Root Development (Annual Plant Reviews, Volume 37)

ANNUAL PLANT REVIEWS VOLUME 37

ANNUAL PLANT REVIEWS VOLUME 37 Root Development

Edited by

Tom Beeckman VIB Department of Plant Systems Biology Ghent University, Ghent, Belgium

A John Wiley & Sons, Ltd., Publication

This edition first published 2010
C 2010 Blackwell Publishing Ltd Blackwell Publishing was acquired by John Wiley & Sons in February 2007. Blackwell’s publishing programme has been merged with Wiley’s global Scientific, Technical, and Medical business to form Wiley-Blackwell. Registered office John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, United Kingdom Editorial office 9600 Garsington Road, Oxford, OX4 2DQ, United Kingdom 2121 State Avenue, Ames, Iowa 50014-8300, USA For details of our global editorial offices, for customer services and for information about how to apply for permission to reuse the copyright material in this book please see our website at www.wiley.com/wiley-blackwell. The right of the author to be identified as the author of this work has been asserted in accordance with the UK Copyright, Designs and Patents Act 1988. 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 the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic books. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. Library of Congress Cataloging-in-Publication Data Root development / edited by Tom Beeckman. – 1st ed. p. cm. – (Annual plant reviews ; v. 37) Includes bibliographical references and index. ISBN 978-1-4051-6150-3 (hardback : alk. paper) 1. Roots (Botany)–Development. I. Beeckman, Tom. II. Series: Annual plant reviews ; v. 37. QK644.R64 2009 575.5 438–dc22 2009020434 A catalogue record for this book is available from the British Library. Annual plant reviews (Print) ISSN 1460-1494 Annual plant reviews (Online) ISSN 1756-9710 R Inc., New Delhi, India Set in 10/12 pt Palatino by Aptara
Printed in Singapore 1 2010

Annual Plant Reviews A series for researchers and postgraduates in the plant sciences. Each volume in this series focuses on a theme of topical importance and emphasis is placed on rapid publication. Editorial Board: Prof. Jeremy A. Roberts (Editor-in-Chief), Plant Science Division, School of Biosciences, University of Nottingham, Sutton Bonington Campus, Loughborough, Leicestershire, LE12 5RD, UK; Dr David Evans, School of Biological and Molecular Sciences, Oxford Brookes University, Headington, Oxford, OX3 0BP; Prof. Hidemasa Imaseki, Obata-Minami 2419, Moriyama-ku, Nagoya 463, Japan; Dr Michael T. McManus, Institute of Molecular BioSciences, Massey University, Palmerston North, New Zealand; Dr Jocelyn K.C. Rose, Department of Plant Biology, Cornell University, Ithaca, NY 14853, USA. Titles in the series: 1. Arabidopsis Edited by M. Anderson and J.A. Roberts 2. Biochemistry of Plant Secondary Metabolism Edited by M. Wink 3. Functions of Plant Secondary Metabolites and their Exploitation in Biotechnology Edited by M. Wink 4. Molecular Plant Pathology Edited by M. Dickinson and J. Beynon 5. Vacuolar Compartments Edited by D.G. Robinson and J.C. Rogers 6. Plant Reproduction Edited by S.D. O’Neill and J.A. Roberts 7. Protein–Protein Interactions in Plant Biology Edited by M.T. McManus, W.A. Laing, and A.C. Allan 8. The Plant Cell Wall Edited by J.K.C. Rose 9. The Golgi Apparatus and the Plant Secretory Pathway Edited by D.G. Robinson 10. The Plant Cytoskeleton in Cell Differentiation and Development Edited by P.J. Hussey 11. Plant–Pathogen Interactions Edited by N.J. Talbot 12. Polarity in Plants Edited by K. Lindsey 13. Plastids Edited by S.G. Moller 14. Plant Pigments and Their Manipulation Edited by K.M. Davies

15. Membrane Transport in Plants Edited by M.R. Blatt 16. Intercellular Communication in Plants Edited by A.J. Fleming 17. Plant Architecture and Its Manipulation Edited by CGN Turnbull 18. Plasmodesmata Edited by K.J. Oparka 19. Plant Epigenetics Edited by P. Meyer 20. Flowering and Its Manipulation Edited by C. Ainsworth 21. Endogenous Plant Rhythms Edited by A. Hall and H. McWatters 22. Control of Primary Metabolism in Plants Edited by W.C. Plaxton and M.T. McManus 23. Biology of the Plant Cuticle Edited by M. Riederer 24. Plant Hormone Signaling Edited by P. Hadden and S.G. Thomas 25. Plant Cell Separation and Adhesion Edited by J.R. Roberts and Z. Gonzalez-Carranza 26. Senescence Processes in Plants Edited by S. Gan 27. Seed Development, Dormancy and Germination Edited by K.J. Bradford and H. Nonogaki 28. Plant Proteomics Edited by C. Finnie 29. Regulation of Transcription in Plants Edited by K. Grasser 30. Light and Plant Development Edited by G. Whitelam 31. Plant Mitochondria Edited by David C. Logan 32. Cell Cycle Control and Plant Development Edited by D. Inz´e 33. Intracellular Signaling in Plants Edited by Z. Yang 34. Molecular Aspects of Plant Disease Resistance Edited by Jane Parker 35. Plant Systems Biology Edited by Gloria M. Coruzzi and Rodrigo A. Guti´errez 36. The Moss Physcomitrella Patens Edited by Celia Knight 37. Root Development Edited by Tom Beeckman 38. Fruit Development and Seed Dispersal Edited by Lars Østergaard

CONTENTS

Contributors Preface 1

Arabidopsis Root Development Marijn Luijten and Renze Heidstra 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8

2

3

Introduction Specification of the apical and basal cell lineage Root stem cell niche specification Radial patterning Stem cell maintenance Meristem maintenance and root zonation Meristem activation, root growth and cell division Concluding remarks

xi xv 1 1 3 10 14 18 22 25 28

Vascular Morphogenesis During Root Development Ana Campilho, Ove Lindgren and Yk¨a Helariutta 2.1 Introduction 2.2 The Arabidopsis root vascular system 2.3 Molecular genetics of the stele: a rapidly developing field 2.4 Vascular genomics – getting the big picture

39

Root Epidermal Development in Arabidopsis Rebecca Horn, Keke Yi, Benoˆıt Menand, Monica Pernas-Ochoa, Seiji Takeda, Tom Walker and Liam Dolan 3.1 Introduction 3.2 Establishment of the epidermis in Arabidopsis 3.3 Establishment of distinct cell fates in the root epidermis 3.4 Root hair initiation and tip growth 3.5 Effects of nutrients on root hair cell differentiation and morphogenesis 3.6 Root hairs and nutrient uptake 3.7 Perspectives

64

39 40 44 57

65 65 68 70 75 76 77

vii

viii
Contents 4 Lateral Root Formation Jocelyn E. Malamy 4.1 Introduction 4.2 How does a single lateral root form? 4.3 How are the number and placement of lateral roots determined? 4.4 Agricultural importance of lateral root formation 5 Adventitious Root Formation: New Insights and Perspectives Gaia Geiss, Laurent Gutierrez and Catherine Bellini 5.1 Introduction 5.2 Role and origin of ARs 5.3 Factors influencing adventitious rooting 5.4 New insights into genetics and molecular mechanisms involved in adventitious rooting 5.5 Conclusion and perspectives 6 Root Gravitropism Ranjan Swarup and Malcolm J. Bennett 6.1 6.2 6.3 6.4 6.5 6.6

Introduction Gravity perception Root gravitropic signal transmission The root gravitropic response Attenuating the root gravitropic response Future directions

7 Molecular and Genetic Dissection of Cereal Root System Development Frank Hochholdinger and Roman Zimmermann 7.1 Introduction 7.2 Morphology of cereal root systems 7.3 Morphological and anatomical comparison of Arabidopsis and cereal root systems 7.4 Molecular and genetic analysis of cereal root formation 7.5 Prospects 8 Fern Root Development Guichuan Hou and Elison B. Blancaflor 8.1 Introduction 8.2 Overview of the fern root system – shoot-borne roots 8.3 Anatomical and structural aspects of fern root development

83 83 85 107 112 127 128 128 131 142 147 157 157 158 160 164 166 167

175 175 176 177 178 187 192 192 195 197

Contents
ix

8.4 8.5 9

10

When Plants Socialize: Symbioses and Root Development Benjamin P´eret, Sergio Svistoonoff and Laurent Laplaze 9.1 Introduction 9.2 Arbuscular mycorrhizae 9.3 Ectomycorrhizae 9.4 Actinorhizal symbioses 9.5 Concluding remarks Legume Root Architecture: A Peculiar Root System Silvina Gonzalez-Rizzo, Philippe Laporte, Martin Crespi and Florian Frugier 10.1 10.2 10.3 10.4

11

Comparison of legume lateral roots and nitrogen-fixing nodules Recent advances in genetics and genomics of nitrogen-fixing nodule development in legumes Evidences for a crosstalk between symbiotic nodule and LR developmental pathways Common signals in nodulation and LR development

203 205 209 209 210 218 223 228 239

240 245 257 263

Effect of Nutrient Availability on Root System Development Alfredo Cruz-Ram´ırez, Carlos Calder´on-V´azquez and Luis Herrera-Estrella

288

11.1 11.2

288

11.3 12

LR formation in ferns Concluding remarks and future prospects

Introduction Regulation of the root system architecture by nutrient availability Conclusions

Studying Root Development Using a Genomic Approach Jose R. Dinneny and Philip N. Benfey 12.1 Introduction: how root development enables a genomic approach 12.2 Genome-scale technologies for understanding gene function 12.3 Building transcriptional networks: an introduction 12.4 Building transcriptional networks I: creating high-resolution spatial maps of gene expression 12.5 Building transcriptional networks II: exploring the role of transcriptional and posttranscriptional mechanisms in regulating TF activity 12.6 Building transcriptional networks III: identifying direct targets of TFS – the SHORTROOT pathway

292 315 325

326 327 330 331

339 341

x
Contents 12.7 12.8

Exploring gene function using genomic variation: quantitative trait locus analysis of root growth Future directions: from single gene biology to systems biology

Index Color plate (between pages 174 and 175)

343 345 352

CONTRIBUTORS

Catherine Bellini Department of Forest Genetics and Plant Physiology, Ume˚a Plant Science Center, Swedish University of Agricultural Sciences, Ume˚a, Sweden and Laboratoire de Biologie Cellulaire, Institut National de la Recherche Scientifique, Versailles, France Philip N. Benfey Department of Biology and Institute for Genome Sciences and Policy Center for Systems Biology, Duke University, Durham, NC, USA Malcolm J. Bennett Centre for Plant Integrative Biology and Plant Sciences Division, School of Biosciences, University of Nottingham, Sutton Bonington Campus, Loughborough, UK Elison B. Blancaflor Plant Biology Division, The Samuel Roberts Noble Foundation, Sam Noble Parkway, Ardmore, OK, USA ´ Carlos Calderon-V´ azquez Nacional Laboratory of Genomics for Biodiversity, CINVESTAV Campus Guanajuato, Irapuato, Guanajuato, M´exico Ana Campilho Institute of Biotechnology, University of Helsinki, Helsinki, Finland Martin Crespi Institut des Sciences du V´eg´etal (ISV), Centre National de la Recherche Scientifique, Gif-sur-Yvette, France Alfredo Cruz-Ram´ırez ´ Nacional Laboratory of Genomics for Biodiversity, Centro de Investigacion y de Estudios Avanzados del Instituto Politecnico Nacional, Irapuato, Guanajuato, Mexico

xi

xii
Contributors Jose R. Dinneny Department of Biology, Institute for Genome Sciences and Policy Center for Systems Biology, Duke University, Durham, NC, USA and Temasek Lifesciences Laboratory, National University of Singapore, Singapore Liam Dolan Department of Cell and Developmental Biology, John Innes Centre, Norwich, UK Florian Frugier Institut des Sciences du V´eg´etal (ISV), Centre National de la Recherche Scientifique, Gif-sur-Yvette, France Gaia Geiss Department of Forest Genetics and Plant Physiology, Ume˚a Plant Science Center, Swedish University of Agricultural Sciences, Ume˚a, Sweden and Laboratoire de Biologie Cellulaire, Institut National de la Recherche Scientifique, Versailles, France Silvina Gonzalez-Rizzo Institut des Sciences du V´eg´etal (ISV), Centre National de la Recherche Scientifique, Gif-sur-Yvette, France Laurent Gutierrez Department of Forest Genetics and Plant Physiology, Ume˚a Plant Science Center, Swedish University of Agricultural Sciences, Ume˚a, Sweden Renze Heidstra Department of Biology, Section Molecular Genetics, Utrecht University, Utrecht, The Netherlands Yk¨a Helariutta Institute of Biotechnology, University of Helsinki, Helsinki, Finland Luis Herrera-Estrella ´ Nacional Laboratory of Genomics for Biodiversity, Centro de Investigacion y de Estudios Avanzados del Instituto Politecnico Nacional, Irapuato, Guanajuato, Mexico Frank Hochholdinger Center for Plant Molecular Biology (ZMBP), Department of General ¨ ¨ Genetics, University of Tubingen, Tubingen, Germany

Contributors
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Rebecca Horn Department of Cell and Developmental Biology, John Innes Centre, Norwich, UK Guichuan Hou College of Arts and Sciences, Microscopy Facility, Appalachian State University, Boone, NC, USA Laurent Laplaze Institut de Recherche pour le D´eveloppement (IRD), UMR Diversit´e et Adaptation des Plantes Cultiv´ees, Equipe rhizogen`ese, Montpellier Cedex, France Philippe Laporte Institut des Sciences du V´eg´etal (ISV), Centre National de la Recherche Scientifique, Gif-sur-Yvette, France Ove Lindgren Institute of Biotechnology, University of Helsinki, Helsinki, Finland Marijn Luijten Department of Biology, Section Molecular Genetics, Utrecht University, Utrecht, The Netherlands Jocelyn E. Malamy Department of Molecular Genetics and Cell Biology, The University of Chicago, Chicago, IL, USA Benoˆıt Menand Department of Cell and Developmental Biology, John Innes Centre, Norwich, UK Benjamin P´eret Institut de Recherche pour le D´eveloppement (IRD), Un´et´e Mixte de Recherche Diversit´e et Adaptation des Plantes Cultiv´ees, Equipe rhizogen`ese, Montpellier Cedex, France Monica Pernas-Ochoa Department of Cell and Developmental Biology, John Innes Centre, Norwich, UK Sergio Svistoonoff Institut de Recherche pour le D´eveloppement (IRD), Un´et´e Mixte de Recherche Diversit´e et Adaptation des Plantes Cultiv´ees, Equipe rhizogen`ese, Montpellier Cedex, France

xiv
Contributors Ranjan Swarup Centre for Plant Integrative Biology and Plant Sciences Division, School of Biosciences, University of Nottingham, Sutton Bonington Campus, Loughborough, UK Seiji Takeda Department of Cell and Developmental Biology, John Innes Centre, Norwich, UK Tom Walker Department of Cell and Developmental Biology, John Innes Centre, Norwich, UK Keke Yi Department of Cell and Developmental Biology, John Innes Centre, Norwich, UK Roman Zimmermann Center for Plant Molecular Biology (ZMBP), Department of General ¨ ¨ Genetics, University of Tubingen, Tubingen, Germany

PREFACE

During the last decade, a remarkable proliferation of interest in root development has taken place. The reason for plant roots to be and remain an exciting and intriguing field of science is not explicitly clear. When compared to flowers, roots distinctly lack the ornamental attractiveness of flowering structures. Pondering over this seeming contradiction, I think the answer might be found in the secret and hidden nature of roots. Scientists are, by default, attracted by mysteries: the bigger the puzzle, the larger the provocation to dig into the enigma. Among all growth processes occurring in plants, the development of roots is without discussion, at the summit of secrecy. Roots reside their entire life in the soil sheltered from daylight, proliferate and colonize the rhizosphere through the continuous development of new meristems that arise deep inside tissues invisible from the surface, interact sneakily with various microorganisms that provoke dramatic growth changes and are capable of reacting upon the ever-changing soil conditions in an unfathomable way. Indeed, plant roots grow and develop while continuously monitoring information from the environment. They are often subjected to periods of soil water deficit and, unfortunately, the frequency of such phenomena is likely to increase in the future even outside today’s arid and semiarid regions. Therefore, it will become essential to understand root development in detail in the next decades, to learn how plants cope with a changing environment, and to translate the fundamental knowledge into applications. No doubt, there is still a long way to go and the question is whether a deeper understanding of fundamental root biology could be implemented on time into applications to ease the essential needs of the future world population. Nevertheless, it is at least hopeful to notice that insight into the subterranean part of plants has seriously made progress in the recent years, especially at the level of its developmental biology. Data have been accumulating exponentially and the entire root development area cannot easily be summarized in the format of one single review paper. Hence, I am glad and proud that representatives of all top laboratories on root development have accepted to contribute to the timely publication of this book and have helped me with this challenge. As, you will see, they all did a great job. In the first four basic chapters, the main root developmental processes are covered. In contrast to previous root development reviews, these chapters are characterized by the enumeration of an increasing number of molecular components that are essential for root development and have been identified recently. This tendency reflects that our discipline has come to maturity. A major player in this evolution has certainly been the selection of and the xv

xvi
Preface restriction to one single model system, namely Arabidopsis thaliana. The root of this species has all it needs to guarantee easy laboratory handling and experimentation. On top of this, its genome is sequenced and a myriad of molecular tools have become available. In the first chapter, the origin of the primary root during embryogenesis and the establishment of a typical cellular pattern are discussed as well as the preservation of this pattern after germination, implying the introduction of the concept of a stem cell niche in the root tip. The second chapter deals with the development of the vascular tissue with references to the involvement of plant hormones and summarizes some large-scale approaches, which collectively have been designated as ’vascular genomics’, a new discipline in root developmental biology. In the third chapter, the focus lies on particular tissue layer of the root, namely the epidermis. It is a border tissue through which roots make contact with the soil. Following a discussion on its development and the specification of the different cell types, an outlook on its nutrient uptake function is provided. Chapter 4 discusses the branching of roots, a process that contributes considerably to their plasticity toward changes in the rhizosphere. It is therefore no surprise that this chapter concludes with a section on the agricultural importance of lateral root formation. In contrast to lateral roots, adventitious roots are part of the root system that are not derived from the primary root; their formation is discussed in Chapter 5. This aspect of root development also incorporates a high economic potential because it represents the essential step in the vegetative propagation of plants. The lion’s share of this chapter deals with the many environmental and endogenous factors that influence adventitious root formation, illustrating the extreme complexity of this process. An intensively studied developmental process occurring in roots is root gravitropism that is by far the most obvious adaptive trait of roots. Several recent breakthroughs of gravitropism are summarized in Chapter 6. In the following four chapters, we step out of the Arabidopsis-dominated world and learn more about root development of cereals and ferns and changes in root architecture induced by symbiotic interactions with soil microorganisms. In these chapters, our view on root development is largely expanded by the treatment of issues that are completely lacking in the Arabidopsis model. Nevertheless, on many occasions, the authors can make interesting comparisons with some fundamental insights provided by the Arabidopsis root community, highlighting the value of a multidisciplinary thinking while studying roots. I hope that this book will contribute to bring the different root scientists together and will help researchers and students to compare their findings with observations made in other plants. Next, the importance of nutrient availability on root development is treated in Chapter 11 from an evolutionary perspective: the availability of mineral nutrients has played a major role in evolution and development of the diverse root system architectures. Finally, in Chapter 12, we return to the Arabidopsis model, but this time to discuss novel molecular tools that have led to the creation of expression maps of over 22 000 genes at a cellular resolution. The importance of such genome-scale

Preface
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data sets is commented and should convince any reader that systems biology approaches can be implemented in and are beneficial for root developmental research. In the coming years, this approach will be extremely valuable to decipher this complex biological process. Before allowing you to descend to the below-ground level and to enter the fascinating world of roots, I want to thank all contributors for their impeccable texts and for their endless patience with the ’not that speedy’ editing process for which I am personally the one to blame. Furthermore, I am indebted for the daily support and motivation of my scientific work to all present and previous members of the Root Development group at the Department of Plant Systems Biology in Gent and, last but not least, to Martine De Cock, the secret weapon of our department, who has been editing uncountable manuscripts over the years and who, this time again, helped me during the preparation of this book. Tom Beeckman

Annual Plant Reviews (2009) 37, 1–38 doi: 10.1002/9781444310023.ch1

www.interscience.wiley.com

Chapter 1

ARABIDOPSIS ROOT DEVELOPMENT Marijn Luijten and Renze Heidstra Department of Biology, Section Molecular Genetics, Utrecht University, Padualaan 8, 3584 CH, Utrecht, The Netherlands

Abstract: In plant development, the basic body plan is laid down during embryogenesis. Development carries on postembryogenically above and below ground with the continuous formation and outgrowth of lateral organs shaping the adult plant. In the past two decades, molecular genetics has been the preferred approach to study Arabidopsis thaliana root development. These efforts have resulted in the identification of numerous genes, involved in as many regulatory processes of root growth and development. Incidentally, conserved mechanisms and genetic factors that act in root and shoot growth have been uncovered, revealing general principles of plant development. Transport-mediated graded distribution of the phytohormone auxin, for example, acts as a global organizer that is locally translated into distinct cellular responses by specific auxin/indole-3-acetic acid–AUXIN RESPONSE FACTOR pairs. In the root, these responses promote expression of the PLETHORA regulators that act dose-dependently in controlling root morphology. The stem cell niche is uniquely defined by the combinatorial activity of the PLETHORA and SHORT-ROOT/SCARECROW transcription factors yet deploy signaling mechanisms that are conserved in root and shoot stem cell maintenance. Perpetual divisions of the stem cells are tightly regulated, interconnecting epigenetic factors, hormonal control and core cell cycle components. In this chapter, we will focus on recent advances in our understanding of Arabidopsis root development. Taking embryogenesis as a starting point, we will describe the genes and mechanisms involved in root meristem and stem cell patterning and maintenance. Keywords: root stem cell niche; PLETHORA genes; embryogenesis; cell patterning; root meristem maintenance

1.1 Introduction A unique property of plants is their lifelong ability to grow and to continuously develop, elaborating on the basic body plan laid down in the embryo. Therefore, plants depend on the incessant activity of confined populations of 1

2
Root Development stem cells located at opposite ends of the apical–basal body axis. With each asymmetric stem cell division, one daughter cell is maintained as a stem cell, while the other will differentiate to form specific tissues. In this way, a limited number of stem cells can generate organs of the size of trees. 1.1.1

Arabidopsis root morphology

In the root of the dicotyledonous model plant Arabidopsis thaliana, a small amount of stem cells generate all the different tissues that can be distinguished along the symmetrical radial axis (Fig. 1.1a). Because their rigid cell walls make it impossible for plant cells to move, the stereotyped division pattern of the root stem cells organizes the separate tissues in concentric columns or cell files. From outside to inside, these layers are designated as lateral root cap, epidermis, cortex, endodermis and pericycle as the cell files that surround the central vascular tissue. Clonal analysis and ablation studies indicate that cell lineage does not necessarily determine cell fate and pattern formation, but that plant cells are flexible and rather rely on positional information for adapting their final fate (Dolan et al., 1993; Scheres et al., 1994; van den Berg et al., 1995). At the basal end, a set of stem cells gives rise to the central portion of the root cap, known as the columella. Internal to and contacting all the stem cells is a small number of mitotically less active cells, the quiescent center (QC). Along the apical–basal axis of the root, stem cell daughters continuously travel through time, crossing the zone of cell division (meristematic zone), the zone of cell expansion and elongation (elongation zone) and, ultimately, meet their destiny in the differentiation zone. In other words, at any time, all developmental stages are present within the root tip. Together with the predictable fate of all individual cells that compose the root, these features make the Arabidopsis root an excellent system to study the genetic control of organ development. First, we will briefly discuss the fundamental principles of auxin transport and action because these processes have become particularly important for our understanding of Arabidopsis root development. 1.1.2

Principles of auxin transport and signaling

During the past century, the phytohormone auxin has been identified and extensively studied for its effects on development in many plant species (reviewed in Srivastava, 2002). Auxin is transported through the plant in a directional cell-to-cell fashion, called polar auxin transport that is mediated by the auxin influx carrier AUX1 (Bennett et al., 1996), the PINFORMED (PIN) family of auxin efflux facilitators (Paponov et al., 2005) and members of the P-glycoprotein subfamily of ATP-binding cassette transporters (Geisler and Murphy, 2006). PIN membrane proteins are frequently localized polarly to the cell membrane and have been shown to act as the main direction-determining factor of polar auxin transport (Wi´sniewska et al., 2006).

Arabidopsis Root Development
3

Blocking the auxin efflux machinery with 1-N-naphtylphtalamic acid (NPA) or application of the auxin analog 2,4-dichlorophenoxyacetic acid (2,4D), disrupted auxin transport and distribution in in vitro-cultured embryos and resulted in embryo defects, such as cup-shaped embryos with no functional root pole or ball-shaped embryos lacking any distinct apical–basal axis (Liu et al., 1993; Hadfi et al., 1998; Friml et al., 2003). These studies suggest an important link between auxin distribution and early embryo development. We will discuss further evidence in support of this hypothesis. The specificity of the response to auxin is thought to be generated by optimized pairs of interacting transcriptional regulators, designated auxin/indole-3-acetic acid (Aux/IAA) proteins and auxin response factor (ARF) transcription factors present in the auxin-responsive cell (Hamann et al., 2002; Tiwari et al., 2004; Weijers et al., 2005). Binding of auxin to the transport inhibitor response 1 (TIR1), an auxin receptor and F-box subunit of the SCFTIR1 E3 ubiquitin ligase (Dharmasiri et al., 2005; Kepinski and Leyser, 2005), promotes degradation of the Aux/IAA proteins via the ubiquitin proteasome pathway and releases the repressive hold on the interacting ARF transcription factor proteins (reviewed in Quint and Gray, 2006). Activity of the synthetic auxin-responsive promoter DIRECT REPEAT5 (DR5) is often used as an indirect marker to visualize auxin distribution (Sabatini et al., 1999; Friml et al., 2003). Although the DR5 reporter activity is merely an output of the cellular auxin response, its activity has been shown to correspond well with auxin levels by using immunolocalization with an anti-IAA antibody (Friml et al., 2003). 1.1.3

Chapter outline

In this chapter, we will discuss the critical stages particularly relevant for setting up the root pattern during the Arabidopsis embryogenesis, followed by the developmental processes important in postembryonic root growth. The main topics discussed are (1) the general importance of auxin transport and signaling in all of the illustrated developmental processes, (2) the combinatorial code utilizing PLETHORA and SHORT-ROOT/SCARECROW genes that positions and specifies the stem cell niche to the auxin machinery with feedback loops, (3) the role of WUSCHEL-RELATED HOMEODOMAIN (WOX) genes extending toward elucidating common factors in stem cell maintenance from root and shoot organizers, (4) radial patterning, (5) regulation of root zonation and (6) briefly, the integration and crosstalk of hormones regulating root growth and development. For reviews on root development of monocots and ferns, see Chapters 7 and 8.

1.2 Specification of the apical and basal cell lineage Embryogenesis starts with the elongation of the zygote followed by an asymmetric division producing two daughter cells with different fate and

4
Root Development characteristics. In Arabidopsis, the small apical cell develops into an eightcell embryo proper after three rounds of stereotypic divisions forming two ¨ layers with four cells each (Jurgens, 2001; Fig. 1.1b). While the apical layer of cells gives rise to the shoot meristem and the larger part of the cotyledons, descendents of the basal proembryo layer form the remaining part of the cotyledons, the hypocotyl, the embryonic root and the proximal root stem cells. The larger basal cell of the divided zygote produces a file of seven to nine cells by repetitive horizontal divisions that embody the extraembryonic suspensor that serves as a conduit for nutrients and growth regulators to support the embryo proper development (Yeung and Meinke, 1993). From this basal cell lineage, the uppermost cell (the presumptive hypophysis) is sequestered by the embryo to adopt an embryonic fate and participate in development (Fig. 1.1b). During heart stage of embryogenesis, the hypophyseal cell divides asymmetrically to generate an upper lens-shaped cell that will form the QC and a basal columella root cap progenitor. When the QC is specified, it induces the surrounding cells to become the root meristem stem cells (Dolan et al., 1993; Scheres et al., 1994; Fig. 1.1b). Postembryonic development initiates from these stem cells that are laid down in the heart of the root meristem. 1.2.1

Zygote and early embryo development

The first step toward root formation is the establishment of the apical–basal axis of the developing zygote. Over the past two decades, several factors involved in this early patterning event have been identified and linked at the genetic level. The early expression dynamics of several members of the WOX transcription factor family coincides with specific cell fate changes during early embryo development (Haecker et al., 2004). Before fertilization, WOX2 and WOX8 are expressed in the egg cell and the central cell of the embryo sac and, thereafter, in the elongating zygote. After division of the zygote, WOX2 mRNA is restricted to the apical cell, whereas WOX8 mRNA is found exclusively in the basal cell accompanied by WOX9 expression. At the octant (8-cell) stage, WOX2 and WOX8 mRNAs remain confined to the corresponding embryo domains of the apical and basal cell derivatives, respectively. WOX9 expression expands into the central domain of the embryo, crossing the clonal boundary established at the first zygotic division, and weakens in the uppermost suspensor cell (Haecker et al., 2004). Although this dynamic WOX9 expression pattern is not always observed, these early expressed WOX genes share redundant functions during embryo development (Wu et al., 2007). Strong wox9 loss-of-function mutants arrest development of two to three divisions after the zygote stage. Eradicating WOX8 function in the wox9 background enhances this defect, arresting embryos after the first division. Likewise, wox8 increases apical embryo defects caused by an insertion mutation in WOX2, resulting in abnormal cotyledon separation (Wu et al., 2007).

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Development of the basal cell lineage has been shown to require a mitogenactivated protein kinase kinase (MAPKK), named YODA (Lukowitz et al., 2004). In yoda loss-of-function mutants, the elongation of the zygote is suppressed and the cells of the basal lineage divide in an unpredictable manner. As a result, these cells fail to form the suspensor and are ultimately incorporated into the developing embryo. The apical cell initially develops normally into a wild-type octant embryo, indicating a specific role for the MAP kinase signaling pathway in the correct specification of the basal cell lineage (Lukowitz et al., 2004), perhaps through the partitioning of fate determinants, such as the WOX genes. After division of the zygote, a profound auxin response maximum in the apical daughter cell is quickly established as visualized by fluorescent DR5 reporter expression (Friml et al., 2003). Expression of PIN1 is restricted to the apical cell and the PIN1 protein is distributed within the cell membrane in a nonpolar fashion. In contrast, PIN7 is polarly localized to the apical membrane of the basal cell facing the apical embryo pole. The asymmetric localization of PIN1 and PIN7 proteins present at the first steps of embryogenesis suggests the auxin transport routes from the maternal tissues to establish the auxin gradient and to initiate polarization (Friml et al., 2003; Fig. 1.1b). In a small, but reproducible, number of early pin7 mutant embryos, the apical cell divides horizontally rather than vertically, accompanied by misexpression of the DR5 marker in the extraembryonic suspensor. At later stages, these pin7 embryos display irregular division patterns in the lower embryo regions and, occasionally, fail to establish a proembryo (Friml et al., 2003). Interestingly, around the globular (32-cell) stage, pin7 embryos start recovering from these defects, eventually resulting in fertile plants with no apparent phenotype. Double, triple and quadruple pin mutant embryos show more severe patterning defects and do not recover, indicating functional redundancy among the PIN genes (Friml et al., 2003; Blilou et al., 2005; Vieten et al., 2005). PIN proteins in overlapping expression domains compensate for the loss of one another, even to the extent of ectopic PIN2 expression in pin3pin4pin7 embryos, whereas PIN2 is normally not expressed at these embryonic stages (Blilou et al., 2005), implying the presence of a flexible compensatory mechanism for the loss of PIN proteins in embryos. Defects in cultured NPA-treated embryos and early pin7 embryos resemble the phenotypes of the gnom/emb30 (gn) mutant (Mayer et al., 1993). GN encodes a Brefeldin A (BFA)-sensitive membrane-associated guanine nucleotide exchange factor for ARF GTPase involved in the regulation of intracellular endosomal trafficking (Steinmann et al., 1999; Geldner et al., 2003). Strong loss-of-function gn mutants display a nearly symmetric division of the zygote, followed by oblique divisions of the apical cell. Unable to compensate these defects, gn mutants fail to specify the hypophysis and, ultimately, develop into seedlings without a functional root meristem. In gn embryos, the DR5 reporter activity was detected ectopically in the suspensor, mimicking the DR5 expression upon auxin efflux inhibition (Friml et al., 2003). Coordinated

6
Root Development localization of PIN1 is perturbed in gn embryos (Steinmann et al., 1999), but with a BFA-resistant form of GN, the PIN1 localization is no longer sensitive to BFA, whereas other trafficking processes remain affected (Geldner et al., 2003). Thus, GN is responsible for mediating the intracellular trafficking of PIN1-containing endosomes. Mutants of the vacuolar protein sorting 29 (VPS29), a member of the retromer complex, display similar embryonic defects as those reported for gn (Jaillais et al., 2007). Moreover, the PIN1 localization is also affected in these mutants. Genetic analysis indicates that the VPS29 function is required downstream of GN for proper PIN1 protein cycling. Although more PIN protein family members have been shown to rapidly cycle between the plasma membrane and endosomal compartments and to internalize upon BFA treatment, the GN action does not seem to mediate all PIN protein trafficking to the same extent (Friml et al., 2003; Geldner et al., 2003; Grebe et al., 2003). Recently, a PIN2-specific endosomal cycling route has been described that depends on the SORTING NEXIN 1 (SNX1) protein, hinting at the existence of at least two different endosomal factors for the trafficking of auxin transport facilitators (Jaillais et al., 2006). As in weak gn mutants, growth is impaired in snx1 roots and the normal auxin distribution is perturbed. Double homozygous mutants for vsp29 and snx1 could not be obtained, suggesting that the loss of function of these two genes is either gametophytic or embryonically lethal (Jaillais et al., 2007). As SNX1 and VPS29 are possible components of the retromer complex in plants, VPS29 might contribute to the SNX1–PIN2 pathway as well. Molecular analysis of the auxin-insensitive mutants monopteros (mp) and bodenlos (bdl) highlighted a role for auxin in embryonic root formation (Berleth ¨ and Jurgens, 1993; Hamann et al., 1999). Initial abnormalities in mp and bdl are manifested in the apical region as early as the two-cell stage. In contrast to wild type, the apical cell in mp and bdl embryos divides horizontally, with twice the number of tiers at the octant stage as a consequence. At later stages, mutant embryos fail to specify the hypophysis correctly (see below) resulting in the complete absence of a seedling root. The abnormal divisions in the proembryo are apparent before any defect can be discerned in the basal cell lineage, suggesting instructive communication across the clonal boundary between hypophysis and proembryo. Interestingly in this respect, the WOX9 expression does not shift into the central embryo domain in mp and bdl mutants, nor is it downregulated in the hypophysis, implying that WOX9 acts as a downstream component of auxin signaling (Haecker et al., 2004). The mpbdl double mutant embryos closely resemble the single-mutant phenotype, indicative for a role of both genes in the same developmental pathway (Hamann et al., 1999). Indeed, the auxin response inhibitor BDL/IAA12 has been identified as an in planta interacting partner of the MP/ARF5 transcription factor (Hardtke and Berleth, 1998; Hamann et al., 2002; Weijers et al., 2006). The bdl mutant phenotype is caused by a dominant mutation that enhances protein stability, thereby constitutively inhibiting MP-dependent responses (Hamann et al., 2002). During early embryo development, MP and BDL transcripts are coexpressed in the apical embryo domain as early as the

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first division of the zygote, and, from the globular stage onward, both mRNA species become gradually restricted to the provascular cells (Hamann et al., ¨ 2002; Weijers and Jurgens, 2005). Recently, a stabilized version of the BDL paralog IAA13 has been demonstrated to cause similar embryonic defects as those seen in bdl (Weijers et al., 2005). Since IAA13 is expressed in the same domain as BDL, both BDL and IAA13 are assumed to need to be degraded in early embryogenesis for MP to promote root specification. Recently, binding of the TOPLESS (TPL) protein to BDL was found to be required for BDL repression of MP activity. In accordance, tpl can suppress the patterning defects of the bdl mutant (Szemenyei et al., 2008). Together, these data suggest that correct auxin homeostasis and signaling are required for axis formation and specification of basal cell fates. The embryonic patterning activity of MP is largely dispensable when the presumptive glutamate carboxypeptidase ALTERED MERISTEM PROGRAM 1 (AMP1) is not functional (Vidaurre et al., 2007). amp1 suppresses the phenotype of mp during embryo development and amp1mp mutants frequently form hypocotyls and roots postembryonically. In amp1 embryos, a failure of basal cell descendants to attain suspensor cell fate leads to abnormal divisions, generating additional cell tiers in the embryo proper. Together with the overlapping expression domains of both genes, MP might interfere locally with AMP1-promoted cell differentiation to maintain the basal meristematic region (Vidaurre et al., 2007). In contrast to mp and bdl mutants, early patterning defects in hobbit (hbt) and auxin resistant 6 (axr6) develop first in the basal cell lineage (Willemsen et al., 1998; Hobbie et al., 2000). Instead of the typical horizontal partitioning, the uppermost derivative of the basal cell displays vertical divisions at the four-cell stage. At later developmental stages, hbt and axr6 fail to develop a functional root meristem and, similarly to mp and bdl, result in seedlings without root. The HBT gene encodes a subunit of the anaphase-promoting complex (APC), a class of ubiquitin protein ligase (Blilou et al., 2002). Unlike CDC27A and core APC/C subunits that are constitutively expressed (Blilou et al., 2002; Capron et al., 2003; Kwee and Sundaresan, 2003), HBT/CDC27B is restricted to mitotically active and elongating cells and is mostly excluded from differentiated tissues. Analysis of weak hbt alleles and loss-of-function clones indicates that the primary function of HBT is to mediate cell division and endoreduplication, contributing to meristem activity and cell expansion (P´erez et al., 2008). Consistent with a role for the APC in protein degradation, the auxin response regulators were found to be stabilized in hbt plants (Blilou et al., 2002). The induced loss-of-function hbt clones, however, indicate that cell division and expansion defects caused by HBT removal are not primarily caused by altered perception or altered auxin distribution in the root meristem (Serralbo et al., 2006). Moreover, it remains to be seen whether Aux/IAA proteins are direct substrates of the APC complex. Mutations in HBT suggest that alterations in cell cycle can interfere with specific signaling events as a consequence of modified cell division patterns. The connection between cell cycle and embryonic patterning is evident from studies of the tilted1 mutant

8
Root Development (Jenik et al., 2005). This mutant carries a viable mutation in the catalytic subunit of DNA polymerase ε that lengthens the cell cycle throughout embryo development, with aberrant hypophyseal cell divisions and ultimately displacement of the root pole from its normal position on top of the suspensor as a consequence. Loss-of-function mutations in AXR6, which encodes the CULLIN subunit of the SCFTIR1 E3 ubiquitin ligase complex, result in accumulation of the AXR2/IAA7 protein and, most probably, other Aux/IAA proteins. Potential Aux/IAA candidates to accumulate in axr6 are the BDL protein and/or its functional paralog IAA13 (Hellmann et al., 2003). 1.2.2

