2002; Liu et al. 2004; Liu et al. 2007) (discussed in greater detail in Section 8.9). Neither approach works for stichotrichous ciliates like Oxytricha and Stylonychia, whose IESs are short, often only dozens of basepairs (Eder et  al. 1993; Prescott and DuBois 1996). These short, precisely excised IESs lack the distinct Paramecium‐like sequence motifs at IES ends to guide their excision, and their tiny sizes make it unlikely for histones, which wrap approximately 150 bp of DNA each (Kornberg 1974) to pro­ vide the ‘missing’ information (Prescott and DuBois 1996). These considerations led several investigators to propose models of template‐guided genome unscrambling, where the templates come from the parental (and already unscrambled) MAC (Prescott et  al. 2003; Angeleska et  al. 2007). In these models, the templating information was suggested to take the form of long, MAC‐derived DNA or RNA molecules capable of basepairing with MIC DNA during unscrambling. Nowacki et al. (2008) experimentally tested and verified an RNA template model in Oxytricha (Figure 8.2F). Endogenous long noncoding putative template RNAs (both sense and antisense relative to the protein coding sequence) were detected for several chromosomes, expressed only during genome rearrangements (Nowacki et al. 2008). Degradation of these putative template molecules by RNAi resulted in a partial stalling of the genome rearrangement process and the accumulation of precursor molecules and partially rearranged products (Nowacki et  al. 2008). Furthermore, cytoplasmic microinjection of alternative template RNAs containing permuted MDS order was able

8.8  Long Noncoding RNA: An Interface for Short Noncoding RNA

to reprogram the endogenous rearrangement pathway to produce matching reordered MAC chromosomes (Nowacki et al. 2008). These aberrantly rearranged products were stably inherited in subsequent generations (Nowacki et al. 2008), proving that their con­ struction is not specified by any germline genomic sequences and highlighting the importance of somatic parent‐to‐progeny influence. The transfer of point substitutions from artificial microinjected RNA templates to endogenous DNA chromosomal sequences was also observed, particularly near gene segment (MDS) boundaries, pro­ viding a mechanism for non‐Mendelian inheritance of RNA polymerase‐induced sequence variation (Nowacki et al. 2008). In addition to controlling the structure of MAC chromosomes, template RNAs in Oxytricha and Stylonychia also regulate MAC chromosomal copy number (Heyse et al. 2010; Nowacki et al. 2010). As discussed previously, Stichotrich MAC chromo­ somes are polyploid, with an average chromosomal copy number in Oxytricha of around 2000 molecules (Prescott 1994). This copy number is fluid, with room for drift and adaptation (Steinbruck 1983; Baird and Klobutcher 1991; Harper et al. 1991). The experimental manipulation of RNA template abundances in both Oxytricha and Stylonychia caused matching changes in the copy number of the corresponding chro­ mosomes (Heyse et al. 2010; Nowacki et al. 2010). In Oxytricha the altered copy num­ bers were stably inherited in the next generation without further experimental manipulation (Nowacki et  al. 2010). Changes in copy number have been linked to altered gene expression (Xu et al. 2012), so the template‐guided changes in either gene sequence or copy number would be predicted to have evolutionarily selectable effects on the ciliate. The heritable influence of somatic (MAC) genome structure on the organism’s physi­ ology is a fascinating blend of Lamarckian acquired characteristics and natural selection à la Darwin: the variants produced are truly random with respect to fitness, but qualify as bona‐fide acquired characteristics in the Lamarckian sense. Any RNA‐driven somatic changes in gene sequence or abundance with positive effects on the fitness of the organ­ ism can be inherited and selected in a classical sense, but will never impact the sequence of the germline genome. In a very real way, the MAC constitutes an RNA‐driven somatic epigenome that has the capacity to sample sequence space more rapidly and efficiently than the germline genome alone, and can supply an overlay of heritable information that dynamically and heritably modulates the coding capacity of the organism at the time of genome rearrangement.

8.8 ­Long Noncoding RNA: An Interface for Short Noncoding RNA An emerging theme in ciliate genome rearrangement mechanisms is the central role of  noncoding RNA in shaping the genome. As discussed in the scanRNA section (Section 8.5), long noncoding RNAs are specifically produced in the maternal macronu­ clear genomes to provide docking and scanning sequences in both Tetrahymena and Paramecium (Chalker and Yao 2001; Garnier et al. 2004; Lepere et al. 2008) (Figure 8.2B); it is also likely that the scanRNA final interaction in the developing nucleus is also with nascent RNA transcripts (Aronica et  al. 2008) (Figure 8.2C), since the structure of double‐­ stranded DNA is likely resistant to pairing interactions with small RNA.

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However, the detection of noncoding RNAs from eliminated sequence in Paramecium or Tetrahymena is fundamentally ambiguous because according to the scanRNA model, they may be produced in the germline as precursors for scanRNAs, or may serve as docking sites for those scanRNAs in the developing (new) somatic nucleus. The nucleus of origin cannot be confidently ascertained from analysis of total RNA isolated during the genome rearrangement process. However, this problem is solved in Oxytricha, which produces no germline scanRNAs, instead generating piRNAs from the somatic genome (Figure 8.2D–F) (Fang et al. 2012; Zahler et al. 2012). This gives us an opportu­ nity to test for the presence of noncoding transcripts arising from eliminated sequence elements, as they cannot be confused with piRNA precursors. The author of this chapter therefore tested Oxytricha for such nascent transcripts (Figure 8.2F, small blue segment). Consistent with the model that noncoding RNAs serving as docking sites for small RNAs, conjugation‐specific transcripts encoding ­germline‐limited 170‐bp satellite repeats were observed (Figure 8.3) (Dawson et  al. 1984; Chen et al. 2014). Since these micronuclear sequences are not made into small RNAs in this organism (Figure 8.2D–F) (Fang et al. 2012; Zahler et al. 2012), the detected Agarose Northern 1

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Figure 8.3  Experimental detection of noncoding RNA (ncRNA) from eliminated micronuclear (MIC) regions during sexual cycle in Oxytricha (corresponding to the nascent ncRNAs shown in Figure 8.2F). An agarose northern membrane was probed with a MIC‐limited 170‐bp satellite repeat revealing noncoding RNAs ranging from several hundred basepairs to over 10,000 bp. The abundance of these ncRNAs peaks at 18 h and 24 h post mixing of complementary mating types JRB310 and JRB510. The membrane was stripped and reprobed with Actin1 as a loading control, and ethidium‐stained ribosomal RNAs (28S and 18S) from the gel prior to transfer are shown as additional load controls due to expression fluctuation in Actin1 during the sexual cycle.

8.9  Short RNA-Mediated Heterochromatin Formation and DNA Elimination

ncRNAs cannot be small RNA precursors and most likely are produced from the new (developing) somatic genome prior to DNA splicing (Figure 8.2F). Consistent with a role in piRNA‐docking, the expression profile of the ncRNAs detected in Figure 8.3 matches the expression of piRNAs precisely: they peak at 18 h and 24 h, declining strongly thereafter (Fang et al. 2012; Zahler et al. 2012). These data strongly support the model that noncoding RNAs are vital platforms for small RNA interaction in Tetrahymena, Paramecium, and Oxytricha—even though the small RNAs themselves arise from different nuclei and target different classes of genomic sequence for different functional outcomes (elimination or retention) (Figure 8.2). Pervasive transcription of the Oxytricha zygotic nucleus during conjugation provides a convenient mechanism to activate the expression of germline‐limited genes (such as telomere‐bearing element (TBE) transposases (Nowacki et al. 2009)) involved in the genome rearrangement pro­ cess, and the elimination of these elements may serve as a parsimonious means to switch them off when their service is no longer required (i.e., when the new somatic genome is fully formed).

8.9 ­Short RNA-Mediated Heterochromatin Formation and DNA Elimination The immediate after‐effects of scanRNAs targeting DNA sequences in ciliates were ini­ tially reported in Tetrahymena. The interaction of scanRNAs with the nascent noncod­ ing RNAs (described above) in the developing somatic genome results in a massive reorganization of chromatin, leading to elimination of the sequences targeted. Eliminated sequences are marked by repressive chromatin modifications including methylated his­ tone 3 lysine 9 (H3K9) and lysine 27 (H3K27) (Madireddi et al. 1996; Taverna et al. 2002; Liu et al. 2004; Liu et al. 2007). The key developmental histone‐marking enzyme is a histone methyltransferase homolog of Enhancer of Zeste (E(z)), known in Tetrahymena as E(z)‐like 1 (EZL1). In contrast to wild‐type cells, ΔEZL1 cells fail to eliminate DNA, do not produce H3K9 or H3K27 methyl modifications, and progeny cells do not survive (Liu et  al. 2007). Recently, these findings were extended to Paramecium, where both H3K27 and H3K9 methylation were found to occur in eliminated DNA, and to be deposited by an Enhancer of Zeste homolog, also named EZL1 (Lhuillier‐Akakpo et al. 2014). Histone methyl marks are also regulated by the Paramecium chromatin assembly factor PtCAF‐1 (Ignarski et  al. 2014). As in Tetrahymena, knockdown of PtCAF‐1 (Ignarski et al. 2014) or EZL1 (Lhuillier‐Akakpo et al. 2014) inhibited histone methyla­ tion (both K27 and K9), disrupted genome rearrangements and DNA elimination, and resulted in death of the progeny. In Tetrahymena two chromodomain proteins, Pdd1p and Pdd3p, recognize the repressive histone methyl marks and localize to electron‐dense chromatin elimination structures in the developing MAC (Madireddi et al. 1996; Nikiforov et al. 2000; Taverna et al. 2002). Confirming the role of Pdd1p in DNA elimination, Taverna et al. (2002) induced elimination of a DNA segment simply by tagging it with Pdd1p. These results elegantly demonstrate that Pdd1p is a key reader of methylated histones and that its presence is sufficient to cause DNA sequence elimination. In both Tetrahymena and Paramecium a number of novel proteins have been iden­ tified that are essential to the genome rearrangement process, some of which are

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chromatin‐associated proteins, though their specific roles remain unknown (Rexer and Chalker 2007; Yao et al. 2007; Matsuda et al. 2010; Arambasic et al. 2014). However, the molecular scissors for cutting the DNA have been identified, consisting of domesticated PiggyBac‐family transposase proteins for both Tetrahymena and Paramecium (Baudry et al. 2009; Cheng et al. 2010) (Figure 8.4C and D). Interestingly, the Oxytricha system utilizes a different transposase, from the Tc1/mariner family in its genome rearrange­ ment process, highlighting yet another lineage‐specific difference (Nowacki et al. 2009) (Figure 8.4B). It is also important to note that the Oxytricha transposase genes are pre­ sent in thousands of copies dispersed throughout the germline genome (Williams et al. 1993; Doak et al. 1994) (Figure 8.4B), while the Tetrahymena and Paramecium enzymes are resident in the somatic genome and occur as domesticated single gene copy genes without traditional hallmarks of transposon structure such as flanking terminal repeats (Baudry et al. 2009; Cheng et al. 2010) (Figure 8.4C and D). The repair of DNA sequences after IES excision in Paramecium and Tetrahymena involves homologs of the non‐homologous end‐joining (NHEJ) pathway such as Ku80, XRCC4, and Ligase IV (Kapusta et  al. 2011; Lin et  al. 2012). However, the molecular machinery used by Oxytricha and Stylonychia to rejoin DNA segments is currently unknown. Cytosine DNA methylation is a component of heterochromatin and is important for imprinting, development, and genome stability in most eukaryotes (Law and Jacobsen 2010). Indeed, in plants, histone 3 lysine 9 (H3K9) methylation is controlled by DNA methylation (Soppe et al. 2002). In ciliates, however, very little is known about the role of DNA methylation in marking eliminated sequences during genome rearrangements, and there have been conflicting reports on the matter. The absence of cytosine methylation has been reported in several early studies of Tetrahymena, Paramecium, Stylonychia, and Oxytricha (Gorovsky et  al. 1973; Cummings et al. 1974; Rae and Spear 1978; Ammermann et al. 1981), but these reports did not examine cells during the genome rearrangement process when developmen­ tally regulated heterochromatin is generated and DNA elimination occurs. DNA meth­ yltransferase inhibitory drugs affect recovery from germline nucleus destruction in Paramecium (Kwok and Ng 1989), but no methylated cytosine has been directly detected in this organism. Another study found that cytosine methylation plays a role in encystment of the distant relative to Paramecium, Tetrahymena, and Oxytricha, Colpoda inflata (Palacios et al. 1994), and a study of Stylonychia reported a low level of de novo cytosine methylation of transposons during genome rearrangements (Juranek et al. 2003). In a recent report, two Stylonychia genes (MDP1 and MDP2) appear to be transcriptionally repressed by a mere three or one methylcytosine residue(s), respec­ tively. The removal of these epigenetic marks allows developmentally regulated gene expression to occur (Bulic et al. 2013). In Oxytricha, we reported a dynamically high level of cytosine methylation and hydroxymethylation that is functionally required for elimination of various genomic elements (Bracht et  al. 2012), and we speculate that methylcytosine might play a role in marking DNA for elimination in other organisms as well (Bracht 2014). Studies of DNA methylation in Tetrahymena and Paramecium are needed to determine whether the link to elimination is conserved in ciliates, but an emerging theme is that several characteristics of heterochromatin (including repres­ sive histone modifications and cytosine DNA methylation) are associated with the ultimate genomic silencing in ciliates: DNA elimination (Bracht 2014). Given the ‘sign

8.9  Short RNA-Mediated Heterochromatin Formation and DNA Elimination

change’ of small RNAs in Oxytricha relative to Tetrahymena and Paramecium, one might predict that the Oxytricha small RNAs antagonize the deposition of methylation on the DNA rather than directing methylation of histones, as reported for scanRNAs in Tetrahymena. A. IES excision in Euplotes crassus Developmental Excision Transposase MIC TA

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IES 2

MAC

Figure 8.4  Overview of autonomous and non‐autonomous transposable elements in ciliates and other eukaryotes. Black boxes indicate telomeres, which are added de novo to ends of macronuclear (MAC) chromosomes, after micronuclear (MIC) chromosome fragmentation. Light‐green regions are either mobile elements or thought to have mobile element origins (e.g., IESs). Arrowheads on both ends of most green sequences represent inverted terminal repeats. Blue arrows indicate ORFs encoding transposase enzyme. Dark‐green triangles indicate direct repeats. Curved arrows indicate excision or mobilization activity of transposase enzymes. A: IES excision in Euplotes crassus. MIC transposase‐encoding Tc1/mariner elements (Tec elements) are thought to control excision of IESs (though experimental proof is lacking so the arrows are dashed). Tec elements and IESs are flanked by TA motifs and the same inverted terminal repeats. B: IES excision in Oxytricha trifallax. MIC‐encoded Tc1/mariner elements (telomere‐bearing elements, or TBEs) produce transposase required for excision of IESs. TBE elements are flanked by ANT motifs and strong (72‐bp) inverted terminal repeats, while IESs have extremely weak inverted terminal repeat tendencies only detectable in aggregate (Chen et al. 2014). IESs are generally flanked by direct repeats of 2–20 bp known as pointers; the 2‐bp‐long instances are biased toward TA (Chen et al. 2014). C: IES excision in Paramecium tetaurelia. A MAC gene (PiggyMac (PGM)) produces a PiggyBac family transposase required for IES excision. IESs are always flanked by TA and have inverted terminal repeats similar to the Euplotes Tec and IES motif. D: IES excision in Tetrahymena thermophila. As in Paramecium, a MAC gene (Tetrahymena PiggyBac 2 (TBP2)) produces PiggyBac family transposase that is required for IES excision. Most IESs are long (>500 bp), lack inverted terminal repeats and direct repeats, and imprecisely excised; however, a small number of IESs (not shown in figure) are short (< 500 bp), flanked by TTAA, and precisely removed (Fass et al. 2011). E: Many eukaryotes have both autonomous and non‐autonomous transposable elements analogous to ciliate transposon/IES systems. Autonomous elements are self‐mobilizing and encode functional transposase enzyme, while non‐autonomous versions (miniature inverted‐repeat transposable elements (MITEs)) lack the coding sequence for functional transposase and therefore rely upon the enzyme produced from autonomous elements to mediate their transposition. Source: Chen et al., 2014; Fass et al. 2011; Smit and Riggs 1996; Casacuberta and Santiago 2003; Wessler 2006; Yang et al. 2009. Reproduced with permission of Springer. (See plate section for color representation of this figure.)

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8  RNA-Mediated Somatic Genome Rearrangement in Ciliates Developmental Excision

C. IES excision in Paramecium tetaurelia MIC

TA

Tc1 / mariner family

TA

TA IES 1 TA

TA

IES 2 TA Transposase

MAC

TA

TA

PiggyMac (PGM) gene PiggyBac family (transposase gene only) Developmental Excision

D. IES excision in Tetrahymena thermophila MIC Tc1 / mariner family

IES 1

IES 2 Transposase

MAC

Mobilize

E. MITE elements

Tetrahymena PiggyBac 2 (TPB2) gene PiggyBac family (transposase gene only)

Transposase Autonomous Element

MITE MITE Nonautonomous Elements

Figure 8.4  (Continued)

8.10 ­Transposable Elements and the Origins of Genome Rearrangements The involvement of transposase proteins in DNA elimination in ciliates is an important clue to an ancient connection between genome rearrangements, transposable elements, and their host genomes (Baudry et al. 2009; Nowacki et al. 2009; Cheng et al. 2010). Jacobs and Klobutcher (1996) noted that the IESs of the ciliate Euplotes crassus contain inverted terminal repeat sequences (5′‐TATrGCRN‐3′) strikingly similar to the sequence observed at the ends of germline‐limited Tec transposons (5′‐TATAGAGG‐3′) (Jaraczewski and Jahn 1993) (Figure 8.4A), suggesting a functional (but currently untested) link. (Note: in this discussion, bolded font signifies the extremely strong con­ servation of the TA dinucleotide embedded in slightly variant and less tightly con­ strained 8‐bp motifs.) Furthermore, a very similar sequence (5′‐TAYAGYNR‐3′) could be found at the ends of Paramecium IESs (Klobutcher and Herrick 1995; Jacobs and Klobutcher 1996) (Figure 8.4C), suggesting broad conservation of this sequence across vast evolutionary distance. Tc1/mariner class transposons also exhibit a similar sequence at their termini (5′‐TACAGTKS‐3′) (Klobutcher and Herrick 1995; Jacobs and Klobutcher 1996). The TA dinucleotide (though not the full inverted motif ), which can be found flanking some IESs in ciliates beyond Paramecium and Euplotes

8.10  Transposable Elements and the Origins of Genome Rearrangements

(see below), may be significant because TA is the preferred target insertion site for Tc1/ mariner class transposases (Plasterk et al. 1999). These observations led to the hypoth­ esis that IESs originated as full transposons which invaded an ancestral ciliate germline genome (Klobutcher and Herrick 1997). Additional support for the transposon model for IES origins has accumulated since its initial proposal. At the time the transposon invasion model was proposed, Tetrahymena did not fit because it was thought that its IESs uniformly lacked TA or other recognizable motifs (Klobutcher and Herrick 1997), leading Jacobs and Klobutcher (1996) to suggest that three separate transposon invasion events may have occurred in ciliates. However, work (Fass et al. 2011) uncovered a new class of small (less than 500 bp) IESs in Tetrahymena that are flanked by TTAA repeats containing the embed­ ded TA dinucleotide. In contrast to most IESs in Tetrahymena (Figure 8.4D), these short IESs are precisely eliminated and can be found interrupting exonic sequence, whereas longer IESs are imprecisely excised and thus are relegated to noncoding regions of the genome. These data suggest that an ancient TTAA‐bound class of IESs may have since diversified into the non‐TA, imprecisely excised IESs that now make up the majority of Tetrahymena’s eliminated sequence elements. As described earlier, Oxytricha does not have a consistent motif at the ends of IESs, relying instead upon short direct repeat sequences known as pointers, which can be from 2 to 20 bp (Chen et  al. 2014) (Figure 8.4B). The two copies of the pointer are located at IES boundaries and one copy is always left in the somatic genome after rear­ rangement is finished (Figure 8.4B). Since pointers vary in sequence and length, the link between IESs and transposons was less clear in Oxytricha (though the functional role of a Tc1/mariner class transposase in IES excision is quite suggestive (Nowacki et  al. 2009)). However, our recent work on analysis of IESs genome‐wide has found that 2‐bp pointers are strongly biased toward TA (Chen et  al. 2014). It seems likely that in Oxytricha, TA‐flanked IESs, consonant with transposable element origins, were ances­ tral and have since diversified into the modern system, perhaps through a relaxation of selective pressures and the requirement that IESs be able to excise, but not necessarily to reinsert (Figure 8.4B). Taken together, these results unify all well‐studied ciliates and extend earlier findings to two difficult cases, Oxytricha and Tetrahymena (Klobutcher and Herrick 1995; Klobutcher and Herrick 1997). The transposon model of IES origins suggests that an ancient invasion of Tc1/mariner transposons in a common ancestor of all (molecularly characterized) ciliates was wildly successful, leading to a rapid invasion of the unsuspecting, unprepared host (Klobutcher and Herrick 1995; Jacobs and Klobutcher 1996; Klobutcher and Herrick 1997). This Tc1/mariner transposon then, for reasons unknown, became largely inactive for inser­ tion and most of the inserted elements decayed to produce excision‐competent IESs (Klobutcher and Herrick 1995; Jacobs and Klobutcher 1996; Klobutcher and Herrick 1997). Excision duties are most likely performed by germline‐encoded Tc1/mariner transposase proteins in Euplotes (Jacobs and Klobutcher 1996) and Oxytricha (Nowacki et al. 2009); however, excision in the lineage leading to Tetrahymena and Paramecium was taken on by a different (somatic) transposase, PiggyBac (Baudry et al. 2009; Cheng et al. 2010) (Figure 8.4C and D). The diversification of ciliates along independent evolutionary trajectories resulted in the wide variety of ciliate systems extant today (Figure 8.4A–D). This long independent evolutionary history has obscured the common origin of IESs as TA‐based, precisely

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excising mobile DNA elements. In support of this model, Tc1/mariner transposons can still be found in the germline genomes of Paramecium (Le Mouel et al. 2003; Arnaiz et al. 2012), Tetrahymena (Eisen et al. 2006), Euplotes (Jahn et al. 1993), and Oxytricha (Williams et al. 1993) (Figure 8.4A–D). It is important to note that the model does not preclude more recent, lineage‐specific transposon invasions: the Tc1/mariner family TBE transposons of Oxytricha appear to be a relatively recent invasion, not ancestral, suggesting a functional replacement by transposons of the same type in this lineage (Klobutcher and Herrick 1997; Zoller et al. 2012). While upon initial inspection the ciliate elaboration of IES excision seems unneces­ sarily baroque, complex transposon‐based systems have been uncovered in other eukaryotes that demonstrate surprising parallels. In both plants and animals, transpo­ son decay has been reported, producing complex systems of full‐length elements as well as truncated, still‐mobile fragments known as miniature inverted‐repeat transposable elements (MITEs) (Wessler et al. 1995; Smit and Riggs 1996; Casacuberta and Santiago 2003; Wessler 2006; Yang et al. 2009). The former encode functional transposase pro­ teins, while the latter truncated, shortened elements that are reliant, for mobilization, upon the transposase encoded in the former (Figure 8.4E). In these systems, the trans­ posons are divided into autonomous (full‐length transposase‐encoding elements) ver­ sus non‐autonomous elements (containing no protein‐coding regions, and often bearing little more than inverted terminal repeats) (Figure 8.4E). Interestingly, MITEs bear some striking similarities to IESs: they are short (less than 500 bp), are utterly dependent on transposase from full‐length elements for their mobility, and have inverted terminal repeats, as is the case for Euplotes and Paramecium IESs. The reason IESs in some ciliate species lack inverted repeats may be tied to the lack of selection pressure: excision might be achieved with very minimal sequence constraint, especially in systems such as Tetrahymena (Figure 8.4D) whose IESs are largely in noncoding regions and can be excised imprecisely. The interplay between transposable elements and genome rearrangements can be quite complex in ciliates. For example, in both Tetrahymena and Oxytricha there are micronuclear‐limited transposable elements containing telomeres at their ends but internal to the micronuclear chromosomes in which they are inserted (Cherry and Blackburn 1985; Herrick et al. 1985). In Tetrahymena these elements are known as Tel‐1 transposons (Cherry and Blackburn 1985), while in Oxytricha they are known as tel­ omere‐bearing elements (TBEs) (Herrick et  al. 1985) (Figure 8.4B). Importantly, the fact that they contain standard ciliate telomeres at the ends of the transposable ele­ ments suggests the existence of each as free linear forms for some period of time, dur­ ing which they are acted upon by the ciliate telomerase before being reintegrated into the germline genome. The details of this process remain only poorly understood, but the telomeres may allow the free linear form to persist in the developing MAC for a time, potentially undergoing amplification to high copy number like other MAC chro­ mosomes, before making a trans‐nuclear migration and reinserting into the MIC genome (Cherry and Blackburn 1985; Herrick et  al. 1985). Alternatively, the pre‐ encoded internal telomeres may serve to allow the transposon to survive transiently as a MAC chromosome without having extra telomeric sequence added to its ends after excision. Regardless, the implication is that these elements have interacted at some point with the cellular telomerase machinery before reintegrating into the germline genome. The existence of telomere‐bearing transposable elements implicates a

8.11  Transposons, Phase Variation, and Programmed Genome Engineering in Bacteria

potential crosstalk between the invasive DNA, the host telomerase machinery, and two different host nuclei.