Specification of the hypophysis

The proembryo induces the uppermost extraembryonic suspensor cell to become the hypophysis that is destined to generate the QC and columella root cap. Based on morphological observations, this hypophysis specification has been speculated to take place between the octant and dermatogen (16-cell) ¨ ¨ stage of embryogenesis (Jurgens and Mayer, 1994; Jurgens, 2001). However, besides the transient expression of WOX9 in the uppermost suspensor cell at the four-cell stage, stable molecular markers have not been identified at these stages to characterize the specification of the hypophysis. The first genetic markers, indicative of hypophysis identity, appear some time between the transition from dermatogen to globular stage with the onset of WOX5 and PIN4 gene expression (Friml et al., 2002a; Haecker et al., 2004). Moreover, between dermatogen and globular stage, the suspensor has reached its final cell number (7–9 cells) and will no longer divide (Mansfield and Briarty, 1991). Based on these morphological and gene expression criteria, specification of the hypophysis might well occur between dermatogen and globular stage rather than before these stages (Fig. 1.1b). Coinciding with hypophysis specification, a dramatic shift in the apical–basal auxin response gradient can be observed (Friml et al., 2003). Expression of the DR5 marker shifts basally into the uppermost suspensor cells and its activity in the proembryo ceases. This event synchronically occurs with a reversal in the PIN protein polarity. From an apparently random distribution, PIN1 becomes basally localized in the provascular cells facing the hypophyseal cell, whereas PIN7 shifts from the apical to the basal membrane of the suspensor cells (Steinmann et al., 1999; Friml et al., 2003; Fig. 1.1b). Concomitantly, the hypophyseal cell boundary becomes marked by the presence of a third PIN family member, PIN4, presumably supporting the action of the PIN1-facilitated auxin transport (Friml et al., 2002a, 2003; Fig. 1.1b). PIN7-mediated efflux of auxin in the suspensor might operate at a lower rate than the auxin transport by PIN1 and PIN4, with the observed reversal of the auxin gradient as a consequence (Friml et al., 2003). Positioning of the auxin maximum in pin4 mutant embryos is delayed and less restricted to the basal domain than that in the wild type (Friml et al., 2002a). pin4 mutants

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display premature divisions of the hypophysis derivatives at the globular stage and occasional supernumerary cell divisions in the QC region at later stages. These aberrant divisions correlate with an expanded expression domain of the QC-specific enhancer trap line QC25, suggesting that cell fate is not properly specified in the root meristem of pin4 mutants because of impaired positioning of the basal auxin maximum (Friml et al., 2002a). The importance of basal auxin accumulation for the hypophysis fate determination is further supported by genetic studies on molecular components involved in the regulation of polar PIN delivery (Friml et al., 2004; Michniewicz et al., 2007). Overexpression of the protein serine/threonine kinase PINOID (Christensen et al., 2000; Benjamins et al., 2001) in the early globular embryo prohibits the normal basal-to-apical shift of the PIN1 localization and, hence, averts the appearance of a basal auxin maximum. Consequently, the hypophysis is misspecified, as demonstrated by its aberrant transverse division and absence of a seedling root (Friml et al., 2004). Loss of function of the A regulatory subunits of the protein phosphatase 2A (PP2A) also leads to a basal-to-apical PIN polarity shift in developing embryos, resulting in unfocused DR5 expression at the basal end and uncoordinated divisions of the presumptive hypophysis (Michniewicz et al., 2007).Biochemical studies demonstrate PP2A and PID act antagonistically on reversible phosphorylation of PIN proteins, thereby regulating the apical-to-basal targeting of these efflux carriers (Michniewicz et al., 2007). Specification of the hypophysis relies heavily on the interacting BDL and MP proteins. In bdl and mp mutants, the presumptive hypophysis fails to undergo the asymmetric division that generates the precursors of the QC and ¨ columella stem cells (Berleth and Jurgens, 1993; Hamann et al., 1999). Interestingly, MP and BDL transcripts accumulate in the proembryo, but the basal part of the early embryo does not express either mRNA. After the asymmetric division of the hypophysis, these transcripts start to accumulate in the lensshaped daughter cell (Hamann et al., 2002). Since at early stages neither MP nor BDL proteins move in the presumptive hypophysis, they probably act noncell autonomously in hypophysis specification through a secondary signal (Hamann et al., 2002; Weijers et al., 2006; Fig. 1.1c). A potential candidate to act as such a messenger might be auxin itself; after all, auxin accumulates in the uppermost suspensor cell at the time of its specification to become the hypophysis (Friml et al., 2003). Moreover, both PIN1 expression levels and the basal auxin translocation toward the hypophysis appear to depend on BDL activity, putting auxin transport and part of its machinery downstream of the MP and BDL action. Exogenous application of the synthetic auxin 2,4-D to mp and bdl mutants is not sufficient to restore the hypophysis specification, signifying that MP/BDL-dependent factors exist other than auxin that act on this specification event. However, the 2,4-D concentration used in the attempt to rescue the mp and bdl root phenotypes has also been reported to interfere with normal development (Friml et al., 2003). It would be interesting to see whether auxin specifically produced in, or supplied to, the hypophyseal

10
Root Development cell could complement the mp and bdl hypophyseal cell defect. Intriguingly, ectopic expression of a stabilized bdl mutant in the hypophysis derivatives at heart stage did not inflict embryonic root defects, suggesting that BDLdependent MP action is only required for hypophysis specification and plays no role in subsequent embryonic root formation (Weijers et al., 2006). Although orc mutant has been identified in a genetic screen for postembryonic root patterning defects, the first phenotypic abnormalities have been traced back to late globular embryos, where divisions of the hypophyseal cell are either absent or irregular (Willemsen et al., 2003). Ectopic DR5 activity suggests aberrant distribution of auxin in basal orc embryo domains. Map-based cloning of the orc mutation has identified a point mutation in STEROL METHYLTRANSFERASE 1 (SMT1), encoding a protein required for sterol homeostasis, indicating that a balanced sterol composition is a major requirement for proper auxin distribution during embryogenesis (Diener et al., 2000; Willemsen et al., 2003). Together, the data establish that an auxin maximum at the basal pole is important for correct hypophysis specification and subsequent root meristem formation. In addition, specification of the hypophysis might also rely on another, MP/BDL-dependent noncell autonomous signal, suggesting that, rather than specifying the hypophysis fate directly, auxin induces a primary response in the adjacent proembryo cells and is then transmitted basally as part of a secondary signal (Weijers et al., 2006; Fig. 1.1c).

1.3

Root stem cell niche specification

At late globular stage, the hypophysis has divided asymmetrically, producing a basal cell from which the root cap cells originate and a smaller lens-shaped ¨ apical cell that generates the four mitotically inactive QC cells (Jurgens and Mayer, 1994; Scheres et al., 1994; Fig. 1.1b). At heart stage, the QC is assumed to recruit the immediately adjacent cells to become the root stem cells. The QC and surrounding stem cells together form the root stem cell niche, a specialized microenvironment conditioned to maintain stem cells. At late heart stage, asymmetric periclinal division in the lowest protoderm cells generate epidermis and lateral root cap, marking the first stem cell division within the newly formed root stem cell niche (Scheres et al., 1994). At midtorpedo stage, a first round of asymmetric divisions of the columella stem cells produces a second layer of columella root cap cells and the ground tissue ¨ stem cell daughters generated the first endodermal cells (Jurgens and Mayer, 1994). 1.3.1

A role for AP2 transcription factors

From the early globular stage on, an auxin maximum is located at the basal tip of the developing embryo that is maintained throughout postembryonic

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root development (Sabatini et al., 1999; Friml et al., 2003; Fig. 1.1b). In postembryonic roots, a lateral shift of the endogenous auxin maximum induced by auxin transport inhibitors respecifies the root stem cell niche fate in ground tissue layers. Former endodermal cells express the QC-specific marker QC46 and the adjacent cortex cells display the columella stem cell marker J2341 and divide. Exogenous application of auxin induces cell fate changes similar to those observed upon polar auxin transport inhibition, suggesting that an auxin maximum functions as a positional cue for the stem cell niche (Sabatini et al., 1999). In the past years, potential effector genes of the instructive auxin maximum have been identified. PLETHORA1 (PLT1) and PLT2 encode auxin-inducible members of the AP2 domain transcription factor family and are redundantly required for the embryonic specification of the stem cell-organizing QC (Aida et al., 2004). plt1plt2 double mutants show aberrant divisions of the lensshaped cell and fail to express the QC25 and QC46 markers throughout embryogenesis. Exogenous application of auxin fails to rescue the stem cell defects in plt1plt2 mutants, indicating that the required PLT activity for root stem cell niche formation cannot be bypassed by auxin. At heart stage, expression of the PLT genes is restricted to the basal stem cell niche, in which MP and its close homolog NONPHOTOTROPIC HYPOCOTYL 4 (NPH4/ARF7; Harper et al., 2000) act redundantly to maintain the PLT expression (Aida et al., 2004). Strikingly, in pin2pin3pin4pin7 quadruple mutants, the PLT1 expression expands throughout the whole embryo. When explanted, these embryos develop reduced cotyledons and root hairs emerge at more apical positions on the seedling. Conversely, in pin1pin3pin4pin7 quadruple mutants, the expanded expression of the shoot meristem identity gene WUSCHEL (WUS) leads to explants with arrested root growth and an expanded shoot domain, implying that directional auxin transport conducted by the PIN proteins regulates the patterning of embryonic stem cell domains (Blilou et al., 2005). In turn, the PIN4 transcript has been found to be severely reduced in plt1plt2 mutants, as well as the expression of PIN3 and PIN7 in the postembryonic elongation zone. All together, these findings suggest a feedback loop for embryonic root primordium formation and stabilization: PIN proteins restrict PLT expression in the basal embryo domain to initiate embryonic root specification, and, in turn, PLT activity regulates PIN transcription to stabilize the position of the root primordium (Blilou et al., 2005). Recently, two additional PLT family members, PLT3 and BABY BOOM (BBM), have been identified that act in concert with PLT1 and PLT2, contributing to embryonic root development and stem cell maintenance (Galinha et al., 2007). From the heart stage of embryogenesis onward, the expression domain of PLT3 and BBM largely overlaps with that of PLT1 and PLT2, with the strongest expression in the basal stem cell niche. Removal of wild-type copies of PLT3 and/or BBM in a plt1plt2 double mutant background increases the patterning defects in the root pole development. These defects culminate in root- and hypocotyl-less seedlings in the progeny of self-fertilized

12
Root Development plt1−/− plt2+/− plt3−/− bmm−/− , indicating that the PLT activities are largely additive and dosage dependent. Aberrant development starts at the early globular stage as revealed by transverse divisions of the hypophysis. Apical embryo development, however, seems largely unaffected in triple mutants. Together with homeotic fate transformation of the shoot meristem toward root fate upon ectopic PLT2 expression, these results suggest that PLT genes act as master regulators for root development (Galinha et al., 2007). 1.3.2

A role for GRAS transcription factors

Other transcription factors involved in stem cell niche specification at early heart stage are represented by members of the GRAS family, SHORT-ROOT (SHR) and SCARECROW (SCR). shr and scr mutants display a disorganized QC/columella region and roots that cease growth prematurely (Benfey et al., 1993; Scheres et al., 1995; Di Laurenzio et al., 1996; Helariutta et al., 2000). Moreover, expression of QC25 and QC46 is perturbed in both mutants and columella stem cells fail to be maintained, indicating a role for SHR and SCR in specification of the QC and maintenance of a functional stem cell niche (Sabatini et al., 2003). The earliest detectable SCR expression is in the hypophyseal cell and precedes its division to generate the QC and the columella lineages (Wysocka-Diller et al., 2000). At later developmental stages, SCR expression gets restricted to the endodermal cell layer, cortex/endodermis stem cell and the QC (Fig. 1.2a). In shr mutants, expression of SCR is severely reduced, indicative that SHR regulates the SCR expression (Helariutta et al., 2000). Recently, Levesque et al. (2006) have identified SCR as a direct target gene that is positively regulated by SHR. SHR is not transcribed in the SCR domain but in the adjacent provascular cells from where regulated and targeted movement transfers the protein into the adjacent cell layers to promote SCR transcription (Nakajima et al., 2001; Gallagher et al., 2004; Fig. 1.2a). In scr mutants, recuperation of the SCR expression in the QC is sufficient for cell-autonomous rescue of the QC identity, maintenance of the surrounding stem cells and root growth. In contrast, ectopic expression of SCR only in the stem cells is insufficient for their maintenance and to restore the QC identity, suggesting that the SCR activity enables the QC to maintain the surrounding stem cells in a noncell-autonomous fashion (Sabatini et al., 2003). These observations are in concordance with earlier studies in which the QC has been identified as a negative regulator of differentiation of adjacent stem cells (van den Berg et al., 1997). Re-expression of SCR in the QC of shr mutants is not sufficient to restore QC and stem cell identity, hinting at a more elaborate role for SHR in QC function and stem cell maintenance than its sole requirement for SCR transcription (Sabatini et al., 2003). In fact, recent findings indicate that SHR and SCR proteins interact and share a common set of transcriptional target genes (Cui et al., 2007, see below). Most likely, the regulatory activity of the SCR/SHR protein complex is required for the proper QC function.

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In the QC, SCR is required for maintaining its own expression and the nuclear localization of SHR (Helariutta et al., 2000; Cui et al., 2007). An additional factor regulating the nuclear localization of SHR has been identified recently as a zinc finger transcription factor, designated JACKDAW (JKD; Welch et al., 2007). In jkd mutants, the QC is misspecified and stem cell activity is compromised, resulting in plants with reduced root length and meristem cell numbers. Moreover, SCR expression is absent from the QC and SHR predominantly localizes to the cytoplasm. In scr mutants, a similar localization pattern for SHR has been observed, suggesting that JKD controls the nuclear localization of SHR mainly through its effect on the maintenance of SCR expression (Welch et al., 2007). 1.3.3

Combinatorial gene activity positions the stem cell niche

Overexpression of PLT during embryogenesis ectopically accumulates QC and associated root stem cells and, in most extreme cases, leads to a complete transformation of the embryo toward root identity (Aida et al., 2004). In accordance with ectopic auxin application studies (Sabatini et al., 1999; see above), ectopic QC cells in the PLT overexpressors appear to originate from cells adjacent to the stele, suggesting that stele/endodermis-residing factors are necessary, in concert with PLT, to specify the root stem cell niche. SHR and SCR are likely candidates because both are expressed in the stele/endodermis and are required for QC specification and maintenance. In addition, transcription of PLT1 is not affected in shr and scr mutants and PLT activity is not required for SHR or SCR expression, indicative of parallel inputs. Accordingly, in plt1plt2shr and plt1plt2scr triple mutant combinations, cells in the root meristem differentiate earlier than any of the shr or scr single or plt1,plt2 double mutants (Aida et al., 2004). Moreover, although the lack of expression of the QC markers QC25 and QC46 in shrscr and plt1plt2 mutants suggests that both pathways share a set of target genes, the PLT and SHR/SCR pathways do not fully converge on a same set of target genes because the expression of QC184 solely depends on the PLT activity (Sabatini et al., 2003; Aida et al., 2004). All together, these data support a model in which the PLT and SHR/SCR signaling pathways commonly regulate the root stem cell niche specification, providing spatial cues and cell fate instructions through the overlap of their expression domains (Aida et al., 2004; Fig. 1.2a). Although genetic studies performed in the developing embryo would be most informative when examining stem cell niche specification, some of the molecular mechanisms of pattern formation and niche specification might also be learned from regeneration experiments in the postembryonic root. Laser-induced ablation of QC cells rapidly respecifies the QC and root cap in the distal provascular tissue (van den Berg et al., 1995), whereas the laserinduced wound probably disrupts the acropetal auxin flow toward the root tip, resulting in a proximal shift of the auxin response maximum as fast as 3 hours after ablation (Sabatini et al., 1999; Xu et al., 2006). In response, the

14
Root Development PLT1 expression domain shifts accordingly to the newly formed maximum. Later, SHR and SCR also reallocate their expression domains proximally, resulting in a respecified QC within 2 days after ablation. The induction of the new cell fates relies on the newly established expression domains and combinatorial activity of PLT, SHR and SCR, because regeneration does not occur in these mutant backgrounds. Furthermore, wild-type PIN4 expression and polarity is re-established only after PLT, SHR and SCR have adopted their new expression domains. Thus, upon ablation, the formation of a new proximal auxin response maximum first induces cell fate changes mediated by the root patterning genes and, only consequently, changes in polarity of the auxin flow facilitated by the PIN proteins (Xu et al., 2006). These findings support the current model of embryonic root stem cell niche formation. In response to a PIN-mediated auxin maximum, the PLT patterning genes become restricted to define the stem cell region in concert with SHR and SCR and, in turn, control root-specific PIN expression to stabilize the maximum (Blilou et al., 2005; Xu et al., 2006; Galinha et al., 2007).

1.4

Radial patterning

The radial organization starts within a group of cells derived from the apical cell at the octant stage of embryogenesis. A first set of periclinal divisions separates three founder cells for the protoderm, ground tissue and procambium whose derivatives will later form the epidermis/lateral root cap, endodermis/cortex and vascular bundle in the root. The vasculature is laid down at the late globular stage when the inner four procambium cells perform a round of periclinal divisions that separate the pericycle from the central provascular tissue (Scheres et al., 1995). Through subsequent periclinal divisions, the central cells generate the xylem and phloem lineage founder cells that will ultimately produce the corresponding conductive vascular bundles (see Chapter 2). At late heart stage, the most basal protoderm cells carry out an asymmetric periclinal division generating two daughter cells with separate fate (Dolan et al., 1993; Scheres et al., 1994). Consecutive anticlinal divisions of both daughter cells will produce one file of lateral root cap and one epidermal cell layer. Recently, the FEZ and SOMBRERO genes have been identified from a markerbased screen that seem specifically involved in orienting the division of this asymmetric epidermis/lateral root cap stem cell division (Willemsen et al., 2008). The developing epidermal cells will differentiate either as root hair cells or nonhair cells, depending on genetic determinants and positional information (Dolan et al., 1993; see Chapter 3). 1.4.1

Epidermis fate initiation and receptor signaling

The onset of epidermis cell differentiation is marked by the asymmetric segregation of the homeodomain-encoding genes Arabidopsis thaliana MERISTEM

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LAYER1 (ATML1) and PROTODERMAL FACTOR2 (PDF2). Expression of ATML1 and PDF2 initiates at the quadrant-stage embryo and segregates to the L1- or protoderm layer during early globular stage (Lu et al., 1996; Abe et al., 2003). atmlpdf2 double mutation results in severe defects in shoot epidermal cell differentiation, but seems not to affect the anatomy of the root apical meristem and root growth (Abe et al., 2003). Recently, the redundant leucine-rich repeat receptor kinases RECEPTORLIKE PROTEIN KINASE1 (RPK1) and TOADSTOOL2 (TOAD2) have been shown to be critical for radial patterning (Nodine et al., 2007). rpk1toad2 double mutants are embryo lethal and arrest as mushroom-shaped embryos at heart stage. Phenotypic defects are first observed at the early globular stage, correlating with the overlapping expression of both proteins and consist of bloated central protoderm cells. These cells initially express the protoderm marker ML1, but fail to maintain its expression as the phenotype becomes apparent. In addition, the vascular primordium markers ZWILLE/PINHEAD and SHR are ectopically expressed in ground tissue and protoderm, whereas the ground tissue marker SCR is absent. Interestingly, also the correct asymmetric division of the hypophysis is impaired, failing to produce the lens-shaped cell. However, frequent failure to express SCR and the ectopic provascular marker indicates that defects in hypophysis specification cause the aberrant asymmetric division. These results indicate that RPK1/TOAD2 signaling is required to maintain the protoderm fate, to restrict provascular fate and to specify hypophysis and ground tissue either directly or indirectly due to misspecification of the surrounding tissues (Nodine et al., 2007). Up and downstream effectors are needed to separate cause from effect. 1.4.2

Ground tissue patterning

Subsequent asymmetric periclinal divisions of the ground tissue founder cell generate the endodermis and cortex tissues in early heart stage embryos (Scheres et al., 1994). SHR and SCR, besides their function as root stem cell regulators, are also important determinants of this radial pattern formation process (Benfey et al., 1993; Scheres et al., 1995; Di Laurenzio et al., 1996; Helariutta et al., 2000; Fig. 1.2b). In shr and scr mutants, only a single layer of ground tissue is present between epidermis and pericycle as a result of increased recalcitrance of the ground tissue precursor to divide periclinally. Whereas the single layer of ground tissue has a mixed cortex/endodermis identity in scr mutants (Di Laurenzio et al., 1996; Heidstra et al., 2004), it expresses cortex fate markers solely in shr mutants, suggesting a role for SHR not only in promoting the asymmetric stem cell division but also the endodermis identity (Benfey et al., 1993). The different functions of SHR in radial pattern formation and endodermis specification, but also in stem cell maintenance, might be reflected by domain-specific expression or function of direct downstream target genes. An extensive study to identify downstream target genes predicted eight candidates positively regulated by SHR of which four genes were able to bind SHR to their promoter sequences in vivo (Levesque et al., 2006).

16
Root Development Two of these genes, MAGPIE (MGP) and NUTCRACKER (NUC), encode closely related C2H2 zinc finger transcription factors that are expressed in the cortex/endodermis stem cell and the lower endodermal lineage. Therefore, MGP and NUC could act redundantly in stem cell fate regulation and/or radial patterning. A third target, the metabolic enzyme tropine reductase (TRI), is mostly produced in the endodermis, suggesting a possible role in endodermis fate establishment and/or radial patterning. Finally, SHR has been found to directly regulate SCR transcription through binding to the SCR promoter region (Levesque et al., 2006). Given that SCR is cell autonomously required for QC specification and asymmetric ground tissue cell division (Sabatini et al., 2003; Heidstra et al., 2004; see below), SHR functions in these two processes partly through direct regulation of SCR (Levesque et al. (2006). Interestingly, a number of transcriptional targets of SHR also appear to be direct targets of SCR, including SCR itself that can bind to its own promoter (Cui et al., 2007). In a scr background, binding of SHR to the promoter of some of its targets is abolished, indicating functional interdependence between these two transcriptional regulators. A molecular basis for this interdependence is provided by the finding that SCR physically interacts with SHR in yeast two-hybrid and reciprocal pull-down experiments (Cui et al., 2007). Clonal deletion of SCR from the ground tissue stem cells results in the single layer of ground tissue typical for the scr mutant phenotype, strongly hinting at a strict cell-autonomous requirement of SCR for the periclinal division of the endodermis/cortex stem cell daughter (Heidstra et al., 2004). Periclinally divided ground tissue clones that segregate activated SCR expression to the ‘outer’ cells continue to express endodermal markers in the ‘inner’ cells that lack SCR expression. Moreover, endodermis and cortex-specific markers are maintained in their respective ground tissue layer upon clonal SCR deletion. These results indicate that SCR is required only transiently for stable and immediate separation of cell fates, possibly by respecification of the chromatin state at mitosis (Heidstra et al., 2004). Although both SHR and SCR are present in the complete endodermis cell layer, the asymmetric periclinal division is restricted to the cortex/endodermis stem cell daughter (Fig. 1.2b), suggesting that the presence of stem cell-specific factors aid the process of asymmetric division and/or the involvement of extrinsic ‘top-down’ signals from mature ground tissue cells to reinforce the asymmetric division. The latter possibility has been deduced from laser ablation experiments that revealed the inability of ground tissue stem cells to perform asymmetric divisions after being isolated from their mature daughter cells (van den Berg et al., 1995). However, clonal induction of SCR in the scr mutant revealed that all ground tissue cells are competent to perform the periclinal division, including the segregation of both fates, in the absence of mature endodermis and cortex acting as a patterning template. Together with the strict cell-autonomous action of SCR, the need for top-down signaling is ruled out to pattern the ground tissue (Heidstra et al., 2004). A possible reason that the stem cell daughters do not divide periclinally after laser ablation of their mature daughter cells, might be linked to

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noncell-autonomous signaling from the QC that prevents progression of stem cell differentiation (van den Berg et al., 1997; Sabatini et al., 2003; Heidstra et al., 2004; Fig. 1.2b). Movement of SHR to the endodermal lineage requires the protein to be localized cytoplasmically in the stele. However, cytoplasmic localization does not automatically imply movement as demonstrated by missense mutations that disrupt nuclear localization, but also movement of SHR, indicating that SHR transport is regulated and targeted (Gallagher et al., 2004; Fig. 1.2b). Moreover, ectopic expression of a complementing GFP-tagged version of SHR was not able to move from phloem companion cells or epidermal cells, suggesting a need for vascular-specific factors to enable SHR movement (Sena et al., 2004). When expressed from the SCR promoter, SHR induces supernumerary ground tissue layers with endodermis characteristics, substantiating a model in which restricted SHR expression in the stele and limited SHR movement only into the adjacent endodermis prevent continued activation of SCR and, subsequently, additional asymmetric periclinal ground tissue divisions (Nakajima et al., 2001). But how is the SHR movement restricted to only one cell layer? Rare periclinal ground tissue divisions in scr mutants maintain GFP:SHR in both layers, suggesting that SCR is required for the asymmetry of the periclinal ground tissue division and fate separation by restricting SHR movement to the endodermis only (Heidstra et al., 2004; Fig. 1.2b). In addition, movement of GFP:SHR in a scr mutant background has only been observed from the epidermis (Sena et al., 2004). Recent findings that SHR and SCR are bound in a complex indicate that indeed SCR sequesters SHR into the nucleus of endodermal cells, thus preventing its movement (Cui et al., 2007), as illustrated in SCR RNAi plants in which residual SCR cannot seize all incoming SHR proteins and develop additional endodermal layers (Cui et al., 2007). The SHR targets MGP and NUC have been isolated also independently in a screen for ground tissue expressed genes together with a third zinc finger family member JKD (Welch et al., 2007). JKD is expressed in the QC, cortex/endodermis stem cell, and, to a lesser extent, in the endodermal lineage. Initiation of JKD expression at early globular stage does not depend on SHR or SCR, but both factors are required postembryonically for JKD maintenance. mgp and nuc single mutants have no apparent phenotype, suggesting a profound genetic redundancy. jkd mutants, besides the QC defects discussed above, display ectopically periclinal divisions in the cortex producing an additional layer with endodermis fate. These extra divisions have not been observed in jkdshr and jkdscr double mutants, indicating that JKD acts in the ground tissue by modifying the activity of SHR and SCR. MGP RNAi lines in wild type reveal no phenotype, but combined with jkd homozygotes largely complement the ground tissue phenotype, suggesting that MGP promotes cell division. In planta, protein–protein interaction studies among SHR, SCR, JKD and MGP indicate that these proteins are capable of nuclear complex formation (Welch et al., 2007), substantiating an elaborate model for the molecular mechanism that controls radial ground tissue patterning. MGP redundantly facilitates the asymmetric cell division by binding to the

18
Root Development SHR/SCR complex and JKD inhibits this activity by competing for binding in the same complex. As the root ages, a new layer of ground tissue can be formed distant from the QC that rapidly takes on the cortex identity (Baum et al., 2002; Paquette and Benfey, 2005). In scr mutants, formation of this extra cortex layer, termed ‘middle cortex’, is observed at a much earlier time point than in the wild type. This phenotype can even be intensified by reducing gibberellic acid (GA) levels in the scr background, either genetically or chemically, indicating that both SCR and GA act independently as negative regulators of middle cortex initiation (Paquette and Benfey, 2005). In contrast to scr, the ground tissue in shr never develops an extra cortical layer and seems insensitive to reduced GA levels. Thus, the maturation of the ground tissue in the root is promoted by a SHR-dependent mechanism that is independently regulated by SCR and GA (Paquette and Benfey, 2005).

1.5

Stem cell maintenance

Following germination, new cells are continuously added by repetitive asymmetric divisions of stem cells in the heart of the root meristem to ensure perpetuation of, and elaboration on the organization set during embryogenesis. Thereupon, stem cell daughters undergo additional divisions in the meristematic zones before rapid expansion and differentiation. To guarantee stem cell action for extensive periods of time, the balance between stem cell proliferation and differentiation must be tightly regulated. 1.5.1

Conserved factors in root and shoot organizer signaling

Laser-induced cell ablation experiments have identified the QC as a source of short-range signals maintaining the immediately adjacent cells as root stem cells (van den Berg et al., 1997). A similar mechanism operates in the shoot meristem where an organizing center (OC) of slowly dividing cells is essential to maintain the adjacent stem cell pool. The OC is defined and maintained by expression of the WUS homeobox gene (Laux et al., 1996; Mayer et al., 1998). WUS regulates nonautonomously the expression of the secreted protein CLAVATA3 (CLV3) in the stem cells that, in turn, interacts with the CLV1 receptor probably in a complex with CLV2 to limit the WUS domain expression and size of the OC (Fletcher et al., 1999; Brand et al., 2000; Schoof et al., 2000; Ogawa et al., 2008). This negative feedback loop between OC and stem cells provides a robust mechanism to balance the stem cell population in the shoot meristem. Whether a similar regulatory mechanism controls the root stem cell population is uncertain. However, some signaling components might be functionally conserved between both meristems. QC-specific expression of WOX5 is nonautonomously required for distal root stem cell maintenance similar to the role of WUS in the shoot (Sarkar

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et al., 2007; Fig. 1.2a). Loss of WOX5 function results in differentiation of the columella stem cells, while overexpression of WOX5 generates an indefinite number of columella stem cells by blocking the differentiation of their daughters or by reverting the fate of the differentiated columella root cap cells. Whereas stem cell maintenance normally depends on QC signaling, laser-induced ablation of the QC does not lead to differentiation of the additional columella stem cells (van den Berg et al., 1997; Sarkar et al., 2007), suggesting that the WOX5 protein itself moves toward stem cells as the long postulated short-range factor (van den Berg et al., 1997). Alternatively, ectopic WOX5 expression might activate downstream signals that normally arise in the organizer cells that now enable self-maintenance of the stem cell population. Expression of WOX5 is predominantly regulated by the SHR/SCR signaling pathway, identifying WOX5 as one of its downstream effectors in stem cell maintenance. Interestingly, promoter swapping experiments have demonstrated that both WUS and WOX5 genes are interchangeable in stem cell control. WOX5 expression under the control of the WUS promoter can rescue the stem cell defects in the wus mutant and, vice versa, QC-specific expression of WUS can compensate for the loss of WOX5 function, demonstrating that some of the central regulators of stem cell identity in the root and shoot meristems are equivalent (Sarkar et al., 2007; Fig. 1.2a). Additionally, ectopic expression of CLV3-like peptides (CLEs) has been shown to affect root meristem maintenance in a CLV2-dependent manner (Fiers et al., 2005). Whereas CLV3 overexpression inhibits stem cell proliferation in the shoot (Brand et al., 2000), CLE overexpression in roots does not seem to interfere primarily with the QC identity nor with stem cell maintenance, but rather to modulate the activity of the stem cell daughters that populate the meristem (Casamitjana-Mart´ınez et al., 2003; Hobe et al., 2003; Fiers et al., 2004). However, it must be noted that the effect on proximal stem cell division rates has not been investigated. The functional product of genes from the CLE family has been identified in planta as a peptide of only 12 amino acids (Ito et al., 2006; Kondo et al., 2006), generated by the posttranslational processing of CLE gene products. In a mutagenesis screen for suppressors of the CLE19 overexpression phenotype, sol1 and sol2 completely rescued the root length and meristem defects up to 1 week after germination. SOL1 encodes a putative Zn2+ carboxypeptidase, suggesting a role in the processing of CLE peptides. The sol1 mutant, however, displays no abnormalities during plant development. These findings suggest that a CLV-like pathway might control the root meristem maintenance (Casamitjana-Mart´ınez et al., 2003). The signals emanating from the shoot and root organizers required to sustain stem cells in their undifferentiated state remain unidentified. The halted root (hlr) mutant has a reduced meristem activity after germination, resulting in retarded root and shoot growth (Ueda et al., 2004). Although the root meristem is properly specified during embryogenesis, SCR expression in the QC and cortex/endodermis stem cells is rapidly lost during postembryonic development. HLR has been found to encode a subunit of the

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Root Development 26S proteasome (Ueda et al., 2004), indicating that the proteasome machinery is needed for stable expression of meristem regulators within the plant stem cell niches. 1.5.2

Stem cell regulation through cell cycle control

Recently, the RETINOBLASTOMA-RELATED (RBR) gene has been identified as a key component in root stem cell regulation (Wildwater et al., 2005). RNAi-induced reduction of RBR transcript levels in the root meristem results in excessive stem cell accumulation of all cell types. The rates of cell division within the stem cell pool, however, are not affected, indicating that the increase in stem cell numbers observed in RBR RNAi lines is due to prolonged maintenance of the stem cell identity. In agreement with the RNAi phenotype, overexpression of RBR primarily affects the stem cells, leading to a rapid loss of their undifferentiated state. As in wild type, laser-induced ablation of the QC in RBR RNAi roots leads to rapid differentiation of the columella stem cells, showing that the supernumerary stem cells are still under QC control. Interestingly, this experiment demonstrates the ability of the QC to signal over multiple cell layers to maintain stem cells, equivalent with the OC in the shoot meristem. Reduction of RBR can compensate for the loss of stem cell activity in a scr mutant background, but not in the shr or plt1plt2 mutants, implying that the role of SCR in stem cell control is to downregulate the RBR pathway in the QC, thereby promoting stem cell maintenance in a noncell-autonomous manner. After all, if a QC-emitted signal was required to suppress RBR in the surrounding stem cells, then stem cell maintenance should be independent of the QC and RBR RNAi background (Wildwater et al., 2005). In mammals, the RETINOBLASTOMA (RB) protein acts as a tumor suppressor and cell cycle regulator. RB inhibits progression of the cell cycle by formation of a repressive complex with the cell cycle-promoting E2F transcription factors (for a review, see Weinberg, 1995). Phosphorylation of RBR by the upstream acting D-type cyclin (CYCD)-cyclin-dependent kinase (CDK) complexes releases the inhibition on E2F action, allowing the cell cycle to progress. CDK/CYCD action is predicted to be inhibited by Kip-related proteins (KRPs) (De Veylder et al., 2001). Overexpression of the Arabidopsis homologs of these cell cycle components resulted in an accumulation (CYCD3 and E2Fa) or a loss (KRP2) of stem cells, in accordance with their postulated roles in the plant RBR pathway (Wildwater et al., 2005), strongly suggesting that RBR controls the population of stem cells in response to D-type cyclins and through the modulation of E2F action. 1.5.3

Auxin and stem cell regulation

Accumulating evidence points toward a role for the stable basal auxin maximum in stem cell maintenance. In PID overexpressors, quantified auxin levels in the root meristem are significantly decreased as a result of

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mislocalization of PIN proteins (Friml et al., 2004). In concurrence, loss of stem cells is followed by terminal differentiation of the meristem. Interestingly, the 35S::PID-mediated stem cell differentiation is delayed in pin2 and pin4 mutant backgrounds. In these pin mutants, auxin accumulates at the root tip presumably as a consequence of the interrupted auxin flow (Friml et al., 2002a; Ottenschl¨ager et al., 2003), suggesting that increased auxin levels at the root tip in 35S::PID plants can partially restore the stem cell function. In accordance, treatment of 35S::PID plants with NPA restores the DR5 marker expression and prevents meristem differentiation (Friml et al., 2004). Alternative evidence comes from the characterization of the redundant ARF10 and ARF16 auxin response factors that have been found to restrict the distal stem cell fate and to promote columella cell differentiation (Wang et al., 2005). The expression of ARF16 is controlled independently by auxin and microRNA160 (miR160), generating its columella and root cap-specific expression pattern. Whereas no root growth phenotype has been reported for the arf10 and arf16 single loss-of-function mutants (Okushima et al., 2005), the arf10arf16 double mutant displays supernumerary root cap layers containing cells with mixed distal, proximal and QC cell fates. Consistent with the arf10arf16 loss-of-function phenotype, miR160-uncoupled overexpression of ARF16 confers a loss of columella stem cells and differentiation of the proximal meristem (Wang et al., 2005). It would be interesting to determine the interplay between QC signaling and ARF10/ARF16 in distal stem cell maintenance. Possibly, the QC acts noncell autonomously to prevent ARF-mediated differentiation of the columella stem cells. 1.5.4

Chromatin and stem cell maintenance

During the last decade, examples in various model organisms point toward an active role of chromatin-remodeling factors in maintaining the balance between stem cell identity and the path to differentiation (Meshorer and Misteli, 2006). Also in Arabidopsis, chromatin-modifying proteins have been linked to meristem stability and stem cell maintenance. The fasciata1 (fas1) and fas2 mutants display severe defects in root and shoot meristem organization, leading to reduced root growth and aberrant shoot development (Leyser and Furner, 1992; Kaya et al., 2001). The premature termination of the root meristem in fas mutants is accelerated by the reduced activity of the QC and stem cells. Columella stem cells are quickly lost after germination, as indicated by accumulation of starch granules, and lack of SCR expression from the QC (Kaya et al., 2001). Formation of the embryonic root, however, is not affected. These developmental defects are also observed in the brushy1(bru1)/mgoun3 mgo3)/tonsoku (tsk) mutant (Guyomarc’h et al., 2004; Suzuki et al., 2004; Takeda et al., 2004), suggesting that the FAS and BRU1/MGO3/TSK genes are specifically required for postembryonic meristem maintenance by regulating stem cell activity, at least partially, through their role in preserving the QC integrity. FAS1 and FAS2 encode two subunits of the Arabidopsis chromatin

22
Root Development assembly factor 1 (CAF-1) complex, a histone chaperone complex thought to be involved in the maintenance of epigenetic information in chromatin during mitosis (Kaya et al., 2001). The genetic relationship between the CAF1 complex and the BRU1/MGO3/TSK protein that is hypothesized to be part of a nuclear protein complex is unclear, although epistatic analysis suggests that they might have common targets in some developmental aspects (Guyomarc’h et al., 2004; Takeda et al., 2004). Two putative histone chaperones, NAP1-RELATED PROTEIN1 (NRP1) and NRP2, have recently been described as required for maintaining postembryonic root growth (Zhu et al., 2006). Although the initial development of npr1npr2 double mutants seems normal, the root ceases to grow 7 days after germination due to a lack of cell division. In contrast with fas and bru1/mgo3/tsk, the nrp1nrp2 double mutant has a root-specific phenotype, suggesting a more restricted role for NRP1 and NRP2. In tebichi (teb) mutants, developmental root meristem has defects similar to those of the bru1/mgo3/tsk and fas mutants (Inagaki et al., 2006). The TEB gene encodes a protein that contains both helicase and DNA polymerase domains. As in fas2, cells expressing CYCB1;1-␤-glucuronidase (GUS) accumulate in the teb and bru1/mgo3/tsk mutants, suggesting that the normal cell cycle progression is impaired. The disorganization and premature differentiation of the root meristem in these mutants might thus be connected with a defect in cell cycle progression at the G2-to-M transition (Suzuki et al., 2005; Inagaki et al., 2006). These mutants also exhibit shoot meristem maintenance defects, indicating a common role for these chromatin-stabilizing factors in stem cell/meristem maintenance, possibly by maintaining the expression state of the meristem regulatory genes. Additionally, components of the CAF complex have been shown to bind RBR in plants (Ach et al., 1997; Kaya et al., 2001), opening up the possibility that RBR mediates stem cell control through the regulation of the chromatin state. This hypothesis is supported by recent results that indicate that known chromatin factor mutants act as enhancers of the RBR RNAi phenotype (Kornet and Scheres, 2009).