8.11 ­Transposons, Phase Variation, and Programmed Genome Engineering in Bacteria Genome rearrangement is a survival strategy that has been observed in all domains of life (Haselkorn 1992; Jahn and Klobutcher 2002; Redder and Garrett 2006; Filee et al. 2007; Smith et al. 2012). Some bacterial species have adapted the strategy of relying upon relatively common (generally reversible) genetic alterations to change gene expression (van der Woude and Baumler 2004; Moxon et  al. 2006; Wisniewski‐Dye and Vial 2008). In these cases, frequently occurring reversible genetic rearrangements are utilized to generate ‘phase variation’ in which one or more genes are tabbed between active and inactive states that are stable for multiple generations. The envi­ ronment acts upon these phase variations to select the lineages best suited to survive and reproduce; the net effect is a type of differentiation within a single‐celled organ­ ism. For example, the pathogen Streptococcus pneumoniae displays phase variation in colony morphology: capsulated or non‐encapsulated forms. Because at least some of this phase variation is linked to stable, reversible alterations in the structure of genes controlling capsid synthesis (Waite et al. 2001; Waite et al. 2003), these bacteria might be thought of as quasi‐multicellular: specialized, stable cell types within the system of host and pathogen, optimized for either invasion and adhesion (non‐capsulated forms) or immune evasion and systemic infectivity (capsulated forms). There are many other examples of this sort of genetic alteration driving phase variation in bac­ teria: a genetic inversion forms a bi‐stable switch involved in flagellar antigen choice in salmonella (Zieg et al. 1977) and a modified transposable element functions as a promoter, controlling neighboring gene expression in E. coli by reversing its orienta­ tion (Berg 1980). In contrast to these randomly occurring rearrangements, more complex cases of pro­ grammed genome rearrangements also occur during development and differentiation in bacteria. The best‐known example concerns formation of heterocysts, specialized compartments for nitrogen fixation, in the filamentous cyanobacteria Anabaena (Golden et al. 1988; Haselkorn 1992; Carrasco et al. 1994; Meeks et al. 1994; Carrasco et al. 1995). Because nitrogen fixation is highly sensitive to oxygen, these bacteria have developed a differentiation program which alters some cells to generate anaerobic chambers, or heterocysts, in which nitrogen fixation can be carried out. This program of development involves genome rearrangement: excision of three IES‐like sequences that interrupt open reading frames of two key operons (two are shown in Figure 8.5). These developmental genome rearrangements are key switches to regulate expression of the nitrogen fixation genes. As in the ciliates, the genetic deletions occur only in specialized ‘somatic’ heterocyst structures that are short‐lived and non‐reproductive. As in Oxytricha and Stylonychia, the Anabaena genome recombination sites are speci­ fied by short direct repeats (Figure 8.5). Recombinase enzymes encoded within the inserts are responsible for excision, thereby removing themselves from the resultant product, similar to germline‐limited transposons that excise themselves during genome rearrangements in Oxytricha (Nowacki et al. 2009).

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8  RNA-Mediated Somatic Genome Rearrangement in Ciliates A. Anabaena Vegetative cell 11kb

55kb

xisA

xisF

= 5’- GCCTCATTAGG -3’ = 5’- TATTC -3’

B. Anabaena Heterocyst

transcription units

Figure 8.5  An example of developmentally programmed genome rearrangement in a prokaryote. The cyanobacteria Anabaena contains two operons with genes necessary for nitrogen fixation (shown in B). Both operons are interrupted by large (11‐ and 55‐kb) insertions that are developmentally excised upon differentiation into heterocysts and resemble ciliate IESs, since they interrupt protein‐ coding regions. A: Anabaena vegetative cell DNA contains the insertion elements. Open triangles represent direct repeats of 5′‐GCCTCATTAGG‐3′ occurring at breakpoint for the 11‐kb programmed deletion; filled triangles represent direct repeats of 5′‐TATTC‐3′ which demarcate boundaries of the 55‐kb element. As with Oxytricha (pointers) and direct repeats in other ciliates, one copy of the repeat is retained in the rearranged DNA molecule. XisA and xisF denote recombinases responsible for removal of 11‐kb and 55‐kb segments, respectively, and which are not present in product (heterocyst) DNA. B: Structure of heterocyst DNA containing two intact operons (shown as arrows underneath the DNA sequence). Both sequence insertions interrupt coding regions and each insertion inhibits expression of one operon in vegetative cells. Source: Redrawn from Haselkorn 1992.

8.12 ­Transposases, Noncoding RNA, and Chromatin Modifications in VDJ Recombination of Vertebrates The most famous example of somatic genome rearrangements in jawed vertebrates is the generation of an enormously diverse set of antigen‐binding molecules, the immu­ noglobulins and T‐cell receptors. These genes are generated by the recombination of specific cassettes that are encoded, and non‐functional, in the germline (Schatz and Ji 2011). The segments, denoted as variable (V), diversity (D), and joining (J), give rise to the nomenclature of V(D)J recombination, reflecting the structure of the final product of the rearrangement. While the final VDJ structure has only one cassette from each category (V, D, or J), there can be over one‐hundred V cassettes and five to ten D and J cassettes for the cell to choose from (Schatz and Ji 2011). The combinato­ rial possibilities are enormous, and account for the vast diversity of antibody recogni­ tion molecules generated by the immune system of jawed vertebrates. Interestingly, one recombinase protein (known as Recombination Activating Gene 1, or RAG1) appears to derive from an invertebrate transposase which has been domesticated by the cell in service to the immune system (Fugmann 2010). As in ciliates, programmed genome rearrangement—coupled with differentiation and development—is mediated by transposases. There are even parallels to bacterial phase variation (some of which involves transposons, also): The randomness of phase variation appears to have a more complex analog in the apparent randomness of the choice of VDJ cassettes in vertebrate immunology.

  References

In another parallel to ciliate biology, chromatin structure is the key to immunoglobu­ lin genome rearrangements. In addition to RAG1, the production of functional immu­ noglobulin genes requires a second protein, RAG2, which appears not to be transposase‐derived but instead comprises a chromatin ‘reader’ enzyme. RAG2 con­ tains a non‐canonical PHD domain that recognizes histone 3 lysine 4 trimethylation (H3K4me3), a hallmark of open chromatin (Matthews and Oettinger 2009; Schatz and Ji 2011). Echoing the role of noncoding RNA in ciliate genome rearrangements, tran­ scription of the VDJ region and production of noncoding RNAs has been widely reported (Abarrategui and Krangel 2007) and is a requirement for the rearrangement to occur, at least at the Jα locus (Abarrategui and Krangel 2006).

8.13 ­Concluding Remarks: Ubiquitous Genome Variation, Transposons, and Noncoding RNA Emergent themes surrounding genome rearrangements include the co‐option of trans­ posons by the host organism for specialized roles in DNA cutting and rejoining; the epi­ genetic marking of chromatin in regions to be rearranged; and the involvement of noncoding RNAs (both long and short) in the deposition of those marks. Evidence is accumulating that DNA damage responses in metazoa generally involve the production of short noncoding RNAs (Francia et  al. 2012). Epigenetic modifications have been reported in homologous recombination, which has been shown to induce localized de novo DNA methylation (Cuozzo et al. 2007). Somatic genome alterations are increasingly being described even in vertebrates (for a particularly dramatic example in sea lamprey see Smith et al. (2012), and his chapter in this book—Chapter 2). Emerging work demon­ strating widespread genome variation between tissues and cells of the same human‐being (Abyzov et  al. 2012; O’Huallachain et  al. 2012; Shibata et  al. 2012) is challenging our notion of ‘the’ human genome. Perhaps it is most accurate to speak of a human consensus sequence that varies in space (between cells, tissues, and individuals) and time (evolu­ tionary history): a human genome‐ome. More studies of the details of genome rearrange­ ment processes are needed, but the outlines of fundamental unifying principles are already visible: transposons, noncoding RNAs, and epigenetic marks have left an indelible stamp on organisms across the tree of life and will continue to dynamically shape organ­ ismal genome‐omes, both somatic and germline, in complex ways far into the future.

­Acknowledgments J.R.B. gratefully acknowledges the mentorship, guidance, and support of L.F. Landweber. In addition, he acknowledges the feedback from A.D. Goldman, J. Khurana, and X. Wang on drafts of the manuscript, as well as countless helpful discussions with all members of the Landweber laboratory. This work was supported by NIH grant 1F32GM099462.

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Wyngaard G.A. and Rasch E.M. (2000) Patterns of genome size in the copepoda. Hydrobiologia 417:43–56. Xu K., Doak T.G., Lipps H.J., Wang J.M., Swart E.C. and Chang W.J. (2012) Copy number variations of 11 macronuclear chromosomes and their gene expression in Oxytricha trifallax. Gene 505:75–80. Yang G., Nagel D.H., Feschotte C., Hancock C.N. and Wessler S.R. (2009) Tuned for transposition: molecular determinants underlying the hyperactivity of a Stowaway MITE. Science 325:1391–1394. Yao M.C. and Gorovsky M.A. (1974) Comparison of the sequences of macro‐ and micronuclear DNA of Tetrahymena pyriformis. Chromosoma 48:1–18. Yao M.C., Fuller P. and Xi X. (2003) Programmed DNA deletion as an RNA‐guided system of genome defense. Science 300:1581–1584. Yao M.C., Yao C.H., Halasz L.M., Fuller P., Rexer C.H., Wang S.H., Jain R., Coyne R.S. and Chalker D.L. (2007) Identification of novel chromatin‐associated proteins involved in programmed genome rearrangements in Tetrahymena. Journal of cell science 120:1978–1989. You Y., Aufderheide K., Morand J., Rodkey K. and Forney J. (1991) Macronuclear transformation with specific DNA fragments controls the content of the new macronuclear genome in Paramecium tetraurelia. Molecular and cellular biology 11:1133–1137. Zahler A.M., Neeb Z.T., Lin A. and Katzman S. (2012) Mating of the stichotrichous ciliate Oxytricha trifallax induces production of a class of 27 nt small RNAs derived from the parental macronucleus. PLoS one 7:e42371. Zieg J., Silverman M., Hilmen M. and Simon M. (1977) Recombinational switch for gene expression. Science 196:170–172. Zoller S.D., Hammersmith R.L., Swart E.C., Higgins B.P., Doak T.G., Herrick G. and Landweber L.F. (2012) Characterization and taxonomic validity of the ciliate Oxytricha trifallax (class Spirotrichea) based on multiple gene sequences: limitations in identifying genera solely by morphology. Protist 163:643–657.

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9 Mitotic Genome Variations in Yeast and Other Fungi Adrianna Skoneczna1* and Marek Skoneczny2 1 Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Laboratory of Mutagenesis and DNA Repair, Warsaw, Poland 2 Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Department of Genetics, Warsaw, Poland

Abstract During evolution, cells elaborated mechanisms responsible for maintenance of the stable genome. However, without mutations in the genome there would be no evolution, and persisting in changing environmental conditions without them would be hard or even impossible. Thus somatic variations appearing in cell populations contribute to adaptation and drive evolution. This chapter focuses on the mechanisms responsible for the two opposite mechanisms: genome maintenance and mutation tolerance in fungi. We refer to mechanisms ensuring fidelity of replication, detecting and repairing DNA damage, and assuring proper cell cycle and division progression. We present possible sources of genetic variation and mechanisms contributing to their appearance in the cell with respect to DNA sequence specificity, proficiency of cellular DNA quality control systems, metabolic stage of the cell, environmental stress, cell cycle phase, and ploidy. Keywords  genome maintenance; replication fidelity; DNA lesion; DNA repair; recom­ bination; spontaneous mutations; DNA rearrangement; chromosome instability; aneu­ ploidy; ploidy maintenance; ploidy shift; ploidy change; Saccharomyces cerevisiae; Schizosaccharomyces pombe; Candida albicans

9.1 ­Introduction Maintaining the stable genome is crucial for proper cell functioning and basically for cell survival. On the other hand, changes printed into the genome sequence are funda­ mental for adaptation to shifting environmental conditions, so they are the driving force of evolution. In every cell, even if not exposed to any genotoxic stress condition, spon­ taneous mutations arise that lead to genome variations. Spontaneous changes in DNA *Corresponding author: [email protected] Skoneczna A. and Skoneczny M. (2017) Mitotic genome variations in yeast and other fungi. In: Li X.‐Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley‐Blackwell, Hoboken, NJ, Ch. 9, pp. 199–250. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

200

9  Mitotic Genome Variations in Yeast and Other Fungi

sequence can be provoked by a wide range of factors, such as replication errors, error‐ prone endogenous DNA lesions repair, and aberrations during cell division. The type and frequency of the resulting mutations depends on the metabolic stage of the cell, its age, its ploidy, the actual phase of the cell cycle, effectiveness and specificity of DNA quality control systems, the DNA sequence in which the mutation occurred, and its chromosomal localization. From this point of view, yeast cells do not differ much from cells of higher eukaryotes. Both in simple unicellular organisms and in metazoan cells, the same or very similar mechanisms exist that assure genome maintenance and there are similar mechanisms that drive its instability. Many of them are found also in plant cells. Since they are evolutionarily conserved, the rules governing preservation of intact genome established for yeast are easily applicable to other cells. As yeast cells have been studied for many years to explore this field, a wealth of knowledge has been collected. Let us then take the lesson from yeast.

9.2 ­The Replication Process as a Possible Source of Genome Instability One of the best‐known sources of mutation appearing in the yeast genome, as well as in the genome of every eukaryotic cell, is the replication process. It is common knowledge that spontaneous mutation frequency is a derivative of fidelity of replication, which can be considered from three aspects: (1) the accuracy in nucleotide selection during DNA synthesis by DNA polymerases (Pol), (2) the removal of mis‐inserted nucleotides by the 3′→5′ exonuclease proofreading activity associated with DNA Pol, and (3) the activity of the mismatch repair (MMR) system, which has the ability of post‐replicative correc­ tion of Pol errors that have escaped proofreading. Yeast (Saccharomyces cerevisiae) has three DNA Pols engaged in replication of the nuclear genome: polymerase alpha‐­ primase (Pol α), polymerase delta (Pol δ), and polymerase epsilon (Pol ε). Pol α makes mistakes with the frequency 10–4 to 10–5, while the other two replicases are much more accurate in DNA synthesis, with an error frequency of 10–6 to 10–7, due to their proof­ reading activity enabling mis‐incorporation correction (Garcia‐Diaz and Bebenek 2007; Burgers 2009). Similar frequency of errors, 10–5 to 10–6, shows polymerase gamma (Pol γ) responsible for replication of the mitochondrial genome. After DNA synthesis, the MMR system can reduce the level of replication errors by another 10–3 per basepair (Kunz et al. 1998). Errors introduced during DNA synthesis are mostly point mutations: base substitu­ tions and frameshifts. Therefore, depending on the marker used to detect various point mutation reversions or the mutagenic events being considered, the observed mutation frequency falls in the range between 10–7 and 10–9 (Fedorova et al. 1998; Halas et al. 2002; McIntyre et al. 2006). Studies of mutagenesis spectra give comparable results. The rate of mutation events, such as transitions, transversions, and frameshifts in wild‐type yeast strains, is in the range of about 10–7 (Kunz et al. 1998; Ohnishi et al. 2004; McIntyre et al. 2007). Similar levels of spontaneous mutagenesis are observed in various species of fungi. When the level of mutagenesis is expressed as a mutation rate per basepair per replication (μb) it is 2.9 × 10–10 in budding yeasts S. cerevisiae, 3.2 × 10–10 in fission yeasts Schizosaccharomyces pombe, and 6.6 × 10–11 in Neurospora crassa cells (Drake 2009). The numbers are even more convincing when expressed as mutation rate per

9.2  The Replication Process as a Possible Source of Genome Instability

genome per replication (μg): 0.0037, 0.0044, and 0.0028, respectively. In fact, when the mutation rate is expressed per effective genome per replication (μeg) the result obtained for human cells, 0.004, does not vary from that of Escherichia coli or Caenorhabditis elegans cells, at 0.0025 and 0.004, respectively (Drake et al. 1998). Indeed, DNA replica­ tion is a quite accurate process as long as everything is going fine. However, when a problem occurs, the mutation rate will rapidly increase. 9.2.1  DNA Polymerases as Guardians of Genome Maintenance

In wild‐type yeast cells, during undisturbed replication, three Pols cooperate at the replication fork to synthesize nascent DNA. Despite their scheduled roles—Pol ε syn­ thesizing the leading strand, and primase Pol α and Pol δ synthesizing the lagging strand (Karthikeyan et al. 2000; Larrea et al. 2010, St Charles et al. 2015)–they work together at the very same foci (Hiraga et al. 2005), being able to substitute for each other (Garbacz et  al. 2015) or to fix each other’s errors (Pavlov et  al. 2004; Burgers 2009). However, replication asymmetry observed in budding as well as in fission yeast influences the fidelity of this process (Kunkel 2011; Miyabe et al. 2011). Errors can be introduced at two stages of DNA synthesis: during selection for correct dNTP by Watson–Crick basepairing and discrimination against each of the possible mismatched bases prior to covalent incorporation (Echols and Goodman 1991), or after incorporation of mis‐inserted bases as a result of balancing between synthesis and excision that compromises the rate of DNA extension (Kunkel and Bebenek 2000). In summary, replication error insertion is determined by: (1) base–base hydrogen bonding between nucleotides in nascent and template strands, (2) basepair geometry and substrate‐induced conformational change in Pol molecule, (3) strength of Pol interaction with the DNA minor groove at and upstream of the active site of Pol, which influences nucleotide selectivity, and (4) the efficiency of exonucleolytic proofreading and strand misalignment. Each particular Pol is influenced differently by each of these factors, which results in unique patterns of errors, or ‘fingerprints,’ imprinted into the area of synthesis (leading strand, lagging strand, gaps to fill, mitochondrial genome, etc.). The same factors influence other, non‐replicative Pols functioning in the yeast S. cerevisiae cells, except that these Pols are devoid of proofreading activity. Rev1, addi­ tionally, does not use canonical Watson–Crick basepairing, which leads to the lowest DNA synthesis accuracy among all yeast Pols (Nair et  al. 2005; Prakash et  al. 2005; Waters et al. 2009). Yeast replicative Pols work in complexes built from several subunits (Table 9.1). The major subunit of the complex is responsible for DNA polymerization and, optionally, for 3′→5′ exonuclease proofreading activity. In S. cerevisiae the accessory proteins con­ tribute to activity of the enzyme, its fidelity and processivity. Moreover, accessory subu­ nits can modulate Pol activities through specific interactions with a variety of cellular components. For example, Pol32, a non‐catalytic subunit of Pol δ, interacts with Pol30 and Pol1. Interaction with Pol30 determines Pol δ processivity. Pol30, called proliferat­ ing cell nuclear antigen (PCNA), forms a homotrimeric structure, the DNA Pol proces­ sivity factor, that works as a sliding clamp encircling DNA strand and tethering the Pol to the template, preventing its dissociation. Interaction of Pol32 subunit of Pol δ with Pol1 subunit of Pol α enables the cooperation between Pol α and Pol δ that is extremely important during lagging strand synthesis (Johansson et al. 2004).

201

S. cerevisiae

S. pombe

Gene

Pol ε

Pol δ

Pol α

SPCC737.07c

HCS1

POL2 / DUN2

POL32

d

pol2

cdm1

cdc27

cdc1

spp2

PRI2

POL31d

spp1

PRI1

cdc6

spb70

POL12

POL3/ CDC2

pol1

POL1/ CDC17

Replication fork DNA polymerases

DNA polymerase

POL2

POL32

HYS2

POL3 / CDC2

HCS1

PRI2

PRI1

POL12

POL1

C. albicans

POLE

POLD4

POLD3

POLD2

POLD1

PRIMA2

PRIM1

POLA2

POLA1

H. sapiens

DNA polymerase and 3′–> 5′ exonuclease, metal ion binding, interacts with Mrc1p

Interacts with PCNA and Pol1p

DNA polymerase and 3′→5′ exonuclease

ATP‐dependent 5′→3′ DNA helicase

Primase

DNA polymerase, interacts with Pol32p and Ctf4p

Molecular function a

Leading strand elongation, chromatin silencing at telomere, mitotic sister chromatid cohesion, MMR, NER, DSB repair via NHEJ

Lagging strand synthesis, MMR, BER, NER, PRR, chromatin silencing at telomere

Lagging strand DNA synthesis

Telomere capping

Replication initiation on leading and lagging strand (Okazaki fragments initiation), RNA‐dependent DNA replication, DSB repair

Cellular processes a

Table 9.1  Description of the genes encoding subunits of fungi DNA polymerase complexes and their orthologs in human cells.

G1/S

G1/S

Expression peaks

B

B

B

10–6– 10–7

10–6– 10–7

10–4– 10–5

Family Fidelity

RAD30

Pol η

MIP1

POL4/ POLX

Pol γ

Pol β

Other DNA polymerases

Pol κ

REV1

POL32d

pol4

pog1

dinB /mug40

eso1

rev1

rev7

REV7

POL31d

rev3

REV3

Rev1p

Pol ζ

MIP1

RAD32 / RAD30

REV1

REV7

REV3

DPB4

dpb4

DPB4

Translesion DNA polymerases

DPB3

dpb3

DPB3

DPB2

dpb2

DPB2

POLB

POLG2

POLG1

DINB1

POLH/ XPV / RAD30A

REV1

REV7

POLZ / REV3L

POLE4

POLE3

POLE2

DNA polymerase, metal ion binding b DNA‐directed RNA polymerase activity, 5′‐deoxyribose‐5‐ phosphate lyase b

DNA polymerase and 3′→5′ exonuclease

DNA polymerase

DNA polymerase, deoxycytidyl transferase, interacts with PCNA and Pol ζ

Interacts with PCNA and Pol1p

DNA polymerase, interacts with Rev1p, Rad30p, and Pol3p

Interacts with Isw2p

DNA binding

DSB repair via NHEJ, BER, gap filling

Mitochondrial DNA replication

X

A

Y

Translesion synthesis b

Y

B

Y

G2/M

G1/S

G1/S

Translesion synthesis, PRR, DSB repair

Translesion synthesis, PRR, DSB repair

Cell cycle control

Translesion synthesis, PRR

Replisome assembly

(Continued)

10–3– 10–4

10–5– 10–6

10–1– 10–2

10–4– 10–5

PAP2/TRF4

Pol σ

cid14

pol5

S. pombe

Gene

TCA5/ POL93

TRF4

POL5

C. albicans

TRF4

MybBP1A

H. sapiens

DNA polymerase activity, RNA‐ directed DNA polymerase, aspartic‐type endopeptidase, exoribonuclease H, structural molecule c

Poly(A) polymerase

Poly(A) polymerase, 5′‐deoxyribose‐5‐ phosphate lyase

DNA polymerase, rDNA binding

Molecular function a

BER

Nuclear polyadenylation‐ dependent RNA, catabolic process, BER

rRNA transcription, not required for replication

Cellular processes a

Expression peaks

B

B

B

Family Fidelity

All information listed in this Table is available at http://www.yeastgenome.org/, http://www.candidagenome.org/, and http://www.pombase.org/ online databases or has been published by Murakumo 2002; Gonzalez‐Barrera et al. 2005; Shcherbakova and Fijalkowska 2006; Garcia‐Diaz and Bebenek 2007; Burgers 2009; Waters et al. 2009; Johnson et al. 2012. The reader is referred to these sources, as well as references therein, for further details. a) Gene ontology annotations for S. cerevisiae protein. Orthologs in other fungi mostly play a similar role in the cell. In some cases more information is available for the gene product from other fungi than for its S. cerevisiae ortholog. b Gene ontology annotations for S. pombe. c) Gene ontology annotations for C. albicans. d) Pol31p and Pol32p are shared subunits for two DNA polymerases Pol δ and Pol ζ.

TRF5

POL5

S. cerevisiae

Pol ϕ

DNA polymerase

Table 9.1  (Continued)

9.2  The Replication Process as a Possible Source of Genome Instability

Accessory subunits of Pol δ complex (Pol31 and Pol32) and also all non‐catalytic subunits of Pol ε (Dpb2, Dpb3, and Dpb4) enable Pols interaction with different pro­ teins engaged in DNA repair pathways or regulating alternative Pols’ access to DNA template. Thereby they contribute to accuracy of DNA synthesis by influencing dNTP selection and/or proofreading efficiency of major catalytic subunits of respective com­ plexes (Huang et al. 2002; Giot et al. 1997; Jaszczur et al. 2008; Aksenova et al. 2010; Kraszewska et al. 2012). However, a growing amount of evidence indicates that acces­ sory subunits of Pols influence genome stability. For example, weaker interaction between Dpb2 and Pol2, as well as between GINS and Pol ε, leads to an increase in the frequency of spontaneous mutations (Kraszewska et al. 2012; Garbacz et al. 2015). Accessory subunits participate also in regulation of chromosome replication. It was shown that Pol1 interaction with chromatin‐binding protein Ctf4 associated with GINS complex couples Mcm2‐7 helicase to Pol α. This interaction is crucial to form proper replisome progression complex on DNA during new strand synthesis (Gambus et  al. 2009). On the other hand, Pol2 (Pol ε catalytic subunit) binds to the S‐phase checkpoint protein Mrc1 associated with Mcm2‐7 helicase, which permits coupling of polymerization and unwinding on the leading DNA strand of the replication fork during un­disturbed DNA synthesis (Lou et  al. 2008). During DNA damage stress, Mrc1 is phosphorylated in a Mec1‐dependent manner. Phosphorylated Mrc1 forms a pausing complex with Tof1 and is required for Pol2 stabilization at stalled replication forks (Osborn and Elledge 2003). Thus interactions of replicative Pols with Ctf4 and Mrc1 proteins protect cells against chronic activation of the DNA damage checkpoint during chromosome replication and permit finishing the cell cycle. In fact, almost all components of the replication fork machinery contribute to assuring replication accuracy. Mutations impeding the function of proteins engaged in initiation, elongation, or control of replication could lead to mutator phenotype. Indeed, genome instability phenotype has been reported for strains carrying defects in Mcm2‐7 helicase complex, origin recognition complex, replication factor C complex (RFC), alternative RFC‐like complexes, and other proteins participating in replication at different stages of this process, for example, Dpb11, Cdc45, Sld4, Ctf18, Csm3, or Pol30 (Stone et al. 2008; Alabrudzinska et  al. 2011; Li and Tye 2011; Stirling et  al. 2011; Cheng et  al. 2012). Destroying the replication machinery leads to replication stall or even fork collapse. Replication block can be passed through if recognized by a checkpoint kinase Mec1 and by subsequent direct stabilization of the replisome, or by activating the checkpoint response to regulate DNA repair engaging strand switching or origin firing, fork restart, and cell cycle progression (Friedel et al. 2009). In contrast to mutations arising because of defects in Pols, which are mainly base substitutions and frameshifts, the errors appearing due to perturbation in replication control mechanisms are mostly rearrangements. 9.2.2  dNTP Cellular Level and their Pool Bias Contribute to Genome Stability

Another source of possible replication errors is the availability of dNTPs in the cell. In S. cerevisiae cells, ribonucleotide reductase (RNR), the enzyme responsible for the rate‐ limiting step in dNTP synthesis, is tightly regulated; therefore the concentration of dNTP is highest in S and lowest in G1 phase. This fluctuation of dNTP concentration

205

206

9  Mitotic Genome Variations in Yeast and Other Fungi

controls cell cycle progression and the initiation of DNA replication (Chabes and Stillman 2007). Following DNA damage, genes encoding RNR subunits are induced and the RNR inhibitor Sml1 is phosphorylated and degraded in an Mec1/Rad53/Dun1‐ dependent manner, so dNTP production necessary for DNA repair and replication increases (Zhao and Rothstein 2002). It was shown that the constitutively high dNTP concentration transiently arrests cell cycle progression in late G1 phase. In G1 phase the dNTPs are usually low to enable assembly of DNA replication origins. High dNTP level in G1 phase results in defective pre‐replicative complex (pre‐RCs) activation based on the slow assembly of Cdc45, a component of the pre‐initiation complex, onto chroma­ tin. Data show that even a moderate increase in dNTP due to RNR overproduction resulted in synthetic sickness with orc2‐1 and orc5‐1 (the origin recognition complex mutants). Moreover, a high level of dNTP pool not only negatively affects activation of late origins of replication, but also inhibits the DNA damage checkpoint (Chabes and Stillman 2007). Exposure to hydroxyurea, an inhibitor of RNR activity, has the opposite effect on the dNTP pool. A reduced dNTP pool triggers cell‐cycle arrest in S phase. Transcription of RNR genes is controlled by checkpoint kinase Rad53 (a homolog of Sch. pombe cds1 and human CHK2; see Table 9.2) activated via its phosphorylation in response to DNA damage or replication block in an Mec1‐, Tel1‐, or trans autophos­ phorylation‐dependent fashion. This Rad53 activation, besides up‐regulation of dNTP levels, results in forks stabilization, inhibition of late‐origin initiation, and delayed entry to mitosis. Hence timing of Rad53 deactivation is important. Dephosphorylation of Rad53 is managed by type 2A‐like protein phosphatase activity of the Pph3–Psy2 com­ plex that allows DNA synthesis resumption. It was shown that Rad53 independently regulates replication fork restart and the late origins firing and that regulation of these processes is mediated by specific Rad53 phosphatases (O’Neill et  al. 2007). Cells deprived of Rad53, when treated with RNR inhibitor hydroxyurea, accumulate unusual DNA structures at replication forks, resulting in genome rearrangements. Therefore Rad53 prevents collapse of the fork and avoids genome destabilization when replication pauses (Lopes et al. 2001). Similarly to Rad53, impaired function of other checkpoint proteins responding to DNA damage or replication block (e.g., Mec1, Rad9, Tel1, Mrc1, and Ctf18) leads to elevated genome instability (Lustig and Petes 1986; Yuen et al. 2007; Lou et al. 2008; Razidlo and Lahue 2008; Alabrudzinska et al. 2011). As already described, changes in dNTP level as well as disturbed bias in the dNTP pool can provoke huge abnormalities in the cellular genome (Ouspenski et al. 1999; Chabes et al. 2003; Fasullo et al. 2010; Kumar et al. 2010; Kumar et al. 2011; Davidson et al. 2012). To avoid genome rearrange­ ments, several cellular processes were employed. In addition to cell cycle‐ and DNA damage‐coupled transcription control of RNR genes and RNR enzymatic activity inhi­ bition by Sml1 (Spd1 in Sch. pombe), as mentioned above, RNR activity is regulated by compartmentation of its subunits and by association of RNR full complex after relocali­ zation of its small subunits Rnr2 and Rnr4 from nucleus to cytoplasm in response to DNA damage signal (Yao et al. 2003). Moreover, it has been shown that thioredoxin and glutaredoxin are physiologically relevant electron donors for RNR during DNA precur­ sor synthesis; thus dNTP production can be influenced also by cytoplasmic thiore­ doxins (Trx1 and Trx2), glutathione reductase (Glr1), and gamma glutamylcysteine synthetase (Gsh1). In fact, trx1Δ trx2Δ cells are unable to accumulate dNTP and display elevated level of glutathione reductase (Koc et  al. 2006). In addition, cells employ

S. pombe

C. albicans

rad24

cdc45

swi3

drc1/sld2

BMH1 BMH2

CDC45/SLD4

CSM3

SLD2

SLD2

orf19.11586

CDC45

BMH1

Replication initiation and elongation factors

S. cerevisiae

Genes

Replication fork pausing

DNA replication origin binding, ssDNA binding, DNA strand annealing activity

RECQ4L b

DNA replication origin binding, chromatin binding

DNA replication origin binding, phosphoserine binding

TIPIN

CDC45

–

H. sapiens

Molecular function a

Mitotic cell cycle DNA replication checkpoint, DNA replication pre‐initiation complex assembly, DNA‐ dependent DNA replication initiation, DSB repair via BIR

DNA replication checkpoint, replication fork arrest, replication fork protection, establishment of mitotic sister chromatid maintenance of DNA repeat elements, meiotic chromosome segregation, mitotic sister chromatid cohesion, DNA repair

DNA replication initiation, pre‐RC assembly, DSB repair via BIR, regulation of chromatin silencing at telomere

DNA damage checkpoint, DNA replication initiation, pre‐RC assembly, negative regulation of ubiquitin‐protein ligase activity, Ras/MAPK signal transduction during pseudohyphal growth

Biological process a

(Continued)

Rothmund– Thomson, RAPADILINO, and Baller–Gerold syndromes

Velocardiofacial and DiGeorge syndromes

Human disease associated with the gene

Table 9.2  Fungal genes encoding proteins involved in genome maintenance that influence normal or disturbed replication and their human orthologs.

ORC complex

SCF complex

ubc15

rbx1

cul1

CDC34

HRT1

CDC53

orc1‐6

skp1

SKP1

ORC1‐6

pop2

hsk1 dfp1

CDC7 DBF4

CDC4

S. pombe

Genes

S. cerevisiae

Table 9.2  (Continued)

SKP1

FBXW7

CDC7 DBF4A or DBF4B

H. sapiens

ORC1‐6

CDC53

RBX1

ORC1/PARC1 ORC3/LATHEO ORC2,‐4,‐6

CUL1

RBX1

CDC34/UBC3 CDC34/UBE2R1

SKP1

CDC4

CDC7

C. albicans

DNA replication origin binding, ATP binding, ATPase activity, chromatin binding

Ubiquitin binding, F‐box protein DNA replication origin binding Ubiquitin‐ protein ligase activity RING finger protein Cullin

DNA replication origin binding Protein serine/ threonine kinase activity

Molecular function a

DNA replication initiation, pre‐RC assembly, chromatin silencing at silent mating‐type cassette

Protein complex assembly, kinetochore assembly, regulation of exit from mitosis, protein neddylation, SCF‐ dependent proteasomal ubiquitin‐dependent protein catabolic process, cytokinesis, septin ring assembly, vacuolar acidification

G1/S and G2/M transition of mitotic cell cycle

Negative regulation of exit from mitosis, DNA replication initiation, DSB repair via BIR, regulation of chromatin silencing at telomere, protein phosphorylation

Biological process a

Defects in ORC1, ORC4, and ORC6 are the cause of Meier–Gorlin syndrome type 1, 2, and 3, respectively

FBXW7 is mutated in breast cancer

Human disease associated with the gene

PSF1‐3 SLD5

RFC1‐5

POL30

RFA1‐3

GINS complex

RFC complex

PCNA

RPA

ssb1‐3

pcn1

rfc1‐5

psf1‐3 sld5

mcm2‐7

RFA1‐2

POL30

RFC1‐5

PSF1‐3 SLD5

MCM2,‐3,‐6, CDC54/MCM4 CDC46/MCM5 CDC47/MCM7

DNA damage and replication block checkpoint proteins

MCM2‐7

MCM helicase

RFA1, RFA3 RFA2 or RFA4

PCNA

RFC1‐5

GINS1‐4

MCM3,‐6 MCM2/CDCL1 MCM4/CDC21 MCM5/CDC46 MCM7/CDC47

Leading and lagging strand elongation, BER, NER, MMR, PRR, chromatin silencing at silent mating‐type cassette and at telomere, mitotic sister chromatid cohesion DNA unwinding involved in replication, NER, DSB repair via HR, reciprocal meiotic recombination, telomere maintenance via recombination and via telomerase, establishment of protein localization, heteroduplex formation, protein ubiquitination

dsDNA binding, ssDNA binding, sequence‐ specific DNA binding

Leading strand elongation, DNA repair, MMR

DNA replication, DSB repair via BIR

Pre‐RC assembly, DNA strand elongation involved in DNA replication, DSB repair via BIR

DNA polymerase processivity factor activity

DNA clamp loader activity

Unknown

DNA helicase activity, DNA replication origin binding, ssDNA binding, chromatin binding

(Continued)

GINS1/PSF1 and GINS4/SLD5 are up‐regulated in aggressive melanomas

S. pombe

cds1

crb2

RAD53

RAD9

Genes

S. cerevisiae

Table 9.2  (Continued)

RAD9

RAD53

C. albicans

BRCA1/RNF53 or TP53BP1 c

CHK2

H. sapiens

dsDNA binding, histone binding

DNA replication origin binding, protein serine/ threonine/ tyrosine kinase activity

Molecular function a

Intra‐S, G1/S, and G2/M transition DNA damage checkpoint, DNA repair, NER, positive regulation of transcription from RNA polymerase II promoter

DNA damage and replication block checkpoint, G1/S, intra‐S, and G2/M cell cycle arrest, deoxyribonucleoside triphosphate biosynthetic process, DNA repair, DNA‐dependent DNA replication initiation, nucleobase‐containing compound metabolic process, protein localization

Biological process a

BRCA1 defects cause breast cancer and pancreatic cancer type 4 susceptibility TP53BP1 aberration is found in myeloproliferative disorder chronic with eosinophilia

Sporadic cancer, familial breast cancer, Li– Fraumeni syndrome

Human disease associated with the gene

rad3

tel1

mrc1

MEC1

TEL1

MRC1

GIN1/MRC1

TEL1

MEC1

CLSPN

ATM

ATR

Interacts with Pol2p and Tof1

Protein kinase activity, telomeric DNA binding

Protein kinase activity

(Continued)

Breast cancer

Ataxia telangiectasia and B‐cell chronic lymphocytic leukemia, B‐cell non‐Hodgkin lymphoma, T‐prolymphocytic leukemia, colorectal adenocarcinoma Cell cycle checkpoint control in response to DNA damage, DNA damage‐induced protein phosphorylation, histone phosphorylation, DSB repair, telomere maintenance

S‐phase DNA replication and DNA damage checkpoint, DNA replication, replication fork protection, DNA repair, maintenance of DNA repeat elements, telomere maintenance, chromatin silencing at silent mating‐type cassette and at telomere, mitotic sister chromatid cohesion

Autosomal recessive disorder Seckel syndrome, cutaneous telangiectasia, and cancer syndrome, familial

Cell cycle checkpoint control in response to DNA damage or replication blocks, DNA replication, DNA recombination, reciprocal meiotic recombination, telomere maintenance, DNA damage‐induced protein phosphorylation, histone phosphorylation, nucleobase‐ containing compound metabolic process

ELG1 RFC2‐5

CTF18 CTF8 DCC1 RFC2‐5

Ctf18 RFC

DNA replication, DSB repair via HR, negative regulation of DNA recombination, negative regulation of transposition, RNA‐mediated, telomere maintenance, mitotic sister chromatid cohesion Response to DNA damage stimulus, DNA replication initiation, DSB repair via HR, maintenance of DNA trinucleotide repeats, mitotic sister chromatid cohesion

DNA clamp loader activity

CHTF18 CHTF8 DSCC1 RFC2‐5

CTF18 CTF8 DCC1 RFC2‐5

ctf18 ctf8 dcc1 rfc2‐5

DNA replication checkpoint, mitotic cell cycle G2/M transition DNA damage checkpoint, DNA replication initiation, leading and lagging strand elongation, MMR, NER, DSB repair via BIR, recombinational repair, mating type switching, positive regulation of protein phosphorylation

DNA clamp loader activity, chromatin binding

Protein kinase activator activity, protein binding

ATAD5 RFC2‐5

TOPBP1

H. sapiens

Biological process a

ELG1 RFC2‐5

DPB11

C. albicans

Molecular function a

elg1 rfc2‐5

rad4/cut5

DPB11

Elg1‐ RFC

S. pombe

Genes

S. cerevisiae

Table 9.2  (Continued)

CHTF8 expression is significantly reduced in renal and prostate tumors

Human disease associated with the gene

RNR

RNR1‐4

dNTP synthesis

cdc22, suc22

RNR1, RNR21/ RNR2 RNR22/ RNR2

–

Protein serine/ threonine kinase activity

Nucleotide binding, Deoxyribonucleotide biosynthetic process ribonucleoside‐ diphosphate reductase activity, thioredoxin disulfide as acceptor

CHK1

RRM1, RRM2 or RRM2B

DNA damage checkpoint, replication fork protection, protein phosphorylation

Transient G2/M arrest after DNA damage cell cycle checkpoint, replication fork protection, protein phosphorylation

chk1

Protein kinase activity

CHK1

DUN1

srk1

DUN1

DUN1

G1/S and G2/M transition checkpoint, intra‐S DNA damage checkpoint, pachytene checkpoint to inhibit cell cycle in response to unrepaired recombination intermediates, recombinational repair, DSB repair, reciprocal meiotic recombination

dsDNA binding

RAD9A RAD1 HUS1

DDC1 RAD17 MEC3

rad9 rad1 hus1

DDC1 RAD17 MEC3

9‐1‐1 complex

DNA damage checkpoint at G1, S, G2/M and meiotic checkpoints that monitor meiotic recombination, NER, NHEJ, reciprocal meiotic recombination, telomere maintenance through stimulation of Ty1 transposition

DNA clamp loader activity

RAD17 RFC2‐5

RAD24 RFC2‐5

rad17 rfc2‐5

RAD24 RFC2‐5

Rad24‐ RFC

(Continued)

Defects in RRM2B are the cause of mitochondrial DNA depletion syndrome type 8B

Overexpression in breast and colon cancers

MMR

msh2/swi8, msh3/swi4, msh6

mlh1 pms1

MLH1 PMS1

adk1

ADK1

MSH2 MSH3 MSH6

S. pombe

Genes

S. cerevisiae

Table 9.2  (Continued)

MLH1 PMS1

MSH2 MSH3 MSH6

ADK1

C. albicans

MLH1 PMS2

MSH2 MSH3 MSH6

ADK1

H. sapiens

ATP binding, ATPase activity, ssDNA, dsDNA, loop DNA, and dinucleotide insertion or deletion binding

ATP binding, ATPase activity, four‐way junction DNA binding, mispair binding, single nucleotide insertion/ deletion binding, recognition of mismatch

Adenylate kinase

Molecular function a

Defects in MSH2 are the cause of hereditary non‐polyposis colorectal cancer type 1 MSH6 mutations cause colorectal/ endometrial cancer and hereditary non‐polyposis colorectal cancer type 5 Mutation in MLH1 causes hereditary non‐polyposis colorectal cancer; mutation in PMS1 causes mismatch repair cancer syndrome

MMR, meiotic MMR, DNA recombination, mitotic recombination, meiotic gene conversion, chromatin silencing at silent mating‐type cassette, removal of nonhomologous ends

MMR, meiotic MMR, meiotic heteroduplex formation, reciprocal meiotic recombination

DNA‐dependent DNA replication initiation, ADP biosynthetic process, nucleotide metabolic process

Biological process a

Human disease associated with the gene

PRR

RNase H2

RER

rph6

rad2/fen1

FEN1/RAD27

RAD6

rnh201

exo1/mut2

RNH201‐203

EXO1

RAD6

orf19.547

RNH35

EXO1

UBE2A/HR6A or UBE2B/HR6B

FEN1/RAD2

AGS3

EXO1/HEX1

Ubiquitin‐ conjugating enzyme Ubiquitin‐ protein ligase activity

5′→3′ exonuclease activity, 5′‐flap endonuclease activity

Ribonuclease H activity

5′→3′ exonuclease activity, 5′‐flap endonuclease activity

(Continued)

Defects in UBE2A are the cause of mental retardation syndromic X‐linked Nascimento‐type

Huntington disease Fragile X disorder

DNA replication, removal of RNA primer, BER, base‐free sugar‐phosphate removal, DSB repair via NHEJ, gene conversion at mating‐type locus, replicative cell aging Mitotic cell cycle G1/S transition DNA damage checkpoint, error‐free PRR, error‐free and error‐prone TLS, chromatin silencing at telomere, DSB repair via HR, meiotic DSB formation, protein monoubiquitination, histone monoubiquitination, protein polyubiquitination, protein ubiquitination involved in ubiquitin‐dependent protein catabolic process, ubiquitin‐ dependent protein catabolic process via the N‐end rule pathway, regulation of dipeptide transport, transcription from RNA polymerase II promoter

Aicardi–Goutieres syndrome

DNA replication, removal of RNA primer

Gene conversion at mating‐type locus, DSB processing (also meiotic), MMR, telomere maintenance, telomeric 3’ overhang formation

S. pombe

rph18

rad8

mms2

RAD18

RAD5

MMS2

Genes

S. cerevisiae

Table 9.2  (Continued)

MMS2

RAD5

RAD18

C. albicans

MMS2/UBE2V2 or UBE2V1

SHPRH or HLTF

RAD18/RNF73

H. sapiens

Ubiquitin‐ conjugating enzyme Ubiquitin‐ protein ligase activity

DNA‐dependent ATPase activity, Y‐form DNA binding, four‐way junction DNA binding, four‐way junction helicase activity, RING finger containing ubiquitin ligase

ssDNA binding, contributes to ssDNA‐ dependent ATPase activity, ubiquitin‐ protein ligase activity

Molecular function a

Response to DNA damage stimulus, free ubiquitin chain polymerization, protein polyubiquitination, PRR

DSB repair, PRR, free ubiquitin chain polymerization, protein polyubiquitination

Error‐free PRR, error‐free and error‐prone TLS, protein monoubiquitination

Biological process a

SHPRH mutations are found in melanoma and ovarian cancer cell lines

Human disease associated with the gene

pli1

hus5

SIZ1

UBC9

UBC9

SIZ1

UBC13

UBE2I/UBC9

–

UBE2N/BLU

SUMO ligase activity

SUMO ligase activity

Ubiquitin‐ conjugating enzyme

G2/M transition of mitotic cell cycle, mitotic spindle elongation, protein sumoylation

Chromosome segregation, protein sumoylation

Free ubiquitin chain polymerization, protein polyubiquitination, PRR

All information listed in this Table is available at http://www.yeastgenome.org/, http://www.candidagenome.org/, http://www.pombase.org/, and http://www.uniprot. org online databases and the reader is referred to these sources, as well as references therein, for further details. a) Gene ontology annotations for S. cerevisiae protein. Orthologs in other fungi mostly play a similar role in the cell. b) Homology between fungi SLD2 and human RECQ4L gene is limited to the N‐terminal part of the protein coding region, but not the RecQ helicase region. c) There is no single ortholog of S. cerevisiae RAD9 gene in human; instead a family of proteins exist that contain tandem BRCT domains, and among them are BRCA1 and TP53BP1.

ubc13

UBC13

218

9  Mitotic Genome Variations in Yeast and Other Fungi

threonine metabolism to buffer deficiency in RNR by enabling a compensatory increase in de novo purine biosynthesis that provides additional rate‐limiting substrates for dNTP production (Hartman 2007). The interaction between PCNA associated with DNA and Spd1 has been documented. Spd1 is the allosteric regulator of RNR in Sch. pombe. Interaction with PCNA is required for Spd1 degradation by ubiquitin ligase CRL4Cdt2 (Salguero et al. 2012). This finding places RNR directly in the replication fork. 9.2.3  Mismatch Repair (MMR) and Ribonucleotide Excision Repair (RER) Are Used to Clean‐up after Replication

When deoxynucleotide mis‐inserted during DNA synthesis endures proofreading activity of replicative Pols, the only way to avoid subsequent mutation fixation is the  activity of the MMR system. The first step of repair in the yeast MMR system (Spampinato et  al. 2009) depends on specialized protein complexes, Msh2–Msh3 or Msh2–Msh6, which recognize structural abnormalities of DNA helix caused either by  mis‐paired bases and one nucleotide insertion/deletion or by a few nucleotide insertions/­deletions, respectively. Then the nascent and template DNA strands are rec­ ognized and marked via asymmetric binding of Mlh1–Pms1, Mlh1–Mlh2, or Mlh1– Mlh3 complexes (nascent strand is discriminated by its discontinuity associated with DNA replication) and the distortion‐containing segment of the nascent strand can be removed by flap endonucleases (mainly Exo1), providing another chance for Pols to synthesize DNA using the original template. The final step of this repair pathway is ligation. Other components that participate in MMR are: RPA, heterotrimeric complex binding DNA, RFC complex, the PCNA loading clamp, and PCNA itself, serving as a platform giving access to DNA for all proteins involved in the repair. It should be noticed that the Msh2–Msh6 complexes and all three Mlh1–heterocomplexes function also in the resolution of recombination intermediates, and bind to a cross‐shaped four‐single strands structure named Holliday junctions, with a high affinity contributing to recip­ rocal meiotic recombination (Marsischky et al. 1999; Argueso et al. 2003). Besides mis‐paired bases, the MMR system can efficiently correct diverse frameshifts. This kind of mistake is difficult to correct using proofreading activities of Pols, espe­ cially at the polynucleotide tracts. Such DNA sequences are hot‐spots for mutagenesis during DNA synthesis. Frameshifts appear frequently at the polynucleotide tracts due to Pol slippage on DNA strand during replication (Kunkel 1990), or due to template misalignment mechanisms: primer–template misalignment (Bebenek and Kunkel 1990) or dNTP‐stabilized misalignment (Efrati et al. 1997; Kobayashi et al. 2002). The likeli­ ness of Pol slippage is correlated with the tract length. Saccharomyces cerevisiae studies suggested a four nucleotide tract length threshold for Pol slippage to occur (Lehner et al. 2012). However, not only mononucleotide tracts or defects in the MMR system can lead to frameshifts, but also some role in initiating primer–template misalignment is played by DNA regions flanking mononucleotide tracts. Furthermore, it was docu­ mented that non‐homologous end‐joining pathway (NHEJ) defects contribute to de novo creation of tandem duplications from non‐iterated sequence (Lehner et al. 2012). The results obtained recently for S. cerevisiae and Sch. pombe cells suggest one more class of replicative errors resulting from ribonucleotide monophosphate (rNMP) incor­ poration to the nascent DNA. These errors are very common, they happen on average once per 2 kb of newly synthesized DNA, and they are so frequent because of high

9.3 ­Post-Replicative Repair (PRR) or Homologous Recombination (HR)

cellular levels of ribonucleotide triphosphates (rNTPs), even though their incorpora­ tion is mostly limited by the selectivity of DNA replicases. However, if such mis‐­ insertions persist after the DNA synthesis step, they cannot be corrected by MMR, but require special enzyme RNase H2 for repair. RNase H2 is normally used for removing RNA primers from Okazaki fragments, but as already shown, it is also responsible for incision of the ribonucleotide, the initiation step of ribonucleotide excision repair (RER). During the RER pathway the incision step is followed by excision by the flap endonuclease Fen1, or with lower efficiency by Exo1, then by strand displacement syn­ thesis carried out by DNA Pol δ or Pol ε, and finally completed by DNA ligase I (Sparks et al. 2012). The PCNA clamp and its loader RFC also participate in RER. PCNA inter­ acts with all proteins engaged in the repair pathways and gives proteins access to the replication foci (Sparks et al. 2012). In RNase H2‐deficient strains, rNMP incorporated into the leading strand can be removed by processing outside the context of replication in a manner that requires Top1 and repetitive sequence which gives rise to misaligned intermediates and results in short deletions (Clark et  al. 2011; Kunkel 2011; Miyabe et al. 2011). Another way to overcome rNMP incorporation into DNA in RNase H2‐ deficient strains is to use post‐replicative repair (PRR) pathways. It has been shown that in such strains, PCNA is constantly mono‐ or poly‐ubiquitinated, so PRR is constitu­ tively activated, and mis‐incorporated ribonucleotides can be efficiently bypassed by Pol ζ or omitted in an Mms2‐dependent template switch manner (Lazzaro et al. 2012).

9.3 ­Post-Replicative Repair (PRR) or Homologous Recombination (HR) Are Responsible for Error-Free and Error-Prone Repair of Blocking Lesions and Replication Stall-Borne Problems All the processes described in section 9.2, when they work properly, contribute to main­ tenance of the stable genome, and only the malfunction of that cellular machinery or particular circumstances can provoke formation of replication‐dependent mutations. Yet there are processes within the cell that actively generate mutations. These error‐ prone processes are involved in replication over DNA damage in the template. DNA lesions appear frequently as a consequence of endogenous or exogenous stresses, but as long as they are recognized and corrected by appropriate repair pathways, they do not cause any problems. They became mutagenic if they persist until the next DNA replica­ tion round. Unrepaired damage in the template strand causes difficulties during DNA synthesis. The type of DNA lesion determines the selection of the appropriate troubleshooting pathway (Figure 9.1). Some of them can be bypassed by replicative Pol, some need spe­ cialized Pol to be overcome (trans‐lesion synthesis), some of them can be bypassed through DNA template change (damage avoidance), and some can block replication completely causing permanent fork stall, then cell cycle arrest, and finally cell death. The repair pathway dealing with these problems is named post‐replicative repair (PRR). In yeast S. cerevisiae two PRR subpathways exist—an error‐prone one and an error‐free one—and PCNA functions as a molecular switch between them by virtue of alternative covalent modifications (Hoege et  al. 2002; Andersen et  al. 2008; Zhang et  al. 2011).

219

220

9  Mitotic Genome Variations in Yeast and Other Fungi Template specificity (sequence, structure) DNA Pol mutation Change in dNTP pool level or balance dNTP damage rNTP access Endogenous or exogenous genotoxic stress Replication disturbance

Misinsertion Misalignment rNMP insertion DNA damage DNA break Replication block Abnormal DNA structures

Problem detection (e.g., DNA Pols, RNaseH2, Msh2-Msh3, Msh2-Msh6, and MRX complexes)

Checkpoint activation (e.g., Dbf4, SId2, Mrc1, Ctf18, Rad24)

Cell cycle delay

Unrepaired damage Unsolved replication fork block Perturbation in cell division

Signaling (e.g., Mec1, Tel1, Rad53, Rad9)

Recruitment of specialized accessory factors to DNA alteration site (e.g., RFC or RFC-like complexes, PCNA, cohesion complex, DNA repair proteins, DNA Pols)

DNA repair (different proteins depending on activated pathway, e.g., MMR, RER, NER, BER)

DNA damage response

Transcription activation (e.g., damage inducible genes) Protein modification (e.g., chromatin remodeling, PCNA ubiquitilation) Protein relocalization (e.g., RNR subunits trafficking)

Trans-lesion DNA synthesis (recruitment of specialized DNA Pol to the cognate substrate at the replication fork where DNA synthesis has been perturbed)

DNA damage avoidance (mechanisms using HR machinery, template switching, or fork regression)

Cell cycle arrest

Mitotic cell death

Figure 9.1  DNA perturbations, triggered by various events, induce a multitude of molecular mechanisms that lead to damage repair, mutagenesis, or cell death.

During undisturbed replication, PCNA is sumoylated, which permits interaction with replicative Pol δ and Pol ε and fast replisome progression at the replication forks. Detection of bulky lesions on DNA template impedes fork progression and induces Rad6–Rad18‐mediated ubiquitination of PCNA, therefore promoting Ubi–PCNA‐ dependent trans‐lesion DNA synthesis (TLS), or allowing further polyubiquitination of Ubi–PCNA performed by the Ubc13–Mms2–Rad5 ubiquitin ligase complex that in turn favors the damage avoidance pathway (Figure 9.2). After passing through the lesion, Ubp10 deubiquitinates Ubi–PCNA, allowing replisome remodeling, switching back to replicative Pol and effective replication resumption (Gallego‐Sanchez et  al. 2012). Desumoylation of PCNA is performed by SUMO proteases, Ulp1 that removes SUMO and Ulp2 that removes poly‐SUMO chains (Stelter and Ulrich 2003; Huang et al. 2007; Drag and Salvesen 2008).