1.6

Meristem maintenance and root zonation

Once a daughter stem cell has been created through an asymmetric division of the stem cell, it must acquire not only radial tissue-specific properties (e.g. cortex, epidermis or endodermis fate), but also be informed how to conduct itself according to its position on the apical–basal axis. Proliferation of stem cell daughters within the meristematic zone produces a reservoir of cells that, once pushed into the elongation zone, will elongate to ensure steady root growth. It is of crucial importance that cells within and between meristematic and elongation zone act in a coordinated fashion because an imbalance between cell division and rate of cell elongation zone will eventually result in retarded root growth. Auxin has been linked to both these processes because

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cell expansion and division can be stimulated upon external application of auxin (for a review, see Srivastava, 2002). 1.6.1

Auxin transport dictates root zonation

During root growth, a continuous flow of auxin produced by the shoot apex is transported toward the root tip in a PIN-dependent manner. Auxin seems to circulate through the meristem by basipetal transport and lateral redistribution as suggested by the polar localization of the PIN proteins produced in the root (Fig. 1.2c). PIN1 mainly resides at the basal membrane of the vascular cells (G¨alweiler et al., 1998; Blilou et al., 2005). PIN2 localizes apically in epidermal and lateral root cap cells and predominantly basally in cortical ¨ cells (Muller et al., 1998; Blilou et al., 2005). PIN3 is active in tier two and three of the columella without any pronounced polarity. In the elongation zone, however, PIN3 has been found at the basal side of vascular cells and the lateral side of pericycle cells (Friml et al., 2002b; Blilou et al., 2005). PIN4 is detected in and around the QC and localizes basally in provascular cells (Friml et al., 2002a; Blilou et al., 2005). PIN7 resides at lateral and basal membranes of provascular cells in the meristem and elongation zone, whereas in the columella it coincides with the PIN3 domain (Blilou et al., 2005). Mediated by combined PIN1, PIN3 and PIN7 action, auxin is transported basally through the provasculature toward the root cap, where PIN3, PIN4 and PIN7 action maintain the position of the auxin maximum and redistribute auxin to the lateral periphery. PIN2 facilitates acropetal transport through the epidermis to the elongation zone where the auxin can be reloaded into the provascular system facilitated by PIN1, PIN2, PIN3 and PIN7 (Blilou et al., 2005; Fig. 1.2c). Consistent with this model of auxin circulation, ectopic induction of auxin biosynthesis in the QC results in enhanced auxin responses, measured by DR5 reporter expression, appearing first in the columella area, subsequently in the lateral root cap and epidermis, and finally in the provascular strand (Blilou et al., 2005). The significance of this auxin reflux loop on cell division and cell expansion is evident from the analysis of pin mutant root tips. Whereas pin single mutants have root and meristem lengths close to those of the wild type, pin1,pin2 double and all triple and quadruple mutants containing pin2 display a more than additive reduction in root and meristem sizes (Blilou et al., 2005). Importantly, these defects can be rescued by exogenous auxin application, suggesting that basipetal auxin transport to the meristematic cells plays a critical role in meristem length regulation, predominantly through the control of cell division. In the elongation zone, the final cell size is reduced in several pin mutants, while the meristematic cell size corresponds to that of the wild type (Blilou et al., 2005). These data indicate that modulating auxin (re)distribution through PIN gene control can regulate both cell division and cell elongation in the root meristem (Fig. 1.2c). Exactly how robust this system of an efflux-driven auxin gradient is, has been demonstrated in a model that describes diffusion and PIN-facilitated

24
Root Development auxin transport in and across cells within a virtual root system (Grieneisen et al., 2007). In this model, a wide range of parameters has been tested for their effect on global auxin distribution, including membrane permeability, auxin production and decay, drastic alterations in influx and efflux (such as root cut and tissue ablation), and cell division and expansion. None of these in silico experiments influenced auxin distribution within the root; an observation that was supported by equivalent in planta experiments. However, when auxin reflux is prohibited by eliminating all laterally oriented PINs, the maximum quickly dissolves as auxin is directed upward and unable to feed back into the system, suggesting that the presence of the auxin maximum and gradient is solely the consequence of the global PIN topology within the root. Moreover, when cells are given the simple instructions to divide at high auxin levels and elongate at low concentrations, a self-organizing root system develops in time with similar zonation and auxin distribution properties as seen in wild-type roots. Thus, the graded auxin distribution, enforced by PIN-mediated transport, is sufficient to explain the maintenance and growth of the meristematic and elongation zone without the need for additional regulatory processes (Grieneisen et al., 2007, Fig. 1.2c). The explanatory power of this model can now be exploited to assess auxin transportregulated development and make precise predictions on the phenotypic outcome of specific genetic and cell biological manipulations regarding root development. 1.6.2

Dose-dependent regulation of root zonation

How the graded distribution of auxin dictates a range of distinct cellular behaviors is currently unknown, but experimental data suggest an important role for the PLT family of transcription factors. As mentioned above, a combination of four PLT genes are redundantly required for root development and stem cell maintenance. In the postembryonic root, PLT protein fusions display graded distribution along the meristem with maximum expression in the stem cell niche that extends into the meristematic zone and, for PLT2 and PLT3 fusions, into the elongation zone (Galinha et al., 2007; Fig. 1.2c). When produced in restricted domains, the PLT proteins can only compensate partially for the loss of plt1plt2 indicating that the PLT concentration gradient instructs a different output in different regions. Expression of PLT2 in the proximal part of the meristem, for example, increases meristem prolongation but fails to maintain the stem cell niche in plt1plt2 mutants. Steepening the slope of the PLT2 expression gradient in the plt1plt2 background rescues the stem cells, but root and meristem sizes are severely decreased when compared to the almost complete complementation when expressed from its full promoter. Moreover, inducible overexpression of PLT2 strengthens this hypothesis as it shifts the meristem boundary upward, promoting continuous growth of the meristematic zone and inhibiting cell expansion at the elongation zone (Galinha et al., 2007).

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Interestingly, the stem cell area in PLT2 overexpression plants is not altered, suggesting a limiting factor is constricting the high PLT2 dosage effects on the stem cell niche. This factor has been found to be RBR, mentioned above as a key component in root stem cell regulation independently from PLT (Wildwater et al., 2005). Overexpression of PLT2 in RBR RNAi background results in an expansion of dividing cells in the root cap area and ectopic stem cell-like divisions in the proximal meristem (Galinha et al., 2007). Together, these observations reveal that the PLT protein gradients define three outputs in the growing root system. High levels of PLT activity promote stem cell identity and maintenance and low levels mitotic activity of stem cell daughters; further reduction in levels is required for cell differentiation at the elongation zone (Galinha et al., 2007). Although a direct molecular link between auxin action and PLT gene activation is lacking, proper auxin distribution and response systems are essential for correct PLT gene transcription (Aida et al., 2004; Blilou et al., 2005), opening up the possibility that the PLT protein gradient dictates zonation of the root as a read-out of the stable auxin maximum and predicted associated auxin gradient (Grieneisen et al., 2007, Fig. 1.2c).

1.7 Meristem activation, root growth and cell division Reactivation of cell division in the root apical meristem at germination and onset of growth are essential for postembryonic development. Germination is triggered by water uptake by the quiescent dry seed and is generally considered to be complete when the radicle penetrates the seed coat. The earliest signs of germination are the resumption of metabolic processes, followed by directional cell expansion, and eventually the activation of cell division in the apical meristems (reviewed in Koornneef et al., 2002). Once activated, an essential part of the coordinated root growth is the integration of various informative signals into a univocal response often requiring extensive crosstalk between distinct hormone signaling pathways. 1.7.1

Redox homeostasis controls meristem activation

In the root meristemless1 (rml1) mutant, the embryonic root develops normally, but a failure to initiate cell division during germination results in seedlings with an extremely short root unable to establish and maintain an active meristematic zone of cell division (Cheng et al., 1995; Vernoux et al., 2000). RML1 has been found to be allelic to CADMIUM SENSITIVE2, encoding the first enzyme in the glutathione (GSH) biosynthesis pathway, ␥ -glutamylcysteine synthetase (Cobbett et al., 1998; Vernoux et al., 2000). Treatment of rml1 mutants with applied GSH is sufficient to restore postembryonic root development, indicating that the absence of cell division in the rml1 root results from GSH depletion (Vernoux et al., 2000). In wild-type plants, inhibiting

26
Root Development GSH biosynthesis has an effect on the mitotic root growth similar to that in rml1, while exogenous application increases the number of meristematic cells going through the mitotic cycle (S´anchez-Fern´andez et al., 1997; Vernoux et al., 2000). Accordingly, treatment of cultured tobacco (Nicotiana tabacum) cells with the GSH biosynthesis inhibitor, buthionine sulfoximine, traps the cells in G1 phase (Vernoux et al., 2000). A role for endogenous GSH in the control of cell proliferation is provided by the mapping of GSH levels in the root meristem. Low levels of GSH are associated with the mitotically inactive QC compared to the surrounding dividing stem cells (S´anchez-Fern´andez et al., 1997; Jiang et al., 2003). The glutathione redox couple GSH (reduced form) and GSSG (oxidized form) act as a homeostatic redox buffer (for a review, see Meyer and Hell, 2005). In recent years, it has become evident that the intracellular redox state plays a critical role in regulating cell proliferation, possibly by controlling the key components of the G1-to-S transition (den Boer and Murray, 2000; Jiang and Feldman, 2005 and references therein). Therefore, the impairment of GSH production in rml1 mutants leads presumably to an overall changed redox state in the root, arresting the cells in the G1/S phase (Vernoux et al., 2000). Exactly how redox homeostasis and cell cycle regulation interconnect at the molecular level is unclear. Interestingly, auxin has been linked to changes in intracellular redox state via its correlation with the production of reactive oxygen species. Possibly, auxin contributes to the quiescence of the QC cells by maintaining these cells in the G1/S phase through redox control (Jiang et al., 2003; Jiang and Feldman, 2005). At the moment, it is still debated whether cell cycle progression is activated before or after radicle protrusion, but early activated core cell cycle genes, such as CYCD, and the formation of a proper microtubule network certainly ˆ et al., 2005; play the key roles in regulating the extent of cell division (Barroco Masubelele et al., 2005). 1.7.2

Hormonal control of root growth and cell division

Low concentrations of brassinosteroids (BRs) have been shown to promote root growth, whereas high BR levels have a negative effect on root length. In accordance, BR-deficient mutants develop significantly shorter roots than ¨ wild-type plants (Mussig et al., 2003). Recent molecular genetic advances have identified a key component mediating the interaction between BR levels and auxin signaling in root growth. The natural loss-of-function allele brevis radix (brx) negatively affects cell division and cell elongation in the root tip (Mouchel et al., 2004). The brx phenotype results from a root-specific deficiency of BR, due to reduced expression of the rate-limiting enzyme in BR biosynthesis CONSTITUTIVE PHOTOMORPHOGENESIS AND DWARF (CPD) (Mouchel et al., 2006). In brx mutants, the auxin response is severely reduced, but can largely be restored by exogenous brassinolide application, suggesting that BR levels in the root influences auxin-induced transcriptional responses (Mouchel et al., 2006). Expression of BRX is strongly induced by

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auxin and repressed by BR, connecting the production of BR and auxin responses through a BRX-mediated feedback loop with root growth. Moreover, BR treatment has been shown to enhance polar auxin accumulation in the root (Li et al., 2005). The observation that modification of endogenous BR levels alter PIN transcript levels hint at a direct link between BR levels and polar auxin transport activities during developmental processes (Li et al., 2005). Furthermore, GA has been identified as a regulator of root growth, as demonstrated by the reduced primary root growth of GA-deficient seedlings (Fu and Harberd, 2003). The interaction between GA and auxin has been studied in detail, revealing at least two levels of regulation. In roots, auxin signaling has been shown to enhance the GA-induced degradation of the GRAS family member RGA, a repressor of GA signaling and root growth (Fu and Harberd, 2003). Additionally, auxins have been reported to regulate GA metabolism by direct upregulation of enzymes performing rate-limiting steps in GA biosynthesis (Frigerio et al., 2006). The inhibitory effect of ethylene on root growth is, at least partially, also mediated by RGA (Achard et al., 2003). Together, these results indicate that the protein RGA plays an important integrative role in the hormone response network for root growth. Another integrator of hormonal pathways and developmental processes in root growth is the 36-amino acid peptide POLARIS (PLS), linking auxin homeostasis to ethylene signaling (Casson et al., 2002; Chilley et al., 2006). Mutant pls seedlings have thick, short roots and exhibit an enhanced ethylene signaling phenotype known as the triple response. Moreover, acropetal transport of auxin is reduced in pls seedlings as well as the free auxin levels in the root tip. All these phenotypic defects could be suppressed by pharmacological or genetic inhibition of ethylene signaling, indicating that PLS-mediated ethylene signaling has a negative impact on auxin transport and accumulation. Transcription of PLS is rapidly upregulated by auxin and negatively by ethylene, suggesting a self-reinforcing mechanism in which auxin stimulates PLS activity to suppress the growth-inhibitory effects of ethylene signaling in the root tip (Chilley et al., 2006). In contrast, various studies have uncovered a positive correlation between ethylene responses and auxin biosynthesis. Three independent complementary studies of ethylene–auxin crosstalk in root growth indicate that the growth inhibitory effect of ethylene is in ˚ ziˇcka et al., 2007; Stepanova fact mediated predominantly through auxin (Ruˇ et al., 2007; Swarup et al., 2007). Synthesis of auxin in the root tip is enhanced in response to ethylene and distributed basipetally in a manner dependent on PIN2 and AUX1 to the cells of the elongation zone where it accumulates to induce local auxin responses that inhibit cell elongation and overall cell growth. Disruption of auxin biosynthesis by the loss-of-function mutants of the WEAK ETHYLENE INSENSITIVE2/ANTHRANILATE SYNTHASE a1 (WEI2/ASA1) and WEI7/ANTHRANILATE SYNTHASE b1 (ASB1) genes that encode subunits of a rate-limiting enzyme of the auxin biosynthesis, prevent the provoked auxin increase upon ethylene treatment, hence fully suppress the ethylene-mediated inhibition of root growth (Stepanova et al.,

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Root Development 2005). These seemingly contradictory data on ethylene–auxin crosstalk illustrate once more the complex order of crosstalk interactions between hormonal pathways (Chilley et al., 2006). Recently, ethylene signaling has been linked with promoting cell division of the QC (Ortega-Mart´ınez et al., 2007), although it remains unclear whether these effects are mediated by auxin. The enlarged root meristem of cytokinin-deficient plants indicates that cytokinins have a negative regulatory function in root growth (Werner et al., 2003). Rather than controlling cell division rates, cytokinins are thought to regulate the number of mitotic cells in the meristem (Beemster and Baskin, 2000; Werner et al., 2003). By selectively reducing endogenous cytokinin levels in the vascular tissue at the meristem transition zone (TZ), Dello Ioio et al. (2007) pinpointed this region as the site of cytokinin action to control the root meristem size. Reduction of cytokinin levels in other parts of the root meristem had no affect on root growth. Interestingly, proper auxin distribution in the meristem was shown to be necessary for mediating the effects of cytokinin on meristem size control (Dello Ioio et al., 2007). From these data, it is clear that hormones play an essential role in regulating root growth, although the molecular effectors are scarcely known. Where hormones account for the input from different developmental and environmental controls, it is the crosstalk between the various signaling pathways that fine-tunes decisions on cell division, meristem size and elongation.

1.8

Concluding remarks

A precondition for embryonic root initiation is the proper specification of the apical embryo domain, a process that is guided by auxin accumulation in the proembryo. In response to the increase in auxin levels, BDL-dependent action of MP noncell autonomously promotes specification of the hypophysis by enabling basally orientated auxin flow and, presumably, another unidentified signal. It would be interesting to see what the specific auxin response in the hypophysis includes. One possibility is that the changes in auxin response focus and maintain expression of the PLT root determinants. Computational modeling of auxin distribution in the root predicts an auxin maximum and highly stable gradient with morphogenic properties. How the auxin gradient is translated into defined cellular outputs is presently unclear, though the PLT genes are likely candidates to be involved. The functional concentration gradient of the PLT genes overlaps with the auxin gradient and requires the auxin response machinery. Analysis of downstream PLT targets will be needed to assess how much of the response to graded activity is due to additive concentration effects on the same targets and to differences in target specificity. The root stem cells are specified and maintained by the combinatorial action of PLT and SHR/SCR genes. Although these pathways act largely independently from one another, it is conceivable that they convey downstream a set of stem cell-promoting target genes. Most probably, target genes control

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subsets of the processes controlled by their activators, resulting in subtle phenotypes, as illustrated by the wox5 mutant acting downstream in the SHR/SCR pathway in which the lack of columella stem cells does not affect root growth and would, therefore, be easily missed in forward genetics screens. In addition, the identification of such target genes might be hampered by their genetic or functional redundancy, as shown by the lack of phenotypes in a knock-out analysis of QC-enriched genes (Nawy et al., 2005). The interesting findings that (1) the QC can signal over multiple layers when RBR levels are low and (2) WOX5 expression in the QC is required for root stem cell maintenance, point out that common mechanisms and genes operate in both root and shoot stem cell niches. These results might indicate that a common origin for the stem cell niche predates the shoot/root separation. It is becoming more and more clear that Arabidopsis root development is also controlled by microRNAs and chromatin remodeling factors; such factors have now been shown to regulate stem-cell identity and future research will have to focus on linking these regulation levels to the existing patterning genes. Plant hormones provoke many responses after application that could, until recently, not be explained at the molecular level. This situation has changed with the identification of the transport machinery, receptors and their interactors, response factors and other downstream effectors of the main classes of plant hormones. These discoveries will further help to understand the crosstalk between the different hormones in development. The understanding of Arabidopsis root development has taken a giant leap since the early nineties, when the root became appreciated as a model to study basic issues in developmental biology. Many of the fundamental questions from that era have now been answered: the clonal origin of the root and root morphology have been described in detail and major players in the genetic control of root initiation, stem cell maintenance and root growth have been identified. Now is the time to create links that connect the available data and generate interactive gene and protein networks. With the advent of genomics (see also Chapter 11) and related ‘omics’ approaches, a vast amount of data is generated that might help in this process.

Acknowledgments We are indebted to Viola Willemsen and Ben Scheres for valuable discussions and critical reading of the manuscript.

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¨ Muller, A., Guan, C., G¨alweiler, L., T¨anzler, P., Huijser, P., Marchant, A. et al. (1998). AtPIN2 defines a locus of Arabidopsis for root gravitropism control. EMBO J. 17, 6903–6911. ¨ Mussig, C., Shin, G.-H. and Altmann, T. (2003). Brassinosteroids promote root growth in Arabidopsis. Plant Physiol. 133, 1261–1271. Nakajima, K., Sena, G., Nawy, T. and Benfey, P.N. (2001). Intercellular movement of the putative transcription factor SHR in root patterning. Nature 413, 307–311. Nawy, T., Lee, J.-Y., Colinas, J., Wang, J.Y., Thongrod, S.C., Malamy, J.E. et al. (2005). Transcriptional profile of the Arabidopsis root quiescent center. Plant Cell 17, 1908–1925. Nodine, M.D., Yadegari, R. and Tax, F.E. (2007). RPK1 and TOAD2 are two receptorlike kinases redundantly required for Arabidopsis embryonic pattern formation. Dev. Cell 12, 943–956. Ogawa, M., Shinohara, H., Sakagami, Y. and Matsubayashi, Y. (2008). Arabidopsis CLV3 peptide directly binds CLV1 ectodomain. Science 319, 294–294 [Err. Science 319, 901]. Okushima, Y., Overvoorde, P.J., Arima, K., Alonso, J.M., Chan, A., Chang, C. et al. (2005). Functional genomic analysis of the AUXIN RESPONSE FACTOR gene family members in Arabidopsis thaliana: unique and overlapping functions of ARF7 and ARF19. Plant Cell 17, 444–463. Ortega-Mart´ınez, O., Pernas, M., Carol, R.J. and Dolan, L. (2007). Ethylene modulates stem cell division in the Arabidopsis thaliana root. Science 317, 507–510. Ottenschl¨ager, I., Wolff, P., Wolverton, C., Bhalerao, R.P., Sandberg, G., Ishikawa, H. et al. (2003). Gravity-regulated differential auxin transport from columella to lateral root cap cells. Proc. Natl. Acad. Sci. U. S. A. 100, 2987–2991. Paponov, I.A., Teale, W.D., Trebar, M., Blilou, I. and Palme, K. (2005). The PIN auxin efflux facilitators: evolutionary and functional perspectives. Trends Plant Sci. 10, 170–177. Paquette, A.J. and Benfey, P.N. (2005). Maturation of the ground tissue of the root is regulated by gibberellin and SCARECROW and requires SHORT-ROOT. Plant Physiol. 138, 636–640. P´erez-P´erez, J.M., Serralbo, O., Vanstraelen, M., Gonz´alez, C., Criqui, M.-C., Genschik, P. et al. (2008). Specialization of CDC27 function in the Arabidopsis thaliana anaphasepromoting complex (APC/C). Plant J. 53, 78–89. Quint, M. and Gray, W.M. (2006). Auxin signaling. Curr. Opin. Plant Biol. 9, 448– 453. ˚ ziˇcka, K., Ljung, K., Vanneste, S., Podhorsk´a, R., Beeckman, T., Friml, J. et al. (2007). Ruˇ Ethylene regulates root growth through effect on auxin biosynthesis and transportdependent auxin distribution. Plant Cell 19, 2197–2212. Sabatini, S., Beis, D., Wolkenfelt, H., Murfett, J., Guilfoyle, T., Malamy, J. et al. (1999). An auxin-dependent distal organizer of pattern and polarity in the Arabidopsis root. Cell 99, 463–472. Sabatini, S., Heidstra, R., Wildwater, M. and Scheres, B. (2003). SCARECROW is involved in positioning the stem cell niche in the Arabidopsis meristem. Genes Dev. 17, 354–358. S´anchez-Fern´andez, R., Fricker, M., Corben, L.B., White, N.S., Sheard, N., Leaver, C.J. et al. (1997). Cell proliferation and hair tip growth in the Arabidopsis root are under mechanistically different forms of redox control. Proc. Natl. Acad. Sci. U. S. A. 94, 2745–2750.

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Root Development Sarkar, A.K., Luijten, M., Miyashima, S., Lenhard, M., Hashimoto, T., Nakajima, K. et al. (2007). Conserved factors regulate signalling in Arabidopsis thaliana shoot and root stem cell organizers. Nature 446, 811–814. Scheres, B., Di Laurenzio, L., Willemsen, V., Hauser, M.-T., Janmaat, K., Weisbeek, P. et al. (1995). Mutations affecting the radial organisation of the Arabidopsis root display specific defects throughout the embryonic axis. Development 121, 53–62. Scheres, B., Wolkenfelt, H., Willemsen, V., Terlouw, M., Lawson, E., Dean, C. et al. (1994). Embryonic origin of the Arabidopsis primary root and root meristem initials. Development 120, 2475–2487. ¨ Schoof, H., Lenhard, M., Haecker, A., Mayer, K.F.X., Jurgens, G. and Laux, T. (2000). The stem cell population of Arabidopsis shoot meristem is maintained by a regulatory loop between the CLAVATA and WUSCHEL genes. Cell 100, 635–644. Sena, G., Jung, J.W. and Benfey, P.N. (2004). A broad competence to respond to SHORT ROOT revealed by tissue-specific ectopic expression. Development 131, 2817– 2826. Serralbo, O., P´erez-P´erez, J.M., Heidstra, R. and Scheres, B. (2006). Non-cellautonomous rescue of anaphase-promoting complex function revealed by mosaic analysis of HOBBIT, an Arabidopsis CDC27 homolog. Proc. Natl. Acad. Sci. U. S. A. 103, 13250–13255. Srivastava, L. (2002). Plant Growth and Development: Hormones and Environment. New York, Academic Press. Steinmann, T., Geldner, N., Grebe, M., Mangold, S., Jackson, C.L., Paris, S. et al. (1999). Coordinated polar localization of auxin efflux carrier PIN1 by GNOM ARF GEF. Science 286, 316–318. Stepanova, A.N., Hoyt, J.M., Hamilton, A.A. and Alonso, J.M. (2005). A link between ethylene and auxin uncovered by the characterization of two root-specific ethyleneinsensitive mutants in Arabidopsis. Plant Cell 17, 2239–2242. Stepanova, A.N., Yun, J., Likhacheva, A.V. and Alonso, J.M. (2007). Multilevel interactions between ethylene and auxin in Arabidopsis roots. Plant Cell 19, 2169– 2185. Suzuki, T., Inagaki, S., Nakajima, S., Akashi, T., Ohto, M.-A., Kobayashi, M. et al. (2004). A novel Arabidopsis gene TONSOKU is required for proper cell arrangement in root and shoot apical meristems. Plant J. 38, 673–684. Suzuki, T., Nakajima, S., Inagaki, S., Hirano-Nakakita, M., Matsuoka, K., Demura, T. et al. (2005). TONSOKU is expressed in S phase of the cell cycle and its defect delays cell cycle progression in Arabidopsis. Plant Cell Physiol. 46, 736–742. Swarup, R., Perry, P., Hagenbeek, D., Van Der Straeten, D., Beemster, G.T.S., Sandberg, G. et al. (2007). Ethylene up regulates auxin biosynthesis in Arabidopsis seedlings to enhance inhibition of root cell elongation. Plant Cell 19, 2186–2196. Szemenyei, H., Hannon, M. and Long, J.A. (2008). TOPLESS mediates auxindependent transcriptional repression during Arabidopsis embryogenesis. Science 319, 1384–1386. Takeda, S., Tadele, Z., Hofmann, I., Probst, A.V., Angelis, K.J., Kaya, H. et al. (2004). BRU1, a novel link between responses to DNA damage and epigenetic gene silencing in Arabidopsis. Genes Dev. 18, 782–793. Tiwari, S.B., Hagen, G. and Guilfoyle, T.J. (2004). Aux/IAA proteins contain a potent transcriptional repression domain. Plant Cell 16, 533–543. Ueda, M., Matsui, K., Ishiguro, S., Sano, R., Wada, T., Paponov, I. et al. (2004). The HALTED ROOT gene encoding the 26S proteasome subunit RPT2a is essential for the maintenance of Arabidopsis meristems. Development 131, 2101–2111.

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Root Development ˚ ziˇcka, K., Blilou, I. et al. (2006). Wi´sniewska, J., Xu, J., Seifertov´a, D., Brewer, P.B., Ruˇ Polar PIN localization directs auxin flow in plants. Science 312, 883–883. Wu, X., Chory, J. and Weigel, D. (2007). Combinations of WOX activities regulate tissue proliferation during Arabidopsis embryonic development. Dev. Biol. 309, 306–316. Wysocka-Diller, J.W., Helariutta, Y., Fukaki, H., Malamy, J.E. and Benfey, P.N. (2000). Molecular analysis of SCARECROW function reveals a radial patterning mechanism common to root and shoot. Development 127, 595–603. Xu, J., Hofhuis, H., Heidstra, R., Sauer, M., Friml, J. and Scheres, B. (2006). A molecular framework for plant regeneration. Science 311, 385–388. Yeung, E.C. and Meinke, D.W. (1993). Embryogenesis in angiosperms: development of the suspensor. Plant Cell 5, 1371–1381. Zhu, Y., Dong, A., Meyer, D., Pichon, O., Renou, J.-P., Cao, K. et al. (2006). Arabidopsis NRP1 and NRP2 encode histone chaperones and are required for maintaining postembryonic root growth. Plant Cell 18, 2879–2892.

Annual Plant Reviews (2009) 37, 39–63 doi: 10.1002/9781444310023.ch2

www.interscience.wiley.com

Chapter 2

VASCULAR MORPHOGENESIS DURING ROOT DEVELOPMENT Ana Campilho,∗ Ove Lindgren∗ and Yk¨a Helariutta Institute of Biotechnology, University of Helsinki, P.O. Box 65 (Viikinkaari 1), FIN00014 Helsinki, Finland

Abstract: The root vascular cylinder consists of two conductive tissues, phloem and xylem. The vascular pattern, the way these two tissues are arranged, is often dynamic during ontogeny. In this chapter, we will first describe the current knowledge on how the vascular pattern is established anatomically. Then we review the most recent advances on understanding the gene functions behind the patterning, cell proliferation and differentiation related to the establishment of the vascular tissue types. It seems that during these processes the hormonal and genetic controls are tightly integrated. Finally, we discuss the power of some genome-wide approaches for further understanding of the integrated view on vascular development. Keywords: vascular tissue development; phloem; xylem; pattern development; vascular genomics

2.1 Introduction Vascular tissues evolved to provide long-distance transport of water and nutrients, and to provide rigidity to the plant body. They enabled early vascular plants to colonize the land. The vascular system of plants is composed of continuous strands, called the vascular bundles. These structures extend through the entire plant, functionally connecting every part of the shoot with the root system (Fig. 2.1). Vascular bundles typically consist of two conductive tissues, the phloem and xylem (Esau, 1977). Phloem is the main path for distribution of photosynthetic products, whereas xylem is the main transporting tissue for water and nutrients. Both xylem and phloem comprise a number of specialized vascular cell types, including conducting elements (tracheary and sieve elements ∗

These authors contributed equally.

39

40
Root Development

(a)

(b)

prim. xylem sec. xylem: cell expansion

(e)

procambium/cambium sec. phloem

(c)

prim. phloem

(d)

Figure 2.1 Organization of the primary and secondary vascular tissues in Arabidopsis. (a) Schematic representation of the whole plant in a longitudinal view. (b) Transverse section of the stem close to the shoot apex. (c) Transverse section of a leaf. (d) Transverse section of the root close to the root tip. (e) Transverse section of the basal region of the inflorescence stem during the secondary phase of vascular development. (Modified from Nieminen et al., 2004; Copyright American Society of Plant Biologists.)

in xylem and phloem, respectively), parenchyma and sclerenchyma cells. A characteristic feature of the cellular pattern of xylem and phloem is that they are quite diverse, depending on whether they occur in the stem, leaf or root. Another feature is the dynamic nature of the vascular patterns. Developmentally, it is convenient to distinguish primary and secondary vascular tissues (Fig. 2.1): the former originate from procambium, a meristematic tissue in apical meristems and young organ primordia and the latter are formed in many plant taxa when plants start to increase their girth as a result of the activity of a lateral meristem – the vascular cambium. Secondary xylem and phloem are produced toward the inside and outside of the organ, respectively. Secondary development is particularly pronounced in trees with secondary xylem (also known as wood).

2.2

The Arabidopsis root vascular system

In this chapter, we will focus on the Arabidopsis root, which is currently the only relatively well-understood system both at a cellular and a molecular level. 2.2.1

Embryonic origin

For a description of the cellular organization of the Arabidopsis root during embryogenesis, see Chapter 1. Nevertheless, we will briefly describe the main

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Figure 2.2 Development of the radial vascular pattern at successive embryonic stages in schematic longitudinal and transverse views highlighting the vascular cylinder. (a) Early globular stage embryo in which the progenitors of the major tissue types are distinguished, the protoderm (pr), the ground meristem (gm) and in gray the procambium (pc). (b) Late globular stage embryo. Notice the periclinal cell divisions generating the pericycle in dark gray (p) and the vascular primordium in lighter gray (vp). (c) Early heart stage embryo. Notice the anticlinal cell division in transverse view of the pericycle in dark gray (p); the vascular primordium (vp) is in lighter gray. (d) Late heart stage embryo in which procambial cell divisions occur to establish the number of vascular stem cells (in lighter colors); pericycle in dark gray and procambium in lighter gray. (Adapted from Scheres et al., 1994; modified with permission of The Company of Biologists.)

events of patterning of the vascular tissues in root and hypocotyl during embryogenesis. First, procambial tissue is established as the most internal radial domain in the lower tier of the embryo. This event occurs at early globular stage when protoderm, ground meristem and procambium become separated by regular periclinal divisions to form the major tissue types in the embryonic axis (Fig. 2.2a) (Scheres et al., 1995). At late globular to early triangular stage, the four procambium cells have performed a periclinal division to generate the future pericycle and the vascular primordium (Fig. 2.2b). At early heart stage, the number of cells in the pericycle increases by anticlinal divisions, reaching the number of pericycle stem cells counted in the seedling root (Fig. 2.2c) (Scheres et al., 1995). From late heart stage to mature embryo, additional irregular procambial cell divisions occur to establish the final number of vascular stem cells present in the seedling root meristem (approximately 11 ¨ vascular stem cells) (Fig. 2.2d) (Scheres et al., 1995; M¨ahonen et al., 2000). It is worth mentioning that vascular pattern in the postembryonic root and hypocotyl has many similarities. Actually, the separation between root and

42
Root Development hypocotyl occurs late during embryogenesis and displays some variability: cells around the border can be incorporated either into the root or into the hypocotyl (Scheres et al., 1994). However, the vascular tissue of the hypocotyl is formed from the preexisting procambial cells, whereas the root vascular tissues, with the exception of a few layers just below the root/hypocotyl junction are derived from the continuous postembryonic activity of the root apical meristem. In the mature embryo, a continuous procambial system extends in an apical-to-basal axis without features of differentiated vascular elements (Dharmawardhana et al., 1992; Busse and Evert, 1999). Furthermore, at least the cellular pattern of phloem is not entirely completed during embryogenesis (Bonke et al., 2003), but vascular tissue-specific markers have been reported to be expressed within the prospective vascular tissue types (Moussian et al., ¨ 1998; Bonke et al., 2003; M¨ahonen et al., 2006a; Bauby et al., 2007) suggesting vascular cell fate decisions occur during embryogenesis. In the first days of germination, the primary vascular pattern emerges, while phloem and xylem differentiate within the procambial tissue. 2.2.2

Primary vascular tissue organization

The vascular cylinder of the Arabidopsis root is derived from the activity of the root apical meristem. The vascular cylinder comprises the vascular tissues and one more layer of nonvascular cells, the pericycle (Esau, 1977). The primary vascular pattern of the Arabidopsis root exhibits a diarch pattern of organization, i.e. it consists of a xylem axis that extends to the pericycle ¨ and alternates with two phloem poles (Fig. 2.3) (Dolan et al., 1993; M¨ahonen et al., 2000; Baum et al., 2002). Three to five metaxylem cell files are located at the center of the axis between two protoxylem cell files. The protoxylem differentiates in parts of the plant body that have not completed their growth and differentiation. The metaxylem is commonly initiated in the still growing primary plant body, but it matures largely after the elongation is completed (Esau, 1977; Steeves and Sussex, 1989). Furthermore, the central cell file domain of the metaxylem differentiates slightly later than the cell files touching protoxylem (Kubo et al., 2005). Generally, protoxylem and metaxylem differ in types of secondary wall thickenings of their conductive cells, the tracheary elements (TEs). In the protoxylem, annular and helical thickenings predominate and in the metaxylem helical, scalariform and pitted are usually present (Esau, 1977). The primary phloem is classified into protophloem and metaphloem on the same basis as the primary xylem (Esau, 1977). Thus, the two phloem poles contain protophloem and a metaphloem cell file associated with companion cells (Fig. 2.3) (Dolan et al., 1993). Between the xylem axis and two phloem poles there are intervening procambial cell files. The root vascular tissue develops acropetally and, consequently, the earliest stages of root vascular development are always closest to the root tip. Cells

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Vascular cells Protoxylem Metaxylem Protophloem Metaphloem Companion cells Pericycle Intervening procambium Vascular stem cells Other Quiescent center Outer layers Root cap

Figure 2.3 Schematic representation of the different tissues within the vascular cylinder of the Arabidopsis primary root. Xylem cell files can be traced down to the quiescent center, whereas phloem cell files and a proportion on intervening procambial cell files originate as a result of asymmetric cell division (arrows). All vascular cell types are originated from the vascular stem cells just above the quiescent center. A proportion of the procambial cell files remain undifferentiated between xylem and phloem to form the intervening procambium, which is later activated to form the vascular cambium. (Modified from M¨ah¨onen et al., 2000; with permission from Cold Spring Harbor Laboratory Press.)

that form the vascular tissues originate from vascular stem cells above the quiescent center (QC). Stem cells divide for self-renewal and production of daughter cells that will differentiate. Thus, vascular stem cells divide to generate daughter cells that remain in contact with the QC, thus maintaining the stem cell status. The other daughters, after proliferative cell divisions, will differentiate into the different vascular tissue types, xylem, phloem and the intervening pluripotent procambial tissue. In conclusion, vascular tissues in the Arabidopsis root exhibit a simple anatomical structure: a xylem axis with two intercalating phloem poles and intervening procambial cells as observed on a transverse section through the root. In a longitudinal view, xylem cell files can be traced back to a vascular stem cell in contact with the QC but phloem cell files are established later as a result of asymmetric cell divisions that separate phloem from the intervening procambial cell files (Fig. 2.3) ¨ (M¨ahonen et al., 2000). 2.2.3

Secondary vascular tissue organization

The Arabidopsis root can undergo a secondary stage of development (Dolan et al., 1993; Dolan and Roberts, 1995; Baum et al., 2002; Chaffey et al., 2002).

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Root Development

Figure 2.4 Primary Arabidopsis root initiating the secondary growth. Transverse section taken 1.5 cm proximal from the root tip from a 4-week-old primary root illustrating the early divisions of the procambial cells (arrowheads) located between the primary xylem axis and phloem poles leading to the vascular cambium formation. Px, protoxylem; Mx, metaxylem. (From Baum et al., 2002; with permission from the American Journal of Botany.)

The production of secondary vascular tissues depends on the activity of the vascular cambium. This lateral meristem originates from divisions of the procambial cells located between the primary xylem axis and phloem poles (Fig. 2.4). A continuous cylinder of narrow cambial cells with thin cell walls is completely formed when pericycle cells positioned next to the protoxylem cell files also undergo division. Repeated cell divisions in the vascular cambium produce cell files that differentiate into secondary xylem inward the root and secondary phloem outward the root. The process of secondary thickening in the Arabidopsis root is correlated with changes in cell surface epitopes (Dolan and Roberts, 1995). In conclusion, Arabidopsis can be considered as a useful genetic model to study vascular development both in primary and secondary phases of development.

2.3

Molecular genetics of the stele: a rapidly developing field

Next, we will focus on the recent understanding of vascular morphogenesis of the root. Three distinct major questions are addressed. What specifies vascular tissue in the central domain of the root? How are the various vascular cell types specified? What controls the extent of cell proliferation during the various processes of vascular morphogenesis? 2.3.1

Specification of the vascular bundle

The molecular control that establishes vascular bundles and their continuity has mainly been studied in the aerial parts of the plant; especially, vein

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Suspensor

Apical

PIN1 Hypophysis

PIN4 PIN7 Auxin

Basal (a)

(b) Two-cell stage

Globular stage

Triangular stage

Figure 2.5 The canalization model and PIN-mediated auxin accumulation during embryogenesis. (a) The canalization model. Initially random signals getting progressively restricted to a narrow channel by a self-regulatory enhancement of certain cells to propagate the signal. (Reprinted from Scheres and Berleth, 1998; with permission from Elsevier.) (b) Expression patterns of PIN1, PIN4 and PIN7 during Arabidopsis embryo development and the resulting areas of auxin accumulation. Notice how the direction of the PAT switches from apical to mainly basal at the globular stage and the elongated procambial cells at the triangular stage (early heart stage). (From Tanaka et al., 2006; with permission from Birkhauser Verlag, Switzerland.)

patterning in the leaf has been analyzed extensively. Before focusing on this process during root development, it is therefore appropriate to describe conceptually the shoot-based work. To explain vein patterning in the leaf, two major theories have been put forward. The reaction–diffusion theory states that vascular strands are specified locally by a system based on an autocatalytic activator that triggers a more quickly diffusing inhibitor that keeps a new activator at a distance (Meinhardt, 1982). Modeling shows that this system can explain the occurrence of both closed and open veins. On the other hand, the canalization theory states that veins arise from a continuous signal flow that becomes gradually more restricted to regions of future vascular differentiation (Fig. 2.5a) (Sachs, 1981; Scheres and Xu, 2006). The role of auxin transport in establishing early vascular or procambial identity has recently gained strong support. So-called pin-formed (pin) mutants (Goto et al., 1987) typically show several developmental abnormalities, such as fused cotyledons and pin-formed inflorescences with no floral buds. The pin1-1 mutant was found to have a decreased ability to transport auxin, and polar auxin transport (PAT) inhibitors have been found to induce similar phenotypes (Okada et al., 1991). By applying PAT inhibitors on developing embryos of Brassica juncea, it was discovered that the developing embryos also attained a pin1-1-like phenotype (Liu et al., 1993). Cloning of the first of a total of eight members in the Arabidopsis genome, PIN1, revealed that it was similar to bacterial and eukaryotic carrier proteins (G¨alweiler et al., 1998).

46
Root Development In accordance to the proposed role in auxin transport, the PIN1 protein was found, through immunolocalization with antibodies, to be polarly distributed in cells and localized to the areas of the cell membrane that corresponds to the direction of auxin transport, as expected from a putative efflux carrier. Recently, protein:GFP translational construct of PIN1 was used in a study on leaf vascular patterning in which the GFP expression was followed during the development of veins (Scarpella et al., 2006). The expression pattern was closely monitored and found to predict the future leaf vascular strands earlier than other known markers. Cells in the newly formed veins developed a polar PIN1 expression pattern in the cells that was directed toward older veins. Especially intriguing was the discovery that higher-order connected strands have a composite origin in which first the lower strand is created from an epidermal so-called convergence point only followed later by the upper strand. Next, the strands are joined in a closed loop by a single cell that exhibits a bipolar PIN1 expression pattern. These results are supportive of the canalization theory and strongly suggest a defining role for auxin in the early formation of leaf vascular strands. Although it may be extrapolated that auxin might establish the procambial cell identity also during root development, currently relatively little evidence is available. For instance, several of the PIN putative auxin efflux carriers have been shown to be expressed in the developing Arabidopsis embryo with varying contributions from individual PIN proteins, depending on the developmental stage (Friml et al., 2003). The synthetic promoter element DR5 that reliably responds to auxin can be used to indirectly monitor auxin gradients in Arabidopsis (Sabatini et al., 1999). By using a GFP construct under the control of this promoter, auxin gradients in the developing Arabidopsis embryo could be measured and the direction of the auxin flow actually switched from apical to basal after the 32-cell stage, leading to an accumulation in the hypophysis (Fig. 2.5b) (Friml et al., 2003). Simultaneously with this switch in the auxin gradient, PIN7 also changed polarity from apical to basal while PIN1 and PIN4 were also expressed and localized basally. Quadruple mutants of pin1, pin3, pin4 and pin7 had embryos that often failed to establish an apical–basal polarity without or with nonfunctional roots. Another piece of evidence for the role of auxin in the establishment of the root vascular bundle comes from studies on auxin signal transduction. The most studied mechanism consists of an intricate regulatory network of molecular components and acts through two different transcriptional regulators, the AUX/IAAs (Gray et al., 2001) and the AUXIN RESPONSE FACTORS (ARFs) (Ulmasov et al., 1999). AUX/IAAs are a group of short-lived auxin-response proteins that are able to repress ARFs. Recently, TRANSPORT INHIBITOR RESPONSE1 (TIR1) was found to be an F-box protein that is able to bind auxin (Dharmasiri et al., 2005; Kepinski and Leyser, 2005) and is part of a ubiquitination complex that can target AUX/IAAs for degradation and thereby releasing the ARFs for transcriptional activation. Auxin signaling has been implicated in vein development in leaves and plants mutated in

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MONOPTEROS (MP) (Mayer et al., 1991) that was found to encode an ARF (Hardtke and Berleth, 1998). mp mutants display a simplified leaf venation and lack the basal structures of the primary root, including the stele (Fig. 2.6c) ¨ (Berleth and Jurgens, 1993), whereas severe mutants often lack the primary root completely. During Arabidopsis embryo development, the typical elongated procambial cells in the middle of the embryo are absent in mp mutants at the early heart stage. It is interesting that the previously described embryonic apical–basal switch in auxin accumulation (Friml et al., 2003) precedes the development of these elongated procambial cells, further suggesting a role for auxin in the establishment of the root vascular cylinder. The existence of disjointed open veins has been used in support of the reaction diffusion model. In a mutant screen, the venation pattern was analyzed in EMS-mutagenized Arabidopsis M3 lines (Koizumi et al., 2000). This screen resulted in six unique, recessive–lethal mutant lines designated vascular network defective (van1 to van6). All of these mutants were affected in secondary and tertiary leaf venation and displayed fractured vein architectures of varying severity, supporting the reaction–diffusion model. Interestingly, all the van mutants also suffered from impaired root growth when compared to wild-type seedlings and their xylem vessel differentiation took place unusually close to the root tip. However, only van1 showed interrupted xylem cell files in the primary root. Cross sections of the root and the hypocotyl revealed disorganized vascular cylinders and in most cases phloem was at least partially absent. Again, the most severe phenotype was found in VAN1, where the vascular bundle of the root and hypocotyl was highly disorganized and entirely lacked phloem poles. Of the six VAN-genes, VAN3 has been cloned and characterized (Koizumi et al., 2005) and has been identified as an adenosine diphosphate (ADP)-ribosylation factor-guanosine triphosphatase (GTPase)-activating protein (ARF-GAP) implicated in vesicle transport related to auxin signal transduction. In support to this idea, EMB30/GN (GNOM) has been shown to encode a guanine nucleotide exchange factor (GEF) on adenosine diphosphate (ADP)-ribosylation factor-GTPase (ARFGEF), which is responsible for the targeted recycling of the PIN1 putative auxin efflux carrier (Shevell et al., 1994; Busch et al., 1996; Steinmann et al., 1999; Geldner et al., 2003). Inhibitors of auxin transport have also been discovered to interfere with PIN1 cycling between the plasma membrane and endosomal compartments and especially the vesicle-trafficking inhibitor brefeldin A gives rise to a similar phenotype, further highlighting the importance of vesicle cycling in PAT (Geldner et al., 2001). However, in the light of the canalization model, Scarpella et al. (2006) also found the same initial pattern of PIN1 expression in the van3 mutant background as in wild type and concluded that the later disjointed veins found in this mutant could be explained by differentiation defects taking place afterward. Thus, even in a strongly discontinuous venation mutant, used as evidence for the reaction–diffusion hypothesis, there is seemingly an initially continuous pattern that would fit with the auxin canalization model. It remains to be seen whether the

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Figure 2.6 Root vascular phenotypes in the mutants apl and monopteros and plants overexpressing VND6 and VND7. (a) Close-up of stele cross sections from wild type (left) and apl mutant (right). Arrows mark the xylem axis. Note the difference in phloem pole differentiation and the ectopic xylem (black star). (Reproduced from Bonke et al., 2003; with permission from Nature.) (b) Confocal microscope images of primary roots from wild type (left) and apl mutant (right), stained with propidium iodide. Note the ectopic formation of xylem in the root of the apl mutant (white star). (Reproduced from Bonke et al., 2003; with permission from Nature.) (c) Cross sections of hypocotyl showing the difference in radial differentiation between wild type (left) and monopteros ‘basal peg’ mutant (right). Note the complete absence of any discernible stele structures in the mutant. (Reproduced from Berleth and J¨urgens, 1993; with permission from Development.) (d) Longitudinal confocal laser scanning microscope images of Arabidopsis roots from wild type, plants transformed with 35S::VND6 and 35S::VND7, respectively. PX, protoxylem; MX, metaxylem. (From Kubo et al., 2005; with permission from Cold Spring Harbor Laboratory Press.)

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mutations in the other VAN loci might also be explained by the auxin canalization model or whether the two mechanisms could indeed somehow operate in parallel. By using a system to induce protoplasts of Zinnia to differentiate into xylem vessels (see Section 2.3.2), a novel proteoglycan important for xylem strand continuity was discovered (Motose et al., 2004). Developing TEs in cell cultures had a tendency to drag neighboring cells into the TE differentiation pathway, suggesting the existence of a diffusible signal. This factor was purified using affinity chromatography from the arabinogalactan protein fraction of the Zinnia cell growth medium. The polypeptide backbone from deglycosylated xylogen was sequenced and used to generate the corresponding cDNA of this novel factor. Based on sequence similarities, two homologous genes were found in the Arabidopsis genome. When these genes, designated ARABIDOPSIS THALIANA XYLOGEN-PRODUCING, AtXYP1 and AtXYP2, were overexpressed under the control of the CaMV 35S promoter in cultured tobacco cells, xylogen accumulated in the growth medium. Single mutants of these genes had no phenotype, but double knockout mutants displayed distinct morphological defects. The venation pattern of rosette leaves was simplified with a markedly reduced number of tertiary veins and disconnected xylem cell files in the primary root of the double mutants. However, the xylem of this mutant is neither entirely disorganized, nor absent and a role for xylogen has been proposed as a coordinator that guarantees the integrity of the vascular network (Motose et al., 2004). In support of this idea, xylogen was found to have a polar localization in differentiating TEs, accumulating on the apical side of the cell walls suggesting that xylogen is excreted to act on neighboring cells directionally. By a function in the cell–cell signaling of developing TEs, the continuity of xylem strands would be maintained. 2.3.2

Root vascular cell identity and cytokinin

Cells in the root vascular cylinder acquire different identities in an ordered spatial arrangement. How is this differentiation achieved? Recent studies show that cytokinin signaling plays an important role in the regulation of cell fate during vascular root development. The wooden leg (wol) mutant was initially identified as a short-root mutant (Scheres et al., 1995). During late stages of embryogenesis, the wol mutant omits proliferative procambial cell divisions resulting in a reduced number of vascular cell initials. Postembryonically, the periclinal procambial cell divisions required for proliferation of vascular cell files are absent as well. The absence of these formative cell divisions results in a reduced number of cell files within the vascular bundle and is associated with specification of all vascular cell files in the root as pro¨ toxylem (Fig. 2.7b) (Scheres et al., 1995; M¨ahonen et al., 2000). WOL encodes a two-component signal transducer and is allelic to CYTOKININ RECEP¨ TOR1 (CRE1)/AHK4 (M¨ahonen et al., 2000; Inoue et al., 2001; Suzuki et al., 2001). CRE1 makes part of a phosphor-relay circuit where it binds directly

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Figure 2.7 Cytokinin signaling during vascular development in the Arabidopsis root. (a) Cytokinin signal transduction pathway. Cytokinin binding initiates autophosphorylation of the three cytokinin receptors, followed transfer of the phosphoryl group (P) to a histidine phosphotransfer protein (AHP). Phosphorylated AHP then translocates to the nucleus and transfers the P to a type A or type B response regulators (ARR) which leads to a cytokinin response. (From Kakimoto, 2003; reprinted with permission from the Annual Review of Plant Biology, Volume 54 © 2003 by Annual Reviews www.annualreviews.org) (b) Schematic representation for transverse sections of the primary root tip: the number of protoxylem files correlate with the level of cytokinin signaling. (Reprinted courtesy of A.P. M¨ah¨onen.) (c) Negative feedback loop model showing the reciprocal interaction of cytokinin signaling and AHP6 in controlling the balance between the maintenance of procambial cell identity (PC) and the differentiation of protoxylem elements (PX). (From M¨ah¨onen et al., 2006a; reprinted with permission from American Association for the Advancement of Science.)

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to cytokinin and initiates a cascade of phosphotransfer events, through the activity of the histidine phosphor (HP) transmitters and culminating in the activation of nuclear localized response regulators (RRs) (Fig. 2.7a) (reviewed in Hwang et al., 2002). WOL/CRE1 is expressed early in the four innermost cells of the globular-stage embryo, which are the precursors of vascular tissues and from the heart stage onward, it is expressed in the procambium of the embryonic root, hypocotyls and cotyledon shoulders. Postembryonically, ¨ it is expressed in the vascular cylinder and pericycle (M¨ahonen et al., 2000). Moreover, combining the triple knockout mutant for all three genes encoding CRE-family receptors (CRE1/WOL/AHK4, AHK2 and AHK3) (Higuchi et al., 2004; Nishimura et al., 2004) or depleting cytokinins specifically in the vascular cylinder (expressing a CYTOKININ OXIDASE under the CRE1 promoter) results in a reduced number of cell files from which all differentiate into protoxylem resembling the wol mutant phenotype (Fig. 2.7b). Thus, cytokinin signaling is necessary for inhibition of protoxylem differentiation and for normal proliferation and differentiation of procambial cells. In contrast, increased cytokinin signaling in ahp6 mutants (see below) or in plants supplied with high levels of exogenous cytokinin eliminates completely protoxylem identity during root development (Fig. 2.7b). These findings demonstrate that cytokinin signaling is required to promote and maintain cell identities other than protoxylem within the vascular cylinder and that protoxylem is a pro¨ cambial ‘default’ cell identity in the absence of cytokinin signaling (M¨ahonen et al., 2006a). ahp6 mutants were identified through a suppressor screen for wol. In the double ahp6 wol mutant vascular bundle, other vascular cell fates besides protoxylem can be observed. In the ahp6 mutant, protoxylem forms erratically (Fig. 2.7b) and the expression of cytokinin signaling reporter genes (the response regulator, ARR15) shows that cytokinin signaling is active at the position where protoxylem would normally differentiate. AHP6 is a pseudophosphotransfer protein that, unlike other phosphotransfer proteins, lacks the conserved histidine residue in the phosphotransfer domain and inhibits phosphotransfer in vitro experiments. Thus, AHP6 acts as an inhibitor of cy¨ tokinin signaling by interacting with the phosphorelay machinery (M¨ahonen et al., 2006a). AHP6 is expressed in the protoxylem files and two adjacent pericycle cell files. Based on the molecular genetic analysis of AHP6, it was concluded that during protoxylem development AHP6 counteracts cytokinin signaling, thereby allowing cells to differentiate with protoxylem identity ¨ (M¨ahonen et al., 2006a). Furthermore, cytokinin signaling appears to negatively regulate the spatial domain of AHP6 expression, as indicated by broader AHP6 expression in wol and cre1 ahk2 ahk3 backgrounds. AHP6 expression can also be downregulated ¨ by cytokinin treatments (M¨ahonen et al., 2006a). In conclusion, the identity of the pluripotent procambial cells and protoxylem is determined by spatially specific regulatory interaction between cytokinin signaling and its inhibitor. Thus, a negative regulatory feedback loop operates where cytokinin signaling counteracts expression of its inhibitor facilitating protoxylem formation

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Root Development (Fig. 2.7c). These results suggest a position-based mechanism mediated by cytokinin signaling for determining vascular cell fate and are well in line with previous studies in the Arabidopsis root that showed that positional control mechanisms are most important for cell fate decisions (van den Berg et al., 1995). 2.3.3

Xylem and phloem patterning

During the last years, several mutants have been discovered that alter the vascular bundle by changing the symmetry and architecture of phloem and xylem (McConnell and Barton, 1998; Zhong and Ye, 1999; Eshed et al., 2001; McConnell et al., 2001). A few genes have also been shown to be directly involved in the morphogenesis of phloem and xylem poles in the primary ¨ root of Arabidopsis (Bonke et al., 2003; Kubo et al., 2005; M¨ahonen et al., 2006a). In lateral organs, such as leaves, xylem and phloem differentiate asymmetrically from procambium with phloem developing on the abaxial (peripheral) side of the leaf and xylem on the adaxial (central) side of the leaf, meaning that in leaf veins, the xylem cell files are always found on top of the phloem cell files, in contrast to the pattern in the radially symmetrical root. Genes coding for five class III HD-ZIP transcription factors are present in the Arabidopsis genome that play a special role in leaf vascular patterning. EMS mutants of PHABULOSA (PHB), PHAVOLUTA (PHV) and REVOLUTA (REV) showed adaxialization of lateral organs where xylem surrounds phloem (Emery et al., 2003) (Fig. 2.8). These dominant mutations were later found to be in a microRNA (miRNA) targeting site within the gene open reading frames, rendering them immune to degradation by miRNA 165/166 (Reinhart et al.,

Figure 2.8 The adaxialization/abaxialization concept in Arabidopsis leaf. In a completely adaxialized tissue, xylem surrounds phloem whereas in a completely abaxialized tissue, phloem completely surrounds xylem. (Reprinted from Carlsbecker and Helariutta, 2005; with permission from Elsevier.)

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2002; Rhoades et al., 2002). Indeed, these gain-of-function mutants exhibit an ectopic expression of the genes that gives rise to their adaxialized phenotypes. On the other hand, loss-of-function triple mutants of PHB, PHV and REV displayed the opposite phenotype with abaxialization of vascular tissue in severe cases where phloem surrounded xylem in the cotyledons. For a schematic visualization of the adaxialization/abaxialization concept, see Fig. 2.8. A similar, but even more pronounced phenotype in which only one radially symmetrical, abaxialized cotyledon could be seen was discovered in the quintuple mutant with loss-of-function mutations in the five class III HD-ZIP genes, including ATHB8 and ATHB14 (Prigge et al., 2005). Another group of genes, KANADI (KAN), belongs to the GARP family of transcription factors is expressed with a pattern complementary to the class III HD-ZIP genes, and has been suggested to act in an antagonistic manner to them perhaps by acting through miRNA regulation (Bowman, 2004). In accordance to this hypothesis, the kan1 kan2 kan3 triple mutant has a radially symmetrical, adaxialized pattern in which phloem surrounds xylem and overexpression of KAN1 gives rise to abaxialization (Eshed et al., 2001; Kerstetter et al., 2001; Emery et al., 2003). Both classes of genes are expressed in roots (Hawker and Bowman, 2004). Especially, PHB expression has been analyzed in detail (Lee et al., 2006): PHB is expressed in the whole of the stele of the Arabidopsis primary root tip, whereas the translational fusion construct (PHB:YFP) has a much more restricted pattern of expression. This result is indicative of posttranslational regulation of PHB taking place during vascular morphogenesis in root as well as in shoot. Although these genes have shown to be of importance in the shoot vascular patterning, no root phenotype has yet been reported. There is evidence though that the miRNA regulation of class III HD-ZIP genes also plays a role in root vascular patterning (Annelie Carlsbecker, Ove Lindgren and Yk¨a Helariutta, personal communication). A knockout mutant of the ALTERED PHLOEM DEVELOPMENT (APL) gene was found to be seedling lethal and to have a terminal root growth without lateral roots and arrested shoot growth after only a few true leaves were visible. Upon closer examination an impaired phloem development phenotype was discovered (Bonke et al., 2003). Cross sections of the apl mutant primary root revealed that it entirely lacked metaphloem sieve elements and companion cells in the two phloem poles and also had defects in the asymmetric cell divisions associated with phloem morphogenesis (Fig. 2.6a). Interestingly, in phloem poles of apl, TE-like cell types were observed instead of phloem cell types (Fig. 2.6b). Furthermore, a phloem marker construct, AtSUC2::GFP, with the promoter from the sucrose transporter ATSUC2 (Imlau et al., 1999) that is normally expressed in companion cells, failed to show any expression in apl mutant background. Similarly, expression was absent with a protophloem-specific marker, JO701 (Jim Haseloff collection). The corresponding APL gene was identified at a molecular level to encode a transcription factor with an MYB-like DNA-binding domain. In the primary root, APL was expressed in all the phloem cell types, protophloem, metaphloem

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Root Development and companion cells. This expression pattern is spatially dynamic, mirroring the development of sieve elements and companion cells. When APL was ectopically expressed in the whole stele under the control of the WOODEN LEG ¨ promoter (WOL; M¨ahonen et al., 2000) transgenic plants did not display any protoxylem formation near the root tip and metaxylem cells tended to develop later. These results suggest that APL, besides being required for phloem development, might also play a role in inhibiting xylem differentiation in the phloem poles. By utilizing an in vitro system for xylem differentiation in Zinnia that was adapted to Arabidopsis (see Section 2.4), a related group of NAC-domain transcription factors was found to be upregulated at the onset of xylem vessel formation and considered to be of special interest for further studies (Kubo et al., 2005). Four of these genes belonging to the same subfamily were present in this data set and an additional three members were selected based on homology to this group. They were designated VND1 to VND7 (VASCULARRELATED NAC-DOMAIN PROTEIN). VND6 was expressed in immature metaxylem lineages, whereas VND7 in immature protoxylem lineages of the primary root. The importance of these two genes was indicated by ectopic expression using the 35S promoter through which plants transformed with 35S::VND6 and 35S::VND7 produced metaxylem and protoxylem vessels, respectively, outside the stele in the root (Fig. 2.6c). This ability to ectopically change the cell fate was detected in several different tissues, including the primary root. T-DNA insertion or RNAi lines of these genes did not show any phenotype, but by fusing VND6 and VND7 to the SRDX strong repression domain under the control of the 35S CaMV promoter, the transformed plants displayed phenotypes with inhibited protoxylem and metaxylem, respectively, indicating that VND6 and VND7 are key regulators for metaxylem and protoxylem identity. 2.3.4

Cell proliferation during vascular development

A proper balance between cell proliferation and cell differentiation is required at all stages of plant development, which is most evident in the equilibrium between cell division versus cell differentiation in the apical meristems. Enhanced cell proliferation affects organ formation (e.g. shoot meristem) and enhanced cell differentiation leads to meristem consumption (for review, see Chapter 1; B¨aurle and Laux, 2003). There is less information on the control of cell proliferation during vascular development. The emerging picture is that phytohormones (auxin, cytokinin and brassinosteroids) play an important role in controlling the balance between cell division and differentiation during primary and secondary phases of vascular development. It was shown above that auxin and cytokinin have major roles during primary vascular development: auxin for specification of the vascular bundle and cytokinin for cell fate decisions within the vascular bundle. Furthermore,

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for secondary vascular development, it has long been recognized from studies with species other than Arabidopsis that auxin is involved in the initiation of the vascular cambium and also in its subsequent growth, i.e. the division of cambial cells and the differentiation of its daughter cells into secondary xylem and phloem (reviewed by Helariutta and Bhalerao, 2003 and references therein). There is an increased awareness of the usefulness of Arabidopsis as a model system to study (pro-)cambial cell divisions (reviewed by Nieminen et al., 2004). Studies in Arabidopsis inflorescence stems indicate that indeed auxin might play an important role in cell proliferation: pin1 mutants overproduce vascular tissue in the bundles adjacent to cauline leaves of the inflorescence stem (G¨alweiler et al., 1998); ifl1 mutants present a reduction in the production of secondary xylem correlated with a decrease in auxin polar transport (Zhong and Ye, 2001); physiological studies (manipulating endogenous and exogenous auxin levels) indicate that auxin is required for initiation of the vascular cambium and stimulates cell proliferation during secondary vascular development (Little et al., 2002); and in an experimental system in which plant body weight was used to induce the transition between primary and secondary growth, it was shown that auxin is a signal mediator in the process (Ko et al., 2004). Moreover, ATHB8 might regulate cell proliferation downstream of auxin signaling and encodes a small homeodomain-leucine zipper transcription factor that is expressed early in procambial cells. It is positively regulated by auxin, and its overexpression leads to an increased proliferation of vascular tissue both in roots and in shoots, suggesting its involvement as an auxin-regulated factor for cell proliferation in vascular development (Baima et al., 1995, 2001). Thus, these data suggest that auxin is involved in the regulation of cell proliferation at the two stages of Arabidopsis vascular development but an overall picture of the genetic mechanisms leading to that regulation is fragmented. Cytokinins, as described above, play an important role in the normal proliferation of procambial cells as showed by the phenotypes of wol and triple knockout mutant of the CRE-family receptors (CRE1/WOL/AHK4, AHK2 and AHK3). The single and double mutants for this class of receptors present ¨ an almost normal vascular pattern (Higuchi et al., 2004; M¨ahonen et al., 2006b). Recently, it has been shown that CRE1 in addition to its kinase activity in the presence of cytokinins, has also a phosphatase activity that dephosphorylates the HPs in the absence of cytokinins. Thus, CRE1/WOL is a bifunctional kinase/phosphatase, whose activity on the phosphoload of the signal ¨ transduction pathway depends on the status of cytokinin binding (M¨ahonen et al., 2006b). Because, the binding of cytokinin to the WOL/CRE1 receptor is abolished in the wol mutant (Yamada et al., 2001), the wol mutation might have a negative activity (i.e. a constitutive phosphatase activity) on cytokinin signaling initiated by the other two cytokinin receptors. This negative activity leads to the wol phenotype by mediating the promotion of protoxylem ¨ differentiation and inhibition of procambial cell proliferation (M¨ahonen et al., 2006b). AHK2 does not appear to have the phosphatase activity.

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Root Development Consequently, by expressing AHK2 ectopically under the CRE1 promoter during procambial development, an increase in cytokinin responsiveness was found, which correlated with additional proliferation of vascular cell ¨ files (M¨ahonen et al., 2006b). A putative role for cytokinin signaling might be regulating the extent of cambial cell divisions during the secondary vascular development. Recently, a high cambial activity (hca) mutation was identified (Pineau et al., 2005) that has an increased cell proliferation of the vascular cambium, resulting in an overproduction of the secondary vascular tissues in the stem and hypocotyl. Interestingly, hca is hypersensitive to cytokinins, suggesting the corresponding gene product might be involved in cytokininmediated control of vascular cell proliferation. In conclusion, both auxin and cytokinins appear to have a dual role in regulating vascular morphogenesis: they are both involved in specifying vascular cell fate and in regulating vascular cell division. Brassinosteroids, another class of plant hormones, besides their role in cell elongation and differentiation, were reported to be implicated in the extent of cell proliferation ratios between phloem and xylem during vascular development in the Arabidopsis stem (Szekeres et al., 1996; Choe et al., ˜ 1999; Cano-Delgado et al., 2004). Brassinosteroids are perceived by a plasma membrane-localized leucine-rich repeat receptor-like kinase, BRASSINOSTEROID INSENSITIVE1 (BRI1; Wang et al., 2001). BRI1-LIKE1 (BRL) and BRL2 were shown to encode two other functional brassinosteroid receptors with specific function in vascular development based on their specific vascu˜ lar expression patterns (Cano-Delgado et al., 2004). The loss of function of brl1 increases phloem and reduces xylem differentiation in the inflorescence stem ˜ (Cano-Delgado et al., 2004). The three genes are also expressed in the root, but ˜ no mutant phenotypes in the root vasculature were reported (Cano-Delgado et al., 2004). A list of genes known to affect cell vascular proliferation is available, but information about the exact physiological or genetic context in which they act is somehow unclear. We will restrict ourselves to a brief description of those genes. Mutants for HY5, which encodes a bZIP-like transcription factor display an enhancement of cell proliferation during the secondary stage of growth in addition to other developmental defects (Oyama et al., 1997). Mutants for VEIN PATTERNING (VEP1) that encode a novel protein with a mammalian apoptotic homologous domain have a less complex leaf venation pattern and both root and stem showed a reduced cell proliferation in primary and secondary vascular development (Jun et al., 2002). Activation tagging of LEAFY PETIOLE (LEP), which encodes a AP2/EREBP-like transcription factor and is expressed in the developing xylem, results in an increase of xylem cell number in the stem and hypocotyl (van der Graaff et al., 2002). Activation tagging VASCULAR TISSUE SIZE (VAS), which encodes a small putative protein with weak similarity to plant lipid transfer proteins and which is expressed in the vascular cylinder, results in an increase of phloem formation and cambial cells (van der Graaff et al., 2002). Mutants for

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LONESOME HIGHWAY (LHW) gene, which encodes a plant-specific protein with sequence similarity to basic helix-loop-helix (bHLH)-domain proteins, positively regulate the size of the stele cell population (Ohashi-Ito and Bergmann, 2007).

2.4 Vascular genomics – getting the big picture A search for genes specific for xylem, phloem and cambium is faced with the challenge to somehow dissect vascular tissues from the rest of a given plant organ. This is especially true when it comes to genomic efforts in which identification of all the genes specific for a certain vascular tissue type is addressed. In recent years several different approaches have been taken to deal with this problem in Arabidopsis. What these approaches have in common is that microarray hybridization of mRNA extracted from the prepared vascular tissues or cells are used to deduce expression specificities. The microarray data are later confirmed by making GUS or GFP constructs for some of the novel genes and by using previously characterized markers. By dissecting the Arabidopsis root–hypocotyl, Zhao et al. (2005) were able to define three different categories of genes: phloem/cambium, xylem and non vascular. The extracted mRNA was hybridized to the 24 K Affymetrix ATH1 GeneChip and the resulting gene expression levels were compared pairwise to each other. The results were visualized in a triangle plot. Among the already characterized genes in this study were IRREGULAR XYLEM genes (IRX1, IRX3 and IRX5; Taylor et al., 2003) that all were found in the xylem category, and APL and SUC2 that according to the microarray analysis were found to be expressed in phloem/cambium. To prove the validity of the data, expression of genes not previously characterized was analyzed by promoter::GUS fusions. In accordance to the microarray data, MYR1 (Thelander et al., 2002) was found to be phloem-specific in root and XYLEM NAC DOMAIN1 (XND1) was localized in the xylem. Another vascular genomics effort has been developed by inducing cells to take on xylem identity. This system, called the in vitro xylem vessel element formation system was originally developed for Zinnia elegans cells (Fukuda and Komamine, 1980). In response to the plant hormones auxin and cytokinin, Zinnia mesophyll cells were found to differentiate into TEs with their characteristic secondary cell wall thickenings. This versatile system has been used to evaluate the effect of plant hormones on TE differentiation and has recently been adapted to Arabidopsis cell cultures (Kubo et al., 2005). By adding brassinolide and boric acid to a 7-day-old cell culture, initially grown in cytokinin and sucrose, 50% of the cells were found to differentiate into xylem vessel elements within 7 days (Fig. 2.9a). By performing microarray analysis on the differentiating cells, using the 24 K Affymetrix ATH1 GeneChip, 1705 genes had a more than eightfold change over time. These genes were divided into several different sets, depending on the nature of their induction (Fig. 2.9) and

Figure 2.9 Expression data from xylem induction in an Arabidopsis cell culture system. (a) Induction of tracheary elements over time in an Arabidopsis cell culture. (From Kubo et al., 2005; with permission from Cold Spring Harbor Laboratory Press.) (b) Expression data from microarray analysis divided into several different sets depending on the nature of induction during sieve element formation. (From Kubo et al., 2005; with permission from Cold Spring Harbor Laboratory Press.)

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a homologous group of transcription factors that was identified from one of the sets (VND1 to VND7), were analyzed in the context of root development (see Section 2.3.3). One of the most powerful methods for extracting a specific tissue makes use of the fluorescence-activated cell-sorting (FACS) technique. By expressing the GFP protein under control of various tissue-specific promoters in Arabidopsis and then sorting the GFP-expressing cells after protoplasting treatment, it was possible to perform microarray analysis on a number of different tissues in the primary root. This method was later successfully used to discover novel tissue-specific transcription factors in the vasculature of the primary root, showing a high degree of resolution (Lee et al., 2006). This method is covered more thoroughly in Chapter 12.

References Baima, S., Nobili, F., Sessa, G., Lucchetti, S., Ruberti, I. and Morelli, G. (1995). The expression of the Athb-8 homeobox gene is restricted to provascular cells in Arabidopsis thaliana. Development 121, 4171–4182. Baima, S., Possenti, M., Matteucci, A., Wisman, E., Altamura, M.M., Ruberti, I. et al. (2001). The Arabidopsis ATHB-8 HD-Zip protein acts as a differentiation-promoting transcription factor of the vascular meristems. Plant Physiol. 126, 643–655. Bauby, H., Divol, F., Truernit, E., Grandjean, O. and Palauqui, J.-C. (2007). Protophloem differentiation in early Arabidopsis thaliana development. Plant Cell Physiol. 48, 97–109. Baum, S.F., Dubrovsky, J.G. and Rost, T.L. (2002). Apical organization and maturation of the cortex and vascular cylinder in Arabidopsis thaliana (Brassicaceae) roots. Am. J. Bot. 89, 908–920. B¨aurle, I. and Laux, T. (2003). Apical meristems: the plant’s fountain of youth. BioEssays 25, 961–970. ¨ Berleth, T. and Jurgens, G. (1993). The role of the monopteros gene in organising the basal body region of the Arabidopsis embryo. Development 118, 575–587. ¨ Bonke, M., Thitamadee, S., M¨ahonen, A.P., Hauser, M.-T. and Helariutta, Y. (2003). APL regulates vascular tissue identity in Arabidopsis. Nature 426, 181–186. Bowman, J.L. (2004). Class III HD-Zip gene regulation, the golden fleece of ARGONAUTE activity? BioEssays 26, 938–942. ¨ Busch, M., Mayer, U. and Jurgens, G. (1996). Molecular analysis of the Arabidopsis pattern formation of gene GNOM: gene structure and intragenic complementation. Mol. Gen. Genet. 250, 681–691. Busse, J.S. and Evert, R.F. (1999). Pattern of differentiation of the first vascular elements in the embryo and seedling of Arabidopsis thaliana. Int. J. Plant Sci. 160, 1–13. Carlsbecker, A. and Helariutta, Y. (2005). Phloem and xylem specification: pieces of the puzzle emerge. Curr. Opin. Plant Biol. 8, 512–517. ˜ Cano-Delgado, A., Yin, Y., Yu, C., Vafeados, D., Mora-Garc´ıa, S., Cheng, J.-C. et al. (2004). BRL1 and BRL3 are novel brassinosteroid receptors that function in vascular differentiation in Arabidopsis. Development 131, 5341–5351. Chaffey, N., Cholewa, E., Regan, S. and Sundberg, B. (2002). Secondary xylem development in Arabidopsis: a model for wood formation. Physiol. Plant. 114, 594–600.

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Annual Plant Reviews (2009) 37, 64–82 doi: 10.1002/9781444310023.ch3

www.interscience.wiley.com

Chapter 3

ROOT EPIDERMAL DEVELOPMENT IN ARABIDOPSIS Rebecca Horn,∗ Keke Yi,∗ Benoˆıt Menand,∗ Monica Pernas-Ochoa,∗ Seiji Takeda,∗ Tom Walker* and Liam Dolan Department of Cell and Developmental Biology, John Innes Centre, Norwich NR4 7UH, United Kingdom

Abstract: The root epidermis is the layer of cells that surrounds the mature root before it undergoes secondary thickening and is an active interface between the plant and the soil environment. The epidermis develops early in embryogenesis along with the rest of the root meristem (radicle). Upon germination, it emerges from below the protective root cap and differentiates into hair and hairless epidermal cells. Hair cells serve to increase the surface area of the root across which water and nutrients can be transported and are the sites where the plant first makes contacts with a range of microorganisms, including the nitrogen-fixing bacteria of the genus Rhizobium. There is an enormous diversity of root epidermis types among the land plants but for the purposes of this chapter, we will focus largely on the development of the epidermis of Arabidopsis. First, we will describe the earliest stages of epidermal development in the embryo. Then, we will define the functions of a number of genes that are required for the maintenance of epidermal identity in the postembryonic seedling root and the mechanism that gives rise to the patterned array of hair and hairless cells in the epidermis. Finally, we will consider the mechanism of root hair morphogenesis and the effect of environmental factors, such as inorganic phosphate and iron, on root hair development in both Arabidopsis and some crop species. Keywords: root hairs; root epidermis; Arabidopsis; pattern formation; cell differentiation

∗

These authors contributed equally to this work.

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3.1 Introduction The root epidermis is the layer of cells that surrounds the mature root before it undergoes secondary thickening and is an active interface between the plant and the soil environment. The epidermis develops early in embryogenesis along with the rest of the root meristem (radicle). Upon germination, it emerges from below the protective root cap and differentiates into hair and hairless epidermal cells. Hair cells serve to increase the surface area of the root across which water and nutrients can be transported and are the sites where the plant first makes contacts with a range of microorganisms, including the nitrogen-fixing bacteria of the genus Rhizobium. There is an enormous diversity of root types among the land plants but for the purposes of this chapter, we will focus largely on the development of the epidermis of Arabidopsis. First, we will describe the earliest stages of epidermal development in the embryo. Then, we will define the functions of a number of genes that are required for the maintenance of epidermal identity in the postembryonic seedling root and the mechanism that gives rise to the patterned array of hair and hairless cells in the epidermis. Finally, we will consider the mechanism of root hair morphogenesis and the effect of environmental factors, such as inorganic phosphate and iron, on root hair development in both Arabidopsis and some crop species.

3.2 Establishment of the epidermis in Arabidopsis The root epidermis develops progressively from early embryogenesis through the mature embryo in the dormant seed. It starts as a simple layer covering an inner mass of cells. Below, we will describe how cells in the early epidermis, or protoderm, acquire different characteristics based on their position, and how this outer layer is maintained in the maturing embryo and later in the life of the root.