9.3 ­Post-Replicative Repair (PRR) or Homologous Recombination (HR) DNA damage avoidance: sister chromatid junction or fork regression

Inhibition of sister chromatid cohesion

Ubp10

Prevention from Ecol binding

polyUbi diUbi

Rad5/ Mms2/Ubc13 Ubp10

SUMO

Ulp1/(Ulp2?)

Ubi

SUMO diSUMO

Rad6/Rad18

Ubc9/Siz1

Specialized Pols binding

Srs2 binding Replicative Pols binding RNR stimulation Trans-lesion synthesis HR inhibition Replication over the lesion

Figure 9.2  Controlling of DNA repair pathways by various covalent modifications of PCNA.

In Sch. pombe the information concerning PRR is limited. It is known that Sch. pombe and S. cerevisiae share the homologous sets of PRR genes (see Tables 9.1 and 9.2), but they are used in different ways. The major distinctions of Sch. pombe cells in the control of lesion tolerance pathways are: existence of pathways that involve several TLS Pols, requirement for TLS of enzymes needed for mono‐ and poly‐ubiquitination of PCNA, and, surprisingly, mostly error‐free bypass of different DNA lesions no matter which Pol or Pols synthesize through them (Coulon et  al. 2010). Although Siz1 and Ubc9 homologs exist in Sch. pombe, PCNA sumoylation has not been documented (see Table 9.2). 9.3.1  Sumoylated PCNA‐, Srs2‐, and Replicative Polymerase‐dependent DNA Synthesis on Damaged Template

In budding yeast, SUMO (Smt3) is conjugated to PCNA by the Ubc9–Siz1 complex. The major SUMO acceptor site, which may be also subject to ubiquitination, is Lys164, and this residue is crucial for DNA damage tolerance. There is an additional, minor sumoylation site at Lys127 which does not appear to affect the DNA damage tolerance

221

222

9  Mitotic Genome Variations in Yeast and Other Fungi

activity, but is required for the establishment of sister chromatid cohesion during S phase (Hoege et al. 2002; Stelter and Ulrich 2003; Moldovan et al. 2006). Since PCNA forms a homotrimeric complex, there are three Lys164 residues that could be poten­ tially modified. It has been shown that sumoylation of one Lys164 facilitates modifica­ tion of the next one in the complex, but the ligand for subsequent modification may not necessarily be the same as the primary one. Therefore, in one complex, different modi­ fications can exist, including poly‐SUMO chains (Windecker and Ulrich 2008). It is also important to mention that differently modified PCNA and all subpathways of PRR could operate not only in the replication fork but also either after replication, on single‐ stranded gaps left behind newly restarted replication forks, or formed during repair of lesions arising in G2/M phase that need to be fixed (Daigaku et al. 2010; Karras and Jentsch 2010; Putnam et al. 2010). The level of PCNA modification with SUMO increases during S phase of the cell cycle and during DNA damage response. It has been recently shown that PCNA sumoylation protects the genome against non‐canonical gene conversions, dependent on HR, but reliant not only on the Rad52‐ and Rad51‐dependent pathway, but also on Rad59 and PCNA polyubiquitination (Halas et al. 2016). Sumoylated PCNA has a high affinity to Srs2 and to replicative Pols, which prevents HR and enables DNA synthesis over the lesion, respectively (Papouli et al. 2005; Pfander et  al. 2005; Kolesar et  al. 2012). The 3′→5′ DNA helicase activity of Srs2 (Rong and Klein 1993) protects the cellular genome against DNA expansions and contractions, by unwinding of hairpin intermediates formed by triplet repeats in DNA and by stimulat­ ing fork reversal at these repetitive sequences (Dhar and Lahue 2008; Kerrest et  al. 2009). On the other hand, the translocase activity of Srs2 (Antony et al. 2009) disrupts Rad51 filaments (Krejci et  al. 2003; Veaute et  al. 2003), which are the checkpoint‐­ activating signals, leading to checkpoint recovery (Hishida et  al. 2010; Yeung and Durocher 2011) and preventing recombination (Broomfield and Xiao 2002). The SUMO interaction motif at the C‐terminus of Srs2 protein was shown to dissociate Pol δ and Pol η from the repair foci, limiting extension of repair synthesis during template switch or HR and decreasing reciprocal DNA strand exchange, therefore contributing to main­ tenance of a stable genome (Burkovics et al. 2013). While binding of Srs2 by sumoylated PCNA inhibits the recombination pathway, binding of replicative Pols stimulates replication over the lesion. During normal DNA synthesis, replicative Pols are not able to progress through the damage on the template, but when DNA lesion blocks replication fork progression, signal about damage can be generated. This initiates checkpoint activation, followed by RNR induction resulting in the increase of dNTP levels that facilitates lesion bypass by the replicative Pols (Lis et al. 2008; Sabouri et al. 2008). It was shown that Rnr4/Pol δ‐dependent ethyl methanesul­ fonate (EMS)‐induced mutations are efficiently inhibited by hydroxyurea, an RNR inhibitor (Lis et al. 2008). Moreover, the high mutation rate observed in pol3‐01 mutants depends on the activity of the S‐phase checkpoint and is significantly reduced in dun1Δ strain (Datta et  al. 2000). These data consistently show that an S‐phase checkpoint‐ mediated up‐regulation of dNTP levels stimulates Polδ‐mediated TLS. It was also proved that yeast replicative DNA Pol α and Pol δ are able to insert nucleotides opposite DNA lesions at high dNTP concentration (Haracska et  al. 2003; Niimi et  al. 2004). When proper dNTP concentrations and correct dNTP pool bias are present in vivo during DNA damage, Polε can replicate over 7,8‐dihydro‐8‐oxoguanine (Sabouri et al.

9.3 ­Post-Replicative Repair (PRR) or Homologous Recombination (HR)

2008). In addition, the insertion of nucleotides by replicative Pols opposite lesions can provide a substrate for the TLS Polζ which is efficient at extending from nucleotides inserted opposite these lesions by the replicative DNA Polδ (Haracska et al. 2001b) or another Pol, for example Polη (Johnson et al. 2001). Thus DNA synthesis on template‐ containing lesion needs the cooperation of two Pols. When cyclobutane pyrimidine dimers or cis‐syn T‐T dimer blocks DNA replication by DNA Pol δ or Pol ε, a switch occurs to allow translesion synthesis by DNA Pol η, followed by another switch that allows normal replication to resume (McCulloch et al. 2004a). Also it has been shown for Pol ζ that while it is ineffective at inserting nucleotides opposite 7,8‐dihydro‐8‐­ oxoguanine and O‐6‐methylguanine, it can efficiently extend from the nucleotides inserted opposite them by Pol δ (Haracska et al. 2003). 9.3.2  Ubiquitinated PCNA‐ and Specialized Pol‐Dependent Translesion Synthesis

Following DNA damage, PCNA is monoubiquitinated at Lys164 in a Rad6–Rad18‐ dependent manner (Hoege et al. 2002). Monoubiquitination of PCNA enables its inter­ action with TLS Pols (in various combinations, e.g. Pol η alone, Pol ζ alone, Pol η then Pol ζ or Pol ζ with Rev1) and gives them access to damaged template leading to TLS (Haracska et al. 2001a). The result of this synthesis can be error‐free or error‐prone, depending on the particular lesion and Pol engaged in synthesis (Lawrence et al. 2000; Haracska et al. 2000a; Haracska et al. 2000b; Haracska et al. 2003; Northam et al. 2006; Parker et  al. 2007; Stone et  al. 2011). Some Pols can pass the damaged site in DNA, perfectly reproducing the initial content of the template, and thereby having high effec­ tiveness to the cognate substrate. The other Pols introduce changes across the lesion in the DNA strand synthesized during lesion bypass, and therefore the final sequence is different than the original one, and these Pols are mutagenic (Table 9.3). Among all TLS Pols the most effective in error‐free translesion synthesis is Pol η. It bypasses a range of DNA lesions, recreating the original sequence information properly. It inserts C oppo­ site 7,8‐dihydro‐8‐oxo‐guanine (8‐oxoG) (Haracska et al. 2000a; McCulloch et al. 2009), A‐A opposite the T‐T dimer (Washington et al. 2000; Gibbs et al. 2005), and so on (for more examples see Table 9.3). The same lesions bypassed by other Pols could lead to mutation; for example, Pol δ inserts A opposite 8‐oxoG (Haracska et al. 2000a). However, also Pol η is somewhat error‐prone, incorporating either T or C residues while bypass­ ing O6‐methylguanine lesions (Haracska et al. 2000b). It should be mentioned here that all TLS Pols (Pol η, Pol ζ, and Rev1) should be removed from the replication fork as soon as possible; otherwise, they will produce a lot of mis‐insertions on undamaged template, because their fidelity of replication is very poor (see Table 9.1). In some cases they can even produce complex mutations (e.g., tandem basepair substitutions and clusters of multiple, closely spaced mutations), as was shown for Pol ζ (Stone et al. 2012). For this reason, TLS Pols are actively removed from the cell, when they are not needed any more, by ubiquitin‐dependent proteasomal degradation. Rad30 (Pol η) and Rev1 are short‐living proteins, with an approximately 20‐minute‐long half‐life (Skoneczna et al. 2007; Wiltrout and Walker 2011). Moreover, it has been shown that Pol η is transiently stabilized after UV irradiation when its cog­ nate substrates appear in DNA (Skoneczna et  al. 2007); and that overexpression of Rad30, Rev3, or Rev7 TLS Pols always results in elevation of mutation rate (Rajpal et al. 2000; Pavlov et  al. 2002). The mutator effect is observed also in the strains bearing

223

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9  Mitotic Genome Variations in Yeast and Other Fungi

Table 9.3  Effect of translesion synthesis carried out by various DNA polymerases in S. cerevisiae cells. DNA lesion DNA polymerase

Error-free bypass

Mutagenic bypass

Pol α

Cyclobutane pyrimidine dimer (Nelson et al. 1996)

Pol δ

O(6)‐methylguanine (Haracska et al. 2000b) 7,8‐Dihydro‐8‐oxo‐guanine (McCulloch et al. 2009) T‐T (6‐4) photoadduct and abasic site (Gibbs et al. 2005) EMS‐induced damage (Lis et al. 2008)

Pol ε

7,8‐Dihydro‐8‐oxo‐guanine and O6‐ methylguanine (Sabouri et al. 2008)

Pol ζ

3‐Methyl‐3‐deazaadenine (Monti et al. 2011)

Rev1 Pol η

Cyclobutane pyrimidine dimer (Nelson et al. 1996; Stone et al. 2011) Thymine glycol (Johnson et al. 2003) Abasic site (Nelson et al. 2000) Acetylaminofluorene‐adducted guanine and T‐T (6‐4) photoproduct (Guo et al. 2001; Gibbs et al. 2005; Stone et al. 2011) DNA interstrand crosslinks (McHugh and Sarkar 2006) T‐T (6‐4) photoproduct and abasic site (Nelson et al. 2000; Gibbs et al. 2005)

Cyclobutane T‐T dimer (Johnson et al. 1999; Washington et al. 2000; Gibbs et al. 2005) C‐C and T‐C photoproducts (Yu et al. 2001) 7,8‐Dihydro‐8‐oxoguanine (Haracska et al. 2000a; Carlson and Washington 2005; McCulloch et al. 2009) 8,5′‐Cyclo‐2′‐deoxyadenosine and 8,5′‐cyclo‐2′‐deoxyguanosine (Swanson et al. 2012) 3‐Methyl‐3‐deazaadenine (Monti et al. 2011)

O(6)‐methylguanine (Haracska et al. 2000b) 7,8‐Dihydro‐8‐oxo‐guanine and T‐T cyclobutane dimer (McCulloch et al. 2009)

mutations in catalytic subunits of proteasome (pre2‐K108R, pup1‐T30A, pre3‐T20A) and in strain with deletion of UMP1 gene encoding proteasomal maturase, in which 20S proteasome activity is diminished (Podlaska et al. 2003; McIntyre et al. 2007). Mutator effect is caused also by chemical inhibition of proteasome activity or by inefficiency of ubiquitinating enzymes in marking Pols for degradation (Skoneczna et al. 2007; Wiltrout and Walker 2011).

9.3 ­Post-Replicative Repair (PRR) or Homologous Recombination (HR)

The DNA Pols’ C‐terminal regions contain domain called PIP (a PCNA interacting protein) responsible for their interaction with PCNA and consequently getting access to the DNA template (Garg et al. 2005; Parker et al. 2007; Acharya et al. 2011). It is believed that access to the perturbed replication fork by a particular Pol is determined simply by competition, so it can be regulated by changing the relative abundance of Pols in the nucleus. However, this view seems to be an oversimplification. Pol switching is influenced by numerous factors. First, the relative expression level of Pols differs and changes along cell cycle; for example, replicative Pol expression is highest at G1/S phase, while the highest expression of Pol ζ falls on G2/M (see Table 9.1) (Verma et al. 1991; Waters and Walker 2006). Second, the stability of Pols, for example Pol η and Rev1, is actively regulated via ubiquitin‐dependent degradation (Skoneczna et al. 2007; Wiltrout and Walker 2011). Third, Pols need to be properly assembled. Forming of holoenzyme encompasses not only assembling an active enzyme from several subunits, which probably needs chaperone assistance, but also installing the necessary cofactors, for example [4Fe‐4S] cluster in the CysB motif of several Pols, like Pol α, Pol ε, Pol δ, and Pol ζ (needed for their stabilization) or Zn2+ ion in CysA motif of Pol δ (required for its PCNA‐mediated processivity) (Netz et  al. 2011). In addition, Pol switching is influenced by: (1) changes in the localization of particular Pols, with respect to replication fork asymmetry and higher affinity of some Pols to operate on leading strand, lagging strand, or specific DNA substrate, such as ssDNA gaps; (2) interactions of Pols with their cognate sliding clamp proteins, as well as with other Pols; (3) post‐translational modification of Pols and clamp proteins (Hoege et al. 2002; Sutton 2010); and (4) the relative affinities of different Pols to the DNA substrate in need of replication (McCulloch et al. 2004a; McCulloch et al. 2004b). Moreover, it has been shown that besides PCNA monoubiquitination, Siz1‐mediated PCNA sumoylation is required for TLS stimulation. The epistatic analysis of relation­ ships between SIZ1, MMS2, and RAD5, with respect to UV sensitivity, demonstrated that in the absence of PCNA sumoylation, Mms2–Ubc13 and Rad5 influence TLS independently (Halas et al. 2011). Other data proved that Pol ζ gets access to the rep­ lication fork and is responsible for highly mutagenic synthesis of DNA during replica­ tion on undamaged template when replisome is defective (Northam et  al. 2006; Northam et al. 2010). 9.3.3  The Polyubiquitinated PCNA‐ and Rad5‐Dependent Damage Avoidance Pathway

After DNA damage, Mms2 and Ubc13 proteins are redistributed from cytoplasm to nucleus, where they form a complex with Rad5 and then polyubiquitinate Lys164 of PCNA (Ulrich and Jentsch 2000). It is a matter of debate whether polyubiquitin chain is built sequentially or is transferred en block on PCNA; both scenarios are possible. Ubiquitins in the chain are linked via Lys63. Polyubiquitinated PCNA stimulates the error‐free damage avoidance pathway of PRR (Branzei et al. 2004; Branzei et al. 2008). Thus DNA damage in the template DNA can be omitted by (1) template switch or sister chromatid junctions, which uses HR proteins Rad51, Rad52, and Rad54, as well as Sgs1–Top3 complex activity that permits resolving cross‐structures arising during repair (Gangavarapu et al. 2006; Branzei and Foiani 2007; Ball et al. 2009), or (2) Rad5‐ dependent fork regression (the ‘chicken foot’ DNA structure), followed by nascent

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strand annealing and DNA synthesis (Blastyak et al. 2007). Both branches of this path­ way are error‐free. 9.3.4  The Alternative PCNA‐, RPA‐, and 5′‐Junction‐Dependent Pathway Involved in Gap Filling and Telomere Maintenance

As previously mentioned, PCNA works at the center of replication fork and takes part in various activities after DNA damage, including DNA repair, as well as cell cycle check­ points. However, another PCNA‐like sliding clamp exists in yeast that is designed espe­ cially for the latter activity. This is the heterotrimeric checkpoint clamp consisting of the Ddc1, Rad17, and Mec3 subunits in S. cerevisiae (and Rad9, Rad1, and Hus1 subunits in Sch. pombe, hence its name the 9‐1‐1 complex). The 9‐1‐1 complex is an early response factor to DNA damage that activates checkpoints (Majka and Burgers 2003). It can be loaded on DNA in two different configurations: at the 5′ end of ssDNA coated with RPA, and at both the 5′ and 3′ junction of naked DNA at gaps left behind replication on lag­ ging strand (Majka et al. 2006). It subsequently stimulates Mec1 kinase simultaneously or in concert with Dpb11, depending on cell cycle phase (Navadgi‐Patil and Burgers 2009; Puddu et al. 2011). The 9‐1‐1 complex plays also a major role in DNA repair by interacting with and stimulating several other proteins. Those include specialized, but not replicative, DNA Pols (Kai and Wang 2002; Cardone et al. 2008) during repair of the gaps left behind replication fork, and both Rad51 and Dmc1 during meiotic DSB repair (Shinohara et al. 2003) and resection of uncapped telomeres (Dewar and Lydall 2012). Similarly to PCNA, the 9‐1‐1 complex is ubiquitinated, but published data disagree whether this modification is dependent on Rad6 or not, and which residue, Lys197 or Lys164, is the major acceptor site for ubiquitin. It is also not clear yet if this ubiquitination contributes to the function of the 9‐1‐1 complex by stimulating its role in DNA repair or cell cycle control, or rather mediates its proteasomal degradation (Fu et al. 2008; Davies et  al. 2010). Anyway, it is perfectly clear that physical interaction between Rad18 and 9‐1‐1 complex subunits exists and that both of them trigger transcription of DNA dam­ age‐inducible genes (Fu et al. 2008). There is also no doubt about the crucial role of the 9‐1‐1 complex in DNA repair and in checkpoint control under DNA damage stress. 9.3.5  Crosstalk between RFC Complexes Adapts Cellular Response to Different Stresses Arising from Genome Perturbations

As described in sections 9.2 and 9.3, PCNA and 9‐1‐1 complexes play a central role dur­ ing DNA replication and repair, serve as a convenient platform enabling access of a num­ ber of proteins involved in these processes to unwounded ssDNA, sense cellular stresses, and decide which repair path will be employed in particular circumstances. However, there are molecules that work behind the frontline, attached to nucleus envelope, allow­ ing PCNA and its alternative, the 9‐1‐1 complex, to stay on DNA (Haas et  al. 2012). Those molecules are chromatin‐associated factors called replication factor C complexes (RFC) that load and unload sliding clamps, which promote DNA replication and repair. In eukaryotic cells, four different sliding clamp loaders exist (Majka and Burgers 2004). All are heteropentamers and, in all of them, four smaller subunits are the same (Rfc2–5) (Yao et al. 2006), whereas the largest subunit is different in each of them. The classical RFC complex contains Rfc1 protein, which can be replaced by Ctf18, Elg1, or Rad24, leading to formation of alternative RFC complexes (Green et al. 2000; Mayer et al. 2001;

9.3 ­Post-Replicative Repair (PRR) or Homologous Recombination (HR)

Kanellis et al. 2003). The canonical RFC complex loads and unloads PCNA on/off DNA during normal replication (Bowman et al. 2004). Ctf18‐RFC and Elg1‐RFC not only con­ tribute to loading/unloading of PCNA on/off DNA in special circumstances, but also control positioning of PCNA on DNA. The 9‐1‐1‐complex has its own clamp loader, Rad24‐RFC. Moreover, the 9‐1‐1‐complex is placed on DNA in response to DNA dam­ age, in a reaction directed by RPA (Majka et al. 2006). The Rad24‐RFC (known also as a DNA damage checkpoint clamp loader) is able to unload PCNA clamps from DNA, which facilitates shutdown of replication during DNA damage response (Yao et al. 2006). The Rad24‐RFC functions are crucial for DNA processing, recombination, DNA repair, and survival of the cells following various stresses (Aylon and Kupiec 2003; Aylon et al. 2003). Ctf18‐RFC unloads PCNA from DNA (Bylund and Burgers 2005) and promotes sister chromatid cohesion, whereas Elg1‐RFC inhibits sister chromatid cohesion (Hanna et al. 2001; Maradeo and Skibbens 2010; Maradeo et al. 2010). Moreover, data show that Elg1 stimulates the recruitment of Ctf18 to chromatin, and suggest that Elg1, Ctf18, and Ctf4 may coordinate the relative movement of the replication fork with respect to the cohesin ring (Parnas et al. 2009). Elg1‐RFC has high affinity to the sumoylated form of PCNA and can diminish its level bound to chromatin (Parnas et al. 2010). It is unclear at this point how Elg1–RFC complex contributes to reduction of the level of sumoylated PCNA on DNA. It is possible that Elg1–RFC actively unloads sumoylated PCNA from chromatin; however, it is also possible that Elg1–RFC recruits the desumoylating enzyme Ulp1 to chromatin to reduce the level of sumoylated PCNA. The elg1Δ strain displays a hyper‐recombination phenotype (Ogiwara et  al. 2007). Furthermore, the elg1Δ deletion has synthetic lethal effect, with mutations causing HR defects (Aroya and Kupiec 2005). Ctf18‐RFC plays various roles at different stages of replication. It is involved in initiation of replication (Ma et al. 2010) as well as in replica­ tion checkpoint activation (Kubota et al. 2011; García‐Rodríguez et al. 2015; Okimoto et al. 2016), and contributes to the maintenance of exceptionally difficult DNA regions, for example, triplet repeats (Gellon et al. 2011) and telomers (Gao et al. 2014). Ctf18‐ RFC is also involved in the maintenance of proper genome copy number during cell divisions (Alabrudzinska et al. 2011). The lack of any of the alternative RFC complexes causes genome instability (Aroya and Kupiec 2005; Banerjee et al. 2007). The alternative RFCs are able to control and precisely channel the cellular response to DNA damage or replication stress. On the one hand, they employ PCNA to enable replication resumption; on the other hand, they employ cohesion complex to maintain chromosome integrity. Moreover, alternative RFCs work in shifts during cell cycle, adjusting the method of repair of DNA damage to the actual phase of cell cycle, as well as to cell ploidy (Delacôte and Lopez 2008; Li and Tye 2011). However, the molecular mechanisms by which the RFCs exert diverse effects on the cell remain to be uncovered. 9.3.6  Break‐Induced Replication (BIR) Is a Vastly Inaccurate Repair Pathway

Among all DNA lesions occurring frequently in every cell, one of the most severe is double‐strand DNA break (DSB). The proper repair of DSBs protects cells against genetic material rearrangements that can lead to genomic instability and, consequently, even to cell death. There are two major mechanisms responsible for DSB repair in the cell: HR and NHEJ. Choice of the repair pathway determines accuracy of the repair. Applying any of

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the HR subpathways restores original sequence information, similar to almost perfect repair at the replication fork via sister chromatid junction or fork regression (Haber 2006; Maher et al. 2011). Only on rare occasions, as, for example, in the case of recipro­ cal recombination operating at heterozygous locus in diploid cells, may it lead to gene conversion. However, when DNA is severely damaged, for example after γ‐irradiation, HR between non‐allelic repetitive elements, primarily Ty retrotransposons, spreads across the genome, resulting in chromosomal aberrations (Argueso et  al. 2008). It is interesting that NHEJ does not contribute to this phenomenon, but instead, as men­ tioned in section 9.2.3, NHEJ is uniquely responsible for producing short tandem dupli­ cations from non‐iterated sequences via a template misalignment mechanism (Lehner et al. 2012) and for creating small deletions by misalignment of unresected DSB ends, as well as larger deletions by misalignment of resected ones (Haber 2006). Homology‐dependent repair of DNA DSBs by gene conversion involves only short tracts of DNA synthesis and results in limited loss of heterozygosity (LOH). On the other hand, a high level of LOH is observed in the break‐induced replication (BIR). BIR is the process that is used to repair DSBs that expose only one strand end formed due to replication fork collapse or by erosion of uncapped telomeres. During BIR, the repair of DSBs occurs by invasion into a homologous sequence followed by replication to the end of the chromosome, resulting in non‐reciprocal translocation and extensive LOH. Thus BIR is an exceptional process that has capacity to duplicate hundreds of kilobases, mim­ icking normal DNA replication, requiring all three major replicative DNA Pols, the replicative DNA helicase (Cdc45, GINS, and Mcm2‐7 proteins) as well as Cdc7 kinase, necessary for both initiation of DNA replication and PRR. The origin recognition com­ plex (ORC) and Cdc6, which are critical to create a pre‐replication complex (pre‐RC), are dispensable (Lydeard et al. 2010a). Thus BIR is initiated at DSBs rather than at rep­ lication origins. However, while DNA synthesis during replication is a high‐fidelity process, synthesis during BIR is extremely inaccurate due to the 2,800‐fold higher rate of frameshift mutations than during normal replication that occur not only near the break, but also at long distances from it (Deem et  al. 2011). Indeed, BIR is the most mutagenic of all repair pathways processing DSB. The Exo1‐ and Sgs1‐dependent DNA ends resection can suppress BIR (Marrero and Symington 2010), while overproduction of Rad51 enhances it. Therefore the extensive 5′ to 3′ resection, which occurs prior to the initiation of synthesis of a new DNA strand during BIR, precludes efficient mainte­ nance of an Rad51 filament near the DSB end. On the other hand, the limitation 5′ to 3′ resection, which also revokes activation of the Mec1‐dependent DNA damage check­ point, is accompanied by a high level of de novo telomere addition (Lydeard et al. 2010b). Another factor that contributes to BIR regulation is modification of PCNA. The data show that mutations in PCNA, which abolish its ubiquitination or sumoylation, lead to BIR inhibition. It means that origin‐independent BIR required crosstalk between fac­ tors engaged in standard synthesis of DNA and proteins involved in PRR (Lydeard et al. 2010a). When the strand invasion fails (e.g., in rad51Δ mutants), or recombination‐­ initiated DNA synthesis breaks down (e.g., in Pol δ complex mutants), both of which leads to malfunction of BIR, the BIR events in diploid S. cerevisiae appear due to half crossovers resulting from abnormal processing of recombination intermediates (Deem et al. 2008; Smith et al. 2009). The repair of severe lesions in DNA such as DSBs or stalled replication forks requires  the  coordinated activities of both HR and DNA replication machineries.

9.4  Ploidy Maintenance and Chromosome Integrity Mechanisms

Growing  evidence indicates that during repair, some molecular activities of proteins participating in each of these processes could be engaged in both of them, and that to restore genome integrity following severe DNA damage, specialized accessory proteins are indispensable for the effective coupling of recombination to replication (Maher et al. 2011).

9.4 ­Ploidy Maintenance and Chromosome Integrity Mechanisms Long evolutionary history of life has brought the mechanisms of genome stability to perfection. Optimizing the fidelity of replication and the balance between the efficiency and costs of DNA repair assures success in raising future generations and adapting to changing environment. An important aspect of this optimization is the number of cop­ ies of genetic material found in various organisms or at various growth stages of any given organism. Having more than one copy of the genome helps to create the ‘reservoir of variability’ beneficial for adaptation, and also is crucial for HR, the most accurate, error‐free DNA repair mechanism. So, it is not surprising that most eukaryotes are diploids or polyploids for most of their lives. Having multiple copies of the genome has also its dark side. It will cause genome rear­ rangements that, especially under genotoxic stress, result in excessive genome shuffling with disastrous rather than beneficial effects on survival. In yeast S. cerevisiae the fre­ quency of spontaneous mutagenesis in diploid cells at heterozygous marker in forward mutation test is two orders of magnitude higher than in haploid cells, about 10–4 versus 10–6, respectively (Hiraoka et al. 2000; Ohnishi et al. 2004). This difference is due to the fact that multiple copies of the genome trigger recombination events that in diploid cells are much more frequent than in haploids. Point mutations such as base substitution or frameshifts remain on the same level in both types of cells. Such a huge difference strongly suggests the existence of the genome stability mechanisms devoted to the diploid genome. First results showing differences in budding yeast cellular response to DNA damage with respect to ploidy have recently been published. The mutation mcm4Chaos3, which leads to MCM helicase defect, causes replication stress in both haploid and diploid cells, whereas only diploid mutants exhibit G2/M delay, severe genetic instability, and reduced viability. These different outcomes are due to differences in repair pathways choice: while haploids use the Rad6‐dependent pathways that resume stalled forks, dip­ loids use the Rad52‐ and MRX‐dependent DSB repair pathways (Li and Tye 2011). The investigation of the nature and frequency of chromosomal rearrangements at the marker gene placed at the telomeric region showed a decrease in chromosome rear­ rangement frequency, along with an increase in complexity of the rearrangements occurring at the target gene with the increase of ploidy level. The presence of short DNA tandem repeat sequences seems to be a key requirement for deletion and reciprocal translocation processes to occur in diploids (Tourrette et al. 2007). The genomic screen for sensitivity to doxorubicin, a DSB inducer, revealed a huge difference in response of haploid and diploid cells to this agent. Exposure of the whole homodiploid and haploid yeast deletion clone collection to doxorubicin caused much higher damage and resulted in death or severely reduced growth fitness in a five‐ fold higher number of diploid mutant clones than in haploid mutant clones. Moreover,

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the toxic effect of doxorubicin was observed in some mutants, for example, bem1Δ, ctf4Δ, ctk1Δ, hfi1Δ, nup133Δ, and tho2Δ, exclusively in diploids. Additionally, strains bearing these mutations displayed severe G1/S phase cell cycle progression defects after treatment and some mutants were significantly enhanced (ctk1Δ and hfi1Δ) or deficient (tho2Δ) in recombination (Westmoreland et al. 2009). 9.4.1  Processes that Affect Aneuploidy in Yeasts

As mentioned above, diploids are oversensitive to DSB induction, DSB repair pathways can lead to changes in DNA sequence, including chromosomal aberrations, and in dip­ loid strains the most frequent groups of mutagenic events are rearrangements. Taking this information together, we can assume that DSBs in diploid cells can provoke aneu­ ploidization. In fact, a lot of damaging agents which induce DSBs in DNA result in chromosome aberrations. Also dysfunction of genes involved in DSB recognition, sign­ aling, or repair display hyper‐recombination phenotype. We have discussed this already. Recent discoveries point also to other causative agents, different from DSBs, that play a role in the formation of aneuploidy, and gain or loss of individual chromosomes. Data indicate that the hydroquinone‐induced delay in G2/M transition checkpoint, which is activated by the Hog1–Swe1 pathway, also results in aneuploidy (Shiga et  al. 2010). What is even more puzzling is that it was proposed that aneuploidy is a form of large‐ scale mutation that is able to confer adaptive phenotypes under diverse stress condi­ tions. This hypothesis recently gained support for Candida albicans cells (Selmecki et al. 2010). Diverse stresses can induce an increase of chromosome instability. It has been shown that the proteotoxic stress, caused by transient Hsp90 inhibition or by heat‐shock condition, markedly increased chromosome instability to produce a cell population with various karyotypes. This induced chromosome instability is probably linked to the Hsp90 chaperone role in the kinetochore complex assembly (Stemmann et al. 2002). Cells exposed to Hsp90 inhibitor end up with changes in chromosome XV, which lead to multi‐drug‐resistant phenotype and aneuploid chromosome stoichio­ metries. These results strongly suggest that aneuploidy is a form of stress‐inducible mutation, capable of fueling rapid phenotypic evolution and drug resistance, and proves the Hsp90‐dependent adaptive mechanism under stress conditions (Chen et al. 2012). However, in the cell additional mechanisms could lead to aneuploidization. Among them are all the processes ensuring proper segregation of genetic material, including quality control of freshly replicated DNA (replication checkpoint), starting of division (mitotic checkpoint), spindle organization and orientation, proper attachment of chro­ mosomes together with correction of mis‐attachment errors (spindle assembly check­ point), with subsequent chromosome segregation mechanisms (anaphase‐promoting complex checkpoint), cytokinesis, and septation (Suijkerbuijk and Kops 2008; McCulley and Petes 2010; Thompson et al. 2010; Silva et al. 2011; Stirling et al. 2012). All molecules engaged in this complex project can influence DNA content in daughter cells. Of course, all abnormalities should be detected and corrected by checkpoint proteins, but in the case of huge DNA damage, spindle element perturbation, or inefficiency of the control system, aberration in DNA content does occur. In such cases, cells with abnormal DNA content will be eliminated during the next mitotic division, as they undergo mitotic arrest triggered by unequal DNA content, which generates signal for elimination by mitotic catastrophe. If the cell escapes the execution it can: (1) undergo senescence, leading

9.4  Ploidy Maintenance and Chromosome Integrity Mechanisms

finally to cell death; (2) carry on the aberrant divisions, which cause an aneuploidization or cell death (as in the case when in daughter cells essential genes are lost); or (3) undergo mitotic slippage or cytokinesis failure, mechanisms that lead to polyploidization. There are many examples of various deficiencies leading to changes in cellular DNA content. Mutations causing defects in proper organization/functioning of cohesion complex (e.g., ctf18Δ deletion or eco1 point mutations) lead to aneuploidization or poly­ ploidization (Spencer et al. 1990; Kouprina et al. 1993; Unal et al. 2007; Bermúdez‐López et al. 2010). Inability to sense a tension on mitotic chromosomes (e.g., in sgo1 mutant) causes aberration in the spindle checkpoint and frequently leads to chromosome loss (Indjeian et al. 2005). Lack of separase Esp1 or an excess of securin Pds1 leads to poly­ ploidization due to sister chromatid separation failure (Lu and Cross 2009). Also dys­ function of components of the mitotic exit network (MEN) or proteins involved in cytokinesis such as Inn1, Cyk3, or Chs2 results in aneuploidization (Meitinger et al. 2010). Sometimes the reasons for improper chromosome segregation are not so obvious. The aft1Δ cells display chromosome segregation defects in meiosis (Hamza and Baetz 2012). Data show that the Aft1 protein not only acts as an iron‐responsive transcription factor, but also associates with the kinetochore complex through Iml3. Likewise, Aft1 stimulates the association of cohesin with the pericentric region of chromatin. This in turn is crucial to resist microtubule tension (Hamza and Baetz 2012). Regardless of the origin of aneuploidy in yeast cells, experimental data consistently show that having more chromosomes does not lead to living a better life. Gaining an additional copy of certain chromosomes initially decreases cellular fitness. It causes increased chromo­ some loss and mitotic recombination, and defects in DNA damage repair. Subsequently, an aneuploidy‐induced genomic instability arises, which facilitates the development of  further genetic alterations and leads to genome instability (Sheltzer et  al. 2011). However, parallel culture of isogenic aneuploid yeast strains, with DNA contents between 1c and 3c and distinct karyotypes, conducted under various experimental con­ ditions (e.g., in the presence of different chemotherapeutic or antifungal drugs), dem­ onstrated that under certain conditions, some aneuploid strains grew significantly better than the control ones. Results of these experiments led to the conclusion that aneuploidy can drive evolution, because it affects gene expression, thereby influencing the proteome level and raising the significant phenotypic variation that could help the cell to adapt to changed environmental conditions (Pavelka et al. 2010). Another set of experiments demonstrated that yeast strains may vary in phenotypes, such as gene expression, cell morphology, and sporulation rate, though even they have a common origin and were obtained from the integration of the same linear DNA construct to the genome. This heterogeneity results from different methods that yeast cells applied in response to a single initial event inducing translocation (Rossi et al. 2010). 9.4.2  Ploidy Changes in Yeasts

Ploidy changes in yeast cells, similarly to aneuploidy, can be provoked by DNA damage, mitotic spindle perturbations, γ‐irradiation, various anticancer drugs, and of course heat shock. Interestingly, it was shown for yeast cells that ploidy changes may occur also as a result of a single gene deletion. Two experimental approaches revealed an outstand­ ing phenomenon, which is exclusive for diploid cells. Saccharomyces cerevisiae diploid cells constitutively exposed to rearrangement stress leading to gross chromosomal

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rearrangements (GCR) as a consequence of gene deletion, e.g. RAD52 (engaged in HR) or CTF18 (regulating PCNA and cohesion complex access to DNA), can reduce their genome to the haploid level. However, rad52/rad52 cells do it sequentially, losing the chromosomes one by one during successive divisions (Song and Petes 2012), whereas ctf18/ctf18 cells reduce their genome in a single step (Alabrudzinska et  al. 2011). Anyway, in both cases, haploidization results in restriction of rearrangement events; as in haploids, these events are less frequent due to scarcity of homologous sequences in their genomes. In wild‐type diploids, GCR rate is 10–4, as opposed to 10–9 in wild‐type haploid cells (Chen and Kolodner 1999; Hiraoka et al. 2000; Ohnishi et al. 2004). It is still not clear if the tendency to reduce the ploidy level observed in diploid strains bear­ ing rad52 or ctf18 mutation is incidental or rather it is a way to avoid further genome destabilization. However, haploid strains that appeared in the process of ploidy reduc­ tion have much lower mutation rate than initial diploids, they are fertile, and they are more vital, so they will dominate the cell population and have better chances to survive, although it should be noted that the frequency of haploidization is not so high and most cells in the initial diploid population drift in an opposite direction, and accumulate sec­ ondary changes in the genome, thus becoming aneuploid or polyploid cells, with all the consequences, such as elevated mutation rate, decreased fitness, and escalating division problems leading to permanent cell cycle arrest and finally cell death. Ploidy change was reported for other yeast mutants as well, but in those cases it was the conversion of haploids into diploids. Such change accompanies deletion of a num­ ber of genes. Haploid cells devoid of ZDS1 gene encoding factor involved in mitotic exit will turn into diploid when exposed to Ca2+ ions (Miyakawa and Mizunuma 2007). The mutations in any of three adjacent residues, L97, Y98, or G99, near the C‐terminus of histone H4 lead to polyploidy (Yu et al. 2011). The deletion of SFH1 or RSC3 coding components of the RSC chromatin remodeling complex required for G1/S transition results in ploidy shift, which, however, is eliminated by knockout of the S‐phase cyclin CLB5 and by transient depletion of replication origin licensing factor Cdc6p (Campsteijn et al. 2007). Similar increase in ploidy phenomena is induced by deletion of the IPL1 gene encoding the component of the Aurora kinase complex (also called the chromo­ somal passenger complex), which is an essential regulator of chromosome segregation, spindle checkpoint, and cytokinesis, or, by deletion of BEM2 encoding protein, which is involved in the control of cytoskeleton organization and required for bud emergence (Chan and Botstein 1993). The ploidy shift was also reported for other fungi. Successive loss of chromosomes, through different stages of aneuploidy, ending with haploid state was described for dif­ ferent species of imperfect fungi after treatment with benomyl (Sthal and Esser 1993; Samsonova et al. 1996), γ‐radiation (Mortimer et al. 1981), and after protoplast fusion (Fournier et al. 1977; Buttner et al. 1990), or in early zygote progeny (Kurishko 1986). The polyploidy and subsequent ploidy reduction via multipolar spindle was observed in tetraploid cells of C. albicans (Suzuki et al. 1986). 9.4.3  Possible Mechanism of Ploidy Change in Yeast

In yeast and in other fungi, cell ploidy changes occur frequently. Analyzing environ­ mental signals and the type of mutations that predispose to this kind of events, one can presume that aneuploidy and/or ploidy change happens when a cell arrested at mitotic

9.4  Ploidy Maintenance and Chromosome Integrity Mechanisms

checkpoint following aberration does not die, but instead overcomes the arrest and continues aberrant divisions. The division arrest may be initiated by severe DNA dam­ age, spindle perturbation, or even heat shock, and the division should not be continued until the problem is solved. If signal concerning irregularities does not disappear and arrest persists, the cell should be marked for removal by mitotic catastrophe, similar to the situation in mammalian cells (Vitale et al. 2011). When something goes wrong, for example, cell cycle control fails or a mutation in genes predisposing to genomic instabil­ ity is present, the cell is allowed to escape from execution and the aberrant division is finished, with all the consequences, like gaining or losing single chromosomes, or undergoing sequential steps of illegitimate polyploidization and depolyploidization, including haploidization (Figure 9.3). Aneuploid Multipolar spindle

Depolyploidization

Inducers of mitotic catastrophe

Tetraploid

Mitotic death

otic Mit age p slip

Abberant mitosis

idi Po ly

tion

iza

Mitotic arrest

oid

plo

Aneuploid

upl

za

tio

n

Ane

Cytokinesis failure

Aneuploid Haploidization

Multipolar spindle

Aneuploid

Abberant mitosis Aneuploid Aneuploid Nucleophagy Haploid

Aneuploid

Haploid

Haploids

Figure 9.3  Causes and consequences of cellular DNA content disturbances.

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9  Mitotic Genome Variations in Yeast and Other Fungi

Mitotic arrested cells can duplicate their genome by two possible mechanisms: ‘slip­ page’ from mitosis, or a round of aberrant mitosis without cytokinesis. Aneuploidization can be provoked by unrepaired DSB, by monotelic, syntelic, or merotelic chromosomes, or by non‐ synchronous and delayed spindle movement. What mechanism may lead to haploidization? One can expect at least three different scenarios: 1) Multipolar spindle formation leading to four haploid cells. This idea corresponds well with the observation that the chromosome loss in ctf18Δ/ctf18Δ diploid S. cerevisiae cells was not random, but rather that full sets of chromosomes were lost at once (Alabrudzinska et al. 2011). 2) Successive loss of chromosomes, through different stages of aneuploidy, ending with haploid state (Song and Petes 2012). 3) Selective elimination of the parental chromosomes from the cell via micronuclei formation, heterochromatinization, and fragmentation of micronucleated chroma­ tin in the final step during haploidization in the process resembling apoptosis and named programmed DNA elimination (Gernand et al. 2005). This phenomenon was observed in different animal (Zelesco and Marshall Graves 1983) and plant hybrid cells, such as during the development of wheat × pearl millet hybrid embryos (Gernand et al. 2005). In mammalian cells the formation of micronuclei containing aggregated DSB DNA, kinetochores, and centromers is widely used to score genomic instability, genotoxic exposure, and replication stress (Norppa and Falck 2003; Huang et al. 2011; Xu et al. 2011). However, it was not checked if assayed cells can reduce ploidy after micronuclei blebing and DNA degradation. In yeasts cells nucle­ oautophagy may function as an equivalent mechanism (Krick et  al. 2008; Robert et al. 2011).

9.5 ­Concluding Remarks During evolution, living cells elaborated the range of mechanisms ensuring genome stability. These mechanisms assure constant genome content by optimizing the fidelity of replication, removing errors and lesions from DNA strands, repairing breaks, resolv­ ing abnormal DNA structures arising during cell cycle, monitoring cell divisions, detecting problems connected with DNA damage or replication block, and adjusting the response to the problem with respect to cell phase, stage, and ploidy. However, sometimes scrupulously protecting the genome against any change could lead straight to death and only taking a risk of mutation permits survival. That is why mutation toler­ ance mechanisms evolved in parallel to those maintaining the genome. Ultimately, the optimal balance between the efficiency and costs of DNA repair assures success in rais­ ing future generations and adapting to changing environment. Thus genome variations contribute to survival and evolution. In this chapter, we focused on the mechanisms leading to genome instability in fungi, providing a comprehensive overview of how all these pathways orchestrate in eukaryotic cells and telling how the cells in some circumstances have to pay a very high mutagenic price for surviving. Since all processes ensuring genome stability are highly conserved, from yeast to humans, significant findings in yeast may be extrapolated to vertebrates, greatly facilitating the molecular analysis of these complex regulatory networks.

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Webliography www.candidagenome.org: The Candida genome database is a resource for genomic sequence data and gene and protein information for Candida albicans. www.pombase.org: PomBase is a comprehensive database for the fission yeast Schizosaccharomyces pombe, providing structural and functional annotation, literature curation, and access to large‐scale datasets.

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www.uniprot.org: The UniProt is designed to provide the scientific community with a comprehensive, high‐quality, and freely accessible resource of protein sequence and functional information. www.yeastgenome.org: The Saccharomyces Genome Database (SGD) provides comprehensive integrated biological information for the budding yeast Saccharomyces cerevisiae, along with search and analysis tools to explore these data, enabling the discovery of functional relationships between sequence and gene products in fungi and higher organisms.

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Part IV General Genome Biology

253

10 Genome Variation in Archaeans, Bacteria, and Asexually Reproducing Eukaryotes Xiu-Qing Li* Fredericton Research and Development Centre, Agriculture and Agri-Food Canada, Fredericton, New Brunswick, Canada

Abstract Prokaryotes and various lower eukaryotes reproduce mainly or solely through various asexual processes. Genetic diversity in these species is therefore mainly dependent on nonmeiotic variation. This chapter reviews and characterizes genome variation in terms of genome size, base composition, and gene directional levels with the aim of clarifying the extent of non-meiotic genetic variation and providing insight into which types of genetic variation have been associated with these lower organisms’ use in evolution. Genomic size variation is greater in gram-positive bacteria and archaeans than in gram-negative bacteria. The average gene size ranged from 0.135 Mb for Nanoarchaeota archaeans to 5.162 Mb for Actinobacteria. There is a 66-fold size difference between the largest and smallest genomes in bacteria. The normal probability plot of genome sizes suggests that the evolution of the largest bacterial genomes was based on different underlying mechanisms than for other bacteria. Fungi that reproduce primarily by asexual means tend to have larger genomes than those that reproduce sexually. Archaeans and bacteria have far greater within-subkingdom variation in GC contents than plants and animals. The average high/ low GC content ratio is approximately 3.9 in prokaryotes and only 1.4 in sexually reproducing eukaryotes. A positive correlation between genome size and GC content is commonly found in Actinobacteria and Proteobacteria; however, such a correlation is not found for most other bacteria such as those in the phyla Firmicutes, Tenericutes, and Aquificae and is not likely to exist in archaeans either. The percentage of neighbor genes on the same direction is higher in bacteria than in archaeans. Prokaryotes and non-meiotic eukaryotes exhibit extensive genomic variation and changes in genome size, GC content, and gene direction. Keywords  chromosome number; genome size evolution; chromosomal size; nucleotide composition; genome size and GC content correlation; gene direction; gram-positive bacteria; gram-negative bacteria; archaeans; fungi; protists; higher organisms; dicot plants; monocot plants; non-mammalian animals; non-primate mammalian animals *Corresponding author: [email protected]; [email protected]; [email protected]

Li X.-Q. (2017) Genome variation in archaeans, bacteria, and asexually reproducing eukaryotes. In: Li X.-Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley-Blackwell, Hoboken, NJ, Ch. 10, pp. 253–266. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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10.1 ­Introduction Non-meiotic genome variation, a broad term of somatic genome variation, is responsible for genome diversity in viruses, gram-negative bacteria, gram-positive bacteria, and archaeans, and also largely responsible for the variation in fungi that reproduce asexually. In addition to undergoing within-genome variation such as mutation, transposition, and recombination, living organisms, from prokaryotes to higher plants, may receive exogenous genetic material through lateral gene transfer. Lateral gene transfer is known to occur through conjugation, transformation, and transduction in bacteria (Narra and Ochman 2006) and through natural Agrobacterium-mediated transformation in plants (Schell and Van Montagu 1977; Joos et al. 1983). However, lateral gene transfer usually involves far less genetic material recombination then meiosis and fertilization-based genetic recombination and segregation. Li (2015a) has recently reviewed/described laboratory protocols in somatic genome manipulation, and Li (2015b) describes various methods in genome sequence analysis. The reader is referred to Li and Du (2014a) for the features and evolution of the RNA poly(A) site region of eukaryote genomes and to Li (2014) for base composition of the three-prime untranslated region (3′UTR), in plants and animals. Therefore this chapter reviews and analyzes genome size, base composition, and gene direction with a view to clarifying the extent of diversity that can result from non-meiotic variation. The findings may also be useful for studying mechanisms of adaptation and evolution in non-meiotic lower organisms.

10.2 ­Chromosome Number in Prokaryote Species It was reported that Haloarcula hispanica has two chromosomes and one megaplasmid (Liu et  al. 2011). Since further research is likely required to clarify the relationship between the two chromosomes, we did not take the sum of the two chromosomes when we calculated genome size in this chapter. It is known that multiple-replication origins of chromosomes exist in H. hispanica and that the activities of two replication origins on the main chromosome are controlled differently (Wu et  al. 2014). It is likely that copy-number control of the minichromosome is independent from that of the major chromosome in Haloferax mediterranei (Liu et  al. 2013). It would be interesting to examine the mechanisms responsible for maintaining two chromosomes in this archaean and, more specifically, whether they include a multiple-chromosome mechanism similar to that found in eukaryotic chromosome systems. We examined all or nearly all the species for which complete genome sequences were available in the NCBI database in summer 2014, and found that all archaeal or bacterial genomes had one chromosome, with the exception of three archaean species that had one large chromosome and one small chromosome in their genome (Li and Du 2014b). These three archaeans species are Haloarcula hispanica, Haloarcula marismortui, and Halorubrum lacusprofundi. The size of the main chromosome in these three species is 2.995 Mb, 3.132 Mb, and 2.735 Mb, respectively, and the size of the second chromosome or minichromosome is 0.489 Mb, 0.288 Mb, and 0.530 Mb, respectively. The minichromosome is larger than the plasmids found in bacteria such as the Agrobacterium tumefaciens str. C58 plasmid Ti, which is 0.214 Mb (AE007871.2).

10.3  Genome Size Variation in Archaeans and Bacteria

10.3 ­Genome Size Variation in Archaeans and Bacteria Bacterial genome size has been examined in several reviews (Trevors 1996; Trevors 1998; Stȩpkowski and Legocki 2001). Bacteria may have facilitated recombination by increasing their genome size and changing the order of their genes (Trevors 1996; Trevors 1998). It was also hypothesized that the same lineage may diverge into species having substantially reduced genomes (presumably losing genes) or species having enlarged genomes (presumably integrating new genes) and that the loss of DNA repair genes may cause mutational bias resulting in a high A+T content (Stȩpkowski and Legocki 2001). Li and Du (2014b) analyzed genome size in most of the prokaryotes for which the complete genome sequences are available in the NCBI database. An estimation of genome-size variation in prokaryotes as a whole is critical for understanding the extent to which non-meiotic variation has shaped the prokaryote kingdom. This may also provide insights into the mechanisms causing the variation in different phyla/ classes of prokaryotes. The average genome sizes of archaeans and bacteria are shown in Figure 10.1(this study). The genome list used in this analysis was largely the same as that in Li and Du (2014b), but a stricter approach to removing redundant sequences was applied (usually due to sequencing of the same strains in different laboratories). Phylum or class level average genome size was ranked as follows: gram-negative bacteria (2.67 Mb) > gram-positive bacteria (2.53 Mb) > archaeans (1.77 Mb) (Figure 10.1). The average genome size of each large group of prokaryotes varies greatly (Figure 10.1). Nanoarchaeota archaeans have smallest average genome size (0.136 Mb) among archaeans and bacteria. Tenericutes have smallest average genome size (0.907 Mb) among bacteria and Actinobacteria have the largest average genome size (5.16 Mb). Three

2.67 Mb

2,000,000

Aquificae Planctomycetes Spirochaetes Fusobacteria Nitrospirae Deinococcus Proteobacteria Verrucomicrobia BacGramN MEAN

3,000,000

2.53 Mb

4,000,000

Tenericutes Firmicutes Actinobacteria BacGramP MEAN

5,000,000

1.77 Mb

Genome or chr size (bp)

6,000,000

1,000,000 Nanoarchaeota Euryarchaeota Thaumarchaeota Crenarchaeota Archaean MEAN

0

Figure 10.1  Average genome size of archaeans, gram-positive bacteria, and gram-negative bacteria. The number of genomes is 61,184, and 235 for archaeans, gram-positive bacteria, and gram-negative bacteria, respectively. The mean genome size order is 1.77 Mb < 2.53 Mb < 2.67 Mb for archaeans, gram-positive bacteria, and gram-negative bacteria, respectively. Note that both bacteria and archaeans showed considerable variation in genome size. Source: This study.

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bacterial phyla—Nanoarchaeota, Tenericutes, and Aquificae—have smaller genomes than archaeans on average. Among the gram-positive bacteria, Mycoplasma genitalium (Tenericutes) has the smallest genome (0.580 Mb on average out of five genomes) and Amycolatopsis mediterranei S699 (CP003729.1, an Actinobacterium) has the largest genome (10.247 Mb)(this study). Nanoarchaeota (0.136 Mb on average) is the archaean with the smallest genome. The  largest sequenced archaeal genome (3.132 Mb or 3,131,724 bp) is from chro­ mosome I (gi|55229667|gb|AY596297.1) of H. marismortui ATCC 43049 (Archaea, Euryarchaeota). The H. marismortui chromosome I is 23 times larger than the average size of Nanoarchaeota archaeal genomes. In the bacterial kingdom, the gram-negative Proteobacteria have both the smallest genome (0.162 Mb; GC: 14.55%; species Candidatus Carsonella ruddii) and the largest genome (10.351 Mb; GC: 62.19%; species Myxococcus stipitatus, with a 66-fold difference between the two (this study). Not only is the genome of M. stipitatus 66 times larger than that of Candidatus C. ruddii, but also its GC or C+G content is five times greater (this study).