3.2.1

Protoderm formation and associated transcription factor expression

The single-cell zygote first divides to produce a larger basal and smaller apical cell. Despite its smaller size, the apical cell produces most of the embryo proper (including the entire epidermis), whereas the basal cell contributes part of the root meristem and forms the extraembryonic suspensor, a filamentous column of cells linking the embryo to the endosperm (Fig. 3.1). The apical cells divide three times to form an eight-cell proembryo. At this stage, no radial pattern is present and each cell is exposed to the egg sac. Each of these eight apical cells divides periclinally (new wall parallel to the surface of the embryo) to form an inner and an outer layer of cells, known

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Embryo Apical cell Protoderm Basal cell Inner cells Suspensor

1-cell stage

8-cell stage

16-cell stage

Globular embryo Figure 3.1 Embryonic origin of the protoderm. Early development of an Arabidopsis embryo showing the formation of the protoderm (shaded grey) as a result of periclinal divisions (the new wall is oriented parallel to the embryo surface) in the cells of the eight-cell embryo.

as the protoderm, or epidermis precursor (Fig. 3.1) (Torres-Ruiz et al., 1996; ¨ Jurgens, 2001). No mutations resulting in a loss of the embryonic epidermis have been identified. Nevertheless, the expression patterns of some transcription factors suggest that they might be involved in early epidermal development. These patterns distinguish the cells in the inner and outer positions. The Arabidopsis MERISTEM LAYER 1 (ATML1) gene encodes a transcription factor that is transcribed in the apical cell formed by the first division of the zygote and remains expressed in all apical cells until the 16-cell stage after which expression is only maintained in the protoderm (Lu et al., 1996). Therefore, ATML1 is expressed only in cells located at the surface of the embryo. Another transcription factor gene, PROTODERM FACTOR2 (PDF2), has the same expression pattern and both have been shown to bind 8-bp cis elements, called L1 boxes in the promoters of target genes (Abe et al., 2001, 2003). These elements are present in the promoters of other protoderm-specific genes such as PDF1 and LIPID TRANSFER PROTEIN1 (LTP1), and suggest a role for ATML1 and PDF2 in specifying the protoderm (Abe et al., 2003). The presence of such boxes in the promoters of both ATML1 and PDF2 themselves suggests that the genes can positively regulate their own expression. Therefore, an active repression mechanism must be present to switch these genes off in the inner cells after the 16-cell stage (Abe et al., 2003). While these genes are expressed in outer cells, other transcription factors are expressed only in inner cells. WUSCHEL (WUS) is expressed in all apical cells up until the 8-cell stage just as ATML1 and PDF2 are; but in contrast to these genes, WUS expression becomes re¨ stricted to the inner, albeit just uppermost, cells (Mayer and Jurgens, 1998). The expression of such genes seems a likely source of protoderm and inner cell identity; however, the question remains how cells perceive themselves as being on the exterior of the embryo and express genes accordingly. There

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are two ways this might be achieved. First, an external signal could promote protodermal development that is received by the outer cells but not by the inner cells. Second, an internally produced signal could promote internal cell identity, in which case the protoderm would receive less of the signal than the inner cells. Such position-dependent signals have not been identified in Arabidopsis, but potential exterior signals might be produced by the endosperm. It is also possible that the remnants of the zygote cell wall, present on the external face of the embryo, might be a source of signalling molecules, as observed in Fucus (Brownlee and Berger, 1995). The periclinal division of each of the eight cells in the apical region of the embryo results in the formation of an inner cell and an outer cell, which go on to develop different identities. The importance of this division can be seen in mutant plants where this division is defective. KNOLLE (KN) is a protein involved in vesicle trafficking required for new cell wall formation at cytokinesis. In kn mutants, cell walls only partially form and, as a consequence, daughter cells remain connected to each other after karyokinesis. In such embryos, protodermal markers can be seen in the inner cells and inner cell markers in the protoderm (Lukowitz et al., 1996), indicating that cell division does not initiate pattern but is required for its maintenance along strict boundaries. 3.2.2

Initiation of the root epidermis

As the radial pattern develops, an apical–basal axis is also forming. At the basal end of this axis, a signal derived from the embryo, thought to be auxin (Sabatini et al., 1999; Friml et al., 2002), induces the top cell of the suspensor and the adjacent cells to initiate the root meristem. The apical-most suspensor cell divides to form a lens-shaped cell that will go on to form the quiescent centre (QC). It is hypothesized that the QC signals back to adjacent cells in the embryo, keeping them in an undifferentiated state as stem cells, or initials (van den Berg et al., 1997). There are separate groups of initials for each of the main tissue layers. The basal-most ring of protodermal cells become epidermal initials and will give rise to the epidermis of the mature seedling root (Figs. 3.2a and 3.2b) (Dolan et al., 1993). Upon completion of root meristem development at the end of the heart stage the epidermal initials divide, periclinally, to produce another outer ring of cells (Fig. 3.2c): the lateral root cap that encloses the epidermis and forms a continuous unit with the columella to produce the root cap (Dolan et al., 1993; Wenzel and Rost, 2001). The future epidermis of the root is derived from a ring of approximately 16 initials in a stereotypical pattern of cell divisions. Anticlinal (new cell wall perpendicular to the root surface) divisions of the epidermal initials generate two cells. The first is a new initial and the second is a cell that will go on to form a clone or a packet of epidermal cells through further anticlinal divisions (Dolan et al., 1993, 1994; Kidner et al., 2000). Starting as the outer layer of a ball of cells in the globular embryo, by the end of embryogenesis the epidermis has formed. While the pattern of cell

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Root Development division that gives rise to the epidermis has been described in detail, we know little about the mechanism by which the epidermis develops. Nevertheless, we have begun to identify genes that are required for the maintenance of epidermal identity in the seedling and these will be discussed in the next section.

3.2.3

Maintenance of the epidermis in the mature root

After the epidermis layer is established in early embryogenesis, it is maintained through mature root development. The identification of mutants with defective epidermal phenotypes in mature roots has revealed pathways involved in maintenance of this epidermis patterns during the postembryonic development of the root. Two types of mutants with abnormal epidermal development have been identified: tornado (trn1 and trn2) and schizoriza (scz). trn mutants (trn1 and trn2) develop lateral root cap cells in the position where epidermal cells develop in wild type, suggesting that TRN1 and TRN2 negatively regulate lateral root cap fate in epidermal cells (Cnops et al., 2000). Interestingly, an allele of trn1, called lopped1 (lop1), has a defect in auxin transport (Carland and McHale, 1996), indicating a role for auxin in the maintenance of epidermis identity. scz mutants develop root hairs from subepidermal cells (cortex), suggesting that the SCZ gene represses epidermal fate in the cortex (Mylona et al., 2002). We are still far from understanding the mechanism of maintenance of epidermis pattern, but it seems to be clear that although the establishment of epidermis identity is an embryonic event, the later maintenance of this identity requires the activity of genes, such as SCZ and TRN.

3.3

Establishment of distinct cell fates in the root epidermis

In mature Arabidopsis roots, epidermal cells are arranged in alternating files of hair-bearing (H) and non-hair-bearing (N) cells. The identity of epidermal cells is determined by their position relative to underlying cortical cells. Epidermal cells located adjacent to two cortical cells develop hairs, whereas those that are adjacent to only one cortical cell remain hairless (Dolan et al., 1994; Galway et al., 1994). This cell fate decision occurs before the emergence of the hair. The future H cells are smaller and have a denser cytoplasm than the cells in N position in the meristematic region (Fig. 3.3a). We will describe the nature of the root epidermis cell fate and how it is established. We will then discuss when the pattern is established. A fundamental question about cell differentiation in multicellular organisms is whether the fate of a cell is dependent on cell lineage or on cell position. Two experiments have shown that cell identity in the epidermis is determined by positional signals. When an N cell is ablated with a laser on

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a confocal microscope, the neighbouring cell in an H cell position divides to form a new cell in the position previously occupied by the ablated N cell (Berger et al., 1998). This new cell develops N identity – that is it develops the identity that corresponds to its new position. This shows that the identity of the epidermal cell is controlled by positional information rather than cell lineage. Confirmation of a role for positional information was obtained by examining the patterns of cell division and the identities of cells that result from these divisions during wild-type development. Occasionally, longitudinal anticlinal cell division takes place in cells in the H position. One of the resulting daughter cells contacts two cortical cells and develops H identity and the other contacts a single cortical cell and develops N identity (Berger et al., 1998). Together, the cell ablation and the analysis of the fates following longitudinal anticlinal cell division indicate that cell position controls cell identity in the root epidermis (Fig. 3.3b). The role of positional information has been reinforced by the recent identification of a putative transmembrane leucine-rich repeat receptor kinase called SCRAMBLED (SCM) that is required for cell patterning in the root epidermis. SCM was identified through a genetic screen for mutants having a patchy expression pattern of a marker of N cells (Kwak et al., 2005). In wild-type plants, the marker is expressed in longitudinal stripes of N cells, whereas in the SCM mutant background the pattern is scrambled. SCM is expressed in the stele, the endodermis, the cortex and in all epidermal cells, suggesting that it is not its expression pattern but rather the localized distribution of its ligand that determines the pattern of cell identities. SCM controls the expression of the transcription factors that define the epidermal cell identity and has thus been proposed to relay the positional information to the transcriptional network that governs epidermal cell differentiation. Negative regulators of H cell identity include the homeodomain Zip-related transcription factor GLABRA2 (GL2), the WD40 protein TRANSPARENT TESTA GLABRA1 (TTG1), the R2R3 MYB-type transcription factor WEREWOLF (WER), and the basic helix-loop-helix transcription factors GLABRA3 (GL3) and ENHANCER OF GLABRA3 (EGL3). On the other hand, the MYB-repeat transcription factor CAPRICE (CPC) and the related proteins TRIPTYCHON (TRY) and ENHANCER OF TRY AND CPC1 (ETC1) promote the H cell fate (reviewed in Schiefelbein, 2003). Some of these proteins (GL3, EGL3, TTG1, TRY, WER and GL2) also control epidermal development in the hypocotyls and leaves, which means that some part of the molecular mechanism controlling cell identity is shared between shoot and roots (Schiefelbein, 2003). The regulatory interaction between most of these transcription factors has been described precisely with reporter gene fusions and multiple mutant analyses. A model to account for the regulatory interactions between components of this pathway is schematized in Fig. 3.3b. WER is the first known protein downstream of SCM (Kwak et al., 2005). When activated predominantly in N cells, it promotes the transcription of CPC by direct interaction with the CPC promoter (Koshino-Kimura et al., 2005; Ryu et al., 2005). Then,

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Root Development CPC moves to neighbouring cells in H positions where it inhibits expression of GL2, a negative regulator of root hair development (Wada et al., 2002; Kurata et al., 2005). Furthermore, CPC inhibits its own expression and that of WER in H cells (Lee and Schiefelbein, 2002). GL3 and EGL3 enhance GL2 expression in N cells, probably through a complex with WER and TTG1 as suggested by yeast two-hybrid experiments (Bernhardt et al., 2003). The expression of GL3 and EGL3 is induced in H cells by CPC but the GL3 protein has been shown to move to N cells (Bernhardt et al., 2005). TRY and ETC1 may act redundantly to CPC in a yet unknown manner (Kirik et al., 2004). Thus, a network of transcription factors that interact through cell-to-cell movement, feedback inhibition and direct interactions controls root epidermal patterning. GL2, WER and CPC are also expressed in the embryo, suggesting that the cell patterning is established during embryogenesis (Lin and Schiefelbein, 2001; Costa and Dolan, 2003). GL2 is first expressed in the protoderm in the heart stage. Later, GL2 expression is restricted to the future N cells in the transition between torpedo and mature stage. This limitation of GL2 expression depends on the interaction with WER and CPC, in a manner similar to that in the mature root. Therefore, the organization of the two different epidermal identities depends on a patterning mechanism that is initially established in the embryo. In summary, positional information relayed by a transmembrane receptor kinase to a cascade of transcription factors establishes and maintains root epidermal cell specification. This pathway is established during embryogenesis and involves cell-to-cell movement of transcription factors.

3.4

Root hair initiation and tip growth

Root hairs are tip-growing projections that arise from specialized root epidermal cells. They initiate at the restricted site on the surface of specialized epidermal cells. In Arabidopsis, a small bulge (swelling) forms at the apical end of hair cells (end of the cell nearest to the root meristem), which then elongates by polarized tip growth (Fig. 3.4). The ease with which even subtle mutants can be identified in genetic screens has led to the use of the root hair as a model system for understanding the mechanism of cell differentiation and morphogenesis. Nevertheless, our current understanding of the mechanism of hair growth comes from a combination of cellular, physiological and genetic approaches that have been carried out in a number of different species. In the next section, we will review the current knowledge of the mechanism controlling the morphogenesis of the Arabidopsis root hair. 3.4.1

Restriction of the initiation site in root hair cells

Root hairs initiate from the end of the cell nearest to the root meristem – that is at the apical end of the apical–basal axis that runs along the root hair cell. This

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Figure 3.4 Arabidopsis root hair development. Root hair development of Arabidopsis imaged with differential interference contrast (DIC) optics. Asterisks mark the position of nuclei in cells and arrowheads the location of a dense cytoplasm at the hair tip in an otherwise highly vacuolated cell. (a) Hair cell before forming a bulge (swelling). (b) Hair cell in the process of forming a small bulge at the end of the cell nearest to the root meristem. (c) Bulge initiating tip growth. (d) Rapidly elongating root hair. After the formation of a bulge, the root hair accumulates dense cytoplasm at its tip. This cytoplasmically dense tip remains in place as long as the hair is growing and disappears once growth stops (b–d). Before the bulge forms, the nucleus is located next to the inner periclinal face of the cell (a), it then moves toward the end of the cell nearest to the meristem during the early phase of hair growth (b, c) and finally enters the growing hair (d) and maintains a more or less constant distance from the growing tip.

asymmetry indicates that the hair-forming cells are already polarized before hair outgrowth. The mechanism that establishes this difference between the two ends of the cell is unknown, but might involve auxin because auxin is known to flux away from the meristem through the epidermis and mutants with defects in auxin signalling often develop hairs in defective locations (Masucci and Schiefelbein, 1994, 1996; Pitts et al., 1998). Moreover, loss-offunction mutations in a STEROL METHYLTRANSFERASE1 gene, which is required for the synthesis of major membrane sterols, results in altered auxin distribution and cell polarity defects in roots, including more randomized formation of root hair bulges over the apical–basal axis in a hair cell (Willemsen et al., 2003). Mutants that develop hairs in the wrong place or at multiple sites along the epidermal cell define genes involved in the spatial control of hair outgrowth. tiny root hair1 (trh1) mutants have multiple hair initiation sites, suggesting

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Root Development that the wild-type activity of TRH1 is required to specify a single site of hair outgrowth (Rigas et al., 2001). TRH1 encodes a member of the HAK family of potassium transporters and has been shown to be capable of catalysing auxin efflux in roots and yeast cells (Vicente-Agullo et al., 2004). The precise mode of TRH1 action is unknown, but its role in auxin transport reinforces the view that auxin is required for the selection of the site of outgrowth in root hair cells. The root hair defective 6 (rhd6) mutant produces fewer hairs than wild type; however, when hairs form in this mutant, they initiate at a more basal position (away from the root meristem) than wild-type hairs (Masucci and Schiefelbein, 1994). This indicates that RHD6 is required to specify the position of hair emergence along the apical–basal axis of the hair cell. Occasionally, two hairs form on a single epidermal cell, indicating that RHD6 is also involved in restriction of emerging site in hair cells as well as bulge formation (Masucci and Schiefelbein, 1994). Since RHD6 is a basic helix-loop-helix transcription factor that accumulates in H cells before but not during hair initiation, it is likely that this protein controls H cell differentiation through the transcriptional control of genes that encode products that are themselves active during the cellular restructuring that occurs during the process of hair initiation (Menand et al., 2007). Mutant plants that lack the SCN1/RhoGDI protein develop multiple sites of hair initiation implying that it restricts the numbers of sites of hair growth to a singularity (Carol et al., 2005). This suggests that GTPases of the Rho family, called ROPs in Arabidopsis, might be involved in the selection of the site of out-growth since Rho proteins are regulated by GDP dissociation inhibitors (GDIs). Consistent with this model is the observation that ROP2 is located at the tip of the root hair and this localization requires SCN1/RhoGDI function. Furthermore, transgenic plants expressing mutant versions of ROP2 develop abnormal hairs (Molendijk et al., 2001; Jones et al., 2002). Together, these observations suggest that Rho GTPases control the selection of the initiation site and that these genes are active throughout the growth of the root hair. 3.4.2

Physiological characteristics of the early phase of root hair growth

During hair bulge initiation, three major physiological changes occur in root hair cells, one of which is the localized change of pH of the cell. At the root hair initiation site, cytoplasmic pH is elevated, whereas cell wall pH decreases simultaneously during bulge formation (Bibikova et al., 1998). This acidification of cell walls is believed to induce cell wall loosening, since extracted wall loosening enzymes from cucumber hypocotyls are active in acidic condition (McQueen-Mason et al., 1992). The second change is the accumulation of Ca2+ at the tip of hair bulges. Using the ratiometric fluorescent Ca2+ indicator Indo-1, Wymer et al. (1997) showed that free cytoplasmic Ca2+ is elevated at the tip of hair bulges, whereas no elevation occurs before the initiation (Wymer et al., 1997).

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Interestingly, the accumulation site of Ca2+ in growing hairs changes upon treatment with the microtubule (MT)-stabilizing drug taxol or physical stimulation of the tip, resulting in aberrant growth direction and production of waving or branching hairs (Bibikova et al., 1997, 1999). This indicates that Ca2+ accumulation is dependent on MTs and that Ca2+ and MT regulate direction and stabilization of tip-growing root hairs. Finally, reactive oxygen species (ROS) have been shown to accumulate at the tip of root hairs (Foreman et al., 2003; Carol et al., 2005). ROOT HAIR DEFECTIVE2 (RHD2) encodes an NADPH oxidase homolog AtrbohC and has been shown to be involved in ROS production in root hairs (Foreman et al., 2003). The rhd2 mutant form bulges, but hair elongation does not occur (Schiefelbein and Somerville, 1990) nor does Ca2+ accumulate in the mutant hair bulges (Wymer et al., 1997). Furthermore, Ca2+ accumulates at the hair tip upon application of ROS to the rhd2 hair bulges (Foreman et al., 2003), indicating that RHD2-derived ROS is required for outgrowth from the bulge (and not the formation of the bulge itself) and also for the accumulation of relatively high levels of Ca2+ at the growing hair tip. In other words, RHD2 regulates the transition from initiation to tip growth. RHD2-derived ROS might control a number of factors required for tip growth. For example, ROS activates the hyperpolarization-activated calcium channel that is required for the accumulation of Ca2+ at the root hair tip. It may control other processes that remain to be identified. As both Ca2+ and ROS continue to accumulate at the tip of growing root hairs (Wymer et al., 1997; Carol et al., 2005), they are involved not only in the early stages of development of the bulge but also in later maintenance of tip growth. Next, we will describe proteins that have been identified as being involved in the elongation of the root hair. 3.4.3

Mechanism of root hair elongation

During tip growth, an axial cytoskeleton targets vesicles to the growing tip. Drug studies have shown that MTs control the direction of growth – root hairs treated with MT-depolymerizing agents continue to grow, but the directionality of their growth is defective. On the other hand, microfilaments are absolutely required for growth, actin2 mutants are hairless and treatment with actin-depolymerizing drugs blocks all hair growth (Bibikova et al., 1999; Ringli et al., 2002; Nishimura et al., 2003). Other components of the tip growth mechanism have been identified through genetic screens and they will be described in the following sections. Screens for mutants with defective hair morphogenesis have identified a plethora of genes involved in tip growth. These include genes involved in root cell wall formation, signalling cascades and GTPases networks. A cellulose synthase-like protein called KOJAK (KJK) and a pair of leucinerich repeat/extensin cell wall proteins called LEUCINE-RICH REPEAT EXTENSIN1 (LRX1) and LRX2 are required for hair growth because kjk mutants

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Root Development and lrx1/lrx2 double mutants arrest hair growth early after initiation (Baumberger et al., 2001, 2003; Favery et al., 2001), suggesting that they are involved in generating new cell wall in the growing hair. Two cell wall loosening proteins, Arabidopsis thaliana EXPANSIN 7 (AtEXP7) and AtEXP18, are expressed in initiating and tip-growing root hairs, suggesting that wall loosening at particular site of cells is important for polarized growth, although loss-offunction mutations have wild-type phenotypes (Cho and Cosgrove, 2002). The characterization of a number of mutants has shown that lipids are important for a number of aspects of root hair elongation. The genes CAN OF WORMS1 (COW1), encoding a phosphatidylinositol transfer protein, and the phospholipase D
1 (AtPLD
1), regulated by GL2, are expressed in root hairs and decrease of their activity causes aberrant hair morphogenesis (Ohashi ¨ et al., 2003; Bohme et al., 2004). Arabidopsis thaliana 3 -PHOSPHOINOSITIDEDEPENDENT KINASE-1 (AtPDK1) regulates the activity of AGC2 (a member of a family comprising cAMP-dependent protein kinases, cGMP-dependent protein kinases and protein kinase C). AGC2 is expressed in root hairs and plants homozygous for agc2 loss of function alleles develop short-root hairs (Anthony et al., 2004), suggesting that lipid signalling is required for hair tip growth. Loss of the TIP GROWTH DEFECTIVE 1 (TIP1) gene that encodes an S-acyl transferase results in branched root hair growth (Hemsley et al., 2005). Although subcellular localization of TIP1 has not been determined, it is predicted that TIP1 acylates residues in regulatory proteins, such as GTPase proteins, calcineurin B-like calcium-sensing proteins and calcium-dependent protein kinases. Two serine/threonine kinase-encoding genes, INCOMPLETE ROOT HAIR ELONGATION (IRE) and OXIDATIVE SIGNAL-INDUCIBLE1 (OXI1), are expressed in root hair cells and loss-of-function mutations in these genes result in the growth of shorter root hairs (Oyama et al., 2002; Rentel et al., 2004). It is thought that OXI1 activates the expression of mitogen-activated protein kinase (MAPK) proteins during root hair growth (Rentel et al., 2004). Although no MAPK proteins have been shown to control root hair development in Arabidopsis, a MAPK from alfalfa (SIMK) has been shown to be localized to hair tip and is required for root hair growth (Samaj et al., 2003). These data suggest that signalling cascade mediated by kinases is crucial for root hair elongation. The cytoplasm at the tip of growing hairs is dense and filled with vesicles that deliver new wall and membrane material to the surface or recycle membrane through an endocytotic pathway. A GTPase protein, called ADPribosylation factor 1 (ARF1), is required for membrane trafficking in the cell and is localized to the Golgi apparatus of root epidermal cells. It is co-localized with an endocytic marker FM4-64, which marks endocytotic vesicles, suggesting that the ARF1 is involved in root hair endocytosis (Xu and Scheres, 2005). Nevertheless, it is unknown if the function of ARF1 is required for growth because mutants have wild-type root hairs. Rab proteins are another class of GTPases; RabA4b is located to the root hair tip and regulates vesicle trafficking in root hair cells (Preuss et al., 2004). This tip localization of RabA4b

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is altered by the addition of actin-depolymerizing agent latrunculin B and root hair growth is inhibited (Preuss et al., 2004), suggesting that actin-based membrane trafficking is crucial for hair growth. Again, the precise requirement for Rab4 in root hair elongation is unknown because a defective root hair phenotype in mutants lacking Rab4 function has not been reported. These data show that cell wall biosynthesis, signal transduction and protein modification are active in growing hairs and that actin-based membrane trafficking is required to maintain the polarized cell growth.

3.5 Effects of nutrients on root hair cell differentiation and morphogenesis In Arabidopsis, deficiency of phosphate and iron both lead to changes in root hair length and density. The increase in density results from increased hair production from H cells and some hair production from N cells (Bates and Lynch, 1996; Schmidt et al., 2000; Ma et al., 2001). Analysis of mutants with defects at different stages of the root hair developmental pathway showed that the deficiency signal is perceived at an early stage of epidermal cell development and that the response to iron and phosphorus deficiency are differentially regulated. In the mature primary root, GL2, TTG and WER are important for forming root hairs in N cells under both stress conditions, while GL2 and WER are involved in hair production from H cells under phosphate deficiency stress only. Another important transcription factor, CPC is required for H cell differentiation under both stress conditions, but only contributes ¨ to N cell hair production during phosphate deficiency (Muller and Schmidt, 2004). Iron deficiency results in root hair branching, and CPC, GL2 and TTG all contribute to the control of this phenotype. Root hair elongation in response to iron and phosphate deficiency is regulated by KJK, LRX1, RHD1, RHD2, RHD4, RHD3, RHD6 and TIP1, while TRH1 is important only under ¨ iron deficiency stress (Muller and Schmidt, 2004). Auxin and ethylene are involved in root hair development (Tanimoto et al., 1995; Masucci and Schiefelbein, 1996). Exogenous application of auxin or ethylene can induce increased root hair density and root hair length (Pitts et al., 1998) in a process that requires the ethylene signalling pathway (Cho and Cosgrove, 2002). These hormones have been shown to act downstream of TTG and GL2 to promote root hair formation in Arabidopsis (Masucci and Schiefelbein, 1996). Investigations into the role of auxin and ethylene in root hair development under both iron and phosphate stress indicate that auxin and ethylene signalling are crucial for root hair development under iron deficiency stress (Schmidt and Schikora, 2001) and that ethylene is required in Arabidopsis for the complete root hair responses to phosphate deficiency (Zhang et al., 2003). In summary, the phosphate and iron deficiency stress can affect the root epidermal cell fate and root hair morphology. Further efforts need to focus

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Root Development on the mechanism for the plant to perceive the stress signal, the regulation system for the root epidermal cell specification under the stress and the crosstalk between hormones and stress on the root hair development.

3.6

Root hairs and nutrient uptake

Nutrient deficiency limits crop growth and yields worldwide. Fertilizer application alone cannot always ameliorate this limitation. Certain nutrients, such as phosphorus, become unavailable to plants by forming insoluble complexes at both high and low pH. Further, as with iron, diffusion rates are generally low because the ions readily bind to soil particles. One mechanism by which the plant can augment uptake of limiting nutrients is to increase the area over which ion uptake is possible. Production of root hairs, which have been shown to increase the total root surface area between two- and fivefold over the root surface area alone, is a function of both hair length and hair density (Gahoonia et al., 1997). Hair length can vary considerably between ecotypes of the same species. In wheat and barley, such variation correlates with zones of phosphorus depletion in the soil around the root (Gahoonia et al., 1997). There is direct evidence that this zone of phosphorus depletion is due to the activity of root hairs. In experiments with the cereal grass rye (Secale cereale), roots were grown against a mesh through which root hairs alone could protrude. As they grew across an air gap they contacted soil labelled with phosphorus (32 P). After 2 days, 32 P was detected in the shoot, showing that phosphorus is absorbed by root hairs and translocated to the shoot (Gahoonia and Nielsen, 1998). Variation in root hair density directly affects the amount of phosphorus a plant can acquire and assimilate (Bates and Lynch, 1996; Gahoonia and Nielsen, 2004). Wild-type Arabidopsis, a mutant with reduced root hair number (rhd6), and a mutant that develops very short hairs (rhd2) were grown at a range of phosphate concentrations. Shoot biomass of all genotypes was higher at increased phosphorus levels, suggesting that hairs were not important under conditions where phosphate was not limiting. When grown in low phosphorus, wild-type plants had significantly more shoot biomass than rhd6, which in turn, had more shoot biomass than rhd2. When grown in increasing phosphate, rhd6 shoot biomass increased to wild-type levels. The biomass of rhd2 mutants also increased to wild-type levels, but required more phosphate than rhd6 plants. At the highest phosphorus concentrations, there was no significant difference in biomass between the three plant types. As root length did not differ significantly between the plants in these experiments, these results suggest that, under limited phosphorus availability, increased root hair length and density lead to enhanced phosphate uptake per unit root length. Such observations in the model Arabidopsis have direct implications for field crops. Cultivars of barley with longer root hairs sustain high grain yields

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under low phosphate conditions – there is a positive correlation between the volume of soil explored by the root and phosphate uptake (Gahoonia and Nielsen, 2004). In field trials, cultivars with longer root hairs sustained grain yield at low phosphorus levels, and yield levels did not significantly increase with phosphate addition. By contrast, cultivars with shorter root hairs had lower grain yield under the P limited conditions, but yield was responsive to phosphate addition. The relative increase in surface area by root hair proliferation thus increases the zone of soil around the root that the plant can mine for limiting nutrients. Root hairs can take up these nutrients directly and, under limiting conditions, longer haired ecotypes can take up and assimilate more nutrients than shorter haired ecotypes.

3.7 Perspectives The Arabidopsis root epidermis has provided insights into the fundamental mechanisms controlling organogenesis, pattern formation and cell morphogenesis. In fact, the discovery that cell-to-cell movement of transcription factors is responsible for the development of epidermal cell pattern is a novel principle for patterning populations of cells that may operate in other multicellular organisms (Wada et al., 1997, 2002). This breakthrough has made the root epidermis a leading model for deciphering how cellular pattern formation takes place in plants. While it is clear that progress in understanding the development of the root epidermis has been made ignorance exceeds knowledge in many areas. For example, we have still to identify the genes that programme epidermal development in the embryo – at the moment we know only about the pattern of expression of genes whose function is unclear. Another challenge will be to define a mechanism by which the root epidermis senses biotic and abiotic components of the terrestrial environment and how these fine-tune the developmental programme. While these discoveries will tell us something about how land plants have coped with extracting nutrients and water from their substrate over the past 470 million years, they might also help humanity deal with the uncertain future – can knowledge of the fundamental mechanisms underpinning root development help us breed crops for a more sustainable world?

Acknowledgements We are grateful to the John Innes Centre, Biotechnology and Biological Sciences Research Council (BBSRC), Natural and Environmental Research Council (NERC), and European Union – Marie Curie programme and the Gatsby Charitable foundation for funding our research programme in epidermal development.

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Root Development Lee, M.M. and Schiefelbein, J. (2002). Cell pattern in the Arabidopsis root epidermis determined by lateral inhibition with feedback. Plant Cell 14, 611–618. Lin, Y. and Schiefelbein, J. (2001). Embryonic control of epidermal cell patterning in the root and hypocotyl of Arabidopsis. Development 128, 3697–3705. Lu, P., Porat, R., Nadeau, J.A. and O’Neill, S.D. (1996). Identification of a meristem L1 layer-specific gene in Arabidopsis that is expressed during embryonic pattern formation and defines a new class of homeobox genes. Plant Cell 8, 2155–2168. ¨ Lukowitz, W., Mayer, U. and Jurgens, G. (1996). Cytokinesis in the Arabidopsis embryo involves the syntaxin-related KNOLLE gene product. Cell 84, 61–71. Ma, Z., Bielenberg, D.G., Brown, K.M. and Lynch, J.P. (2001). Regulation of root hair density by phosphorus availability in Arabidopsis thaliana. Plant Cell Environ. 24, 459–467. Masucci, J.D. and Schiefelbein, J.W. (1994). The rhd6 mutation of Arabidopsis thaliana alters root-hair initiation through an auxin- and ethylene-associated process. Plant Physiol. 106, 1335–1346. Masucci, J.D. and Schiefelbein, J.W. (1996). Hormones act downstream of TTG and GL2 to promote root hair outgrowth during epidermis development in the Arabidopsis root. Plant Cell 8, 1505–1517. ¨ Mayer, U. and Jurgens, G. (1998). Pattern formation in plant embryogenesis: a reassessment. Semin. Cell Dev. Biol. 9, 187–193. McQueen-Mason, S., Durachko, D.M. and Cosgrove, D.J. (1992). Two endogenous proteins that induce cell wall extension in plants. Plant Cell 4, 1425–1433. Menand, B., Keke, Y., Jouannic, S., Hoffmann, L., Ryan, E., Linstead, P. et al. (2007). An ancient mechanism controls the development of cells with a rooting function in land plants. Science 316, 1477–1480. Molendijk, A.J., Bischoff, F., Rajendrakumar, C.S.V., Friml, J., Braun, M., Gilroy, S. et al. (2001). Arabidopsis thaliana Rop GTPases are localized to tips of root hairs and control polar growth. EMBO J. 20, 2779–2788. ¨ Muller, M. and Schmidt, W. (2004). Environmentally induced plasticity of root hair development in Arabidopsis. Plant Physiol. 134, 409–419. Mylona, P., Linstead, P., Martienssen, R. and Dolan, L. (2002). SCHIZORIZA controls an asymmetric cell division and restricts epidermal identity in the Arabidopsis root. Development 129, 4327–4334. Nishimura, T., Yokota, E., Wada, T., Shimmen, T. and Okada, K. (2003). An Arabidopsis ACT2 dominant-negative mutation, which disturbs F-actin polymerization, reveals its distinctive function in root development. Plant Cell Physiol. 44, 1131–1140. Ohashi, Y., Oka, A., Rodrigues-Pousada, R., Possenti, M., Ruberti, I., Morelli, G. et al. (2003). Modulation of phospholipid signaling by GLABRA2 in root-hair pattern formation. Science 300, 1427–1430. Oyama, T., Shimura, Y. and Okada, K. (2002). The IRE gene encodes a protein kinase homologue and modulates root hair growth in Arabidopsis. Plant J. 30, 289–299. Pitts, R.J., Cernac, A. and Estelle, M. (1998). Auxin and ethylene promote root hair elongation in Arabidopsis. Plant J. 16, 553–560. Preuss, M.L., Serna, J., Falbel, T.G., Bednarek, S.Y. and Nielsen, E. (2004). The Arabidopsis Rab GTPase RabA4b localizes to the tips of growing root hair cells. Plant Cell 16, 1589–1603. Rentel, M.C., Lecourieux, D., Ouaked, F., Usher, S.L., Petersen, L., Okamoto, H. et al. (2004). OXI1 kinase is necessary for oxidative burst-mediated signalling in Arabidopsis. Nature 427, 858–861.

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Rigas, S., Debrosses, G., Haralampidis, K., Vicente-Agullo, F., Feldmann, K.A., Grabov, A. et al. (2001). TRH1 encodes a potassium transporter required for tip growth in Arabidopsis root hairs. Plant Cell 13, 139–151. Ringli, C., Baumberger, N., Diet, A., Frey, B. and Keller, B. (2002). ACTIN2 is essential for bulge site selection and tip growth during root hair development of Arabidopsis. Plant Physiol. 129, 1464–1472. Ryu, K.H., Kang, Y.H., Park, Y.-H., Hwang, I., Schiefelbein, J. and Lee, M.M. (2005). The WEREWOLF MYB protein directly regulates CAPRICE transcription during cell fate specification in the Arabidopsis root epidermis. Development 132, 4765–4775. Sabatini, S., Beis, D., Wolkenfelt, H., Murfett, J., Guilfoyle, T., Malamy, J. et al. (1999). An auxin-dependent distal organizer of pattern and polarity in the Arabidopsis root. Cell 99, 463–472. Samaj, J., Ovecka, M., Hlavacka, A., Lecourieux, F., Meskiene, I., Lichtscheidl, I. et al. (2003). Involvement of MAP kinase SIMK and actin cytoskeleton in the regulation of root hair tip growth. Cell Biol. Int. 27, 257–259. Schiefelbein, J. (2003). Cell-fate specification in the epidermis: a common patterning mechanism in the root and shoot. Curr. Opin. Plant Biol. 6, 74–78. Schiefelbein, J.W. and Somerville, C. (1990). Genetic control of root hair development in Arabidopsis thaliana. Plant Cell 2, 235–243. Schmidt, W. and Schikora, A. (2001). Different pathways are involved in phosphate and iron stress-induced alterations of root epidermal cell development. Plant Physiol. 125, 2078–2084. Schmidt, W., Tittel, J. and Schikora, A. (2000). Role of hormones in the induction of iron deficiency responses in Arabidopsis roots. Plant Physiol. 122, 1109–1118. Tanimoto, M., Roberts, K. and Dolan, L. (1995). Ethylene is a positive regulator of root hair development in Arabidopsis thaliana. Plant J. 8, 943–948. ¨ Torres-Ruiz, R.A., Lohner, A. and Jurgens, G. (1996). The GURKE gene is required for normal organization of the apical region in the Arabidopsis embryo. Plant J. 10, 1005–1016. Van Den Berg, C., Willemsen, V., Hendriks, G., Weisbeek, P. and Scheres, B. (1997). Short-range control of cell differentiation in the Arabidopsis root meristem. Nature 390, 287–289. Vicente-Agullo, F., Rigas, S., Desbrosses, G., Dolan, L., Hatzopoulos, P. and Grabov, A. (2004). Potassium carrier TRH1 is required for auxin transport in Arabidopsis roots. Plant J. 40, 523–535. Wada, T., Kurata, T., Tominaga, R., Koshino-Kimura, Y., Tachibana, T., Goto, K. et al. (2002). Role of a positive regulator of root hair development, CAPRICE, in Arabidopsis root epidermal cell differentiation. Development 129, 5409–5419. Wada, T., Tachibana, T., Shimura, Y. and Okada, K. (1997). Epidermal cell differentiation in Arabidopsis determined by a Myb homolog, CPC. Science 277, 1113–1116. Wenzel, C.L. and Rost, T.L. (2001). Cell division patterns of the protoderm and root cap in the “closed” root apical meristem of Arabidopsis thaliana. Protoplasma 218, 203–213. Willemsen, V., Friml, J., Grebe, M., Van Den Toorn, A., Palme, K. and Scheres, B. (2003). Cell polarity and PIN protein positioning in Arabidopsis require STEROL METHYLTRANSFERASE1 function. Plant Cell 15, 612–625. Wymer, C.L., Bibikova, T.N. and Gilroy, S. (1997). Cytoplasmic free calcium distributions during the development of root hairs of Arabidopsis thaliana. Plant J. 12, 427–439.

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Annual Plant Reviews (2009) 37, 83–126 doi: 10.1002/9781444310023.ch4

www.interscience.wiley.com

Chapter 4

LATERAL ROOT FORMATION Jocelyn E. Malamy Department of Molecular Genetics and Cell Biology, The University of Chicago, 1103 East 57th Street, Chicago, IL 60637, USA

Abstract: The plant root system is formed through continuous production of lateral roots. Lateral root formation can be divided into several discrete developmental stages. Correct progression through each stage is regulated by intrinsic developmental pathways and environmental response pathways. Similarly, the spacing and position of lateral roots in the root system are constrained by intrinsic developmental pathways, but are also responsive to environmental cues. Studies in Arabidopsis and other plants have provided extensive information about the hormones and genes that regulate lateral root formation, and these studies are reviewed, with an emphasis on the role of the phytohormone auxin. The overall architecture of the root system, which is primarily determined by the number and placement of lateral roots, is a key determinant of plant survival and crop yield, particularly under conditions of water or nutrient stress. Therefore, understanding the regulatory mechanisms that control lateral root formation has important applications to agriculture. Our progress in leveraging our molecular understanding of lateral root formation to improve crops is discussed. Keywords: lateral root development; plant hormones; auxin; root architecture; crop improvement

4.1 Introduction A review on lateral root development actually encompasses two very different scientific inquiries. First, researchers have asked how a single lateral root is formed from the cells of a parent root, and how the steps in the process are regulated. In studying this question, we want to know how lateral root founder cells differentiate and divide to form a lateral root primordium (LRP), how and when developmental patterning occurs in the LRP and how an LRP becomes a lateral root with its own lateral root apical meristem. Second, researchers have worked to understand the signals that determine where and when lateral roots form within the root system. Are the sites of lateral root founder cells predetermined, or are certain cells triggered by signals that 83

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Root Development occur during plant growth, or both? What determines the spacing of the lateral roots? What regulates the distribution of overall root mass in the soil? Each of the above questions must take into account the plant’s responses to its environment, as we know that even genetically identical plants make very different developmental decisions when grown under different conditions. This developmental plasticity is a hallmark of plant growth, and is clearly seen in the regulation of lateral root formation (for review, see Malamy, 2005). A role for environmental signaling in regulating lateral root formation makes intuitive sense, as this allows plants to optimize the placement of roots in the root system in accordance with the complex and frequently changing soil environment. Hence, to truly comprehend lateral root formation we must understand (1) the developmental and environmental cues that contribute to the regulation of this process and (2) the way in which these cues are integrated. A model for developmental plasticity in the root system, or indeed in any other plastic organ has been proposed (Malamy, 2005). First, developmental signaling pathways can be considered to be ‘hard-wired’ into a plant. They determine the range of possible root system phenotypes for that plant genotype, and therefore should be consistent among genetically similar individuals. The existence of such pathways defines, for example, the maximum size and ‘branchiness’ that can be attained by the root system of a given plant species. These ‘hard-wired’ pathways are referred to as ‘intrinsic’ pathways (Malamy, 2005). In contrast, environmental signaling pathways reflect the growth conditions of the plant, altering lateral root formation within the constraints permitted by the intrinsic pathways. Environmental response pathways might influence lateral root formation by modulating components of intrinsic pathways (Malamy, 2005). In this model, these components act as nodes to integrate environmental signals with intrinsic developmental programs and to coordinate root system morphology with growth conditions. In this review, each of the above topics will be addressed. The rate and orientation of lateral root growth are beyond the scope of this chapter, but are considered in a recent review (Malamy, 2005). Instead of a comprehensive survey of all genes suggested to play a role in lateral root formation, the seminal, recent work in the field will be described and the major models that have been derived from these studies will be presented. For excellent overviews of recent literature, the reader is also directed to Casimiro et al. (2003); Lopez-Bucio et al. (2003); Hochholdinger et al. (2004a); Malamy (2005); De Smet et al. (2006a); Fukaki et al. (2007); Osmont et al. (2007); and Zhang et al. (2007). For reviews of older physiological and histological studies, the reader is referred to Charleton (1986, 1996); Peterson and Peterson (1986); Torrey (1986); and Malamy and Benfey (1997a) (and references therein). When addressing any question in plant developmental biology, it is important to consider its agricultural implications. Often, there is little conversation between plant developmental biologists and agronomists. Although both are using similar molecular tools, the developmental biologist hopes to explain

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the science underlying plant traits while the agronomist strives to optimize plant traits to improve crop plants. There is an obvious overlap of interests between the two disciplines in the case of lateral root formation, as the number and placement of lateral roots determine the size and architecture of the root system. Root system architectural traits have a high impact on crop yields (see below; de Dorlodot et al., 2007). Therefore, to facilitate interactions between developmental biologists (such as myself) and agronomists, this chapter concludes with a review of the literature demonstrating the importance of lateral root formation to agriculture and highlighting some areas where cross-disciplinary interactions would be mutually beneficial.

4.2 How does a single lateral root form? 4.2.1

Anatomy of lateral root formation

Lateral root formation is generally considered to occur in a series of distinct stages that can be independently regulated. This model arose early on when researchers noticed that after LRPs are formed, some remain internal to the parent root while others emerge to become lateral roots. Hence, lateral root formation was divided into an ‘initiation’ stage and an ‘emergence’ stage (for review, see Malamy and Benfey, 1997a). The multistage model was supported and extended by the isolation of Arabidopsis mutants that were specifically inhibited at various steps in lateral root formation. For example, the Arabidopsis affected lateral root formation 4 (alf4) and the solitary root (slr/IAA14) mutants fail to show any sign of LRP initiation, while root meristemless (rml) and affected lateral root formation 3 (alf3) form LRP that arrest immediately after emergence, but before apical meristem activation (Celenza et al., 1995; Cheng et al., 1995; Laskowski et al., 1995; Fukaki et al., 2002). Furthermore, environmental cues have been shown to differentially affect initiation and LRP emergence (Malamy, 2005), and auxin has been shown to play specific roles in multiple stages of LR formation (Malamy and Benfey, 1997a; Benkov´a et al., 2003; Casimiro et al., 2003). The stages currently defined in LR formation are LRP initiation, LRP development and patterning, LRP emergence, apical meristem activation and subsequent growth. Each of these stages is described below. 4.2.1.1 LRP initiation In the vast majority of plants, LRPs are formed from specific files of pericycle cells in the parent root. The pericycle is the outermost cell layer of the stele. In dicotyledonous plants, LRP initiation is usually restricted to the pericycle cells adjacent to the xylem poles (XPs) (see Section 4.3.1.1). Lateral root initiation in the XP pericycle cells requires the presence of the xylem elements, as an Arabidopsis mutant that develops only one XP is only able to produce

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Root Development lateral roots adjacent to that pole (Parizot et al., 2008). This would seem to suggest that there is a permissive or stimulatory factor delivered by the xylem. However, lateral root initiation does not require that the xylem element be fully or correctly differentiated (Parizot et al., 2008). The consistent association of lateral root initiation with xylem elements in Arabidopsis might therefore reflect coordinated development of diarch symmetry in the xylem and the pericycle (Parizot et al., 2008). Two XP pericycle-specific promoters were recently described (Parizot et al., 2008). Reporter-gene expression patterns driven by these promoters were not affected by growth conditions or hormones, indicating that the XP pericycle cells possess a stable and distinct identity (Parizot et al., 2008). This notion is supported by other inherent differences between XP and non-XP pericycle cells. XP pericycle cells differ from the rest of the pericycle cells in size (Lloret et al., 1989; Beeckman et al., 2001) and in a number of structural and ultrastructural characteristics (Parizot et al., 2008 and references therein). XP pericycle cells replicate their DNA and are seen at the G2 stage (4C) of the cell cycle after leaving the root apical meristem, while all other pericycle cells appear to arrest at the G1 stage (2C) (Beeckman et al., 2001; De Smet et al., 2006a). Mature XP pericycle cells show patterns of gene expression that are different from the rest of the pericycle cells; the auxin signaling gene SOLITARY ROOT (SLR) is expressed only in the XP pericycle cell files (Fukaki et al., 2005; Vanneste et al., 2005), as is the cyclin CYCA2;1 (Beeckman et al., 2001), while XP pericycle cells fail to express Kip-related protein 2 (KRP2), a cell cycle inhibitor that is expressed in the rest of the pericycle cylinder (Himanen et al., 2002). Treatment with exogenous auxin causes all XP pericycle cells (but not pericycle cells in other positions) to participate in the formation of confluent LRP (Himanen et al., 2004). All of these observations suggest that different signaling events occur in the XP pericycle cells than in the rest of the pericycle, and that XP pericycle cells remain uniquely competent to respond to triggers of cell division. XP pericycle differentiation from non-XP pericycle cells must occur very early, as a pericycle-specific marker line, Rm1007, shows reporter-gene expression in XP pericycle cell initials (Parizot et al., 2008). While mitotic activity/competence in the XP pericycle cells is essential for the initiation of LRP, these cell files are not all LRP precursors during normal plant development. Consequently, differentiation of the XP pericycle cells from the rest of the pericycle is not the primary committed step in LRP initiation. Instead, initiation is first identified by changes in gene expression in the subset of XP pericycle cells that will give rise to the new LRP. These cells, termed ‘founder cells’, display auxin-induced gene expression, as evidenced by GUS staining in a transgenic reporter line that carries the auxin-responsive promoter DR5 fused to uidA (Ulmasov et al., 1997). They also express the mitotic cyclin CYCB1;1 and the cyclin-dependent kinase CDKA;1 (Cdc2a) (Dhooge et al., 1999; Beeckman et al., 2001). These changes in gene expression occur before any anatomical evidence of an LRP (i.e. new cell divisions) can be perceived.

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The idea that the initiation process begins prior to cell division is supported by studies in which initiation was arrested by application of the auxin transport inhibitor naphthylphthalamic acid (NPA), and then stimulated throughout the XP pericycle in a synchronous manner by application of exogenous auxin (Himanen et al., 2004). In this study, anticlinal divisions in the XP pericycle were observed at 6 hours after auxin application. In contrast, changes in gene expression, including cell cycle regulators and signal transduction components, were seen at 2 and 4 hours after auxin application. Support for the concept that commitment to initiation occurs before cell division is also provided by experiments in which cytokinesis was blocked with colchicine. After this treatment, subsets of XP pericycle cells underwent radial expansion; these ‘primordiamorphs’ became LRP when colchicine was removed (Torrey, 1986). Experimentally, founder cells are most easily identified by the occurrence of the first cell divisions and, consequently, the term LRP initiation is often used synonymously with the appearance of these divisions, despite our knowledge that earlier events determine founder cell identity. The first cell divisions in LRP formation have been most completely described in the model plant Arabidopsis. In Arabidopsis, the founder cells in the XP pericycle undergo anticlinal divisions as they leave the apical meristem (Casimiro et al., 2001), an average of 1.4 mm from the root tip (Beeckman et al., 2001). Estimates based on clonal lineage experiments suggest that the founder cells arise in three adjacent XP pericycle files for a single Arabidopsis lateral root, with the majority of LRP cells derived from the central of these files (Kurup et al., 2005). The total number of founder cells for a single LRP in Arabidopsis has been estimated to be 3–11, depending on the study (Dubrovsky et al., 2001; Casimiro et al., 2003). Detailed analyses of the early divisions in a single pericycle file in Arabidopsis showed that one cell first undergoes an asymmetric anticlinal division, followed by a similar asymmetric division in an adjacent cell in the same file (Casimiro et al., 2001; Dubrovsky et al., 2001). Asymmetric anticlinal divisions in adjacent pericycle cells have also been observed in many (but not all) other plants (for example Peterson and Peterson, 1986; Casero et al., 1993). A second type of lateral root initiation in Arabidopsis has also been described in which a single founder cell undergoes a series of anticlinal divisions to produce several cells of equal size (Dubrovsky et al., 2001). Interestingly, lateral roots in the fern Marsilea quadrifolia initiate via divisions in a single endodermal cell adjacent to the XP (Lin and Raghavan, 1991). A detailed comparison of lateral root formation in ferns and angiosperms could provide fascinating insights into the evolutionary origins of lateral root initiation mechanisms (see also Chapter 8). 4.2.1.2 LRP development and patterning The anticlinal divisions in the founder cells lead to the formation of a Stage I primordium (Fig. 4.1; Malamy and Benfey, 1997a, 1997b). The subsequent cell divisions in the LRP occur very quickly. In Arabidopsis, cell doubling

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Root Development times of 2.7–4.9 hours were observed, as opposed to 13.6 hours for average pericycle cells in this region (Dubrovsky et al., 2001 and references therein). In Arabidopsis, the cells of the Stage I LRP expand radially and divide periclinally to form a two-layered (Stage II) structure (Fig. 4.1; Malamy and Benfey, 1997a, 1997b; Casimiro et al., 2001). Subsequently, the outer layer undergoes another periclinal division to create a three-layered (Stage III) LRP, and the inner layer follows with another periclinal division to create a four-layered (Stage IV) structure (Fig. 4.1). The next divisions are more complex, as the layers become subdivided. The central cells of the outermost layer divide periclinally to form the root cap and epidermis (Fig. 4.1). At approximately the same time, the peripheral cells of the next internal layer divide periclinally to form the cortex and endodermis. The identification of these cell types is based on two observations: (1) the relative position of the cells in the LRP, which at this stage closely resembles a primary root tip; and (2) examination of cell-type-specific markers. The latter study confirmed that a radial pattern is established in the LRP that is very similar to that seen in the primary root tip. Root cap, epidermis, endodermis, cortex and stele cells can all be identified in the developing LRP (Malamy and Benfey, 1997b). Indeed, the specific expression of cell-type markers in the LRP as early as Stage II indicates that radial patterning occurs concurrent with development, in contrast to earlier ideas that a proliferation stage occurs first and is followed by patterning (Malamy and Benfey, 1997a, 1997b). The development of the LRP is accompanied in some plants by cell divisions in underlying stelar parenchyma and overlying endodermis and cortex (Peterson and Peterson, 1986). In maize and tomato, the endodermis contributes a temporary root cap to the LRP, which is replaced by a cap of pericycle origin (Charleton, 1986; Ivanchenko et al., 2006). In some cases, the tissues outside the pericycle have been suggested to contribute to LRP development by producing digestive enzymes, essentially destroying themselves to create a cavity in which the LRP can develop (Peterson and Peterson, 1986). 4.2.1.3 LRP emergence The patterned Arabidopsis LRP emerges from the parent root by passing through the overlying parental cell layers (endodermis, cortex and epidermis). This occurs through cell expansions at the base of the LRP (Malamy and Benfey, 1997a, 1997b). The idea of expansion-driven emergence is consistent with earlier work in other plants that demonstrated a low mitotic index in ‘just emerged’ LRP followed by an increase in cell proliferation as the new apical meristem is activated (Peterson and Peterson, 1986). The cells overlying the LRP in the parent root have been described variously as ‘crushed’ or ‘separated’ by passage of the developing LRP (Peterson and Peterson, 1986; Laskowski et al., 2006). In many plants, it has been hypothesized that LRP emergence is facilitated by the production of cell wall digesting or loosening enzymes, either by the LRP itself or by adjacent cells (Peterson and Peterson, 1986 and references therein). In maize, for example,

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a ‘poche digestive’ arising from overlying endodermal cells has been postulated to produce hydrolytic enzymes such as acid phosphatases (Charleton, 1996). More recently, Laskowski et al. (2006) demonstrated that the pectate lyase (PL) genes PLA1 and PLA2 are induced by auxin treatment, and showed that the timing of PL gene induction is consistent with their playing a role in LRP emergence. 4.2.1.4 Apical meristem activation Once emergence has taken place, new growth occurs via cell divisions at the tip of the LRP, indicating that a new apical meristem is now responsible for further growth (Malamy and Benfey, 1997a, 1997b). This is consistent with earlier work showing that there is an increase in mitotic index in LRP postemergence (Peterson and Peterson, 1986). Furthermore, the quiescent center, identified by decreased incorporation of DNA precursors and indicative of an organized meristem, is only present postemergence (Torrey, 1986). Activation of the apical meristem is the definitive measure for when an LRP transitions into a lateral root. 4.2.1.5 Apical meristem maintenance For continued growth, the lateral root apical meristem must maintain its integrity. If stem cells in the meristem are not maintained, as for example in scarecrow (scr), shortroot (shr) or plethora (plt) mutants (see below), lateral root growth ceases and the cells at the tip terminally differentiate. In alf3-1, lack of meristem activation and/or maintenance apparently leads to cell death (Celenza et al., 1995). 4.2.2

Similarity to embryonic root formation

The similarities of lateral root formation to embryonic root formation have not been overlooked. Indeed, the formation of the LRP and the embryonic root in Arabidopsis follows a nearly identical series of cell divisions. Detailed observation of the expression of the SCARECROW gene in the embyro and LRP highlights these similarities (Fig. 4.2). SCARECROW is essential for radial patterning in the embryonic, primary and lateral roots. It is expressed in basal cells in the early embryo, and its pattern is refined through embryogenesis to the embryonic endodermal cells and the endodermis/cortex initials. The divisions that create this pattern in the LRP are remarkably similar (Fig. 4.2). Emergence of the lateral root from the parent root via cell expansion is paralleled by the emergence of the radical from the seed, which also occurs by cell expansion. In both cases, emergence is then followed by meristem activation. It remains to be determined whether the same regulatory signals guide each step of the developmental process (see below). Despite the similarities in their developmental programs, the regulation of lateral root initiation is genetically distinct from primary root initiation. For example, the alf4-1 and slr mutants fail to initiate any lateral roots, but

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Root Development nevertheless form normal primary roots (Celenza et al., 1995; Fukaki et al., 2002). Furthermore, mutants have been isolated in maize that lack specific types of lateral roots but show no defects in primary root formation (Hochholdinger et al., 2004a, 2004b). 4.2.3

Regulation of lateral root formation

4.2.3.1 Auxin It is impossible to discuss the regulation of lateral root formation without discussing auxin, which clearly plays a central role at many stages. As it is not currently possible to directly visualize auxin within the plant, research into the role of auxin in lateral root formation has mostly focused on the effects of exogenous auxin and the defects observed in auxin-response and auxintransport mutants. Recently, new advances in auxin detection have given us a better idea than ever before of the sites of auxin synthesis in Arabidopsis and the flow of that auxin to the various regions of the root. Therefore, it is worthwhile to summarize our understanding of auxin production and movement in the root at this point, so that this information can be integrated into the subsequent discussions of LRP initiation, development and emergence. The primary auxin, indole-3-acetic acid (IAA), is synthesized in the young tissues of the shoot system (Ljung et al., 2001). The timing of IAA movement from shoot to root has been modeled for young Arabidopsis seedlings. IAA is present at low levels in the cotyledons and high levels in the first pair of true leaves of Arabidopsis seedlings by 3 days postgermination. Levels in these leaves drop precipitously from day 3 through day 12. In the seedling root, IAA content reaches a maximum at 5–7 days postgermination and then declines. This transient ‘pulse’ of auxin might be the result of export from the first true leaves (Bhalerao et al., 2002). To directly test that shoot IAA contributes to the auxin pool in the root, intact seedlings or excised roots were labeled with deuterated water, and newly synthesized IAA was quantitated in shoot and root. These studies confirmed that the shoots are a major source of root auxin in young seedlings (Ljung et al., 2001, 2005). The extent to which shootderived IAA contributes to the root auxin pool at later developmental times is less clear (see below). Polar transport has been demonstrated to move auxin from the shoot to the root base, and from the root base to the root tip, via the vasculature (toward the root tip, therefore referred to as ‘acropetal movement’ within the root) (for review, see Teale et al., 2006). Mutants with decreased transport, including pin1 and aux1, revealed the mechanics of polar auxin transport. The polarlocalized PIN family of auxin efflux carriers mediates polar flow of IAA from cell to cell. PIN1 is a major player in shoot-to-root movement of auxin, and is located in the basal membrane of vascular cells (xylem parenchyma and protophloem cells) (Teale et al., 2006). PIN1-mediated vascular transport is not the only mechanism for auxin movement from shoot to root tip. When NPA, a specific inhibitor of

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PIN1-mediated polar auxin transport, was present in the growth media, the amount of newly synthesized IAA in the root of 8-day-old seedlings was unaffected, although excision of shoot tissue caused a strong reduction in levels of newly synthesized IAA in the root (Ljung et al., 2001, 2005). The above data are consistent with the idea that auxin is transported from shoot to root tip in an NPA-resistant stream. This stream may become more important as the seedling ages, as NPA significantly reduced auxin content in the root tip of 4-day-old but not 8-day-old seedlings (Ljung et al., 2005). One such auxin transport stream is the phloem, where source:sink relations move leaf metabolites to the root by bulk flow. Indeed, significant levels of auxin have been identified in the phloem (i.e. Hoad, 1995). AUX1, an auxin influx carrier (Bennett et al., 1996), has been implicated in loading auxin into the phloem in the leaves (Marchant et al., 2002) and unloading auxin from the phloem at the root tip (Swarup et al., 2001). Indeed, the AUX1 protein is located throughout developing leaves and in the upper membrane of protophloem cells at the root tip, consistent with the above model. aux1 mutants have reduced auxin transport from shoot to root. Furthermore, auxin accumulates in the leaves of 6-day-old aux1 mutant seedlings and is reduced in the roots (Marchant et al., 2002). In sum, the data lead to a model in which auxin moves from the shoot to the root tip via multiple pathways in the vascular cells (passively in the phloem, mediated by AUX1, and actively in the xylem parenchyma, mediated by PIN1). Recent studies have utilized highly sensitive tools to re-assess the idea that the shoot is the sole source of root auxin. These experiments showed that all regions of the root accumulate newly synthesized auxin even in the absence of the shoot tissues (Ljung et al., 2001, 2005). Biosynthetic capacity is relatively low in the roots of 3-day-old seedlings, but greatly increases by 10 days (Bhalerao et al., 2002), consistent with earlier reports of auxin ¨ synthesis capability in the root (Muller et al., 1998). In isolated roots incubated in deuterated water, IAA was synthesized de novo with the highest rates in the 2 mm region containing the root apical meristem (Ljung et al., 2001, 2005). This could explain the observation that Arabidopsis seedlings grown on NPA for 10 days actually accumulate auxin in the root tip (Casimiro et al., 2001): IAA accumulates in the root tips of NPA-treated plants because it is synthesized there and fails to be exported. Root tip synthesis of IAA might also explain the high levels of IAA observed in the root tip at 3–6 days, before shoot-derived auxin is predicted to reach this location (Bhalerao et al., 2002). Root tip-derived auxin might contribute to auxin in other regions of the root, as there is polar transport of auxin backwards from the root tip to the more distal regions via the lateral root cap and epidermis (basipetal), mediated by AUX1 and PIN2 (Swarup et al., 2001; Friml, 2003). The complex PIN-mediated movements of auxin within the root tip and root are discussed in detail elsewhere in this volume (see Chapters 1 and 6). Tips of lateral roots were also shown to have high rates of IAA synthesis (Ljung et al., 2005). Although it has been impossible to directly determine

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Root Development the stage at which lateral roots begin to synthesize auxin, indirect evidence suggests that onset occurs in late stage LRP. Mutations in MDR4, which encodes a protein essential for acropetal transport of auxin in the root, lead to a dramatic reduction of auxin in early stage LRP, as assessed by activity of the DR5 auxin-inducible promoter (Wu et al., 2007). Hence, auxin is apparently not synthesized in these early LRP, but is transported to them in an MDR4dependent manner. Laskowski et al. (1995) performed seminal experiments in which Arabidopsis LRPs were excised and cultured in the presence or absence of auxin. This study showed that young LRP (3–5 cell layers or smaller) required exogenous auxin to develop further, while older LRP developed into lateral roots in the absence of exogenous auxin. Since auxin is believed to be essential for an LRP to develop into a lateral root (see below), this suggests that LRPs begin to synthesize their own auxin at some point after Stage IV or V. This idea is consistent with the speculation by Bhalerao et al. (2002) that the increase in IAA biosynthesis seen in 10-day-old versus 3-day-old seedling roots might reflect the increased presence of developing LRP and lateral roots. The production and movement of IAA is of course only part of the story of root auxin. IAA modification, conjugation and cellular localization also affect the functional concentrations present in any tissue. Furthermore, it appears that levels of synthesis are tightly monitored and controlled in the plant; for example, NPA treatment results in feedback inhibition of IAA synthesis in the root tip (Ljung et al., 2001, 2005). In addition, it is important to note that other major endogenous auxins exist. Indole butyric acid (IBA) moves from shoot to root by a mechanism that is apparently distinct from IAA transport, based on the observations that polar movement of IBA is insensitive to NPA and does not appear to require AUX1 or PIN2 (Rashotte et al., 2003). Consistently, a rice mutant in IBA transport, arm2, was not affected in IAA transport (Chhun et al., 2005). 4.2.3.2 Regulation of initiation 4.2.3.2.1 Auxin. Auxin clearly plays a key role in LRP initiation. The application of exogenous auxin is sufficient to stimulate the formation of LRP throughout the XP pericycle in Arabidopsis (Himanen et al., 2004) and in other plants (Torrey, 1986 and reference therein). Furthermore, mutants in auxin response such as transport inhibitor response 1 (tir1) and auxin resistant 4 (axr4) (for review, see Casimiro et al., 2003), and in auxin movement such as aux1 (Marchant et al., 2002) and transport inhibitor response 3 (tir3)/big (Ruegger et al., 1997; Lopez-Bucio et al., 2005) have significantly reduced levels of LRP initiation, while mutants with increased auxin production such as rooty ((rty)/aberrant lateral root formation (alf1)/superroot1 (sur1)) and superroot2 (sur2) have increased numbers of LRP (Boerjan et al., 1995; Celenza et al., 1995; King et al., 1995; Delarue et al., 1998). Activation of the auxin responsive promoter DR5 in the lateral root founder cells is one of the earliest known markers of LRP initiation, preceding the first anticlinal divisions.

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4.2.3.2.1.i Auxin transport. Since initiation occurs in the region of the primary root just basal to the meristem, research has focused on how auxin might be delivered to pericycle cells in this region. The application of auxin transport inhibitors to either the entire root or specifically to the root:shoot junction was reported to block the initiation of LRP (Reed et al., 1998). Based on this result, the general dogma for many years has been that shoot-derived auxin is essential for initiation. More recently, it has been suggested that basipetal transport of root tip auxin might be responsible for LRP initiation instead (Casimiro et al., 2001). This hypothesis is based on an examination of NPA inhibition of LRP initiation; in the presence of NPA, IAA levels actually increase in the root tip, leading to a model in which NPA-sensitive transport of auxin from the root tip to the more basal tissues is essential for LRP initiation (Casimiro et al., 2001). This model argues that initiation may not involve shoot-derived auxin at all, based on the finding that the stm1 mutant, which lacks a shoot apical meristem and leaves (the major sources of apical auxin), still initiates normal numbers of LRP (Casimiro et al., 2001). Another possibility, endorsed by Marchant et al. (2002), is that the shoot-derived auxin essential for lateral root initiation moves via the phloem, and its transport to the root tip is therefore NPA resistant. In this case, AUX1 mediates loading of auxin at the cotyledons and/or young leaves into the phloem and unloading of this auxin at the root tip. From the root tip, NPA-sensitive basipetal transport of shoot-derived auxin is necessary for initiation. Alternatively, the NPA sensitivity of LRP initiation may be explained by much more localized events, such as the requirement of PIN proteins for import of auxin into XP pericycle cells (see below). A final model is that redundant mechanisms might be present, such that either shoot- or root tip-derived auxin can independently induce LRP initiation. Root tip auxin would need to be transported to the basal meristem by PIN2 proteins, while shoot auxin would be transported both passively in the phloem and acropetally via PIN1. This model would be consistent with the fact that neither the pin1 nor the pin2 mutant, defective in acropetal and basipetal IAA transport in the root, respectively, has a reported LRP initiation defect (Benkov´a et al., 2003) (although the PIN1 may compensate for the loss of PIN2 in a pin2 mutant (Blilou et al., 2005)). Support for the existence of redundant mechanisms for lateral root initiation may be provided by recent studies in rice. Shoot excision had no effect on lateral root formation under normal growth conditions. In contrast, lateral root numbers and density were stimulated when rice plants are exposed to high humidity conditions, although it is not clear whether this reflects a change in LRP initiation or in later steps (Chhun et al., 2007), and the high humidity stimulation of lateral root formation was eliminated by excision of the shoot tissue. Stimulation was restored by application of auxin to the decapitated shoot. These experiments are interesting because they support the notion that shoot-derived auxin and polar auxin transport from the shoot are not required for LRP initiation. However, they also show that auxin transport from the shoot to the root can be modulated, perhaps by environmental

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Root Development response pathways, and that this can enhance lateral root formation. Auxin transport from the shoot to the root was shown to increase under high humidity conditions, and the speed of transport and lack of inhibition by NPA suggested that auxin was moving through the phloem rather than through PIN-mediated transport pathways. The authors speculate that high humidity leads to an increase in AUX1-mediated loading of auxin into the phloem at the shoot and subsequent transport of this auxin to the root (Chhun et al., 2007). After delivery of auxin to the appropriate region of the primary root, it must be restricted to the founder cells in the XP pericycle. Whatever the source of the auxin, accumulation in specific pericycle cells probably triggers their division. De Smet et al. (2006a) suggested that PIN proteins in pericycle cells might allow canalization of auxin away from neighboring cells. This would create foci of auxin accumulation which would predict the future sites of cell division and specification of founder cells. In fact, NPA may inhibit initiation by preventing this PIN-mediated accumulation of auxin, rather than inhibiting long-distance auxin flow. It is worth noting that many components of the auxin transport machinery affect lateral root initiation. Although the role of each protein is poorly understood, this emphasizes the importance of correct auxin movement and distribution. The pin3 mutant has reduced LRP initiation, which is even more pronounced in pin1/pin3 double mutants (Benkov´a et al., 2003). PIN3 is expressed at the basal side of vascular cells and the lateral side of young pericycle cells (Blilou et al., 2005). Hence, PIN3 may be involved in delivery or localization of auxin in the pericycle. pin7, in contrast, shows increased initiation, as does ectopic expression of PIN1. These findings are consistent with the idea that the PIN proteins are able to redistribute auxin, and their altered expression can increase or decrease local auxin pools, somehow affecting key events in LRP initiation. The redundancy of the PIN proteins and the alteration of PIN gene expression in pin mutants (Blilou et al., 2005) make it difficult to decipher the exact interaction between auxin localization and specific PIN proteins at this time. In addition to the PIN proteins, the phosphoglycoproteins (PGPs) have also been shown to be auxin transporters. The mutant pgp4 has decreased basipetal auxin transport and increased LRP initiation (Santelia et al., 2005; Terasaka et al., 2005). pgp1 and pgp19 (mdr1) have reduced lateral root formation, although analyses to determine whether the defect is in initiation or in later stages was not reported (Lin and Wang, 2005). Finally, BIG mutations, which reduce polar auxin transport, also reduce LRP initiation under all growth conditions (Lopez-Bucio et al., 2005). 4.2.3.2.1.ii Auxin signaling. How auxin regulates lateral root initiation once it reaches the appropriate pericycle cells has long been a fundamental question in the field. AUX/IAA and ARF proteins are known to mediate many auxin-stimulated processes by regulating gene expression (for review, see Teale et al., 2006). Auxin acts by binding to an F-box protein such as TIR1.

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This interaction leads to the ubiquitination of AUX/IAA proteins, which in the absence of auxin repress the activity of ARF transcription factors through heterodimerization. Ubiquitination of the AUX/IAA proteins leads to their degradation, hence de-repressing the ARF proteins (for review, see Teale et al., 2006). Recent studies show that the AUX/IAA gene SLR/IAA14 is a likely transducer of the auxin signal in the pericycle. Dominant mutations in domain II of AUX/IAA proteins stabilize them by preventing ubiquitination, leading to insensitivity to auxin. A dominant mutation of this type in SLR/IAA14 completely inhibits LRP initiation, even in the presence of exogenous auxin, suggesting that the wild-type SLR/IAA14 protein represses auxin responses that are crucial for initiation (Fukaki et al., 2002). SLR/IAA14 is expressed and acts in the XP pericycle cells, as expression of the stabilized form of the SLR/IAA14 protein only in this location is sufficient to inhibit initiation in wild-type plants (Fukaki et al., 2005). SLR/IAA14 has been shown to interact with ARF7 and ARF19 in yeast two-hybrid assays, consistent with the fact that the arf7/arf19 double mutant resembles the slr/iaa14 mutant (Fukaki et al., 2005; Wilmoth et al., 2005). This suggests that stabilized SLR/IAA14 represses initiation by inactivating ARF7/ARF19 function. This idea is supported by the finding that exogenous auxin fails to stimulate lateral root initiation in the arf7/arf19 mutant background (Okushima et al., 2007). A model has been proposed that places SLR as a primary transducer of the auxin signal for LRP initiation. In this model, auxin-mediated degradation of SLR/IAA14 leads to the de-repression of genes essential for initiation. This model is supported by microarray analyses that revealed a large number of putative auxin signaling molecules that are probably regulated by SLR/IAA14 (Vanneste et al., 2005). The auxin-induced genes LBD16 and LBD19 may be two of the genes that mediate SLR, ARF7 and ARF19 effects on lateral root initiation. Expression of both the LBD16 and LBD19 genes is activated by ARF7 and ARF19, and overexpression of either LBD gene partially rescues the lateral root initiation defect seen in the arf7/arf19 double mutant. Interestingly, over-expression of LBD16 also restores the ability of arf7/arf19 to increase lateral root initiation in response to exogenous auxin (Okushima et al., 2007). This implicates LRB16 as a positive regulator of auxin signaling that may itself be regulated by SLR and ARF7/ARF19. Other genes have also been implicated in SLR-mediated regulation of lateral root initiation. ARF7 physically interacts with the transcription factor MYB77, and an arf7/myb77 double mutant had strongly reduced lateral root formation compared to arf7 (although it was not confirmed microscopically that the reduction was at the level of initiation) (Shin et al., 2007). The slr/iaa14 phenotype is suppressed by mutations in PICKLE, which encodes a putative chromatin remodeling factor (Fukaki et al., 2006), suggesting that chromatin remodeling is essential for SLR-induced repression of ARF7/ARF19. In summary, the downstream mechanism of SLR and ARF7/ARF19 regulation of lateral root initiation is complex and remains to be fully elucidated.

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Root Development AUX/IAA genes other than SLR (e.g. MSG2/IAA19, AXR3/IAA17, AXR5/IAA1, SHY2/IAA3 and IAA28) are involved in lateral root formation, although analyses did not determine whether initiation or a later stage in development were affected in the mutants (Leyser et al., 1996; Tian and Reed, 1999; Rogg et al., 2001; Tatematsu et al., 2004; Yang et al., 2004). Furthermore, microarray analyses predicted involvement of seven other AUX/IAA genes and two other ARF genes in lateral root initiation (Vanneste et al., 2005). In addition, other auxin signaling genes have been implicated in lateral root initiation. For example, expression of an unrelated transcriptional activator, NAC1, is rapidly induced by auxin. Over-expression of NAC1 leads to increased LRP initiation (Guo et al., 2005). The NAC1 transcript is cleaved in a process mediated by binding of the small RNA miR164. Over-expression of an altered NAC1 gene in which the miR164 complement is mutated resulted in further increases in initiation, while increased expression of miR164 reduces initiation. Interestingly, miR164 is induced by prolonged auxin exposure, potentially creating a regulatory system in which auxin first induces and then limits NAC1 expression. NAC1 protein is also unstable because it is ubiquitinated by SINAT5 E3 ligase and thereby targeted to the proteosome (Xie et al., 2002). The high turnover rate of both NAC1 protein and mRNA makes NAC1 highly responsive to auxin signaling. It is presently unclear how NAC1-mediated transcriptional activation integrates into the complex auxin signaling pathways previously described to mediate LRP initiation. A mutant in tomato, diageotropic (dgt), displays auxin insensitivity and pleiotropic phenotypes including the strong inhibition of LRP initiation (Ivanchenko et al., 2006). XP pericycle cells are 40% shorter in dgt than in the wild type, but these smaller cells are not associated with LRP initiation (Ivanchenko et al., 2006). In dgt, application of NAA leads to divisions in differentiated XP pericycle cells but no formation of founder cells, implying that DGT is essential for auxin stimulation of founder cell formation in the XP pericycle. SLR promoter activity is reduced in dgt, placing DGT upstream of SLR (Ivanchenko et al., 2006). The DGT gene encodes a cyclophilin (Oh et al., 2006). Cyclophilins are predicted to have peptidyl-proline cis-trans isomerase activity, thus potentially stabilizing folding of their target proteins. They are inhibited by cyclosporin A. Consistently, cyclosporin A inhibits the ability of tomato hypocotyls to produce lateral roots in response to exogenous auxin (Oh et al., 2006). All of the above studies have focused on the role of IAA in LRP initiation. It is important to remember that other endogenous auxins, such as IBA, might play distinct but critical roles in initiation. Exogenous IBA is a potent ¨ inducer of lateral root initiation (Ludwig-Muller, 2000; Poupart et al., 2005). Consistently, a mutant in the rice arm2 gene is defective in IBA transport, has reduced lateral root numbers, and is rescued by IBA (Chhun et al., 2005). An Arabidopsis mutant, resistant to IBA 1 (rib1) has increased lateral root formation, NPA insensitivity, and increased acropetal IBA transport (Poupart and Waddell, 2000; Poupart et al., 2005). IBA might be a direct inducer of lateral

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root initiation, or might act through its conversion to IAA (Bartel et al., 2001). Alternatively, IBA might interfere with the proteins responsible for correct IAA distribution. This idea is consistent with the demonstration that IAA transport is NPA-resistant in the rib1 mutant (Poupart et al., 2005). 4.2.3.2.2 Cell cycle regulators. Since one of the key events in LRP initiation is the stimulation of cell divisions in the XP pericycle cells, it is natural to postulate that cell cycle regulatory proteins might play a key role in initiation. However, activation of the cell cycle was shown to be necessary but not sufficient to trigger LRP initiation. The cell cycle is regulated by cyclindependent kinases (CDKs), whose activity is modulated by cyclins and other interacting proteins such as kinase inhibitor protein (Kip)-related proteins (KRPs). CDKA;1 is expressed throughout the XP pericycle, while CycB1;1 is specifically induced in lateral root founder cells, even before cell divisions occur (Ferreira et al., 1994). Constitutive expression of KRP2 reduces lateral root numbers by more than 60%, presumably by inhibiting CDKs, indicating that competence for cell division is essential for lateral root initiation (Himanen et al., 2002). However, expression of CYCB1;1 (Cyc1At) throughout the XP pericycle (under the control of the CDKA;1 (CdkA) promoter) did not lead to an increase in LRP initiation (Doerner et al., 1996), indicating that activation of the cell cycle is not sufficient for LRP initiation in XP pericycle cells. Several other studies support this idea. The dgt mutant in tomato undergoes divisions in the XP pericycle but fails to initiate LRP. Indeed, application of the auxin NAA increased the occurrence of these divisions in dgt without triggering other hallmarks of initiation (Ivanchenko et al., 2006). Similarly, expression of CYCLIN D3;1 (CYCD3;1) induced cell divisions in XP pericycle cells of the slr/IAA 14 mutant but did not repair the LRP initiation defect (Vanneste et al., 2005). Hence, the XP pericycle can be thought of as an extended meristem, retaining the capacity to proliferate but requiring additional cues to determine the fate of the daughter cells (Beeckman et al., 2001; Didonato et al., 2004). The alf4 mutant provides unique insights into regulation of cell cycle in the XP pericycle. alf4 mutant plants have little or no LRP initiation and the phenotype is only partially rescued by exogenous auxin. alf4 still expresses a cell-type-specific marker for XP pericycle cells, as well as the CDKA;1 gene that correlates with competence for cell division. In contrast, CDKB1;1, which is associated with G2-to-M transition, shows increased expression in alf4 while CYCB1;1, which marks LRP initiation sites, is not expressed in the alf4 pericycle. This suggests that alf4 pericycle cells arrest at an early stage in the cell cycle and therefore that ALF4 is necessary for the activation/maintenance of cell divisions in the pericycle. The authors suggest that ALF4 might act to prevent terminal differentiation of the XP pericycle cells. ALF4 encodes a large, nuclear-localized protein with no clear homology to other known proteins (DiDonato et al., 2004). Definition of its role awaits further experiments. It is possible that key regulators of the cell cycle are modulated by auxin. As mentioned above, KRP2 is expressed in all pericycle cells except those at the

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Root Development XP. Exposure to exogenous auxin downregulates KRP2 in XP pericycle cells (Himanen et al., 2002). In addition, systematic analysis of gene expression in the auxin signaling mutant slr/IAA14 revealed that many cell cycle genes are not induced in the presence of a stabilized SLR/IAA14 protein, including CYCA2;4 and CDKB2;1 (Vanneste et al., 2005). Based on these findings, Vanneste et al. (2005) propose that a primary role of auxin in LRP initiation is to trigger degradation of SLR/IAA14, leading to expression of specific cell cycle regulators. However, as pointed out by those authors, it seems unlikely that activation of the cell cycle alone can explain the ability of auxin to activate LRP initiation (Vanneste et al., 2005). 4.2.3.2.3 Other regulatory genes. An elegant microarray experiment compared gene expression in wild-type and slr/IAA14 plants after synchronous induction of LRP initiation (Vanneste et al., 2005). Using cluster analysis, 913 genes were defined that required SLR/IAA14 degradation for their expression in response to auxin. These genes are all candidate regulators of lateral root initiation. Interestingly, many auxin transport, synthesis and modification genes are on this list, and slr/IAA14 showed increased accumulation of auxin, suggesting complex feedback regulation in auxin signaling in LRP initiation. A similar expression profiling experiment was performed in maize. Gene expression in dissected pericycle cells from wild-type seedlings at 2.5 days after germination, and hence before LRP initiation was detectable, was compared to gene expression in similar cells from rum1, a mutant lacking initiation (Woll et al., 2005). These researchers also performed a proteomic comparison between 2.5-day-old wild-type and rum1 seedlings (Liu et al., 2006). The data sets generated in these studies should provide a good starting point for modeling initiation pathways and planning biological experiments to validate these models in plants other than Arabidopsis. 4.2.3.2.4 Other regulatory hormones. Many hormones other than auxin have been implicated in lateral root initiation (see Malamy, 2005). Often, other hormones are proposed to act by modulating auxin signaling. The hormones listed below have the most compelling experimental support for involvement in lateral root initiation, although it is highly probable that other hormones play at least indirect roles (i.e. ABA: Chen et al., 2006; De Smet et al., 2006b and ethylene: Aloni et al., 2006). 4.2.3.2.4.i Brassinosteroids. Brassinolide application induces the expression of many auxin signaling pathway components implicated in lateral root initiation, such as AXR3/IAA17, AXR2/IAA7, SLR/IAA14 and IAA28, while AXR3/IAA17 expression is decreased in a brassinosteroid biosynthesis mutant (Kim et al., 2006). Consistently, brassinolide application increases lateral root initiation, but this effect is suppressed by co-application of NPA (Bao et al., 2004).