10.4 ­Archaeal and Bacterial Genome Size Distribution Li and Du (2014b) analyzed the genome size distribution in archaeans and bacteria and found considerable variation. They also found that different biological mechanisms likely explain the difference between most genomes and some outlier genomes such as large genomes (Figure 10.2) (Li and Du 2014b). Archaean genomes can be classified into two distinct groups according to their genome size: a smaller group with very small genomes (smaller than 0.53 Mb; mainly Nanoarchaeota) and a larger group of much larger genomes (larger than 1.2 Mb) (Figure 10.2). To date, no archaeal species has been identified that has a genome size that is intermediate between the two groups. The genome sizes of 93% of bacteria are smaller than 6 Mb and their distribution is  approximately normal (approximately linear on the normal probability plot) (Figure  10.2). However, some bacteria (approximately 7%) have very large genomes (approximately 6–10 Mb) and their distribution can be seen as an outlier on the normal probability plot, suggesting that different mechanisms are involved in the size variation Bacteria

Archaea

12,000,000

3,500,000

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0

0 0

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Sample Percentile

100

120

0

20

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120

Sample Percentile

Figure 10.2  Normal probability plots of genome/chromosome size (bp, the Y-axis). Note that the large genomes/chromosomes (on the right side of each plot) do not fit a normal distribution and that archaeal genomes form two large groups. Source: Adapted from Li and Du 2014b.

10.5  Genomic GC Content in Archaeans, Bacteria, Fungi, Protists, Plants, and Animals

in the two groups of bacteria. Generation of the large genomes is likely driven by both the mechanisms that are common to most genomes and the specific mechanisms used only by these large genomes (Li and Du 2014b).

10.5 ­Genomic GC Content in Archaeans, Bacteria, Fungi, Protists, Plants, and Animals DNA contains four types of bases: adenine (A), cytosine (C), thymine (T), and guanine (G). The amounts of A and T are approximately equal, and the amounts of C and G are also approximately equal in DNA according to the Chargaff rule (Chargaff et al. 1952) and Watson and Crick’s pair theory (Watson and Crick 1953). The G-C pair (three hydrogen bonds) is stronger than the A-T pair (two hydrogen bonds). Therefore, in genome analysis, a critical base composition parameter is C+G content (also called GC content, CG content, or G+C content). The expression GC content or GC-richness is widely used; however, the term GC can be confused with the dinucleotide GC. Therefore it is more appropriate to talk about C+G content. In this chapter, GC means C+G content and does not refer to the GC dinucleotide. GC content has been estimated by melting–annealing profiles, CsCl gradient buoyant densities, flow cytometry, and genome sequencing. Each of these methods has its weakness and strength. In plants, base compositions were usually estimated using annealing temperatures (Šmarda and Bureš 2012). Flow cytometry after staining with DAPI (4’,6-diamidino-2-phenylindole) is consistently higher than the GC content in the sequencing data (Šmarda et  al. 2012). Before the second-generation sequencing data were available, the base composition in animals mainly used gene segments and transcribed sequences (Presber et al. 1976; Gardiner 1996; Mooers and Holmes 2000). The GC content is not perfectly correlated with the melting–annealing speed of DNA, because the degree of repetition of satellite sequences can also affect the DNA annealing speed (Appels et al. 1981; Cionini et al. 1985; Nellåker et al. 2009). In recent years, a great many genomes have been sequenced and assembled, making whole-sequence-level evaluation of GC content possible. A general comparison of genomic base composition in major kingdoms (domains of life) was carried out recently (Li and Du 2014b). The accuracy of genome sequence-based GC evaluation varies depending on the completeness of genome sequencing. Gap areas cannot be evaluated using the reference genome unless some unplaced contigs in the assembly are from the gaps. To date, the genome-sequencing approach has the highest resolution and provides the most precise evaluation of GC content. The within-kingdom (domain) variation of GC content is much higher in archaeans and bacteria than in plants and animals (Figure 10.3) (Li and Du 2014b). The C+G content in prokaryotes is much more variable than in higher organisms. The C+G content is highest in monocot plants and lowest in dicot plants. C+G content does not vary greatly among mammal species. The GC content in archaeans varies between 23.11% (APJZ01000005.1, Candidatus Nanopusillus sp. Nst1, a Nanoarchaeota) and 66.72% (CP001365.1, H.  lacusprofundi). Gram-positive bacteria have GC content ranging from 21.39% (NC_011047.1| Candidatus Phytoplasma mali) to 73.32% (CP004370.1, Streptomyces albus). The gram-negative bacterial GC content varies between 13.98% (CP003541.1,

257

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40

30

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70

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10

0

Animal mam. pri. AVERAGE Pongo abelii Macaca mulatta Homo sapiens Pan troglodytes Callithrix jacchus Animal mam. nonPri.AVERAGE Rattus norvegicus Oryctolagus cuniculus Equus caballus Sus scrofa Mus musculus Bos taurus Canis lupus familiaris Animal nonmam.AVERAGE Danio rerio Gallus gallus Taeniopygia guttata Apis mellifera Drosophila melanogaster Caenorhabditis elegans Plant monocot AVERAGE Zea mays Sorghum bicolor Oryza sativa Brachypodium distachyon Plant dicot AVERAGE Solanum lycopersicum Solanum tuberosum Vitis vinifera Populus trichocarpa Medicago truncatula Arabidopsis thaliana Protists: Alveolata Protists: Cryptophyta Fungi: Ascomycota Fungi: Basidiomycota Fungi: Microsporidia Bac GramN: Verrucomicrobia Bac Gram N: Proteobacteria Bac Gram N: Deinococcus Bac Gram N: Nitrospirae Bac Gram N: Fusobacteria Bac Gram N: Spirochaetes Bac Gram N: Planctomycetes Bac Gram N: Aquificae

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Bac GramP: Actinobacteria Bac GramP: Firmicutes Bac GramP: Tenericutes Archaeans: Crenarchaeota Archaeans: Euryarchaeota Archaeans: Thaumarchaeota Archaeans: Nanoarchaeota

Figure 10.3 The genomic base compositions and C+G contents of different kingdoms and large groups. The information is provided by phylum/class for archaeans, gram-positive bacteria, gram-negative bacteria, fungi, and protists and by species for dicot plants, monocot plants, non-mammalian animals, non-primate mammalian animals, and primate animals. Note that (1) C+G content is much more variable in prokaryotes than in higher organisms, (2) among plants and animals, C+G content is highest in monocot plants and lowest in dicot plants, and (3) mammals have very similar C+G contents between species. Source: Modified from Li and Du 2014b.

10.6  Correlation between GC Content and Genome or Chromosome Size

5.2 5.0 3.9

4.0 2.9

1.6

1.7

1.6

1.0

1.1

1.3

1.4

Eukaryote MEAN

2.0

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3.0

3.4

Dicot plants

Highest/Lowest GC (fold)

6.0

Prokaryotes (genomes)

Mammals

Non-mammals

Fungi

Prokaryote MEAN

Bacteria GramN

Bacteria GramP

Archaeans

0.0

Eukaryotes (chromosomes)

Figure 10.4  Ratio (fold) of highest genomic GC content and lowest genomic GC content in prokaryotes and of the highest chromosomal GC content and lowest chromosome GC content in eukaryotes. GramP, Gram-positive bacteria; GramN, gram-negative bacteria. Average GC-content difference between prokaryotes and eukaryotes is highly significant (t-test, P < 0.01). Note that the genomic (or chromosomal for eukaryotes) GC-content ratio between the highest GC content and the lowest GC content is significantly higher in prokaryotes. Source: This study.

Candidatus C. ruddii) and 73.29% (AP012338.1, Phycisphaera mikurensis; a species in Planctomycetes). The variation of the average GC content in prokaryotes is generally higher than that in sexually reproducing eukaryotes (Figure 10.4). The GC-content difference between the GC-richest phyla and the GC-poorest phyla ranges from approximately 2.9-fold (archaeans) to 5.2-fold (gram-negative bacteria) (Figure 10.4). In contrast, the difference is only 1.1-fold to 1.7-fold in eukaryotes (Figure 10.4). This difference between prokaryotes and eukaryotes suggests that genomic variation of base composition is an important source of genetic variation during the adaptation and evolution of prokaryotes. In contrast with the situation for prokaryotes, it is likely that meiotic segregation and recombination are the main mechanisms of genetic variation in plants, animals, and most fungi, and that GC-content variation is less important.

10.6 ­Correlation between GC Content and Genome or Chromosome Size C+G content is positively correlated with genome size in bacteria (Musto et al. 2006; Nishida 2012). A relationship was found between genome size and GC content for aerobic, facultative, and microaerophilic species, but not for anerobic bacteria, in a study encompassing all prokaryotic genomes available in the NCBI database (Musto

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10  Genome Variation in Archaeans, Bacteria, and Asexually Reproducing Eukaryotes

Table 10.1  Correlation between chromosome size and chromosome C+G content in archaeans, gram‐positive bacteria, and gram‐negative bacteria.

Kingdom or subkingdom

Number of chromosomes

Average chromosome size (Mb)

Average chromosome C+G (%)

Spearman correlation between chr size and C+G (R)

Archaeans

 61

1.77

43.09

  0.21 NS

Gram‐positive bacteria

184

2.53

40.05

0.75 **

Gram‐negative bacteria

246

2.67

46.45

0.50 **

Fungi

139

 1.46 a

45.09

0.46 **

a) Chromosome size estimation is based on genome assemblies, which may differ from the chromosomes in real cells. Highly significant (**) (P < 0.01) and not significant (NS) (P > 0.05) according to the ‘algorithm AS 89’ test (Li and Du 2014b).

et al. 2006). In a study using pooled data for one plant, four animals, three protists, and seven fungi, no significant correlation was found between the genomic GC contents and genome sizes (Mitchell 2007). When the base composition of each of the major kingdoms and phyla was analyzed separately using complete and nearly complete genomes from the NCBI database (http://www.ncbi.nlm.nih.gov/nucleotide/), genome size and GC content were found to be positively correlated in various groups of bacteria; however, no correlation was found for other bacteria or for archaeans (Table 10.1; Figure 10.5) (Li and Du 2014b). When all the bacterial genomes were examined together, there was a significant positive correlation between genome size and GC content (Figure 10.5). Archaeans, protists, and fungi do not show this strong correlation (Figure 10.5). When all the gram-positive bacteria were analyzed together, a strong positive correlation was found between genome size and genomic GC content. A similar correlation was found for gram-negative bacterial species (Table 10.1) (Li and Du 2014b). However, when each phyla/class was analyzed separately, a positive correlation between genome size and GC content was found mainly for Actinobacteria and Proteobacteria. There was either no correlation or the correlation was very weak in most other bacterial phyla/classes, such as Firmicutes, Tenericutes, and Aquificae as well as in archaeans. The biology underlying genome evolution for the genome size-GC correlated group and non-correlated group must be different (Li and Du 2014b).

10.7 ­Genome Size and GC-Content Variation in Primarily Asexually Reproducing Fungi Fungi can be classified into a group consisting of species characterized primarily by asexual reproduction and species for which a sexually reproducing stage has not been identified, and a group of species known to have both asexual and sexual reproduction (Groth and Roelfs 1982). We analyzed the genome size and GC contents of various

80

70

Bacteria

Archaea

65

70

60 55

(C+G) (%)

(C+G) (%)

60 50 40

50 45 40 35

30

30

20

25

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20 0

2,000,000

4,000,000

6,000,000

8,000,000

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Genome size (bp)

70

1,000,000

1,500,000

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2,500,000

3,000,000

Genome size (bp) 45

Fungi

Protists

40

60

35 30

(C+G) (%)

50

(C+G) (%)

500,000

40

25 20 15

30

10

20

5 0

10 0

2,000,000

4,000,000

6,000,000

8,000,000

Chromosome length (bp)

10,000,000

12,000,000

0

500,000 1,000,000 1,500,000 2,000,000 2,500,000 3,000,000 3,500,000 4,000,000 Chromosome length (bp)

Figure 10.5 Distribution of values of chromosome size and C+G content in bacteria, archaeans, fungi, and protists. The chromosome number for bacteria, archaea, fungi, and protists is 430, 61, 139, and 21, respectively. Note that C+G content and chromosome or genome size is positively correlated in bacteria, but there is no such correlation or linear relationship in other kingdoms. Source: Modified from Li and Du 2014b.

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10  Genome Variation in Archaeans, Bacteria, and Asexually Reproducing Eukaryotes

T. terrestris

Sexual MEAN Sexual MEAN

S. stipitis

T. terrestris

ASEXUAL

S. sp. 'A. aceri'

E. cymbalariae

A. gossypii

Asexual MEAN

T. inflatum

N. gansuense 2

N. gansuense 1

N. aotearoae

M. anisopliae

M. thermophila

M. acridum

I. farinosa

H. siamensis

H. thompsonii

Genome size (Mb)

(a) 45 40 35 30 25 20 15 10 5 0

SEXUAL

GC content (%)

(b) 70 60 50 40 30 20 10

ASEXUAL

S. stipitis

S. sp. 'A. aceri'

E. cymbalariae

A. gossypii

Asexual MEAN

T. inflatum

N. gansuense 2

N. gansuense 1

N. aotearoae

M. anisopliae

M. thermophila

M. acridum

I. farinosa

H. siamensis

H. thompsonii

0

SEXUAL

Figure 10.6  (A) Genome size and (B) GC contents of primarily asexual fungi and the fungi that are known to have a sexual reproductive stage. NCBI BioProjects ID for the primarily asexually producing fungi: PRJNA192877, PRJNA242986, PRJNA242987, PRJNA245139, PRJNA245140, PRJNA79339, RJNA221524, PRJNA67299, PRJNA174039, PRJNA73163, PRJNA10623, PRJNA78153, PRJNA39551, PRJNA18881, and PRJNA79337, for the species listed in this figure, from left to right, respectively. Species name (in order from left to right): Hirsutella thompsonii, Hypocrella siamensis, Isaria farinose, Metarhizium acridum, Metarhizium anisopliae, Myceliophthora thermophila, Neotyphodium aotearoae, Neotyphodium gansuense (1), Neotyphodium gansuense var. inebrians (2), Tolypocladium inflatum, Ashbya gossypii, Eremothecium cymbalariae, Saccharomycetaceae sp. ‘Ashbya aceri’, Scheffersomyces stipites, and Thielavia terrestris. Note that genome size is larger on average in the primarily asexually reproducing fungi than in the partly sexually reproducing fungi. Source: This study.

species for which the genomes have been sequenced and assembled (http://www.ncbi. nlm.nih.gov/genome/). The results are presented in Figure 10.6. Genome size is significantly greater in the primarily asexually reproducing fungi than in partly sexually reproducing fungi (Figure 10.6A). There was no significant difference in GC content

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10.9  Concluding Remarks

between the two groups (Figure 10.6B). More detailed discussion on DNA mutation mechanisms and somatic genome instability in yeast and other fungi can be found in Chapter 9 (this study). Further research is required to determine whether this genome size increase is a mechanism that drives genetic variation in primarily asexually reproducing fungi. Asexual species lack the opportunity of generating variation through meiosis and therefore may rely on genome rearrangements and result in large genomes.

10.8 ­Variation of Gene Direction In some sequenced genomes, certain genes that have similar function or coordinated expression appear to be clustered (Hurst et al. 2004). Gene arrangement was found to be conserved in various bacteria (Junier et  al. 2012) and eukaryotic organisms (McDonagh et al. 2000; Hurst et al. 2004; Ivens et al. 2005; Peacock et al. 2007; Michalak 2008; Tungsuchat-Huang et al. 2010; Downing et al. 2011). The range of DNA neighborhood that facilitates biochemical interactions might therefore be defined by DNA topology (Junier et  al. 2012); however, gene expression was not significantly altered after the member of one well-defined gene expression neighborhood in Drosophila was separated (Meadows et al. 2010). An unknown mechanism must exist to ensure animal fitness after members of previously defined gene blocks are physically split off (Li and Du 2012). Li and Du (2012) conducted a large-scale analysis of genomes/chromosomes and detected clear differences in gene direction between kingdoms, including archaeans and bacteria, two kingdoms that do not have meiotic segregation. At the chromosome level, all the archaeal and bacterial chromosomes have more same-direction genes than opposite-direction genes; however, at the gene neighborhood level, there is a clear difference between archaeans and bacteria. In archaeans, 66% of the 185,703 neighbor genes analyzed are in the same direction; whereas, in bacteria, 71% of 7,036,353 neighbor genes are in the same direction. Since archaeans and bacteria share the same ancestor but never have had meiotic cell divisions, this general difference in gene direction on chromosomes between the two kingdoms must be variations created though non-­ meiotic approaches.

10.9 ­Concluding Remarks Considerable variation in genome size, GC content, and gene direction in living organisms that do not reproduce through meiosis is attributable to non-meiotic approaches, which likely include multiple molecular mechanisms. The large genome size of certain bacterial species is likely associated with various mechanisms that operate in most other bacterial genomes as well. Similarly, the extensive variation in C+G or GC content characterizing prokaryotes is probably also controlled by multiple mechanisms, since GC content and genome size are strongly correlated in Actinobacteria and Proteobacteria, but not in most other bacteria. The underlying biology responsible for this correlation difference is still unclear. Since genes are usually GC-rich, it is interesting that some

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bacteria have extremely small genome and extremely low GC content. For example, the Candidatus C. ruddii genome is only 0.162 Mb with a GC content of 14.55%. The genome size is even smaller than a plasmid Ti (0.214 Mb, AE007871.2) in A. tumefaciens. Some bacteria have GC content up to 73.28% (C. ruddii, AP012338.1). By what mechanisms do these bacterium ensure its introns and three-prime gene regions work effectively under such an overall high genomic GC content? Are the mutation system and the evolutionary selection system different between these extremely high GC- and extremely low GC-content bacteria? In addition to genome size and GC content, the percentage of neighbor genes on the same direction is higher in bacteria than in archaeans. What non-meiotic mechanisms led archaeans and bacteria to generate the overall difference between the two kingdoms in neighbor gene directions on chromosomes during evolution? Answering these questions may help us to understand how nonmeiotic (or somatic in the broad sense) approaches create these extensive variations, identify the mechanisms of adaptation and evolution of prokaryotes, improve molecular breeding of useful prokaryotes, predict the genome characteristics of future strains (particularly for pathogens), and assist the design of novel species in synthetic biology. ­Acknowledgments We sincerely thank Dave Bartlett, Dan LeBlanc, and Martin Lague for their IT support and general management of computer servers, and Professor Donglei Du for his support in mathematical analysis and programming.

­References Appels R., Dennis E.S., Smyth D.R. and Peacock W.J. (1981) Two repeated DNA sequences from the heterochromatic regions of rye (Secale cereale) chromosomes. Chromosoma 84:265–277. Chargaff E., Lipshitz R. and Green C. (1952) Composition of the desoxypentose nucleic acids of four genera of sea-urchin. J. Biol. Chem. 195:155–160. Cionini P.G., Bassi P., Cremonini R. and Cavallini A. (1985) Cytological localization of fast renaturing and satellite DNA sequences in Vicia faba. Protoplasma 124:106–111. Downing T., Imamura H., Decuypere S., Clark T.G., Coombs G.H., Cotton J.A., Hilley J.D., De Doncker S., Maes I., Mottram J.C., et al. (2011) Whole genome sequencing of multiple Leishmania donovani clinical isolates provides insights into population structure and mechanisms of drug resistance. Genome Res. 21:2143–2156. Gardiner K. (1996) Base composition and gene distribution: critical patterns in mammalian genome organization. Trends Genet. 12:519–524. Groth J.V. and Roelfs A.P. (1982) Effect of sexual and asexual reproduction on race abundance in cereal rust fungus populations. Phytopathology 72:1503–1507. Hurst L.D., Pál C. and Lercher M.J. (2004) The evolutionary dynamics of eukaryotic gene order. Nat. Rev. Genet. 5:299–310. Ivens A.C., Peacock C.S., Worthey E.A., Murphy L., Aggarwal G., Berriman M., Sisk E., Rajandream M.A., Adlem E., Aert R., et al. (2005) The genome of the kinetoplastid parasite, Leishmania major. Science 309:436–442.

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Joos H., Timmerman B., Van Montagu M. and Schell J. (1983) Genetic analysis of transfer and stabilization of Agrobacterium DNA in plant cells. EMBO J 2:2151–2160. Junier I., Hérisson J. and Képès F. (2012) Genomic organization of evolutionarily correlated genes in bacteria: limits and strategies. J. Mol. Biol. 419:369–386. Li X.-Q. (2014) Comparative analysis of the base compositions of the pre-mRNA 3′ cleaved-off region and the mRNA 3′ untranslated region relative to the genomic base composition in animals and plants. PLoS ONE 9:e99928. Li X.-Q. (2015a) Laboratory methods for investigating nuclear and cytoplasmic genomes and transcriptome. In: X-Q Li, DJ Donnelly, TG Jensen, editors. Somatic genome manipulation: advances, methods and applications. New York: Springer, pp 323–352. Li X.-Q. (2015b) Bioinformatic approaches for analysis of gene direction, chromosome base composition, mRNA polyadenylation, and protein network. In: X-Q Li, DJ Donnelly, TG Jensen, editors. Somatic genome manipulation: advances, methods and applications. New York: Springer, pp 353–363. Li X.-Q. and Du D. (2012) Gene direction in living organisms. Sci. Rep. 2(982). Li X.-Q. and Du D. (2014a) Motif types, motif locations and base composition patterns around the RNA polyadenylation site in microorganisms, plants and animals. BMC Evol. Biol. 14:162. Li X.Q. and Du D. (2014b) Variation, evolution, and correlation analysis of C+G content and genome or chromosome size in different kingdoms and phyla. PLoS ONE 9:e88339. Liu H., Wu Z., Li M., Zhang F., Zheng H., Han J., Liu J., Zhou J., Wang S. and Xiang H. (2011) Complete genome sequence of haloarcula hispanica, a model haloarchaeon for studying genetics, metabolism, and virus–host interaction. J. Bacteriol. 193:6086–6087. Liu X., Miao D., Zhang F., Wu Z., Liu J. and Xiang H. (2013) Characterization of the minimal replicon of pHM300 and independent copy number control of major and minor chromosomes of Haloferax mediterranei. FEMS Microbiol. Lett. 339:66–74. McDonagh P.D., Myler P.J. and Stuart K. (2000) The unusual gene organization of Leishmania major chromosome 1 may reflect novel transcription processes. Nucleic Acids Res. 28:2800–2803. Meadows L.A., Chan Y.S., Roote J. and Russell S. (2010) Neighbourhood continuity is not required for correct testis gene expression in Drosophila. PLoS Biol. 8:e1000552. Michalak P. (2008) Coexpression, coregulation, and cofunctionality of neighboring genes in eukaryotic genomes. Genomics 91:243–248. Mitchell D. (2007) GC content and genome length in Chargaff compliant genomes. Biochem. Biophys. Res. Commun. 353:207–210. Mooers A.Ø. and Holmes E.C. (2000) The evolution of base composition and phylogenetic inference. Trends Ecol. Evol. 15:365–369. Musto H., Naya H., Zavala A., Romero H., Alvarez-Valín F. and Bernardi G. (2006) Genomic GC level, optimal growth temperature, and genome size in prokaryotes. Biochem. Biophys. Res. Commun. 347:1–3. Narra H.P. and Ochman H. (2006) Of what use is sex to bacteria? Curr. Biol. 16:R705–R710. Nellåker C., Li F., Uhrzander F., Tyrcha J. and Karlsson H. (2009) Expression profiling of repetitive elements by melting temperature analysis: variation in HERV-W gag expression across human individuals and tissues. BMC Genomics 10:532. Nishida H. (2012) Evolution of genome base composition and genome size in bacteria. Front. Microbiol. 3:420.

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Peacock C.S., Seeger K., Harris D., Murphy L., Ruiz J.C., Quail M.A., Peters N., Adlem E., Tivey A., Aslett M., et al. (2007) Comparative genomic analysis of three Leishmania species that cause diverse human disease. Nat. Genet. 39:839–847. Presber W., Schroeder C. and Krueger D.H. (1976) The evolution of mutation rates via DNA base composition and mutator genes: a proposition. J. Theor. Biol. 59:353–360. Schell J. and Van Montagu M. (1977) The Ti-plasmid of Agrobacterium tumefaciens, a natural vector for the introduction of nif genes in plants? Basic Life Sci. 9:159–179. Šmarda P. and Bureš P. (2012) The variation of base composition in plant genomes. In: JF Wendel, J Greilhuber, J Doležel, IJ Leitch, editors. Plant genome diversity. Vienna: Springer, pp 209–235. Šmarda P., Bureš P., Šmerda J. and Horová L. (2012) Measurements of genomic GC content in plant genomes with flow cytometry: a test for reliability. New Phytol. 193:513–521. Stȩpkowski T. and Legocki A.B. (2001) Reduction of bacterial genome size and expansion resulting from obligate intracellular lifestyle and adaptation to soil habitat. Acta Biochim. Pol. 48:367–381. Trevors J.T. (1996) Genome size in bacteria. Antonie van Leeuwenhoek 69:293–303. Trevors J.T. (1998) Molecular evolution in bacteria: genome size, cell size, restrictionmodification and recognition. Bull. Inst. Pasteur 96:25–33. Tungsuchat-Huang T., Sinagawa-García S.R., Paredes-López O. and Maliga P. (2010) Study of plastid genome stability in tobacco reveals that the loss of marker genes is more likely by gene conversion than by recombination between 34-bp loxP repeats. Plant Physiol. 153:252–259. Watson J.D. and Crick F.H.C. (1953) Molecular structure of nucleic acids: a structure for deoxyribose nucleic acid. Nature 171:737–738. Wu Z., Liu J., Yang H., Liu H. and Xiang H. (2014) Multiple replication origins with diverse control mechanisms in Haloarcula hispanica. Nucleic Acids Res. 42:2282–2294.