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4.2.3.2.4.ii Cytokinin. Cytokinin has long been reported to inhibit lateral root formation (Aloni et al., 2006 and references therein), perhaps by antagonizing the action of auxin (Coenen and Lomax, 1997; Lloret and Casero, 2002). Indeed, in tissue culture, the ratio of auxins to cytokinins determines whether shoots or roots are produced from undifferentiated callus tissue (Skoog and Miller, 1957). Cytokinin is primarily produced in the root cap, and it has long been suggested to inhibit the initiation of lateral roots close to the root tip (Lloret and Casero, 2002). Exogenous cytokinin application inhibits lateral root initiation and counteracts the stimulatory effect of exogenous auxin on lateral root initiation (Li et al., 2006). Furthermore, transgenic Arabidopsis and rice plants with decreased cytokinin production form more lateral roots closer to the root tip (Werner et al., 2003; Debi et al., 2005). Recently, Li et al. (2006) demonstrated that cytokinin blocks the G2-to-M transition in the LRP founder cells. 4.2.3.2.4.iii Nitric oxide. In tomato, nitric oxide (NO) appears to mimic the affects of auxin on LRP initiation. NO released from sodium nitroprusside increases the formation of LRP during normal development. This effect is reversed by application of the NO scavenger 2-(4-carboxyphenyl)-4,4,5,5tetramethylimidazole-1-oxyl-3-oxide (CPTIO) (Correa-Aragunde et al., 2006). Furthermore, CPTIO alone strongly inhibits LRP formation during normal development or in response to exogenous auxin application. The role of NO seems to be quite early in development, as in a synchronized system CPTIO only represses LRP formation when added 0–2 days after NAA induction, but not at later time points. NO, like auxin, induces transcription of CYCD3 and downregulates the expression of KRP2. More research is needed to uncover the physiological significance of these intriguing findings. 4.2.3.2.5 Environmental signals. Many environmental signals, including phosphate, water and nitrate have been shown to impact LRP initiation (for review, see Malamy, 2005). In one case the molecular mechanism of environmental regulation has been characterized in detail. When Arabidopsis plants are grown on nutrient media containing high ratios of carbon:nitrogen and limiting concentrations of nitrogen (0.1–0.01 mM), initiation is strongly repressed (Malamy and Ryan, 2001; Little et al., 2005). This phenomenon might be related to auxin transport, as GUS staining in the DR5 reporter line was strong in the hypocotyls of plants grown under the repressive conditions, suggesting that there might be a bottleneck in shoot-to-root transport (Malamy and Ryan, 2001). Inhibition of initiation on these repressive media requires the high-affinity nitrate transporter NRT2.1, as an nrt2.1 mutant (lin1) restored LRP initiation. The increased LRP initiation phenotype was demonstrated to be independent of nitrate uptake, leading to the suggestion that NRT2.1 acts as a nitrate sensor or signal transducer to coordinate initiation with nutritional cues (Little et al., 2005). Interestingly, Remans et al. (2006) reached the same conclusion to explain a different finding: under experimental conditions

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Root Development in which plants were transferred from high to low nitrate media, mutants in NRT2.1 showed decreased LRP initiation that could not be accounted for by a decrease in nitrate uptake. Therefore, there is a consensus on the role of the nitrate transporter NRT2.1 in uptake-independent regulation of LRP initiation, even though it appears that in different assays NRT2.1 can act either as a repressor or promoter. 4.2.3.3 Regulation of LRP development and patterning As a key player in cell division and expansion, it is not surprising to find that auxin plays a role in the development of the LRP as well as its initiation. The auxin importer AUX1 is expressed in Stage I LRP, and the aux1 mutant is delayed in the transition from Stage I to Stage II (Marchant et al., 2002). This finding suggests that AUX1 might mediate auxin uptake in the lateral root founder cells to facilitate the first periclinal divisions (Marchant et al., 2002). Auxin also appears to be involved in establishing tissue and cell polarities essential for morphogenesis in the LRP. IAA localization was assessed in the developing LRP using the DR5 reporter line (Benkov´a et al., 2003). As the LRP develops, auxin becomes progressively localized to the LRP tip, with highest expression in the columella precursors (as in the primary root tip). This pattern was seen in all LRP that developed normally; however, occasionally LRPs were seen that did not develop normally, and in these cases the DR5 staining pattern was disrupted. Interestingly, a recent report showed that a dramatic reduction of auxin (as assessed by DR5 activity) in the LRP in the mdr4 mutant had no discernible effect on LRP development (Wu et al., 2007). This suggests that a very low level of auxin, below the sensitivity of GUS staining, is sufficient to sustain correct LRP development. Correct distribution of auxin in the developing LRP appears to be mediated by the PIN family of proteins. PIN1, PIN2, PIN3, PIN4, PIN6 and PIN7 efflux carriers showed distinct but overlapping patterns of expression in the LRP (Benkov´a et al., 2003). Mutation or ectopic expression of PIN1 led to a greater percentage of LRP with retarded or aberrant development. These LRP lacked the tip-to-base gradient of auxin (visualized using the DR5 promoter). In contrast, LRP in the pin2 mutant developed more rapidly, and demonstrated an increased auxin gradient. In double and triple pin mutants, NAA-induced LRP formation was aberrant or completely absent, although divisions were seen in the pericycle layer. Hence, multiple PIN genes regulate auxin distribution in the LRP and correct distribution is essential for LRP development. Auxin distribution during LRP development seems to be a dynamic process, as PIN1 was shown to re-localize from the anticlinal sides of founder cells (Stage I) to the periclinal of cells in later developmental stages, correlating with the progressive localization of auxin to the LRP tip. Indeed, when PIN1 re-localization was prevented by addition of BFA or NPA, no auxin gradient was established (Benkov´a et al., 2003). Combined analyses of PIN expression patterns and DR5 staining patterns lead to the ‘fountain model’, in which auxin in the LRP is transported from

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the primary root vasculature to the LRP tip via the interior of the LRP and then retrieved by PIN2 in the LRP epidermal cells (Benkov´a et al., 2003). This idea is consistent with the demonstration of an ‘auxin transport loop’ that has been postulated to maintain meristem size in the primary root tip (Blilou et al., 2005). Cell identities within the LRP are established very early (see above), and patterning precedes the emergence of the LRP or the activation of the apical meristem (Malamy and Benfey, 1997a, 1997b). It is worth noting that the accumulation of auxin in the LRP tip occurs while patterning takes place. An auxin maximum has been shown to be sufficient to induce patterning of meristematic tissues, such that cells immediately adjacent to the maxima re-differentiate to form endodermal, cortex, epidermal and cap cells (Sabatini et al., 1999). Hence, a mechanism that explains the localization of an auxin maximum in the developing primordia might be sufficient to explain the concomitant patterning of the developing LRP. Few genes have been found that are specific regulators of LRP development. The newly identified auxin-induced PUCHI gene is therefore important, as the puchi mutant is disrupted in the early stages of cell division and expansion in the LRP. PUCHI encodes an AP2/EREBP family transcription factor. The mechanism of PUCHI’s action awaits further studies. 4.2.3.4 Regulation of LRP emergence and meristem activation LRP emergence is the process by which the LRP leaves the parent root by expansion of basal cells (Malamy and Benfey, 1997b). For a lateral root to form, the lateral root apical meristem must then become active. Since both of these processes are essential for an LRP to turn into a lateral root, emergence is often expressed as the total number of lateral roots/total number of LRP initiation events × 100 (i.e. x% of all initiation events result in the formation of a lateral root). We will use this definition herein, unless specific anatomical events are clearly specified. Meristem activation is detected in Arabidopsis by increases in cells near the tip of the emerged LRP. The activated meristem then becomes responsible for elongation of the new lateral root. 4.2.3.4.1 Regulatory hormones 4.2.3.4.1.i Auxin. It is clear that auxin plays a critical role in LRP emergence and meristem activation. The Arabidopsis alf3 mutant forms an apparently normal LRP that emerges from the parent root but fails to develop further. This mutant is rescued by a continuous supply of IAA or the IAA precursor indole. Shifting seedlings away from the IAA source at various times demonstrated that indole was required both for activation of meristems and meristem maintenance. Interestingly, a 3–5 cell layer LRP in an explant can develop into a lateral root in culture in the absence of auxin (Laskowski et al., 1995), implying that the auxin required for emergence and meristem activation must be produced at or very near to the LRP.

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Root Development Auxin transport has been strongly implicated in regulation of LRP emergence. Aerial tissues are essential for LRP emergence in young (7-day-old) seedlings (Bhalerao et al., 2002) and stm1, which lacks the aerial tissues that are the major sources of shoot auxin, has an altered pattern of LRP emergence (Casimiro et al., 2001). Results consistent with this idea were obtained by growing seedlings on growth media that represses all LRP emergence (Deak and Malamy, 2005). Upon transfer to permissive conditions, the preformed LRP rapidly emerged in the absence but not in the presence of NPA or TIBA (Deak and Malamy, 2005), suggesting that NPA-sensitive transport of auxin, either from the shoot or within the LRP, is essential for LRP emergence. Wu et al. (2007) showed that mutation of the MDR1 gene, which specifically regulates acropetal but not basipetal auxin transport, results in emerged LRPs that fail to elongate. In contrast, mutations in the MRD4 gene, which specifically regulates basipetal auxin transport, had no effect on lateral root formation in this study (although numbers of lateral roots were altered in other mdr4 alleles in other studies (Santelia et al., 2005; Terasaka et al., 2005)). This supports the hypothesis that auxin from the shoot plays a central role in the regulation of LRP emergence and meristem activation. Interestingly, Bhalerao et al. (2002) also showed that aerial tissues are no longer essential for emergence in slightly older seedlings. The time window in which LRP emergence is shoot-dependent coincides with an observed pulse of auxin, apparently moving from shoot to root. This observation lead to a model in which shoot auxin is essential for LRP emergence early in development but emergence subsequently becomes root autonomous. 4.2.3.4.1.ii ABA. Another hormone that has been strongly implicated in the regulation of LRP emergence is ABA. In wild-type plants, emergence of LRP is strongly inhibited by the addition of osmotica to growth media (Deak and Malamy, 2005). In contrast, two ABA-deficient mutants, aba2-1 and aba3-1, were able to overcome the osmotic inhibition of LRP emergence (Deak and Malamy, 2005; Xiong et al., 2006). Interestingly, the ABA-deficient mutants also showed a higher rate of LRP emergence than wild-type plants in the absence of osmotic stress, suggesting that ABA might be involved in repressing the emergence of LRP under all growth conditions. It is possible that increases in ABA in response to osmotic stress might serve to coordinate osmotic conditions with lateral root formation. Exogenous auxin overcame the osmotic repression of LRP emergence, suggesting that ABA and auxin signaling may have antagonistic effects on this process. High concentrations of exogenous nitrate have also been shown to repress LRP development immediately after emergence, and this repression is significantly reduced in several ABA-deficient mutants and some but not all ABA-insensitive mutants (Signora et al., 2001). Consistent with a role for ABA in meristem activation, exogenous application of ABA caused LRP to arrest development immediately following emergence (De Smet et al., 2003). In the arrested LRP there was a marked reduction in expression from the

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auxin-responsive promoter DR5, initially suggesting that ABA might block meristem activation by downregulation of auxin accumulation or response. However, neither increases in endogenous or exogenous auxin could overcome the arrest, while ABA-grown plants were still able to respond to exogenous auxin in terms of primary root elongation and DR5 promoter activation, implying that the ABA-mediated arrest postemergence was unrelated to auxin signaling. The ABA signaling pathways that are involved in the regulation of lateral root formation are unclear. Recently, FCA has been proposed to act as an ABA receptor. Interestingly, fca mutants showed little reduction of lateral root formation in response to exogenous ABA, as compared to a strong reduction in wild-type plants (Razem et al., 2006). Since the development of LRP was not closely examined in these mutants, the stage at which FCA mediates ABA effects is still unknown (De Smet et al., 2006b). Some of the genes whose mutation leads to ABA insensitivity are expressed in the LRP (i.e. ABI3 and ABI5), and mutations in ABA signaling components such as era1 and abi8 show defects in lateral root formation. However, the significance of these results is not yet clear (Brocard et al., 2002; Brady et al., 2003; De Smet et al., 2006b). Osmotic or ABA repression of LRP emergence was compromised in abi1-1, but not in abi2, abi3 or abi5 (Xiong et al., 2006). Hormonal signaling might not have the same developmental consequences in all plants. Indeed, in the latd mutant isolated in Medicago, LRPs form and emerge from the parent root but then arrest (reminiscent of the Arabidopsis alf3 mutant). Interestingly, the latd mutant phenotype could not be rescued by auxin, but was completely rescued by ABA (Liang et al., 2007). The promotive effect of exogenous ABA on LRP meristem activation is opposite to that predicted by experiments in Arabidopsis, and suggests distinct roles for ABA in Medicago and Arabidopsis. 4.2.3.4.2 Regulatory genes. Very few genes have been definitively shown to affect LRP emergence, and in no cases has the regulatory mechanism been demonstrated. The LATERAL ROOT DEVELOPMENT 2 (LRD2) gene appears to act as a repressor of LRP emergence based on the ability of the lrd2 mutant to overcome osmotic conditions that repress LRP emergence in wild type (Deak and Malamy, 2005). A similar screening approach lead to the isolation of the drought inhibition of lateral root growth 3 (dig3) mutant (Xiong et al., 2006), that is insensitive to ABA or osmotic inhibition of LRP emergence. Cloning of the LRD2 revealed that it is allelic to LACS2, a long chain acyl coA synthetase (Macgregor et al., 2008). The role of LACS2 in LRP emergence is described further in the following section. The cloning of the DIG3 gene has not yet been reported. The CEGENDUO (CEG) gene encodes a novel auxin-inducible F-box protein. Mutations in this gene do not affect initiation, but do affect total lateral root number, suggesting that this gene acts at the emergence/meristem activation stage (Dong et al., 2006).

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Root Development Cells overlying emerging LRP separate, suggesting that degradation of the pectin-rich middle lamellae is associated with LRP emergence (Laskowski et al., 2006). Indeed, the pectin in LRP is methylated while that in the overlying cells is demethylated, suggesting that there may be different susceptibilities to degratory enzymes such as pectate lyases (PLs). Microarray studies demonstrate that specific PL genes (PLA1 and PLA2) are auxin induced, that the timing of auxin-induction is consistent with a role in LRP emergence, and that the SLR auxin-response pathway is essential for induction of these genes (Laskowski et al., 2006). Closely related plants often show phenotypic variability. In Arabidopsis, different ecotypes have different root system morphologies. In the case of the ecotypes Columbia (Col) and Landsberg erecta (Ler), the differences are due to increased LRP initiation and emergence in Ler (Fitz Gerald et al., 2006). Two strong QTLs have been identified that account for a large proportion of the variation in root system development between Col and Ler. Analysis of near-isogenic lines in which each QTL region was isolated in the opposite genotypic background demonstrated that both QTLs play a role in LRP initiation and emergence. A QTL for the variation in lateral root number/density between the Bay-0 and Shadara ecotypes was also mapped, although the developmental stage affected by the QTL was not examined (Loudet et al., 2005). Although QTL studies have been done in many plants, Arabidopsis offers the most straightforward opportunity for cloning the underlying genes. It will be very exciting to see whether the gene variability that leads to phenotypic variability in wild plant can be anchored to known signaling pathways. 4.2.3.4.3 Environmental cues. Environmental cues appear to play important roles in emergence, distinct from the environmental regulation of lateral root initiation. Regulation of LRP emergence might be an extremely effective way to optimize the distribution of roots in the soil. If ‘dormant’ LRPs were formed throughout the root system, the plant could respond to the appearance of a water or nutrient supply by rapidly activating LRP in that region. 4.2.3.4.3.i Osmotic stress. Mild osmotic stress represses the emergence of LRP and subsequent activation of the lateral root meristems. This was shown using equimolar concentrations of mannitol, NaCl or nitrogen salts in the media of culture-grown plants (Deak and Malamy, 2005). As stated above, osmotic repression of LRP emergence requires ABA, because in ABA-deficient mutants the percentage of LRP that emerge is significantly increased. Osmotic repression could be overcome by addition of IAA or NAA. ABA and auxin appear to have opposing effects on LRP emergence, suggesting that osmotic stress affects emergence by altering the balance between ABA and auxin signaling. Since osmotic stress is known to lead to increases in ABA accumulation, this offers one possible mechanism for osmotic repression of LRP emergence (Deak and Malamy, 2005).

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New data have lead to the surprising model that the effects of osmotic stress on LRP emergence in plants grown in laboratory experiments reflects altered access to sugars in the growth media. Growth on osmotic stress conditions in culture makes leaves less permeable to external substances, as evidenced by increased permeability to a small molecule dye (Macgregor et al., 2008). Further experiments revealed that physical contact between leaves and growth media is essential for LRP emergence in this system, and that contact between leaves and sucrose (a component of the media) is sufficient to stimulate LRP emergence. This suggests that osmotic stress represses LRP emergence in this system by reducing access of the aerial tissues to sucrose (Macgregor et al., unpublished). This model is supported by the recent cloning of lrd2 as LACS2, which encodes a long chain acyl coA synthetase essential for leaf cutin formation. The lrd2/lacs2 mutant has highly permeable leaves; furthermore, other mutants in cutin formation resemble lrd2 in that they show a dramatic increase in LRP emergence (Macgregor et al., unpublished). In sum, these results suggest that osmotic stress imposed by high levels of nitrogen salts or mannitol represses LRP emergence in culture only indirectly, through its effect on access of the leaves to media sucrose. Hence, the true effect of osmotic stress on lateral root formation in Arabidopsis has yet to be described. 4.2.3.4.3.ii Sugar/light/photosynthate. The above finding raises the possibility that sucrose or a sucrose metabolite is an endogenous regulator of LRP emergence. The effect of exogenous sucrose on lateral root formation was previously demonstrated (Takahashi et al., 2003; Cluis et al., 2004; Karthikeyan et al., 2007). Significantly, in a screen for mutants that overcome osmotic repression of LRP emergence, Qi et al. (2007) isolated a mutant in AtCyt-INV1, a cytoplasmic, neutral, sucrose-specific invertase. Invertases cleave sucrose to form glucose and fructose, and the mutant plant showed an increase in endogenous sucrose and a decrease in endogenous glucose. Finally, nutrient deficiencies (phosphorus and nitrogen) increase the root:shoot ratio and alter root system architecture in many plants, and these responses are accompanied by accumulation of sugars and starches in leaves and altered carbon allocation to the root (Hermans et al., 2006). Consistent with the above studies of the roles of sugars in LRP emergence, there is considerable precedent for the idea that increases in photosynthate leads to increased lateral root formation. Increased ambient CO2 levels lead to increased lateral root formation (Crookshanks et al., 1998). Light stimulates lateral root formation (Reed et al., 1998; Freixes et al., 2002 and references therein) and phytochromes A, B, D and E also appear to play a role, perhaps by affecting the transport of IAA from shoot to root (Salisbury et al., 2007), although the specific effects on LRP initiation and emergence were not distinguished in these studies. HY5 and COP1, components of the light signaling, were also specifically shown to mediate the LRP emergence (Oyama et al., 1997; Cluis et al., 2004). Hence, light, photosynthesis and sugar accumulation in the leaves are probably an important determinant of lateral root

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Root Development formation/LRP emergence, through alterations in carbon/energy partitioning and/or sugar signaling pathways. This should be a fertile area for further research. 4.2.3.4.3.iii Nitrate. High levels of nitrate have also been shown to repress LRP meristem activation in an ABA-dependent fashion. This repression is reversible (Zhang et al., 1999). Sensitivity to nitrate was increased in a nitrate reductase mutant, suggesting that the nitrate itself rather than nitrate metabolites is responsible for repression of these meristems (Zhang et al., 1999). The fact that nitrate inhibition of LRP emergence was not completely abolished in ABA biosynthesis and signaling mutants led the authors to hypothesize that there are both ABA-dependent and ABA-independent pathways involved in this regulatory process. 4.2.3.4.3.iv Phosphate. Phosphate (P) deficiency stimulates LRP emergence while decreasing growth of the primary root (Lopez-Bucio et al., 2005; Nacry et al., 2005). The resulting redistribution of root mass is advantageous for P uptake because P is preferentially found in shallow soils. Low P responses are altered by exogenous auxin, auxin transport inhibitors and in auxin response mutants, suggesting an interaction of P responses with auxin signaling (Lopez-Bucio et al., 2002; Nacry et al., 2005). Low P stimulation of LRP emergence is accompanied by increased GUS staining in the LRP in the DR5 auxin response reporter line and decreased GUS staining in the primary root tip (Lopez-Bucio et al., 2005; Nacry et al., 2005). These findings suggest that low P responses may be mediated by a redistribution of auxin. Indeed, a mutant in low P response was found to carry a mutated allele of BIG/TIR3, a gene encoding a protein known to be essential for polar auxin transport (Lopez-Bucio et al., 2005). Interestingly, growth on low P was able to rescue the arrested LRP in the alf3 mutant, mimicking the rescue by addition of exogenous auxin (Nacry et al., 2005). Together, these results suggest that overaccumulation of auxin in the LRP of plants grown under low P, perhaps due to modulations in auxin transport, is responsible for increases in LRP emergence. 4.2.3.5 Regulation of meristem maintenance The PLETHORA (PLT) genes are AP2 transcription factors that have been shown to play a critical role in organization of the primary root tip. They are localized by the PIN proteins, and in turn regulate expression of PIN genes in the primary root tip. plt1/plt2 double mutants have reduced meristem size in both primary and lateral roots, which terminate growth shortly after emergence. However, total lateral root numbers are increased, indicating that the PLT genes are not essential for LRP initiation, development or emergence, but are required for meristem maintenance (Aida et al., 2004). Similar results are seen with other genes essential for meristem maintenance in the primary root, such as SCARECROW and SHORTROOT, suggesting that meristem maintenance programs are similar in all root types. The

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Pi deficiency response 2 (pdr2) mutant produces short primary and lateral roots with arrested meristems under low P conditions (Ticconi et al., 2004), again suggesting that meristem maintenance in primary and lateral roots are coordinately regulated. The fact that the pdr2 mutant phenotype is only seen under low P conditions suggests that there are genes that coordinate meristem activity with environmental signals in addition to the intrinsic regulators described above. Auxin, required in all previously described steps of lateral root formation, is also needed for maintenance of the apical meristem. Although most of the work on this topic focuses on the primary root meristem, Celenza et al. (1995) showed that the lateral roots of the indole-requiring alf3-1 mutant arrest when indole is removed, demonstrating a continuing need for this hormone. The death of alf3-1 LRP, which lack active apical meristems, further suggests that cell death is a default process at the root tip in the absence of an active apical meristem. Similar cell death is observed in arrested meristems of pdr2 (Ticconi et al., 2004).

4.3 How are the number and placement of lateral roots determined? Lateral roots are formed along the length of the seedling primary root. More lateral roots are formed from these lateral roots in turn, such that the roots in a complex root system are often referred to as secondary, tertiary, quaternary and so on. Having discussed the events that occur during the formation of a single lateral root, we now turn to the problem of how lateral root formation is regulated at the root system level. This includes the questions of site selection, spacing between lateral roots, and modulation of whole root architecture. How many sites will be selected? Will the root system be highly branched or un-branched? Will the lateral roots be evenly distributed in the root system or will the root system be primarily shallow or deep?

4.3.1

Anatomical observations

4.3.1.1 Radial position As described above, in most higher plants lateral roots originate from the pericycle, which is the outmost layer of cells in the root stele, adjacent to the XPs (Peterson and Peterson, 1986; Charleton, 1996). In plants with a small number of xylem arms (diarch, triarch, tetrarch and pentarch symmetry), lateral roots generally initiate at or immediately adjacent to the poles (e.g. Arabidopsis, tomato, onion and pea), although there are exceptions where LRPs form at the protophloem poles (e.g. carrot). In monocots, where there are numerous XPs, lateral roots generally arise adjacent to the protophloem between two XPs (e.g. maize) (Charleton, 1986; Torrey, 1986).

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Root Development 4.3.1.2 Longitudinal position and spacing The earliest divisions in LRP initiation are seen very near to the root tip, suggesting that pericycle cells become LRP founder cells very rapidly after their creation at the apical meristem. Dubrovsky et al. (2006) estimate the developmental window in which initiation takes place in the life of a pericycle cell to be very short, 10–16 hours at the most in Arabidopsis. Regular initiation of lateral roots as the primary root grows would imply a highly predictable pattern in the position of lateral root formation. Indeed, overall there is a clear trend of lateral roots appearing in an acropetal sequence, creating an inverted Christmas tree pattern in many (but not all) seedling root systems (Torrey, 1986; Charleton, 1996). However, longitudinal positioning of lateral roots in most plants is imprecise (Torrey, 1986) and in Arabidopsis the position of lateral roots is extremely hard to predict. This suggests either that LRP initiation does not occur in a strictly acropetal sequence (i.e. new LRPs form between existing LRP), or that LRPs do not develop into lateral roots in a predictable order. In many plants, lateral root formation between existing lateral roots has been reported (Charleton, 1996), but initiation and emergence were not distinguished in most cases. A detailed examination of Arabidopsis seedlings from 7 to 15 days after germination revealed that no new LRPs were formed between existing LRP, implying a strictly acropetal pattern of initiation (Dubrovsky et al., 2006). However, the distance between LRP was quite variable and did not follow any discernible pattern, although significant differences in the average inter-lateral distances between closely related Arabidopsis accessions suggest the existence of some intrinsic patterning. In contrast, De Smet et al. (2007) found spacing between LRP to be relatively constant. LRP emergence is even more variable than initiation. In Arabidopsis, less than 40% of LRP form lateral roots in 2 weeks under certain experimental conditions (for example, Deak and Malamy, 2005; Dubrovsky et al., 2006). When emergence was observed, it did not follow an acropetal (or any other discernable) pattern (Dubrovsky et al., 2006). It is important to note that although the overall pattern of LRP initiation or emergence may appear imprecise, several studies show that within a single vertical file of pericycle cells (all in the same radial relationship to the xylem), spacing may be extremely regular (Charleton, 1996). It is possible that such trends have been missed in other studies, as it is often difficult to group LRP into common files. Longitudinal spacing of LRP appears to follow certain clear trends. LRP do not generally initiate in close proximity to each other or to lateral roots. In Arabidopsis, Dubrovsky et al. (2006) confirmed a minimum distance between successive LRPs, and measured this distance as 5–14 times the diameter of a single LRP. However, as stated above, no consistent pattern of inter-LRP distances could be discerned (Dubrovsky et al., 2006). Arabidopsis lateral root initiations tend to alternate between the opposing XP pericycle files (Dubrovsky et al., 2006). This trend was previously observed in other plants (Charleton, 1996). Finally, in many plants there is a fixed minimum distance between the

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parent root tip and the most apical LRP (Charleton, 1996; Dubrovsky et al., 2006). In Arabidopsis, the first lateral root is initiated approximately 1.4 mm from the root tip (Beeckman et al., 2001). 4.3.1.3 Overall architecture There is tremendous diversity in developmental patterns of root systems among plant species and varieties. Root systems can be highly branched or un-branched, shallow or deep, and can vary in their developmental responses to environmental cues. Differences in root system architecture can be the result of variation in the number or distribution of lateral root initiation events, the rates of development of LRP into lateral roots, lateral root elongation rates, lateral root tropisms and all these parameters in the context of secondary and tertiary lateral roots (Malamy, 2005). 4.3.2

Regulation of the sites of lateral root formation

One of the primary questions in studying root system architecture is whether the sites of lateral root formation are predetermined or are selected during growth and development of the root system. Using the definitions set forth in the introduction, we can say that ‘intrinsic’ pathways would result in predetermination of the sites of LRP and lateral root formation. The involvement of environmental signaling, in contrast, might be expected to cause variations in patterns depending on growth conditions. The interplay of intrinsic and environmental signaling pathways is considered below for each proposed regulatory event. 4.3.2.1 Regulation of radial position The radial position of lateral roots is extremely regular in plants and does not change with growth conditions (Charleton, 1996) or with the addition of stimulatory concentrations of auxin (Himanen et al., 2004). The consistent development of LRP in relation to XPs suggests that intrinsic pathways regulate the radial position of LRP/lateral roots. 4.3.2.2 Regulation of longitudinal position and spacing The imprecise placement of lateral roots has led to the suggestion that a combination of intrinsic pathways and environmental response pathways influence longitudinal positioning (Charleton, 1996). The demonstration that environmental signals can influence both LRP initiation and emergence is consistent with this idea (see above). However, neither the intrinsic nor the environmental response mechanisms that ultimately lead to lateral root site selection are well understood. There is minimum distance that is maintained between LRP/lateral roots. This might be due to inhibitory substances coming from the LRP themselves or to the diminished nutrient or hormone supply in the immediate vicinity of an LRP (Torrey, 1986; De Smet et al., 2006a). In either case, this aspect of

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Root Development spacing should be intrinsic to the system, and therefore could be considered to be part of the predetermined pattern of spacing. Consistent with the idea that spacing is regulated by the LRP themselves, the alf3-1 mutant forms LRP almost on top of each other. alf3-1 LRPs arrest, the cells die shortly after emergence from the parent root, and apical meristems are never activated (Celenza et al., 1995). The phenotype suggests that spacing is maintained by an inhibitory substance that arises only from viable LRP. The inhibition of LRP formation near the primary root tip might reflect the fact that pericycle cells in this region have not reached a maturation state at which they are competent to become founder cells. However, the apparent initiation of new lateral roots at the cut site after root tip decapitation supports either the presence of an inhibitory substance or the lack of a stimulatory substance at the root tip, or both (Torrey, 1986). It is unclear whether the rules for the distance from the root tip to the first LRP is intrinsic to the plant or can be altered in response to growth conditions. The nature of the putative root tip inhibitor is also not known, although the formation of basal lateral roots after tip excision is often suggested to result from accumulation of auxin (Torrey, 1986). This would suggest that the root tip either acts to prevent auxin accumulation, or produces a signal that antagonizes the auxin induction of lateral roots. Recently, an exciting new model was proposed to explain intrinsic mechanisms for site selection and spacing in the Arabidopsis root system (De Smet et al., 2007). Arabidopsis primary roots grow in a regular waving pattern on the surface of the growth media on vertically oriented Petri dishes. LRPs are preferentially formed at the tip of the convex side of each wave. Given that the primary root grows at a constant rate, this means that in this assay lateral roots form with a reasonably regular periodicity of 15 hours. Physiological phenomena with similar periodicity thus became candidates to explain lateral root positioning. Interestingly, GUS staining in the DR5 auxin reporter line revealed an oscillating auxin response maximum in the basal region of the root meristematic zone (basal meristem) with a periodicity of 15 hours. The basal meristem is where specification of LRP would be expected to occur. Indeed, marking the basal meristem with dye particles in live roots during the maxima in the auxin oscillations lead to a significant correlation between subsequently formed LRP and dye particles. These results suggest a tight correlation between auxin in the basal meristem and lateral root initiation. If auxin oscillations, even less precise than those investigated here, really do occur during primary root growth, these findings would offer a compelling mechanism to explain the longitudinal positioning of lateral roots. The regulation of lateral root spacing by gravistimulation provides a good example of the interplay between intrinsic and environmental response pathways. Gravistimulation results in lateral root initiation at the convex side of the zone responding to the cue (Lucas et al., 2008) (explaining the placement of LRP in ‘waving’ roots described above (De Smet et al., 2007)). Altering

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the time period between changes in the gravity vector revealed that there are intrinsic maximum and minimum intervals between initiation events, such that LRPs never form immediately adjacent to each other even if the interval between gravistimulations is very short, and new LRPs initiate in between those induced by gravistimulation if the interval is too long. Hence, an intrinsic patterning of LRP spacing is observed, and this intrinsic pattern can be modified by environmental cues such as gravity. Perhaps because of the complex combination of environmental and intrinsic stimulation of LRP initiation and intrinsic constraints on LRP initiation, the total number of LRP initiations was found to be unrelated to the number of gravistimulations. However, by making the assumption that gravistimulation leads to a local maximum in auxin or auxin response and that LRP initiation leads to a depletion of this maximum, the authors were able to design a mathematical model of the dynamic changes in auxin levels that correctly predicted LRP initiation (Lucas et al., 2008). The question of why LRP initiation occurs on the opposite side of the root than the predicted auxin accumulation following gravistimulation remains to be answered. Environmental cues do not always have global effects on lateral root spacing. For example, responses to nutritional cues show that there can be fine spatial regulation of lateral root site selection. When barley roots were exposed to unevenly distributed nutrients, LRP initiation increased in response to localized P, nitrate, or ammonium, but only in the region where the nutrient levels were elevated (Drew, 1975). An increase in local lateral root density in a nitrate- or P-rich patch in Arabidopsis was also seen in some but not all cases (Zhang and Forde, 1998; Linkohr et al., 2002). 4.3.3

Regulation of overall architecture

Auxin is a critical regulator of many steps in the lateral root formation process and is believed to be a long-distance signal, arriving at LRP founder cells and LRP either from the shoot or the root tip (see above). Therefore, on a root system level, it is plausible to imagine that alterations in the amount of auxin produced at the source or transported to the root system might affect overall levels of LRP initiation, development and/or emergence. It is also possible that the alteration in architecture in response to environmental cues represents changes in these parameters. For example, P deficits inhibit primary root elongation and promote lateral root formation, creating a shallower, wider root system. These changes appear to be accompanied by a redistribution of auxin from the primary root tip to the LRP (Lopez-Bucio et al., 2003, see also Chapter 11). It is important to note that root architecture is also strongly influenced by rates of lateral root elongation (Malamy, 2005). Although not discussed here, lateral root elongation is promoted by localized nitrate patches (Drew, 1975; Zhang et al., 1999), by nitrogen deficiency (Remans et al., 2006), by P (Drew, 1975) and by P deficiency (Lopez-Bucio et al., 2003).

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4.4

Agricultural importance of lateral root formation

The root system is essential for the uptake of water and nutrients from the soil, for micorhizzal interactions, for nodulation and nitrogen fixation in legumes, and for mechanical support of the plant. Hence, the critical role that root system architecture plays in crop yield is not unexpected. Indeed, only root system traits showed a consistent positive contribution to yield in simulation studies of the effects of altering various plant traits over 20 seasons (Sinclair and Muchow, 2001). 4.4.1

Root systems and drought stress

Drought is a major factor limiting yield in crop plants. Estimated losses in yield due to drought stress are over 50%, and 70–80% of the world’s water resource is consumed in irrigation (Chaves and Oliveira, 2004; Condon et al., 2004). In developing countries, where rainfall is often irregular and access to irrigation may be limited, the problem of yield loss due to drought stress is most severe. Furthermore, current predictions are that the severity and duration of drought stress will increase globally (Karl and Trenberth, 2003), and water available for farming in the United States and elsewhere will decrease in the coming years due to climate changes and increased population pressure (Barnett et al., 2004; Service, 2004). Despite tremendous efforts, the development of high-yielding crop varieties suited to water-limited environments has been slow due to the complexity and low heritability of the trait. At present, most of the highest-yield crop varieties do not even survive in harsh environments, severely limiting the economic potential of many developing world nations (Araus et al., 2002; Chaves and Oliveira, 2004). Increased root system size, through increased lateral root formation and/or growth, extends the area explored by the root system. Increased density of shallow roots allows plants to obtain surface water that is subject to rapid evaporation, while a root system that penetrates to greater depths can access water deep in the soil profile. Indeed, studies confirm that increases in root system growth and development play a critical role in yield under drought stress (Champoux et al., 1995; Ali et al., 2000; Shen et al., 2001; Tuberosa et al., 2002, 2003). This correlation has been extensively studied in maize and rice. In maize, rooting density and depth have been demonstrated to be of primary importance in plant water status and yield under drought stress (Araus et al., 2002; Tuberosa et al., 2003 and references therein). There was a significant overlap of QTL’s influencing yield and those influencing root traits (Tuberosa et al., 2002, 2003). In tropical maize selected for higher yields in stress-prone environments root system biomass was shifted to deeper soil layers (Bruce et al., 2002). In rice, similar trends have been reported. Rice can be divided into upland and lowland varieties. Upland rice varieties, adapted for growth in non-flooded areas, generally have deeper root systems. In addition, upland rice varieties with the deepest root systems perform

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better under prolonged drought stress (Champoux et al., 1995; Fukai and Cooper, 1995; Kondo et al., 1999; Price et al., 2002b, 2002c) and increased root depth, density and number positively correlates with water acquisition (Price et al., 2002c). Even in lowland rice planted into flooded soil, studies show that early development of a vigorous root system has a positive effect on plant growth during subsequent water stress, with deeper roots leading to improved water extraction (Kamoshita et al., 2002). Basal root thickness and maximum root length in mature rice plants positively correlates with both vegetative and reproductive stage drought resistance, again emphasizing the adaptive nature of a deep root system (Champoux et al., 1995; Pantuwan et al., 1997; Li et al., 2005a). Root depth has also been positively correlated with yield in soybean (Cortes and Sinclair, 1986), soybean cultivars with a greater root density appeared to have increased water uptake (Carter et al., 1999), and a soybean line with an unusually fibrous and extensive surface root system was shown to be more drought tolerant (Pantalone et al., 1999 and references therein). Large and/or deep root systems have similarly been correlated with water-stress tolerance in other plants, including coffee (Pinheiro et al., 2005) and bean (Ho et al., 2004 and references therein). These consistent correlations allow us to hypothesize that in any given plant and environment, root system size contributes positively to water uptake and drought-stress tolerance. Despite the wealth of studies showing positive correlations of root traits with drought avoidance and yield, it has been difficult to establish an aspect of root system development that is consistently beneficial. In both maize and rice there is a lack of correlation or even a negative correlation between specific root system traits and yield under drought stress (Bruce et al., 2002; Price et al., 2002a; Lilley and Fukai, 1994a, 1994b; Li et al., 2005a). Similarly, QTL identified for root traits and drought avoidance did not overlap in one study (Price et al., 2002a), whereas in another study root depth and drought avoidance were correlated positively but root number and drought avoidance negatively (Li et al., 2005b). One explanation for these contradictory findings is that each study examines a different rice population. In each genetic background, increases in root system number or depth might represent very different physiological and regulatory events. For example, increases in root system size might sometimes represent a reallocation of resources at the expense of yield, but in other cases might not. A further complexity is that root system development is highly plastic and responsive to environmental cues (Cooper and Somrith, 1997; Cooper et al., 1999a, 1999b; Tuberosa et al., 2002; Malamy, 2005). Therefore, some correlations of root system traits and drought resistance might depend heavily on the testing environment. Finally, root systems of both maize and rice are made up of multiple types of roots (lateral, seminal and adventitious) that are independently regulated (Hochholdinger et al., 2004a, 2004b), respond differently to drought conditions and might play different roles in drought avoidance (Yang et al., 2003). These differences are usually ignored in the scoring of root system traits. In short, it is not difficult to see why there are discrepancies

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Root systems and low phosphorus stress

Although there are many types of nutrient stress, P deficiency is one of the most prevalent, with P limiting growth and crop yield in over 30% of the world’s arable land (Vance et al., 2003). Organic P fertilizers can be used to supplement soils, but only a small percentage of the added P is utilized by the plants. The remainder contaminates waterways with catastrophic effect. Furthermore, inexpensive rock P reserves, used for fertilizer, are estimated to be depleted at the current usage rates in 60–80 years (Vance et al., 2003). Therefore, increasing low P tolerance is a critical agricultural goal. P acquisition is highly dependent on root traits. Low P-tolerant plants sense P limitation and respond by increasing lateral root formation, up-regulating expression of high-affinity transporters in the roots and increasing root exudates (LopezBucio et al., 2003; Vance et al., 2003). P has a low mobility in the soil, and larger root systems take up greater amounts of soluble P due to increased surface area. Increased production of root exudates, specifically acid phosphatases and organic acids, release P from bound or precipitated forms in the soil. Organic acid exudates also encourage certain microbial interactions; microorganisms associated with roots produce phytase; phytase releases P from phytate, the primary form of P in the soil. P acquisition is inextricably linked to root system traits. In Arabidopsis, varieties with the highest P acquisition efficiencies have the highest root:shoot mass ratios (Narang et al., 2000). Similar results were obtained in soybean, rice, common bean (Phaseolus vulgaris) and maize: varieties with more lateral roots showed higher P uptake under P limiting conditions (Silberbush and Barber, 1985; Lynch and Brown, 2001; Wissuwa and Ae, 2001a; Zhu et al., 2005). In both Arabidopsis and bean, genotypes that increase lateral root growth and decrease primary root growth in response to low P showed better P acquisition (Narang et al., 2000; Liao et al., 2001; Nielsen et al., 2001; Williamson et al., 2001). This adaptation creates a shallower root system overall and is beneficial because P tends to be concentrated near the surface in P-limited soils (Lynch and Brown,

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2001). In soybean, shoot P content in a segregating RIL population correlated significantly with increased root system size (Kuang et al., 2005). Similarly, Wissuwa and Ae (2001b) found that in rice two of four QTL associated with increased P uptake under P limitation were related to maintenance of root growth. Mathematical simulations in lupin and soybean have predicted that root system architecture is a major determinant of P uptake (Silberbush and Barber, 1983; Dunbabin et al., 2003). All of the above findings make root system traits, especially lateral root density, a very attractive target in attempts to develop crop plants that can tolerate low P conditions. However, as with drought stress, obtaining plants with altered root system traits is essential to test their effects on P uptake and yield under field conditions. 4.4.3

Exploiting root system research for crop improvement

Markers for major-affect QTLs are valuable tools in breeding, and taking this approach to improve root system architecture shows great promise (de Dorlodot et al., 2007). In addition, plant developmental biologists can synergize with agronomists to improve root traits through rational design approaches. In such approaches, candidate genes demonstrated to alter root system architecture are manipulated in crop plants. These genes can be identified by molecular dissection of QTLs, or through forward and reverse genetic approaches in model systems such as Arabidopsis (de Dorlodot et al., 2007). In the first proof of concept that Arabidopsis research can be exploited to manipulate root system architecture, Li et al. (2005a) demonstrated that overexpression of a H+ -pyrophosphatase AVP1 in Arabidopsis leads to an increase in root system development, and Park et al. (2005) showed that this phenotype could be recapitulated in tomato. The increased root system size was accompanied by increased water uptake and drought tolerance in both Arabidopsis and tomato (Li et al., 2005a; Park et al., 2005). Although the mechanism for AVP1-mediated root system regulation is not yet known, this provides a valuable tool to manipulate root system architecture in crop plants. Such a tool will allow agronomists to study the effects of altered root system architecture on yield or stress tolerance in the field, and compare the performance of the transgenic plant to an isogenic control plant – this kind of comparisons has not previously been possible, as morphological traits are usually compared between genetically heterologous crop varieties. The AVP1 studies make it clear that there is sufficient conservation of regulatory mechanisms between Arabidopsis and crop plants to make collaborations between Arabidopsis researchers and agronomists a necessity for future crop research.