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11 RNA Polyadenylation Site Regions: Highly Similar in Base Composition Pattern but Diverse in Sequence—A Combination Ensuring Similar Function but Avoiding Repetitive-Regions-Related Genomic Instability Xiu-Qing Li1* and Donglei Du2 1 2

Fredericton Research and Development Centre, Agriculture and Agri-Food Canada, Fredericton, New Brunswick, Canada Quantitative Methods Research Group, Faculty of Business Administration, University of New Brunswick, Fredericton, Canada

Abstract The genomes of eukaryotes, such as higher plants and animals, may contain tens of thousands of genes. The coding sequences are expected to be very diverse because almost every gene has a specific function. However, the genic region corresponding to the 3′ end of the precursor mRNA performs a very similar function: it serves as a signature for polyadenylation (poly(A) or polyA) of the gene’s transcripts. How can a genome have so many repeated regions with a similar function but still be stable without undergoing excessive homologous DNA recombination in somatic and germline cells? From analyses of the poly(A) site regions in microorganisms, plants, and animals, we found that these regions are highly similar in base composition pattern and, to a certain degree, in motifs, but low in overall sequence similarity. This suggests that the short motifs and base composition patterns enable proper folding, poly(A) complex protein binding, and polyadenylation in the RNA region. Poor overall sequence conservation can minimize repetitive region-caused DNA homologous recombination. Most of the RNAs and reference genomes (e.g., the potato genome) that we analyzed were from diverse somatic cells. The distances from the base abundance peaks to their poly(A) sites were found to be highly similar between plant and animal kingdoms and among their subkingdoms. The results suggest that the similarity in base composition pattern in the poly(A) site region ensures similar function and the poor overall sequence similarity helps to avoid recombination-caused genome instability. Keywords  poly(A) site; genome stability; genome evolution; RNA cleavage; repetitive function; sequence divergence; gene expression regulation; RNA structure; upstream A-rich ­elements; upstream U-rich elements

*Corresponding author: [email protected]; [email protected]; [email protected]

Li X.-Q. and Du D. (2017) RNA polyadenylation site regions: highly similar in base composition pattern but diverse in sequence—a combination ensuring similar function but avoiding repetitive-regions-related genomic instability. In: Li X.-Q., editor. Somatic Genome Variation in Animals, Plants, and Microorganisms: Wiley-Blackwell, Hoboken, NJ, Ch. 11, pp. 267–290. Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

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11  RNA Polyadenylation Site Regions: Highly Similar in Base Composition Pattern but Diverse in Sequence

11.1 ­General Introduction to Gene Number, Direction, and RNA Polyadenylation All genomes rely largely on genes for their functioning. Genome size and base composition vary greatly as life complexity increases (Li and Du 2014a). The average number of genes per genome ranges from 7,152 in protozoan protists to 32,720 in plants (Li and Du 2012). In some species such as protists, most of the genes show same-direction dominance. For example, there are 391 genes in tandem in Leishmania infantum (a protozoan ) (Li and Du 2012). It is known that homologous DNA sequences can undergo recombination in both reproductive cells (through meiosis) and somatic cells (somatic recombination). If genes have sufficient sequence similarity for recombination between them, these same-direction genes would serve as same-directional DNA repeats in the genome and cause frequent gene deletion or duplication, greatly destabilizing the genome. The coding sequence regions of genes are expected to be diverse because each gene or each small group of genes has its own specific function. However, all or nearly all of the RNA poly(A) site regions of the genes have one main function—to serve as a signature for proper polyadenylation of transcripts (mRNA) of the genes. How can a genome have poly(A) site regions with a very similar function but still avoid homologous DNA recombination-caused genome instability? This chapter attempts to address this question through an analysis of the DNA sequence motifs, motif distribution, base composition pattern, and DNA sequence conservation of the poly(A) site regions in comparison with the whole genome. In gene expression, genes are transcribed into precursor mRNAs. The 3′-end fragment of pre-mRNA is usually cleaved off, and the newly formed 3′ end is polyadenylated to generate a polyadenine tail (poly(A) tail) (Chambers and Old 1988; Yamanaka et al. 2010; Yang and Doublié 2011). The polyadenylated mature mRNA is then translated into peptide(s), which directly affects phenotype. The process of RNA cleavage and polyadenylation greatly influences RNA stability and therefore regulates the RNA lifespan and consequently gene functions. Knowledge of the characteristics and evolutionary selection of the poly(A) site is therefore important for both basic research and genetic engineering. The RNA poly(A) site region includes the three-prime untranslated region (3′UTR), the poly(A) site, and the three-prime cleaved-off region (3′COR) (Figure 11.1) (Li 2014). The exact poly(A) site is the phosphodiester bond that connects the last nucleotide of cleaved RNA and the first adenosine of the poly(A) tail, which means it is the bond between the two nucleotides. The 5′ nucleotide and the 3′ adenosine of this poly(A) site bond are called the poly(A) tail attachment position and the poly(A) tail starting position, respectively. However, since the exact bond location is not known for many transcripts, the poly(A) tail starting position is assigned to the genomic or pre-mRNA nucleotide that corresponds to the first A of the poly(A) tail (Li and Du 2013). If the phosphodiester bond that marks the poly(A) site is called position zero, the bond­connected upstream nucleotide can be called position −1 and the corresponding downstream nucleotide can be called position 1. In this chapter, we attempt to explain how poly(A) site regions can have a similar function without an adverse effect on genome stability by describing base composition selection at the poly(A) site and in the 3′UTR and 3′COR, along with the polyadenylation signal motifs in microorganisms, plants, and animals, and by drawing comparisons with the whole genome wherever possible.

11.2  Base Selection at the Poly(A) Tail Starting Position

Figure 11.1  Poly(A) site region, presented using a UC poly(A) site as an example. In this illustration, the poly(A) tail attachment position (position −1) is a thymine. The poly(A) tail starting position (position 1) is a cytosine (C). In this chapter, we focus on the region from −100 to 100, which is a 201-base region. The poly(A) tail starting position is often loosely called ‘the poly(A) site’ in the literature. Note that although this illustration uses a C for the poly(A) tail starting position, in most species an adenine (A) occupies this position.

11.2 ­Base Selection at the Poly(A) Tail Starting Position The poly(A) tail starting position is usually an adenosine in most species (Li and Du 2013). All of the species listed in Table 11.1 have A predominance at the poly(A) tail starting position, except that this A predominance is absent (or weaker than G Table 11.1  Most-frequent hexanucleotide motifs within 50 nucleotides upstream of poly(A) tail starting positions in each species. Mapped unique mRNA (no.)

Most frequent 6mer motif within 50 bases upstream a

Percentage

209

AAUAAA

19.6

Blastocystis hominis

1 717

AAGAAG

10.34

Chlamydomonas reinhardtii

176

UGUAAC

31.3

Phytophthora infestans

63

AAAAUA c

20.6

Trypanosoma cruzi

52

UGUUUU

38.5

Arabidopsis thaliana

4 431

AAUAAA

13.4

Medicago truncatula

136

UAUUUU

17.7

Populus trichocarpa

1 371

UAUUUU

12.6

Solanum lycopersicum

450

UAUUUU

12.9

Species/group

Microorganisms Fungi b

Protists

Dicotyledonous plants

(Continued)

269

270

11  RNA Polyadenylation Site Regions: Highly Similar in Base Composition Pattern but Diverse in Sequence

Table 11.1  (Continued) Mapped unique mRNA (no.)

Most frequent 6mer motif within 50 bases upstream a

Percentage

Oryza sativa

693

AAUAAA

15.3

Sorghum bicolor

1 685

AAUAAA

11.2

Zea mays

10 491

AAUAAA

9.0

Apis mellifera

187

AAUAAA

53.5

Bos taurus

2 679

AAUAAA

63.6

Caenorhabditis elegans

389

AAUAAA

50.6

Callithrix jacchus

79

AAUAAA

55.7

Canis lupus familiaris

108

AAUAAA

68.5

Ciona intestinalis

287

AAUAAA

48.4

Danio rerio

7 246

AAUAAA

61.7

Drosophila melanogaster

954

AAUAAA

59.0

Equus caballus

97

AAUAAA

46.4

Gallus gallus

788

AAUAAA

58.8

Homo sapiens

30 499

AAUAAA

58.2

Mus musculus

8 709

AAUAAA

61.2

Oryctolagus cuniculus

224

AAUAAA

59.8

Pongo abelii

1 965

AAUAAA

58.5

Rattus norvegicus

14 263

AAUAAA

56.0

Sus scrofa

8 114

AAUAAA

40.5

Taeniopygia guttata

808

AAUAAA

59.9

Animal average

4 116

AAUAAA

53.9

Species/group

Monocotyledonous plants

Animals

a) The AAUAAA motif could be found in 11.8% of unique poly(A) sites in monocotyledonous plants and in 11.9% of unique poly(A) sites in all plants (average of all four dicotyledonous plants and all three monocotyledonous plants together). Frequencies of these motifs are all significantly higher than the mean frequencies of hexanucleotide motifs in the region according to the Chi-squared test (P 500 bp), lack inverted terminal repeats and direct repeats, and imprecisely excised; however, a small number of IESs (not shown in figure) are short (< 500 bp), flanked by TTAA, and precisely removed (Fass et al. 2011). E: Many eukaryotes have both autonomous and non‐autonomous transposable elements analogous to ciliate transposon/IES systems. Autonomous elements are self‐mobilizing and encode functional transposase enzyme, while non‐autonomous versions (miniature inverted‐repeat transposable elements (MITEs)) lack the coding sequence for functional transposase and therefore rely upon the enzyme produced from autonomous elements to mediate their transposition. Source: Chen et al., 2014; Fass et al. 2011; Smit and Riggs 1996; Casacuberta and Santiago 2003; Wessler 2006; Yang et al. 2009. Reproduced with permission of Springer.

18

mRNA (C/G)

16

Attach. C/G of non-A sites

C/G ratio

14 12 10 8 6 4 2 0

1

2

3

4

5

6

7

8

9

12 animal species

10 11 12 13 14 15 16 17 18 3 dicots, 3 monocots

Figure 11.2  C/G ratios at the poly(A) tail attachment position of non-A-type poly(A) transcripts. Species were sorted from smallest to largest (from left to right) by C/G ratio at the poly(A) tail attachment position, giving the following ranking: dog, rabbit, rat, zebrafish, mouse, cattle, zebra finch, orangutan, chicken, human, pig, fruit fly. The three dicot plants were ranked as follows: M. truncatula, A. thaliana, poplar. The order for the three monocot plants was rice, maize, and sorghum. Note that the C/G ratio at the poly(A) tail attachment position clearly separates the three groups: animals < dicots < monocots (Li and Du 2013).

7 6

AAUAAA AAUAAA, –21, 6.12% CAAAAA, –1, 5.22%

mRNA (%)

5

UAAAAA, –1, 4.61%

UAAAAA AAAAAU AAAAUA AUUUUU

UUAAAA, –2, 3.32%

2

AAAUAA AUAAAA

UUAAAA

4 3

UUUUUU

UUUUUA

AAAUAA, –21, 2.80%

UAUUUU

AUAAAA, –20, 2.53%

UUUUAA

UUUAAA, –3, 2.11% AUAAAU, –20, 1.62% UUUUUU, 19, 1.12%

1

UUUAAA UUAUUU UUUAUU UGUUUU UAAAAU UUUUCU

0 –100 –95 –90 –85 –80 –75 –70 –65 –60 –55 –50 –45 –40 –35 –30 –25 –20 –15 –10 –5 1 6 11 16 21 26 31 36 41 46 51 56 61 66 71 76 81 86 91

AUAAAU CAAAAA

Position

Figure 11.3  Most frequent 20 hexamer motifs in the poly(A) site region of 30,499 unique poly(A) sites mapped to the human genome. Order of peak point labeling: motif sequence, position, and frequency at a single specific position in the mapped poly(A) site regions (pre-mRNA). The poly(A) sites are unique because each group of identical poly(A) sites was counted as one, which means that each of the 30,499 poly(A) sites was unique. Note that the most frequent motifs are mainly in three places: the −21 position region (upstream A-rich element; mainly AAUAAA), the poly(A) site, and the downstream U-rich region with peak location approximately at +19. Note also that the five most abundant motifs in the 201 bases (including 3′UTR, the poly(A) site, and 3′COR) in terms of values of highest peaks were AAUAAA, CAAAAA, UAAAAA, UUAAAA, and AAAUAA (Li and Du 2014b).

os Ca oph A en ila pis or me m ha la el bd no life itis ga ra ele ste ga r ns Da Ta en Ga nio No io llu re py s r ni m am gia gall o gu us m tta al ta AV ER AG E Ra Mu Bos ttu s m tau s u ru no sc s P rv ulu Ho ong egic s m o a us M o b am sa el m pie ii al ns AV An ER im AG al Ar AV E a ER M bid ed op AG s i Po ca is E g So pulu o t thal la s t run ian Di nu ri ca a co m cho tu tyl tu ca la ed be rp on ro a ou su sA m VE RA O GE So r M on rg yza oc hu s ot m at yle iv Ze bico a do a lor no m us ay AV s ER Pl an AG tA E O VE ve RA ra ll A G E VE RA G E

Dr

3ʹUTR/Genome Ratio

ro s C oph A ae i p no la m is rh e m ab la ell di no ife tis ga ra el ste eg r an Ta D s en G an N on io allu io py s re -m am gia ga rio gu llus m al t AV tata ER AG R Mu Bos at s t E tu m au s u r n s u P orv cu s H ong eg lus o M o ic m am o ab us sa e m pi lii al en An AV ER s im a A Ar lA G E a M bid VER ed o A p Po ica si G E s So pu go th D la lus tru alia ic nu tr n n ot m ich cat a yl ed tu oc ula on be a ou ro rpa s s AV um ER M AG So O on rg ryz E oc hu a ot m sa yl ed bi tiva Z on ea co ou m lor s AV ays Pl E an R t A AG O E VE ve R ra A ll AV GE ER AG E

D

Base Composition (%) 40

45

35 3ʹUTR A

30 3ʹUTR C

3ʹUTR G

25 3ʹUTR U

20 Genome A, T

15 Genome C, G

10

5

1.4

(a)

1.3

1.2

1.1 A 3ʹUTR/Genome Ratio

1.0

0.9 C 3ʹUTR/Genome Ratio

0.8

0.7 G 3ʹUTR/Genome Ratio

0.6

0.5

U 3ʹUTR/Genome Ratio

(b)

Figure 11.5  RNA base compositions of the 3′UTR region, represented by 100 bases upstream of the poly(A) site. (a) Base compositions of 3′UTR and whole genome of each species. (b) 3′UTR/genome ratios of base composition. Note that the U-content difference between dicotyledonous and monocotyledonous plants is smaller in the 3′UTR region than in the whole genome, likely due to a higher overrepresentation of T content in terms of UTR/genome ratio in monocotyledonous plants (Li 2014).

90

Plants

Base composition (%)

80 70 60 50

A

40

C

30

G

20

U

10 –100 –95 –90 –85 –80 –75 –70 –65 –60 –55 –50 –45 –40 –35 –30 –25 –20 –15 –10 –5 1 6 11 16 21 26 31 36 41 46 51 56 61 66 71 76 81 86 91 96 101

0 Base position Animals

Base composition (%)

90 80 70 60 50

A

40

C

30

G

20

U

10 –100 –95 –90 –85 –80 –75 –70 –65 –60 –55 –50 –45 –40 –35 –30 –25 –20 –15 –10 –5 1 6 11 16 21 26 31 36 41 46 51 56 61 66 71 76 81 86 91 96 101

0 Base position

Figure 11.10  Average base abundance at each position of the 201-nucleotide poly(A) site region, showing the U-A-U-SiteA-U (U-A-U-A-U for short) base abundance pattern in plants and animals. Species for plants: Arabidopsis thaliana, Medicago truncatula, Oryza sativa, Populus trichocarpa, Solanum lycopersicum, Sorghum bicolor, Zea mays. Species for animals: Bos taurus, Danio rerio, Drosophila melanogaster, Gallus gallus, Homo sapiens, Mus musculus, Pongo abelii, Rattus norvegicus, Taeniopygia guttata. Position 1 is the poly(A) tail starting position (usually called ‘the poly(A) site’). Position −1 is the poly(A) tail attachment position. Note that base composition analysis showed several clear differences between plants and animals. For example, the upstream A peak was lower but the U peak was higher in plants in comparison with animals. The shape of the downstream U peak was flat in plants but more pointed in animals. A small A peak at position −3 in plants and at −5 in animals exists, in addition to the U-A-U-SiteA-U (or U-A-U-A-U) base abundance pattern. Clearly, the U-A-U-A-U base abundance pattern is more conserved than any known motifs (Li and Du 2014b). (a)

(b)

(c)

Figure 14.1  Karyotype showing maize cell ploidy levels under a microscope. (a) A diploid root cell (2n = 2x = 20); (b) a diploid leaf cell (2n = 2x = 20); and (c) a polyploid leaf cell (2n = 4x = approx. 40). The arrow in (a) points to two B‐chromosome‐like structures. Source: From Ma and Li 2015.

405

Index a acytokinetic mitosis  4 Aegilops speltoides 316 Aegilops umbellulata   315 agricultural implications of somatic genome variation (SGV)  385 bud mutation  389 cellular‐level variation  385–386 dedifferentiation‐ and redifferentiation‐ induced variation  388 DNA damage  389 DNA exchange  391 DNA transfer  390–391 endoploidy variation  387 epigenetics 389 gene mutation  389 intercompartmental interaction  391 intra‐ and interchromosomal variation 387–388 mitochondrial genome sequence or DNA amount variation  390 organelle genome segregation  390–391 organelle transmission  390–391 plastid genome sequence or DNA amount variation  389 ploidy and chromosome number variation 386–387 Agrobacterium tumefaciens  254, 264, 343, 368, 390–391 AKT (serine kinase)  292 diseases caused by somatic mutations 293–294 alternative non‐homologous end‐joining (alt‐NHEJ) 142

Alzheimer’s disease  342 amyloplasts 120 aneuploidy  4, 26 cancerous masses  316 chromosomal instability (CIN)  59, 60–61 plants 315–316 embryonic stem cells (ESCs)  56 mesenchymal stem cells (MSCs)  59 yeasts 230–231 Anillin 25 animals base composition of  3′COR 277–278 base composition of  3′UTR 276 compared with whole genome 276–277 base composition of poly(A) site region 280 compared with plants  280–284 endopolyploidy 314 genomic GC content  257–259 antibody DNA mutation  382 APC/C  8, 11, 12, 14, 16, 20 Apis mellifera  258, 270, 277, 282, 325, 381 APOBEC3B DNA cytosine deaminase 383–384 Arabidopsis thaliana  120, 124, 146, 147, 258, 269, 277, 281, 314 chloroplast genome  318–319 DNA damage  389 DNA repair  341, 354 DNA transposition  343 somatic crossover suppression  388 telomere degeneration  344

Somatic Genome Variation in Animals, Plants, and Microorganisms, First Edition. Edited by Xiu-Qing Li. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

406

Index

archaeans chromosome data  260 genome size distribution  256–257 genome size variation  255–256 genomic GC content  257–259 Armillaria tabescens 315 array‐based comparative genomic hybridization (aCGH)  9 ascorbate peroxidases (APXs)  136 Ashbya gossypii 262 Astasia longa 318 Atriplex confertifolia 311 avirulence (Avr) genes  108–109

b B chromosomes  314, 386 Babesia bovis 278 bacteria genome size distribution  256–257 genome size variation  255–256 genomic GC content  257–259 ploidy 313 base excision repair (BER)  93, 137 Beta vulgaris  312, 389 Blastocystis hominis  269, 270, 274, 275, 276, 278–279 Brachycome dichromosomatica 387 Brassica juncea 324 Brassica napus  110, 111, 321, 322–323, 386, 390 Brassica rapa  107, 110, 323, 386 break‐induced replication (BIR) repair pathway 227–229 breeding criteria  368–370 bud mutation  389

c Caenorhabditis elegans  8, 201, 258, 270, 272, 273, 277 cell fusion  4 EFF‐1 protein  380 endoreplication 10–11 polyploidization 6 Canavalia ensiformis 295 Candida albicans  202, 204, 207, 208, 210, 212, 214, 216, 230 haploidization 315

Candidatus C. ruddii  256, 259, 264 Candidatus Phytoplasma mali 257 cardiomyocytes 23 polyploidization mechanisms 24–25 response to tissue damage  25 upstream control  23–24 Cdc20/fizzy  8, 30 Cdc25/string  8, 13 Cdh1/fizzy‐related  8, 20 Cdt1/double‐parked  9, 18 cell cycle machinery  8–9 cell fusion  4, 378, 380, 385 cellular‐level variation  385–386 centrosomes 61–62 Checkpoint Kinase‐1 (Chk1)  20 Chlamydomonas reinhardtii  121, 129, 139, 141, 143, 146 motifs  269, 270, 272, 275, 279, 284 chloroplast mutator  1 (CHM1) 144 chloroplast RECA (cpRECA)  143, 145 chloroplasts  120, 133, 135 Chorion genes  7, 17, 18 chromatin  9, 19, 29, 51 elimination 168 modifications 186–187 chromosomal passenger complex (CPC) 61 chromosome instability (CIN) in stem cells (SCs) 55–56 mechanisms of CIN  59–60 kinetochore microtubule attachment defects 60–61 sister chromatids cohesion  62 spindle assembly checkpoint (SAC) dysfunction 60 supernumerary centrosomes 61–62 mechanisms of CIN in SCs  63 pluripotent stem cells (PSCs) mouse ESCs  57 parthenogenetic ESCs  57–58 primate ESCs  56–57 somatic stem cells (SSCs) mesenchymal stem cells (MSCs) 58–59 neural stem cells (NSCs)  59

Index

chromosome integrity mechanisms 229–230 aneuploidy in yeasts  230–231 ploidy changes in yeasts  231–232 mechanisms 232–234 chromosome number in prokaryote species 254 Chrysanthemum morifolium 389 ciliates, RNA‐mediated somatic genome rearrangement  168, 187 background 168–170 chromatin modifications  186–187 long noncoding RNA templates as interface for short noncoding RNA 177–179 long noncoding RNA templates in genome rearrangement  176–177 non‐Mendelian inheritance in Paramecium 171–173 nuclear dimorphism  170–171 phase variation  185 programmed genome engineering  185 scanRNA model in Tetrahymena 173–175 short RNA‐mediated heterochromatin formation 179–182 small RNA in Stylonychia and Oxytricha 175–176 transpoases 186–187 transposable elements and genome rearrangments 182–185 transposons 185 VDJ recombination  186–187 ciprofloxacin  123, 143, 144 clonal variation  315, 338, 351, 364 CLOVES syndrome  294 Coffea arabica 386 copy number variation (CNV)  144 CoREST transcription cofactor  13 cryptochromes 140 C‐value  3, 9 cyclin A (CycA)  8, 12, 14, 17, 20–21 cyclin B (CycB)  8, 12, 20–21 cyclin E (CycE)  8, 10, 12, 15–16 cyclin‐dependent kinase (CDK) complexes  8, 9, 12, 14 cyclins  8, 12

cyclobutane pyrimidine dimers (CPDs)  138, 139–140 cytokinesis 8 cytoplasmic male sterility (CMS)  105, 108–109, 321–322 mitochondrial sublimons  109–111

d Dacapo (Dap)  8, 12–13, 17 Datura innoxia 312 dedifferentiation‐ and redifferentiation‐ induced variation  388 Deinococcus radiodurans  301, 341 developmental age  353 developmental variation in nuclear genome primary sequence  299–300, 304–305 antibody gene generation through somatic genome variation  303 developmental variation in DNA sequences 303 DNA damage  300–302 DNA protection  300–302 gene conversion in somatic cells 300–302 genetic mutation  300–302 heritability and stability of developmentally induced variation 303–304 programmed large‐scale somatic genome variation 302 Dianthus broteri 313 D‐loops (displacement loops)  126–129 DNA amplification mechanisms  17–19 DNA cytosine deaminases  340, 381–384 DNA damage  92, 300–302, 389 somatic genome variation (SGV) 338–339 protective pigments  340–341 RNA‐templated repair  341 UV‐induced 138–140 DNA disturbances causes and consequences  233 DNA elimination  180–181 DNA exchange  391 DNA instability, repetitive  342 DNA lesions  219–221

407

408

Index

DNA methylation  92, 93–94, 180 DNA polymerases (DNA Pols)  200 as guardians of genome maintenance 201–205 DNA protection  300–302 DNA recombination  141–145 DNA repair  92–93, 133 double‐strand breaks (DSBs)  141–145 errors in DNA repair  339, 341–342, 345 oxidative stress, photo‐adaptation and ROS detoxification  133–138 recombination 141–145 UV‐induced DNA damage  138–140 DNA transfer  390–391 DNA transportation  343 DNA, extracellular  343 double‐strand breaks (DSBs) in DNA  93, 122, 132 repair (DSBR)  141–145, 227–229 double‐strand DNA (dsDNA)  341 Dp  21, 22 Drosophila amplicons in follicle cells (DAFCs) 17–19 Drosophila melanogaster  6, 7, 258, 270, 273, 277, 282, 283 endocycle oscillations alternative endoreplication modes  17 salivary gland  15–16 endoreplication 10–11 endoreplication initiation endocyte entry in ovarian follicle cells 11–13 endocyte entry into other tissues 14–15 mitotic‐to‐endocyte switch regulation 13–14 gene amplification in follicle cells endocycle‐to‐amplification switch  19 molecular mechanism  17–19 polyploidization 6 somatic mutations  352, 359

e E2F  8, 14–19, 21–22, 27 E2F‐mediated repression  14 E2F1 transcriptional activator  15

embryonic stem cells (ESCs)  55–56 PCSs and iPSCs  56–57 Encephalitozoon cuniculi 321 endocycle‐to‐amplification switch  19 endomitosis 5 endoplasmic reticulum stress  344 endoploidy variation  381, 387 endopolyploidy 313–315 endoreplication  5–7, 10–11 endosymbiosis  138, 316, 318 endosymbiont gene transfer (EGT)  104 Entamoeba histolytica 321 environmentally induced genome instability  91–92, 97 stress, effects of  92 DNA repair  92–93 epigenetic changes  93–95 epigenetic changes, link with genetic changes 95–96 genetic changes  92 transgenerational inheritance  96 epigenetic changes stress response  93 DNA methylation  93–94 histone modification  95 link with genetic changes  95–96 epigenetic variation  365–367 epigenetics 389 Escargot (esg)  14–15, 20 ethanol effects and oxidative stress  340 etioplasts 120 euchromatin 49 evolutionary implications of somatic genome variation (SGV)  370–371 executer mutant of Arabidopsis 135 expressed sequence tags (ESTs)  270, 271–272

f FAK‐related non‐kinase (FRNK)  24 fibroadipose overgrowth (FAO)  294 flavin adenine dinucleotide (FAD)  139 flax, induced inheritable genome variation  77–79, 87 adaptive genomic changes  83–84 environmental stress response  83

Index

genome restructuring  79–80 similar responses in other plants  84–87 specific genomic changes  80–82 flu mutant of Arabidopsis 135 focal adhesion kinase (FAK)  24 follicle cells  9, 11, 15–16 endocyte entry  11–13 gene amplification  17–19 signaling pathways  13–14 fungi chromosome data  260 DNA polymerase complexes  202 genomic GC content  257–259 genomic GC content and size in asexually reproducing fungi 260–263 mitotic genome variation

g Geminin  9, 16, 21 gene amplification  7 gene conversion  343 somatic cells  300–302 Gene definition  367 gene mutation  389 genes, nature of  367–368 genetic mutation  300–302 genetic theories  363‐372 genetic variation  363‐372 Mendelian genetic variation  366 Somagenetic variation  366 Epigenetic variation  366 Meiotic genetic variation  366 genome cycle  368–370 genome generation cycle  353–354 genome instability, environmentally induced  91–92, 97 stress, effects of  92 DNA repair  92–93 epigenetic changes  93–95 epigenetic changes, link with genetic changes 95–96 genetic changes  92 transgenerational inheritance  96 genome maintenance  201–205 genome rearrangements

transposable elements  182–185 genome restructuring in flax  79–80 genome variation  254, 263 chromosome number in prokaryote species 254 gene direction variation  263 genome size distribution in archaeans and bacteria  256–257 genome size variation in archaeans and bacteria 255–256 genomic GC content (genomic C+G content) 257–259 genomic GC content and size in asexually reproducing fungi  260–263 genomic GC content correlation with genome or chromosome size 259–260 genomic organization of polyploid cells 9–10 genotrophs 77–79 germline polyploidy  4 germline restricted chromosome (GRC)  48, 49 Giardia intestinalis 321 Ginkgo biloba 387 glycogen synthase (GYS)  292 glycogen synthase kinase  3 (GSK‐3) 292 gram‐negative bacteria  255 chromosome data  260 gram‐positive bacteria  255 chromosome data  260 guanine 137 guanine + cytosine content in genomes 257–259 correlation with genome or chromosome size 259–260 guanine nucleotide exchange factors (GEFs) 30

h Haber–Weis cycle  136 hagfish 46–47 content of eliminated DNA  47 deletion mechanisms and results  47 Haplochromis obliquidens 316 haploidization 315

409

410

Index

health implications of somatic genome variation (SGV) cellular‐level variation  380 differential treatments of beneficial or harmful SGVs  385 DNA cytosine deaminases  381–384 endoploidy variation  381 human immunodeficiency virus (HIV) 382 immunoglobulin diversity  381–384 mitochondrial genome sequence 384–385 nuclear or ooplasmic transfer‐based therapy 385 overgrowth 381‐383 ploidy and chromosome number variation 380–381 somatic mutation  381–384 tumors 381–384 viral mutation  382 hemihyperplasia multiple lipomatosis (HHML) 294 hepatectomy, partial  27 hepatocytes polyploidization liver regeneration  26–28 mechanisms 25–26 ploidy conveyor model  26 heterochromatin  9, 10, 47, 49 elimination 50 formation 179–182 locus‐specific 93 heteroplasmy  105–107, 112–113 histone acyltransferase (HAT)  19 histone deacetylases (HDACs)  19 histone modification  49, 95 histone octamer  310 histone tetramer  310 histones  123, 176, 179–181 Homo sapiens  6, 140, 202, 204, 207–216, 258, 270 motif sequences  273, 275, 277, 282, 283 homologous recombination (HR)  93, 141, 219–221 double‐strand break repair  227–228 Hordeum bulbosum 387 horizontal DNA transfer  390–391

human embryonic stem cells (hESCs)  56 PCSs and iPSCs  56–57 human immunodeficiency virus (HIV) 382 human polyploidization  6 hybrid varieties  369 hydrogen peroxide  92, 134, 135, 136 hydroxyl radicals  92, 136 hyperplasia 23

i immunoglobulin diversity  381–384 induced inheritable genome variation in flax  77–79, 87 adaptive genomic changes  83–84 environmental stress response  83 genome restructuring  79–80 similar responses in other plants 84–87 specific genomic changes  80–82 induced pluripotent stem cells (iPSCs) mouse ESCs  57 parthenogenetic ESCs  57–58 primate ESCs  56–57 Inner Centromere Protein (INCENP)  61 insulin receptor for substrates (IRS)  291 insulin receptor tyrosine kinase (INSR) 291 insulin signaling pathways  291–292, 295–296 diseases caused by somatic mutations of PI3K, PTEN and AKT  293–294 key protein ranking  293 plant growth regulation  295 plant insulin and medical use  295 intercompartmental interaction  391 internal eliminated sequences (IESs)  170, 176, 182–184 intra‐ and interchromosomal variation 387–388 inverted repeats (IRs)  121, 129

j Jack bean  295

k kinetochore microtubule attachment defects 60–61

Index

l Lactuca sativa 388 Leishmania donovani 368 Leishmania infantum 268 leucine‐rich repeats (LLRs)  112 Linum Insertion Sequence  1 (LIS‐1) 80, 82, 84 Linum usitatissimum 389 liver regeneration  26–28 long interspersed elements (LINEs)  381 loss of heterozygosity (LOH)  228 Lycopersicon esculentum 388

m Macronuclear Development Protein  1 (MDP1) 175 macronucleus (MAC)  170–171, 172, 176–177 Magnolia stellata 390 mammalian microhomology‐mediated end‐joining pathway (MMEJ)  142 mammalian target of rapamycin (mTOR) 292 Marchantia polymorpha 318 marconuclear‐destined sequences (MDSs)  170, 177 maternal transmission of organelles 358–359 MCAP syndrome  294 Medicago truncatula  258, 269, 274, 275, 277, 283 megakaryocytes (MKCs)  28 mechanisms 28–30 abortive division cycles  29 megalencephaly syndrome  294 Mendelian genetic variation  365–367 mesenchymal stem cells (MSCs)  56, 58–59 mesenchymal–epithelial transition factor (MET) 382 methylerythritol cyclodiphosphate (MEcPP) 135 microhomology‐mediated break‐induced replication (MMBIR)  144 microhomology‐mediated recombination (MHMR) 144 micronucleus (MIC)  170, 171 microRNA (miRNA)  276, 344, 345

microtubule–kinetochore coupling  60–61 miniature inverted‐repeat transposable elements (MITEs)  184 mismatch repair (MMR)  200 clean‐up after replications  218–219 mismatch‐repair protein (MUTS)  144 mitochondrial DNA (mtDNA)  104, 105–106, 107, 384 mitochondrial genome  103–105 health implications  384–385 recombination among repeated sequences 321–322 size 320–321 stoichiometric variation and effects 323 sublimon   111, 310 sugenomic molecules  310 mitochondrial genome sequence  390 mitochondrial sublimons and cytoplasmic male sterility (CMS)  109–111 mitogen‐activated protein kinase (MAPK) 293 mitosis  8–9, 30–31 acytokinetic mitosis  4 cardiomyocytes 23 polyploidization 24–25 response to tissue damage  25 upstream control of polyploidization 23–24 endocycle entry in trophoblast lineage 19–22 endocycle oscillations mechanisms in Drosophila 15–17 mechanisms in trophoblast giant cells (TGCs) 22–23 endomitosis 5 endoreplication 10–11 initiation in Drosophila 11–15 gene amplification follicle cells Drosophila 17–19 hepatocytes liver regeneration  26–28 mechanisms of polyploidization 25–26 ploidy conveyor model  26 megakaryocytes (MKCs)  28 mechanisms of polyploidy  28–30

411

412

Index

mitotic genome variations in yeast  199–200, 234 ploidy maintenance and chromosome integrity mechanisms  229–230 aneuploidy in yeasts  230–231 ploidy changes in yeasts  231–232 ploidy changes in yeasts, mechanisms 232–234 post‐replicative repair (PRR) and homologous recombination (HR) 219–221 alternative pathway  226 break‐induced replication (BIR) repair pathway 227–229 crosstalk between RFC complexes 226–227 polyubiquitinated PCNA  225–226 sumoylated PCNA  221–223 ubiquitinated PCNA  223–225 replication as source  200–201 DNA polymerases as guadians of genome maintenance  201–205 dNTP levels and pool bias in genome stability 205–218 fungal proteins involved in genome maintenance 207–217 genes encoding fungal DNA polymerase complexes  202–204 mismatch repair (MMR) and ribonucleotide excision repair (RER) 218–219 mitotic‐to‐endocyte switch  11–13 endocyte entry into other tissues  14–15 regulation signaling pathways  13–14 molecular heterosis  344 modifier of mitochondrial transcripts (mmt) 322 mouse embryonic stem cells (mESCs)  57 Muller’s ratchet  121 Mus musculatis polyploidization 6 MutS homolog  1 (MSH1) 144 myofibrils 23

n neural stem cells (NSCs)  56, 59 Nicotiana glutinosa 324

Nicotiana sylvestris  110, 301, 302, 304, 322, 323, 324, 353 genetic variation  357 mitochondrial genome  390 Nicotiana tabacum  94, 124, 143, 321, 324, 339, 385–386 non‐chromosomal stripe (NCS) maize mutants 106–107 noncoding RNA (ncRNA)  174, 187 genome rearrangement  176–177 long noncoding RNA templates as interface for short noncoding RNA 177–179 VDJ recombination  186–187 non‐homologous end‐joining (NHEJ)  93, 141, 142–143, 218 double‐strand break repair  227–228 non‐human primate embryonic stem cells (nhpESCs) 57 non‐meiotic genome variation  254 non‐Mendelian inheritance  171–173 Notch signalling network  12, 13–14, 19 Nothobranchius furzeri 352 nuclear B chromosomes  316–317 nuclear DNA replication (NDR)  130 nuclear fertility restoration  105, 322 nuclear gene diversity  111–112 nuclear transfer‐based therapy  385 nuclear‐encoded polymerase (NEP)  130, 131 nucleomorph 391 nucleotide excision repair (NER)  93 nucleotide mismatch repair (NMR)  93 nurse cells  10–11, 12, 17

o Oikopleura dioica endoreplication 10–11 polyploidization 6 Oncorhynchus mykiss 312 ooplasmic transfer‐based therapy  385 open reading frames (ORFs)  108, 110 organellar DNA replication (ODR)  130 organelle genomes  324 integrity under light  318–319 maternal transmission  358–359

Index

nuclear genome ploidy and effects 325–326 segregation 390–391 transmission 390–391 origin recognition complex (ORC)  9, 228 oxidative stress  133–138, 300 protective pigments  340–341 oxoguanine [8‐]  137 Oxytricha  169–171, 342 small RNA  175–176 Oxytricha trifallax  169, 181, 342

p p21/Cip1 8 p27/Kip1 8 p57/Kip2 8 Paralichthys olivaceus 313 Paramecium  171–173, 179–180 parthenogenetic embryonic stem cells (pESCs) 57–58 Pediculus humanus 322 Pennisetum glaucum 387 pentatricopeptide repeat (PPR)  111 peroxidase (POD)  301 somatic genome variation (SGV) 339–340 Petunia hybrida 390 Phalaenopsis aphrodite subsp. formosana (Orchidaceae) 314 phase variation  185 phosphatase and tensin homolog (PTEN) 293 diseases caused by somatic mutations 293–294 3′‐phosphoadenosine 5′‐phosphate (PAP) 135 phosphoinositide  3‐kinase (PI3K) 291 diseases caused by somatic mutations 293–294 phosphoinositide  3‐kinase alpha catalytic subunit (PI3KCA) 293, 294 photo‐adaptation 133–138 photocarcinogenesis 300 photorepair 139–140 photosynthetic electron transport (PET) 133–134 Phycisphaera mikurensis 259

Phytophthora infestans  269, 270, 272, 279 PI3K‐activated 3‐phosphoinositide‐ dependent protein kinase 1 (PDK1) 292 Piper magnificum 315 Pisum sativum  143, 388 plants aneuploid cells  315–316 base composition of  3′COR 277–278 base composition of  3′UTR 276 base composition of  3′UTR compared with whole genome 276–277 base composition of poly(A) site region 280 compared with animals  280–284 endopolyploidy 313–315 genomic GC content  257–259 plastid‐encoded polymerase (PEP) 130–131 plastid genome sequence or DNA amount variation 389 plastid genome stability and repair  120–121, 147 characteristics of plastid genome general composition  121–123 structure 123–124 DNA repair  133 double‐strand breaks (DSBs) 141–145 oxidative stress, photo‐adaptation and ROS detoxification  133–138 recombination 141–145 UV‐induced DNA damage  138–140 influence of replication and transcription in stability  131–133 outcomes of DNA rearrangements 145–147 replication of plastid DNA content during development 124–125 coordination during DNA replication 130 origins of replication  129–130 replication machinery  125–126 replication mechanisms  126–129 transcription in plastid  130–131

413

414

Index

plastid genome variation copy number variation  319–320 organelle genome integrity  318–319 plastid genome size  317–318 plastid types  317 recombination among repeated sequences 318 plastid transcriptionally active chromosome (pTAC)  123–124, 131 ploidy and chromosome number variation 386–387 ploidy maintenance  229–230 aneuploidy in yeasts  230–231 ploidy changes in yeasts  231–232 mechanisms 232–234 ploidy reversal  7–8 ploidy variation in somatic cells  310–311, 326 mitochondrial genome recombination among repeated sequences 321–322 size 320–321 stoichiometric variation and effects 323 nuclear genome in somatic cells aneuploid cells in cancerous masses 316 aneuploid cells in cells  315–316 bacteria 313 effects on growth  312–313 endopolyploidy 313–315 individuals or species  311–312 nuclear B chromosomes  316–317 somatic cell haploidization  315 organelle genomes  324 nuclear genome ploidy and effects 325–326 plastid genome variation copy number variation  319–320 organelle genome integrity  318–319 plastid genome size  317–318 plastid types  317 recombination among repeated sequences 318 pluripotent stem cells (PSCs) mouse ESCs  57 parthenogenetic ESCs  57–58 primate ESCs  56–57

poly(A) site (see “RNA polyadenylation site regions”) 269–271 base composition in animals  280 base composition in plants  280 base composition in protists  278–279 low conservation of overall nucleotide sequence 286 poly(A) tail attachment position  269 poly(A) tail starting position  269 RNA structure  286 Signal motifs, AAUAAA, UGUUUG, UGUAAC, UUACUU  275 stability and somatic genome variation  286–287, 358 top  20 hexamer motifs in the poly(A) site region in human 273 polymerase I‐like A (POLIA)  125–126 polymerase I‐like B (POLIB)  125–126, 132 polyploid cells  3 polyploidy  3–4, 30–31 cardiomyocytes 23 mechanisms 24–25 response to tissue damage  25 upstream control  23–24 core cell cycle machinery  8–9 endocycle entry in trophoblast lineage 19–22 endocycle oscillation mechanisms in trophoblast giant cells (TGCs) 22–23 endocycle oscillations mechanisms in Drosophila 15–17 endoreplication 10–11 initiation in Drosophila 11–15 gene amplification follicle cells Drosophila 17–19 genomic organization of cells  9–10 germline polyploidy  4 hepatocytes liver regeneration  26–28 mechanisms 25–26 ploidy conveyor model  26 mechanisms 4–8 acytokinetic mitosis  4 cell fusion  4 endomitosis 5

Index

endoreplication 5–7 gene amplification  7 ploidy reversal  7–8 megakaryocytes (MKCs)  28 mechanisms 28–30 somatic polyploidy  4 polytene cells  3, 354 Populus tremuloides 355 post‐replicative repair (PRR)  219–221 pre‐replication complexes  8–9 programmed genome engineering  185 programmed genome rearrangements (PGR) in vertebrates  45–46 emerging themes and directions biological function of PGR  49–50 deletion mechanisms  50–51 other vertebrates  51 hagfish 46–47 content of eliminated DNA  47 deletion mechanisms and results  47 sea lamprey  48 content of eliminated DNA  48 deletion mechanisms and results 48 zebra finch  48 content of eliminated DNA  49 deletion mechanisms  49 proliferating cell nuclear antigen (PCNA) 201 clean‐up after replications  218–219 DNA lesions  219–221 polyubiquitination 225–226 sumoylated PCNA  221–223 ubiquitination (Ubi‐PCNA)  220, 223–225 proplastids 120 protists base composition of poly(A) site region 278–279 genomic GC content  257–259 Prunus persica 389 PTEN 293–294 pure lines  368–370 purpurin 340 pyrimidine‐pyrimidone (6‐4) photoproducts 138–139 Pyrus communis 389

r radical‐scavenging enzymes  339–340 Rafflesia lagascae 317 Ranunculus bulbosus  316, 386 Rb  21, 22 Rcho‐1 cells   20 reactive oxygen species (ROS)  92, 120 hydrogen peroxide  92, 134, 135, 136 hydroxyl radicals  92, 136 scavenging ROS  124, 134, 135–137 singlet oxygen  134, 135 superoxide anion  92, 134, 135 redifferentiation‐induced variation  388 replication factor C complex (RFC)  205 crosstalk 226–227 replication fork stalling  132, 219 re‐replication 4 restorer gene evolution  111–112 Retinoblastoma‐like pocket proteins  22 retrograde signaling  131, 134, 135 ribonuclease reductase (RNR)  205–206 ribonucleotide excision repair (RER) 218–219 ribonucleotide monophosphate (rNMP) 218 ribonucleotide triphosophates (rNTPs) 219 ribosomal DNA (rDNA)  83, 84, 79, 80, 83 ribulose‐1,5‐bisphosphate carboxylase/ oxygenase (RuBisCo)  320 R‐loops 132 RNA‐interference (RNAi) pathway  175 RNA‐mediated somatic genome rearrangement  168, 187, 342 chromatin modifications  186–187 ciliates 168–170 long noncoding RNA templates 176–177 long noncoding RNA templates as interface for short noncoding RNA 177–179 non‐Mendelian inheritance in Paramecium 171–173 nuclear dimorphism in ciliates  170–171 phase variation  185 programmed genome engineering  185

415

416

Index

RNA‐mediated somatic genome rearrangement (contd.) scanRNA model in Tetrahymena 173–175 short RNA‐mediated heterochromatin formation 179–182 small RNA in Stylonychia and Oxytricha 175–176 transposable elements and genome rearrangments 182–185 transposases 186–187 transposons 185 VDJ recombination  186–187 RNA polyadenylation site regions  268–269, 287–288 alternative polyadenylation  275–276 base composition of  3′COR 277–278 base composition of  3′UTR 276 compared with whole genome 276–277 base composition of poly(A) site region in animals  280 in animals compared with plants 280–284 in plants  280 in protists  278–279 base selection as poly(A) tail starting position 269–271 difference between most frequent motifs 284–285 most frequent upstream motifs 271–272 poly(A) site RNA structure  286 stability and somatic genome variation 286–287 signal motif distribution  273–275 top  20 hexamer motifs in poly(A) site 273, 274 upstream A‐rich region and upstream U‐rich region  284 U–A–U–A–U base abundance pattern 284 whole genome motif frequencies  273 RNA‐templated DNA repair  341 ROS detoxification  133–138 Rubia tinctorium  301, 340

s S phase in mitosis  8–9 Saccharomyces cerevisiae 219–221 effect of translation synthesis  224 ploidy changes  231–232 Saccharomyces cerevisiae × S. bayanus 311 salivary glands  5, 9–10, 14, 15–16, 21 Salvador–Warts–Hippo (SWH) tumor‐ suppressor pathway  14 scanRNA model  173–175 Schizosaccharomyces pombe  200, 202–204, 206, 207–216, 221, 226, 272 motif frequencies  273 Sclerotinia trifoliorum 312 sea lamprey  48 content of eliminated DNA  48 deletion mechanisms and results  48 Separase 27 single nucleotide polymorphisms (SNPs)  303, 382 single‐strand annealing (SSA)  141 single‐strand breaks (SSBs) in DNA  93 single‐stranded DNA (ssDNA)  132 singlet oxygen  134, 135 small non‐coding RNA (sncRNA)  93, 97 small RNA  19, 86, 167–181, 342, 383 Solanum chacoense 324 Solanum demissum 311 Solanum tuberosum  258, 277, 324, 325, 386, 387, 391 Solanum tuberosum group phureja 353 Solanum verneï  324, 391 somatic cell haploidization  315 somatic crossover (mitotic crossover) 343 somatic gene conversion  300 somatic genome variation (SGV) impact on genetic theories and breeding concepts  364, 371–372 breeding criteria  368–370 epigenetic variation  365–367 evolution 370–371 genes, nature of  367–368 genome cycle  368–370 Mendelian genetic variation  365–367 pure lines  368–370

Index

somatic genome  365 somatogenetic variation  365–367 variety stability  368–370 Weismann barrier hypothesis revision 370 somatic genome variation (SGV) implications for agriculture and health agricultural impacts  385 bud mutation  389 cellular‐level variation  385–386 dedifferentiation‐ and redifferentiation‐induced variation 388 DNA damage  389 DNA exchange  391 DNA transfer  390–391 endoploidy variation  387 epigenetics 389 gene mutation  389 intercompartmental interaction 391 intra‐ and interchromosomal variation 387–388 mitochondrial genome sequence or DNA amount variation  390 organelle genome segregation 390–391 organelle transmission  390–391 plastid genome sequence or DNA amount variation  389 ploidy and chromosome number variation 386–387 differential treatments of beneficial or harmful SGVs  385 DNA cytosine deaminases  381–384 somagenetic variation  365–367 somatic genome variation (SGV) implications for agriculture and health 378 endoploidy variation  381 human and animal health  380 cellular‐level variation  380 immunoglobulin diversity  381–384 mitochondrial genome sequence 384–385 natural attributes of SGV  378–379

nuclear or ooplasmic transfer‐based therapy 385 ploidy and chromosome number variation 380–381 somatic mutation  381–384 tumors 381–384 somatic genome variation (SGV) interpretation hypotheses  352, 360 cell‐specific accumulation of variation  352 costs and benefits of SGV  354–357 developmental age  353 genome generation cycle  353–354 genome stability through structural similarity and sequence dissimilarity 358 heterosis created by SVG  357–358 human ability to deal with SGV and diseases 359–360 in vitro culture propagation  357 maternal transmission of organelles 358–359 tissue‐specific requirements  354 somatic genome variation (SGV) molecular mechanisms  338, 344–345 DNA cytosine deaminases  340 DNA damage  338–339 DNA repair errors  341–342 DNA transportation  343 endoplasmic reticulum stress  344 extracellular DNA  343 gene mutation  338 molecular heterosis  344 protective pigments  340–341 radical‐scavenging enzymes  339–340 repetitive DNA instability  342 RNA‐mediated somatic genome rearrangement 342 RNA‐templated DNA repair  341 somatic crossover and gene conversion 343 teleomere degeneration  344 somatic genetic changes  111–112 somatic genome  365 somatic genome rearrangement in ciliates  168, 187 background 168–170 chromatin modifications  186–187

417

418

Index

somatic genome rearrangement in ciliates (contd.) long noncoding RNA templates as interface for short noncoding RNA 177–179 long noncoding RNA templates in genome rearrangement  176–177 non‐Mendelian inheritance in Paramecium 171–173 nuclear dimorphism  170–171 phase variation  185 programmed genome engineering  185 scanRNA model in Tetrahymena 173–175 short RNA‐mediated heterochromatin formation 179–182 small RNA in Stylonychia and Oxytricha 175–176 transposases 186–187 transposable elements and genome rearrangments 182–185 transposons 185 VDJ recombination  186–187 somatic mutation  381–384 somatic polyploidy  4 induction mechanisms  4–8 acytokinetic mitosis  4 cell fusion  4 endomitosis 5 endoreplication 5–7 gene amplification  7 ploidy reversal  7–8 somatic stem cells (SSCs)  55–56 mesenchymal stem cells (MSCs)  58–59 neural stem cells (NSCs)  59 spindle assembly checkpoint (SAC) dysfunction 60 Squalius alburnoides 311 stem cells, (SCs), chromosome instability (CIN) 55–56 mechanisms of CIN  59–60 kinetochore microtubule attachment defects 60–61 sister chromatids cohesion  62 spindle assembly checkpoint (SAC) dysfunction 60 supernumerary centrosomes  61–62

mechanisms of CIN in SCs  63 pluripotent stem cells (PSCs) mouse ESCs  57 parthenogenetic ESCs  57–58 primate ESCs  56–57 somatic stem cells (SSCs) mesenchymal stem cells (MSCs) 58–59 neural stem cells (NSCs)  59 stoichiometric shifting  107 Streptomyces albus 257 Stylonychia 170–171 small RNA  175–176 sublimons 107 mitochondrial sublimons and cytoplasmic male sterility (CMS) 109–111 subperineural glia (SPG)  11 substoichiometric shifting (SSS)  107, 109, 110, 111 superoxide anion  92, 134, 135 superoxide dismutases (SODs)  124, 136, 301 somatic genome variation (SGV) 339–340 suppressor of under‐replication (Su(UR)) 9–10 symbiogenesis 104 synthesis‐dependent strand annealing (SDSA) 141

t TA dinucleotide  182–184 teleomere degeneration  344 telomere‐bearing elements (TBEs)  179, 181–182 Tetrahymena  173–175, 179–180 tetraploidy 25 three‐prime cleaved‐off region (3′COR) 268–269 base composition  277–278 three‐prime untranslated region (3′UTR)  268–269, 358 base composition  276 compared with whole genome 276–277 thymine 137 thymine glycol  137

Index

tobacco mosaic virus (TMV)  94 Trachystoma ballii 324 transgenerational inheritance  96 transposable elements  181–182 genome rearrangements  182–185 transposases 186–187 transposons  185, 187 Trithuria inconspicua 318 Triticum aestivum  122, 124, 388 trophoblast giant cells (TGCs)  7, 10 endocycle entry  19–22 endocycle oscillation mechanisms 22–23 trophoblast stem cells (TSCs)  20 Trypanosoma cruzi  269, 272, 274, 275, 275, 278–279 TTAA repeats  183 tumors 381–384

u ubiquitous genome variation  187 untranslated region (UTR)  358 Uroleptus 170 U‐turn‐like inversion  145

v variegation  144, 145–146 variety stability  368–370 VDJ recombination  186–187 vertebrates programmed genome rearrangements (PGR)  45–46 emerging themes and directions biological function of PGR  49–50 deletion mechanisms  50–51 other vertebrates  51 hagfish 46–47 content of eliminated DNA  47 deletion mechanisms and results  47 sea lamprey  48 content of eliminated DNA  48 deletion mechanisms and results  48 zebra finch  48 content of eliminated DNA  49 deletion mechanisms  49 Vigna unguiculata 387

w Watson–Crick basepairing  201

Weismann barrier theory  364 revision 370 Whirly proteins  144–145

y yeast, mitotic genome variations  199–200, 234 ploidy maintenance and chromosome integrity mechanisms  229–230 aneuploidy 230–231 ploidy changes  231–232 ploidy changes, mechanisms 232–234 post‐replicative repair (PRR) and homologous recombination (HR) 219–221 alternative pathway  226 break‐induced replication (BIR) repair pathway 227–229 crosstalk between RFC complexes 226–227 polyubiquitinated PCNA  225–226 sumoylated PCNA  221–223 ubiquitinated PCNA  223–225 replication as source  200–201 DNA polymerases as guardians of genome maintenance  201–205 dNTP levels and pool bias in genome stability 205–218 fungal proteins involved in genome maintenance 207–217 genes encoding fungal DNA polymerase complexes  202–204 mismatch repair (MMR) and ribonucleotide excision repair (RER) 218–219

z Zea mays  130, 258, 270, 275, 277, 281, 283 DNA transposition  343 mitochondrial genome  320 ploidy 386 somatic gene conversion  388 zebra finch  48 content of eliminated DNA  49 deletion mechanisms  49

419