Acknowledgments The author thanks S.R. McCouch and Y. Helariutta for assistance with Section 4.4 and with Fig. 4.2, respectively. This material is based on the work

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Root Development supported by the National Science Foundation under Grant No. 0238529. Any opinions, findings and conclusions or recommendations expressed in this material are those of the author, and do not necessarily reflect the views of the National Science Foundation.

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Annual Plant Reviews (2009) 37, 127–156 doi: 10.1002/9781444310023.ch5

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Chapter 5

ADVENTITIOUS ROOT FORMATION: NEW INSIGHTS AND PERSPECTIVES Gaia Geiss,1,2,∗ Laurent Gutierrez1 and Catherine Bellini1,2,∗ 1

Department of Forest Genetics and Plant Physiology, Umea˚ Plant Science Center, ˚ Sweden Swedish University of Agricultural Sciences, SE-90183 Umea, 2 Laboratoire de Biologie Cellulaire, Unit´e de Recherche 501, Institut National de la Recherche Scientifique, F-78000 Versailles, France

Abstract: The root system of a plant consists of the primary, lateral and adventitious roots. Lateral roots always develop from roots whereas adventitious roots form from stem or leaf-derived cells. Adventitious rooting is an essential step in the vegetative propagation of economically important horticultural and woody species. It allows clonal propagation and rapid fixation of superior genotypes prior to their introduction into production or breeding programs. Problems associated with rooting of cuttings frequently result in significant economic losses. Development of adventitious roots is a complex process that is affected by multiple factors including phytohormones, light, nutritional status, associated stress responses such as wounding, and genetic characteristics. How endogenous and environmental factors interact to control adventitious root formation is still poorly understood, although significant progress has been made in the understanding of the molecular control of root and lateral root development. In this review, we will summarize the current knowledge on the physiological aspects of AR formation and highlight the recent progress made in the identification of putative molecular players involved in the control of adventitious rooting. Keywords: adventitious roots; clonal propagation; quantitative trait; phytohormones; light; molecular markers

∗

These two authors have contributed equally to the work.

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5.1

Introduction

The root system of a plant consists of the primary, lateral and adventitious roots (ARs). The primary root is initiated during embryogenesis and will later elongate after germination. The lateral and the ARs are initiated and developed postembryonically from differentiated cells. Lateral roots always develop from roots whereas ARs are formed from stem or leaf-derived cells. AR become specialized to serve adaptive purposes besides the three primary functions in plant species: acquisition of water and nutrients from the soil, anchorage of the plant to a substrate and storage of food reserves. AR formation is part of the normal development of the plant and occurs naturally, in most monocotyledonous species for which they constitute the main root system or in many dicotyledonous species that propagate vegetatively. They can also be induced artificially either as an adaptative phenomenon to environmental changes or by wounding or hormone application on explants for artificial vegetative propagation. Although significant progress has been made in the understanding of the molecular control of root and lateral root development, AR formation is a biological developmental process that is still poorly understood. The actual accessibility to new sophisticated tools to undertake genetic and molecular biology analysis opens new possibilities of investigation that will certainly help in identifying genes involved in the control of AR formation. In this review, we will summarize the current knowledge on the physiological aspects of AR formation and highlight the recent progress made in the identification of putative molecular players involved in the control of adventitious rooting. We will mainly focus on ARs of dicotyledonous species because the molecular and genetic dissection of the root system in monocots, such as cereals will be described in Chapter 7.

5.2 5.2.1

Role and origin of ARs ARs allow vegetative propagation in both monocots and dicots

Natural vegetative reproduction is a process found mostly in herbaceous and woody perennials, and typically involves structural modifications of the stem, although any horizontal, underground part of a plant (either stem or root) can contribute to vegetative reproduction of a plant. In a few dicotyledonous species, such as Bryophyllum, leaves are involved in vegetative reproduction and support development of new plants arising from foliar embryos in the notches at the leaf margins (Fig. 5.1a). Many other species, similarly to strawberries (Fragaria) or blackberries (Rubus), mainly propagate vegetatively from stolons or stems from which ARs regenerate to anchor the new plants to the ground (Fig. 5.1b). Among monocotyledonous plants, such as onion

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Figure 5.1

Natural (a, b, c) and induced (d) adventitious roots.

(Allium cepa), garlic (A. sativum) or tulip (Tulipa) bulbs will be used as the unit for vegetative propagation (Fig. 5.1c). AR process is crucial to amplify valuable plants or plants whose final yield, whether fruit or dry matter, is influenced by the presence of ARs. This second group of plants plays an essential role in the world’s food supply, such as maize (Zea mays L.), wheat (Triticum aestivum L.) and rice (Oryza sativa L.). Adventitious rooting is an essential step in artificial vegetative propagation of plants. In horticulture, agriculture and forestry, vegetative or clonal propagation is widely used to multiply elite genotypes obtained in breeding programs or selected from natural populations (Hartmann et al., 1990). AR formation is a prerequisite for successful clonal regeneration of propagules and in many woody species, including most fruit and nut plants, the low ability to regenerate ARs has led to technologies, such as grafting and budding onto rootstocks, that are often themselves propagated through cuttings. AR formation is a limiting step for in vitro propagation that is part of breeding programs of some major trees such as Eucalyptus, Populus, Salix or Cryptomeria in which defects in AR formation can lead to important economical losses. 5.2.2

Origin of the ARs

While the primary root meristem is established during embryogenesis, lateral and AR meristems are formed postembryonically (Casson and Lindsey, 2003). In contrast to lateral roots, which are commonly formed from mature pericycle

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Root Development cells of main roots, ARs develop from different tissues and consequently from different cell types. ARs are also formed after tissue culture regeneration of shoots with hormones applications. Cells, unable to produce ARs in planta, are capable of doing so in cuttings when stimulated by specific inducers either in vitro or not (Altamura, 1996). Most of the time, at the onset of the AR formation in tissue culture, no cells are specified to form ARs. They first dedifferentiate, i.e. acquire competence for cell proliferation and organ regeneration (Ozawa et al., 1998), then initiate cell proliferation, and form AR primordia. Nevertheless, in some plants, such as willow (Salix), preformed AR initials are already present in the stem but remain dormant (Hartmann and Kester, 1983) and develop as ARs only when the stem has been cut and placed in water (Fig. 5.1d). A similar process has been observed in species such as poplar (Populus), jasmine (Jasminun sp.), currant (Ribes sp.) and lemon (Citrus medica) (Hartmann and Kester, 1983). In some other cases, a callus will develop at the basal end of the cutting before the formation of ARs (Hartmann and Kester, 1983). The process of AR development consists of three successive, but interdependent, physiological phases with different requirements: induction (period preceding any histological event), initiation (cell divisions leading to the formation of internal root meristems) and expression (internal growth of root primordia and root emergence) (Kevers et al., 1997). However, according to De Klerk et al. (1999), there is a supplementary step allowing the AR formation in cuttings and shoots, namely cell dedifferentiation that occurs before the induction phase. Using the model system Arabidopsis thaliana, Konishi and Sugiyama (2003) isolated mutants defective in the various stages of the rooting process and could identify a few genes involved in the control of these different steps (see below in this review). 5.2.3

ARs from cuttings

Anatomical processes of AR formation could be analyzed in various species thanks to studies performed on cuttings from vegetative portions of the plant, such as stems (rhizomes, tubers, corms and bulbs), leaves, or roots. Once the cutting is made, ARs emerge from a group of cells able to dedifferentiate and to become meristematic. This group of cells, named the root initials, located in different tissues of the stem depending on the species, will differentiate into a root primordium and subsequently form the AR. For example, in herbaceous species, root initials are located outside and between the vascular bundles. A vascular system develops in the new root primordium, becomes connected to the adjacent vascular bundle, and the root tip continues to grow outward, through the cortex, emerging from the epidermis of the stem. In red raspberry (Rubus strigosus) tip cuttings, root initials arise in the primary rays besides vascular bundles of previously removed leaves. In cuttings of woody perennials, ARs originate next to and outward from the central core of vascular tissues and because of the presence of numerous layers of secondary xylem

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and phloem usually in the young secondary phloem tissue. In white pine (Pinus strobus) cuttings, root initials form in association with rays and leaf traces (Hartmann and Kester, 1983). In apple tree (Malus domestica) cuttings, ARs are initiated from interfascicular cambium adjacent to the phloem cells, and roots emerge from the stem epidermis (De Klerk et al., 1995; J´asik and De Klerk, 1997). In tobacco (Nicotiana tabacum) cuttings, AR primordia are located along the outer perimeter of the vascular cylinder, in close proximity to the stem phloem and xylem (Vera et al., 1994). In contrast to the invariant positioning of primordia across the protoxylem poles of the stele during lateral formation in tobacco roots, AR primordia are initiated along the whole perimeter of the stem vascular tissue. After emerging, the AR behaves like a normal root, giving rise to new aerial laterals. In tobacco leaf explants, Dhaliwal et al. (2003) showed that root primordia arose from the rib parenchyma cells, near the existing vascular bundles.

5.3 Factors influencing adventitious rooting 5.3.1

Environmental factors

5.3.1.1 Abiotic factors 5.3.1.1.1 Mineral nutrition. Mineral nutrition that plays essential and specific roles in plant metabolism is a key factor determining root morphogenesis. Mineral nutrients can function as constituents of organic structures, activators of enzymatic reactions, as charge carriers, and osmoregulators. Adventitious rooting and mineral nutrition are also intimately related, although only few studies have attempted to characterize the effects of specific minerals on each of the three phases of the adventitious rooting process. Indeed, quite a few mineral nutrients are able to influence adventitious rooting, either by inhibiting or increasing the number of ARs or by modulating the root length. Recently, the effect of various concentrations of different minerals on the adventitious rooting response in cuttings of Eucalyptus globulus Labill. has been reported (Schwambach et al., 2005). Calcium (Ca), nitrogen (N) source and zinc (Zn) affected significantly the root number and that phosphorus (P), iron (Fe), manganese (Mn) and nitrogen source influenced the root length, whereas the average rooting time and the percentage of rooted cuttings were more closely related to auxin availability. In addition, instead of having a cumulative effect, these minerals in solution were found to interact in a complex way. The different requirements of each nutrient at each adventitious rooting phase have to be considered when attempting to optimize the vegetative propagation of E. globulus. Cuttings that were rooted in an optimized mineral nutrient medium and acclimatized to ex vitro conditions for 2 months had a significantly higher survival than cuttings rooted in basal medium after transplanting and drought stress.

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Root Development Nevertheless, there is no general rule and a positive or a negative effect of one particular nutrient depends on the species and the growth environmental conditions. For example, P deficiency in hydroponically grown horse gram (Macrotyloma uniflorum) induces AR elongation (Anuradha and Narayanan, 1991) and similarly AR formation in common bean (Phaseolus vulgaris L.) (Miller et al., 2003). On the contrary, Schwambach et al. (2005) showed that P deficiency reduces root length in E. globulus cuttings. Calcium is also one of the few minerals that can markedly modulate adventitious rooting (Druart, 1997; Bellamine et al., 1998; Schwambach et al., 2005). It has recently been shown to act as a secondary messenger at the crosstalk of auxin and nitric oxide (NO) signaling pathways in the activation of processes leading to AR formation (Lanteri et al., 2006) and to be involved in cell division and root primordia elongation process, which occurs during the late rooting phases (Imaseki, 1985). An essential role for Ca, either as a simple mineral or as a secondary messenger in the action of auxin, has also been suggested during the expression phase of rooting of poplar (Populus) cuttings (Bellamine et al., 1998), whereas in E. globulus Ca seems to act earlier during the induction and formation phases (Schwambach et al., 2005). Recently, the plasma membrane (PM) calcium channel blocker, LaCl3 , was shown to significantly compromise the indole-3-acetic acid (IAA) and NO-induced AR formation in cucumber cuttings (Lanteri et al., 2006). The requirement of extracellular Ca2+ to promote AR formation in cucumber explants was corroborated by the use of the membrane-impermeable Ca2+ chelator EGTA. All these data indicate that both intracellular and extracellular Ca2+ pools are required for the action of IAA and NO to trigger AR formation in cucumber (Lanteri et al., 2006). The two most important nutrients required for plant growth and development are nitrogen (N) and carbon (C). The N/C ratio is mainly considered and it is difficult to analyze the effects of these two nutrients separately. A modification in nitrogen supply strongly influences the processes of carbon assimilation, allocation and partitioning, and therefore affects carbohydrate distribution within a plant. Both the nitrogen and carbohydrate status have a great impact on the postharvest performance of plants (Druege et al., 2000). Such relations are even more complicated not only when high survival and continuous development are expected, but also when the harvested product is a starting material that requires a particular subsequent response, such as the regeneration of AR. AR formation can be substantially affected by the initial status of both nitrogen and carbohydrate (Blazich, 1988; Veierskov, 1988). For example, because pelargonium (Pelargonium) cuttings are produced under the high light intensities of low latitudes, their survival and rooting are affected and they are not adapted to the prevailing low light conditions during the winter rooting period in Central European greenhouses (Forschner and Reuther, 1984; Kadner et al., 2000). Analysis of the carbohydrate partitioning in high-light-adapted cuttings of pelargonium in relation to survival and AR formation under low light and evaluation of a graduated supply of mineral

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nitrogen to stock plants, indicated that unstored low-nitrogen cuttings developed a reduced number of ARs and that AR formation in pelargonium is sensitive to nitrogen deficiency (Druege et al., 2004). However, the initial amount of nitrogen reserves is not a predominant characteristic determining the rooting capacity of pelargonium since a significant decrease in root number was observed when excessive nitrogen was supplied to the stock plants. In contrast to the variable relationship with nitrogen, AR formation in pelargonium cuttings depends predominantly on the initial reserves of sugars in the cuttings. Adventitious rooting relies on an adequate supply of carbohydrates in the zone of root regeneration, where they can promote root initiation and development (Li and Leung, 2000; Calamar and de Klerk, 2002). In addition, carbohydrates also modulate gene expression (reviewed in Gibson, 2005) and ´ and Sheen, interact strongly with plant hormone signaling (reviewed in Leon 2003; Gibson, 2004). The influence of sugars in AR formation has also been demonstrated in several species. In E. sideroxylon explants AR formation was increased by sucrose and 2–6% sucrose concentrations in the medium favored root development, whereas 4–6% sucrose induced more callus formation and 8–10% levels of sucrose were detrimental to explants cultures (Cheng et al., 1992). In Arabidopsis, the effects of sugars on the development of ARs have been confirmed in hypocotyls of seedlings grown in continuous darkness (Takahashi et al., 2003). Sucrose concentrations of 0.5–2.0% have been shown to be more effective to induce AR and 5% sucrose suppresses AR induction. Sucrose, glucose and fructose greatly stimulated the induction of ARs when applied exogenously in direct contact with the hypocotyl, but mannose or sorbitol did not. In continuous darkness, elongation growth of the hypocotyl to a certain length seemed to be necessary for the development of ARs because they did not develop under long-day conditions where hypocotyl growth is suppressed even in the presence of sucrose. To check whether both elongation growth of the hypocotyl and sugars were essential for AR formation, Takahashi et al. (2003) analyzed AR formation on hypocotyls of the Arabidopsis hy4 mutant that elongate in the light as well as in darkness because of a defect in blue light perception (Ahmad and Cashmore, 1993). They observed, however, that no ARs were induced under long-day conditions in presence of 1% sucrose, although the hypocotyls were of sufficient length. This result indicated that light might have an inhibitory effect on AR initiation and that an elongated hypocotyl was not of primary importance. 5.3.1.1.2 Light. A few studies have shown that light conditions might affect AR formation. Fett-Neto et al. (2001) have studied the distinct effects of auxin and light on AR development in E. saligna Smith (an easy-to-root genotype) and E. globulus Labill. (difficult to root). Root development was scored as rooting percentage, root density (roots per rooted cutting), mean rooting time and root length. In both species, rooting time was reduced in the presence of auxin. Cuttings from 2-month-old E. saligna seedlings were

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Root Development responsive to lower auxin concentrations than those from E. globulus seedlings. Cuttings from 3-month-old E. saligna seedlings rooted rapidly and they were not significantly affected by light conditions. In contrast, cuttings from 3-month-old E. globulus seedlings were recalcitrant to rooting and did not root at all when grown in the light during the root formation phase. Effective root regeneration of E. globulus cuttings was obtained when exposed for 4 days to 10 mg/L indole-3-butyric acid (IBA) and grown in the dark during the rooting phase. In the absence of exogenous IBA, light exposure during the root formation period had no effect on rooting of E. saligna cuttings, but inhibited rooting of E. globulus cuttings. Since supplementation of auxin abolished the inhibition of rooting caused by light exposure throughout the experiment, light-induced inhibition of rooting in E. globulus might be related to changes in auxin activity. In P. aureus Roxb. rooting of cuttings was promoted by high irradiance in the presence of exogenous auxin, but was inhibited when auxin was not supplied (Jarvis and Shaheed, 1987). Recently, a potential interaction between light and auxin in the regulation of adventitious rooting has been investigated studying Arabidopsis mutants altered in their capacity to form ARs, namely superroot2 (sur2) and argonaute1 (ago1) mutants (Sorin et al., 2005). They used an allelic series of ago1 alleles and ago1sur2 double mutants. The defect in AR formation in ago1 mutants correlated with an alteration of auxin homeostasis in the apical part of the seedling and a hypersensitivity to light. Although the ARGONAUTE1 (AGO1) gene could regulate the expression of genes acting at the crosstalk of auxin and light signaling pathways, it is still not clear whether light induces the production and/or accumulation of an inhibitor in the aerial parts of the plant that would counteract auxin or whether light signaling pathways are involved or both. For example, Picea abies (L.) cuttings grown in high light (difficult-to-root) displayed a higher concentration of endogenous cytokinins than cuttings grown in low light (easy-to-root) (Bollmark and Eliasson, 1990). Fett-Neto et al. (2001) made the hypothesis that light could favor cytokinin accumulation in Eucalyptus, which could explain the inhibition of rooting in E. globulus cuttings grown in light without exogenous auxin. Addition of auxin in the culture medium should overcome the light-mediated, cytokinin-induced inhibition of rooting, but this hypothesis still has to be demonstrated. Some Eucalyptus species have a specific type of putative rooting inhibitors called G inhibitors (Milborrow, 1984). These inhibitors have been isolated from mature leaves and bark of E. grandis W. Hill ex Maiden, shown to suppress rooting in cuttings of E. grandis and Vigna radiata (mung bean) and are present in high concentrations in the mature parts of E. grandis trees (Milborrow, 1984). It is possible that a similar type of inhibitors could contribute to the recalcitrant rooting of cuttings derived from E. globulus seedlings. Ectomycorrhizas (ECMs) fungi are often used to promote adventitious rooting in difficult-to-root species (see below). Changes in light fluxes and spectral quality can also affect mycorrhizal interactions. Niemi et al. (2005)

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studied the effects of broad-spectrum light quality on the interaction between the ectomycorrhizal fungus Pisolithus tinctorius (Pers.) and Scots pine (P. sylvestris L.) hypocotyl cuttings cultured in vitro. They showed that irrespective of light quality, inoculation with P. tinctorius increased formation and growth of ARs on Scots pine hypocotyl cuttings. Light sources with different spectra had no effect on control cuttings, but altered the fungus–cutting interactions. Red-rich daylight inhibited fungal-induced rooting of the cuttings. Since the radial growth of P. tinctorius was not influenced by red light, inhibition of AR and mycorrhiza formation by red light was unrelated to fungal growth. The authors suggested that long-term exposure to different broad-spectrum lights might affect compounds essential to the interaction between Scots pine and P. tinctorius. 5.3.1.1.3 Temperature. Temperature can potentially influence adventitious rooting capacity in many aspects, such as water and nutrient uptake and metabolism in general, promoting or inhibiting enzymatic action. Very little information can be found in the literature regarding the effect of temperature on AR formation. Nevertheless, it is known that it can influence the physiological status of the mother plant and later affect AR development on cuttings or have a direct effect during the adventitious rooting process. Druege et al. (2000) investigated the relationship between internal nitrogen and carbohydrate distribution in chrysanthemum (Chrysanthenum sp.) cuttings and adventitious rooting after different periods of dark cold storage. The main effect of cold storage was a decrease in carbohydrate concentrations in the cutting part, which modified the N/C ratio and led to an increase of rooting. Similar effects have been observed in conifers (Berhens, 1988). Important temperature effects on adventitious rooting of E. saligna Smith and E. globulus Labill. could be observed by treating donor plants or cuttings (Corrˆea and Fett-Neto, 2004). The two species showed temperature preferences and different sensitivities to higher and lower temperature in relation to rooting. Rooting percentage, root density and elongation were increased in E. saligna in warmer temperatures and root growth was inhibited by 15◦ C; moreover, cuttings treated with constant low temperatures were slower to root. On the contrary, cooler temperatures increased E. globulus rooting and constant higher temperatures seemed to reduce rooting percentage. Moderate heat shock on donor plants could be beneficial for rooting of non-auxininduced cuttings of E. saligna, whereas heat shock had deleterious effects on similarly treated E. globulus rooting. Auxin treatment improved the tolerance of microcuttings from both species to more extreme temperature treatments, yielding better rooting responses. The authors suggested a relation between temperature and auxin, probably through a modulation of endogenous auxin metabolism, transport and uptake. Nine temperature-sensitive Arabidopsis mutants defective in various stages of AR formation from hypocotyl explants have been identified confirming that temperature is likely to influence the different phases of the AR formation

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Root Development (Konishi and Sugiyama, 2003). Nevertheless, the role of temperature in controlling the rooting process is not clear yet and has still to be elucidated. 5.3.1.2 Biotic factors 5.3.1.2.1 Ectomycorrhizas. ECMs are symbiotes formed between certain fungal and plant species. Inoculation with specific ECM fungi can enhance root formation and/or subsequent root branching of in vitro cuttings and in vitro adventitious shoots. Specific symbiotic fungi have been studied as rooting agents both in vitro and ex vitro. Depending on the study, inoculation results in higher rooting frequency, a greater number of adventitious and lateral roots, or both (Gay, 1990; Supriyanto and Rohr, 1994; Normand et al., 1996; Fortuna et al., 1998; Karabaghli et al., 1998; Niemi et al., 2000). In vitro AR development on Norway spruce hypocotyl cuttings inoculated with ECM fungi has been associated with the IAA produced and released by the ECM fungus (Karabaghli et al., 1998). However, short treatments with filtrates from Paxillus involutus culture (containing less IAA) were more efficient in promoting rooting than the corresponding treatments with P. tinctorius (containing more IAA), suggesting the presence of another AR-inducing component (Niemi et al., 2002b). P. involutus produced and released, in addition to IAA, a high concentration of putrescine, a polyamine known to support the rooting process (Tonon et al., 2001), whereas P. tinctorius contained traces of the diamine cadaverine. Both fungi produced spermidine known as well to favor rooting frequency in Virginia Pine (P. virginiana P. Mill.) (Tang and Newton, 2005). Both fungi accelerated AR formation and increased subsequent root growth in hypocotyl cuttings of Scots pine (P. sylvestris L.) in vitro. Exogenously applied cadaverine enhanced AR formation caused by P. tinctorius and also promoted mycorrhiza formation by the fungus. Putrescine and P. involutus had a synergistic effect on root initiation, but not on subsequent root growth (Niemi et al., 2002a). The study of the interaction between ECM fungi and plant species appear to be a fine model to highlight numerous factors interfering with the rooting process (Niemi et al., 2004, 2005). 5.3.1.2.2 Agrobacterium rhizogenes. To overcome the rooting deficiency during vegetative propagation of economically important species, many attempts to use A. rhizogenes have been carried out on fruit trees and woody species. A. rhizogenes is a soil Gram-negative bacterium that induces AR formation at the site of infection in a large number of plants (Chilton et al., 1982). Root induction is due to the integration and expression of the T-DNA from the root-inducing Ri plasmid in the plant genome. Four loci involved in root formation have been identified in the T-DNA, and designated root loci (rol) A, B, C and D (Spena et al., 1987). The rolB gene was identified as the critical rol gene for the induction of roots (White et al., 1985; Spena et al., 1987; Capone et al., 1989), because among all the Ri genes and open reading frames (ORFs), it is the only one that can individually induce AR formation in different plant species. Nevertheless, maximum adventitious rooting occurs when the rolB

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gene is combined with either rolA and/or rolC, suggesting that the three genes synergistically control the rooting response (Spena et al., 1987). A. rhizogenes-mediated transformation has been used successfully to enhance AR in fruit trees, such as almond (Prunus dulcis), apple (M. domestica), kiwi (Actinidia sinensis), walnut (Juglans regia) and in forest trees, such as the genus Pinus, Larix and Eucalyptus (reviewed in Damiano and Monticelli, 1998; Li and Leung, 2003). Recently, a significant enhancement of adventitious rooting has been reported in Monterey pine (P. radiata D. Don), a species of considerable economic importance in several southern-hemisphere countries, and that is still, like most conifers, particularly difficult to root (Li and Leung, 2003). Similarly, rooting ability of the M9/29 apple (M. domestica) rootstock, that is recalcitrant to propagation through cuttings because of its poor rooting efficiency, has been improved after introduction of the rolB gene (Zhu et al., 2001). Although it is well established that rolB is responsible for the induction of AR, the molecular mechanisms are still not understood. An appropriate level of rolB expression is necessary for an active growth of hairy roots because either high or low levels correlate with impaired growth of roots (Tanaka et al., 2001). Tobacco (N. tabacum) cells transformed with the rolB gene show an alteration of the auxin-induced PM hyperpolarization (Maurel et al., 1991). Despite the correlations between rolB function and regulation or perception of auxin (Filippini et al., 1994, 1996) no real conclusion for rolB function exists. The RolB protein of the plasmid pRi1724 (1724RolB) has been shown to associate with the N. tabacum 14-3-3-like protein II (Nt14-3-3 II) in BY-2 cells, to localize in the nucleus and to show a decreased capacity to induce AR when compromised to interact with the Nt14-3-3 II protein due to mutation (Moriuchi et al., 2004). Further characterization of these proteins might shed light on the new mechanisms regulating AR formation in higher plants. 5.3.2

Endogenous factors

5.3.2.1 Aging The loss of competence to form AR is one of the most dramatic effects of plant maturation, an age-related developmental process. The rate and the extent of the loss of rooting ability are species dependent. For example, in eastern larch (Larix decidua), the rooting frequency of cuttings decreases from 100 to 50% during the course of 20 years. In contrast, a rapid loss in rooting ability is observed in loblolly pine (P. taeda) (Greenwood et al., 1989). Hypocotyl cuttings from 20- or 50-day-old loblolly pine seedlings rapidly form AR whereas epicotyl cuttings from 50-day-old seedlings root poorly only after 2–3 months (Diaz-Sala et al., 1996). In chestnut (Castanea sp.) and oak (Quercus robur L.), loss of rooting capacity observed in adult plant material could be associated with the differential expression of genes affecting one or all of the process phases (Ballester et al., 1999; Vidal et al., 2003). Arabidopsis could be a valuable system to study the age-related decline of AR formation because de-rooted

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Root Development juvenile hypocotyls from 12-day-old plants rooted readily within a week of culture whereas hypocotyls from adult (26-day-old) plants rooted poorly and only after long periods of time (Diaz-Sala et al., 2002). Rooting capacity does not depend on transition to flowering, because root induction decreased in a similar manner when flowering was delayed. Hopefully, such an experimental system will be helpful to better understand the molecular mechanisms related to the aging-related loss of rooting ability. 5.3.2.2 Polyamines An extensive literature gives evidence that polyamines play an important role in primary, lateral and AR development. An increase in endogenous polyamine levels following IBA-induced rooting and prior to root primordia development has been measured in Phaseolus sp. (Jarvis et al., 1985), Vigna hypocotyls (Shyr and Kao, 1985), tobacco roots and thin cell layers (Tiburcio et al., 1989; Altamura et al., 1991) and P. avium shoots (Biondi et al., 1990). Polyamines seem to either promote or inhibit the adventitious rooting process depending on the developmental phase or the species. A peak of putrescine, independent of exogenous IAA application, during the root induction phase on Fraxinus angustifolia shoots, has been observed, supporting a positive effect of putrescine on AR formation (Tonon et al., 2001). However, putrescine inhibits rooting when applied during the expression phase, whereas an increase of putrescine and IAA levels increase simultaneously at the induction and initiation phase of AR formation in mung bean (V. radiata) (Nag et al., 2001). In some others species, such as Virginia pine (P. virginiana Mill.) rooting frequency increases by exogenously added putrescine and spermidine, but decreases by exogenously added spermine (Tang and Newton, 2005). On the other hand, spermidine is reported to inhibit AR formation in many plants, including cherry (P. avium L.) (Biondi et al., 1990), walnut (J. regia L.) (Kevers et al., 1997) and poplar (Populus tremula L. × P. tremuloides L.) (Hausman et al., 1994). Different cellular compounds that control AR formation during the different phases of the process in mung bean hypocotyls have been dissected (Nag et al., 2001). Polyamines, in particular oxidized products of putrescine (H2 O2 and ␥ -amino butyric acid), play a regulatory role together with auxin during the cell division stage and initiation of the root primordium within the induction phase. Nevertheless, the interrelationship between auxin and polyamines through the rooting process is still unclear. 5.3.2.3 Enzymatic activities of peroxidases Classical plant peroxidases (PODs) are heme-containing enzymes that catalyze the oxidation of diverse organic compounds (Dawson, 1988). Significant roles of PODs have been suggested in plant growth and cell differentiation (Gaspar et al., 1985), in hormone catabolism (Haissig et al., 1992; Kevers et al., 1997; Nag et al., 2001), such as decarboxylative catabolism of IAA (Ljung et al., 2002). Changes in peroxidase activity (PA) and peroxidase isoforms patterns have been proposed as biochemical markers of the successive

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adventitious rooting phases. In some cases, the induction phase is characterized by a rapid decrease in PA, the initiation phase by an increased PA and the expression phase by a gradual PA decrease (Gaspar et al., 1992), such as in yew (Taxus baccata), strawberry tree (Arbutus unedo) (Metaxas et al., 2004), and in the common gardenia (Gardenia jasminoides Ellis) (Hatzilazarou et al., 2006). In narrowleaf ash (F. angustifolia), the root expression phase is characterized by increased PA and low concentrations of polyamines (Tonon et al., 2001). These changes were associated specifically with the rooting process and did not depend on the presence of exogenous IBA. By analyzing qualitative and quantitative changes in isoperoxidase expression (basic and acidic peroxidases) during in vitro rooting of Nothofagus antarctica microshoots, 17 peroxidase bands have been detected, 4 basic and 13 acidic (Martinez-Pastur et al., 2001). Four of the acidic peroxidases appear during the induction stage of rooting. Number and relative activity of the isoperoxidases vary during different stages of adventitious rooting, which make them a good indicator of the processes involved in adventitious rooting of N. antarctica. 5.3.2.4 Phenolic compounds The role of the phenolic compounds on the AR formation seems to depend on the nature of the compound and the plant species (Kling and Meyer, 1983). In some cases, phenolic compounds have been shown to enhance adventitious rooting (Ueda, 1989; Curir et al., 1990). De Klerk et al. (1999) have studied the effect of various phenolic compounds on rooting of apple (M. domestica) stem slices. Ferulic acid turned to be the most active compound. It improved rooting in the presence of auxin and so did the phloroglucinol, but to a less extent. Phenolic compounds might act as antioxidants, inhibiting the auxin oxidation and, thus, allowing the AR formation. In other cases, phenolic compounds have been shown to inhibit adventitious rooting. In walnut, decreasing flavonoid content increases adventitious rooting. In that case the auxin concentration did not vary, at least in the latter phase of the processes (El Euch et al., 1998). In Arabidopsis, the mutant transparent testa (tt4), which has the equivalent mutation in the chalcone synthase encoding gene (CHS), also displays increased lateral and AR development when compared to wildtype plants. In these plants, the auxin transport increases more specifically in the hypocotyl and the inflorescence (Brown et al., 2001).

5.3.3

Phytohormones

To improve rooting, the effects of different growth regulators have been investigated. Auxin is the hormone commonly used to promote adventitious rooting in different species. Auxin and ethylene are more often described as activators, whereas cytokinins and gibberellins usually as inhibitors of AR formation, even when some positive effects have been observed.

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Root Development 5.3.3.1 Auxin The auxin IAA was the first plant hormone to be used to stimulate rooting of cuttings (Cooper, 1935). In the same period, it was discovered that a second auxin IBA also promoted rooting and was even more effective than IAA (Zimmerman and Wilcoxon, 1935). IBA is now used commercially worldwide to root cuttings from many plant species and is more efficient than IAA ¨ (Epstein and Ludwig-Muller, 1993). Exogenous IBA stimulated rooting of epicotyl cuttings of conifers such as P. sylvestris (Flygh et al., 1993) and induces rooting better than IAA on two Eucalyptus species microcuttings differing in their ability to regenerate AR (Fogac¸a and Fett-Neto, 2005). At each step of the AR formation, auxin (endogenous or exogenously applied) plays a central role. At the beginning of the rooting process, high endogenous auxin concentration is normally associated with a high rooting rate (Blaˇckov´a et al., 1997; Caboni et al., 1997). When exogenous auxin is applied to induce rooting, the endogenous auxin concentration usually reaches a peak some hours or days after wounding (Gaspar et al., 1996; Gatineau et al., 1997), coinciding with the initiation of the rooting process. In some cases, the peak of auxin concentration was not observed (Label et al., 1989). The importance of the auxin during the expression phase was demonstrated in poplar (Populus sp.) (Bellamine et al., 1998). Rice (O. sativa) mutants affected in the expression of the O. sativa PINFORMED1 (OsPIN1) gene potentially involved in auxin polar transport are affected in AR emergence and tillering confirming that not only the auxin concentration is important but also its distribution in the different tissues (Xu et al., 2005). Despite its crucial role in AR development, the pattern of auxin action, either IAA or IBA, is still poorly understood. In Arabidopsis, the superroot (sur1 and sur2) mutants accumulate IAA and develop numerous AR on the hypocotyl and cuttings of different organs in the case of sur1 (Boerjan et al., 1995; Delarue et al., 1998). Recently, differential roles for IAA and IBA have been found in the regulation of AR formation from stem segments of Ara¨ bidopsis (Ludwig-Muller et al., 2005). Indeed, exogenous IBA, but not IAA could induce rooting and this effect was inhibited by 3,4,5-triiodobenzoic acid (TIBA), an auxin polar transport inhibitor. They suggested that endogenous IAA and exogenous IBA might interact to promote adventitious rooting in Arabidopsis stem segments. The performance of IBA versus IAA can be explained by several possibilities: higher stability, differences in metabolism, differences in transport and IBA as a slow release source of IAA. The conver¨ sion of IBA to IAA occurs in many plant species (reviewed in Ludwig-Muller et al., 2005). However, in microcuttings of Malus sp., IBA induced more roots than IAA although it was converted to IAA only at very low levels, suggesting that either IBA itself was active or that it modulated the activity of IAA (Van der Krieken et al., 1992). 5.3.3.2 Ethylene Zimmerman and Hitchcock (1933) were the first to show that ethylene plays a major role in AR formation in many plant species, before the discovery that

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auxin could promote it. Since then, a wide array of experiments has been conducted to determine its role in root initiation and development. Results from these experiments were highly variable and, for one particular species, ethylene could promote, inhibit, or have no effect on AR formation. Since the initial report showing that auxin induces ethylene synthesis (Zimmerman and Wilcoxon, 1935), there have been numerous attempts to determine how auxin and ethylene interact during AR formation and development. Inhibitors of ethylene biosynthesis, such as aminoethoxyvinylglycine, have been shown to reduce the number of ARs of mung bean (V. radiata) cuttings (Jusaitis, 1986) and so did inhibitors of ethylene perception, such as silver thiosulfate and 2,5-norbornadiene (Robbins et al., 1985). Support for this type of interaction between auxin and ethylene came from experiments with Rumex palustris plants (Visser et al., 1996), in which higher tissue ethylene concentrations increased the sensitivity of root-forming tissues to endogenous IAA. The ethylene-insensitive never ripe (NR) mutant tomato plants produced more belowground root mass but fewer aboveground AR than wild-type plants (Clark et al., 1999). Applied IBA increased AR formation on vegetative stem cuttings of wild-type plants, but had little or no effect on NR plants. Reduced AR formation was also observed in ethylene-insensitive transgenic petunia plants. Applied 1-aminocyclopropane-1-carboxylic acid (ACC), an ethylene precursor, increased AR formation on vegetative stem cuttings from NR and wild-type plants, but NR cuttings produced fewer ARs than wild-type cuttings. These data suggested that the positive effect of auxin on adventitious rooting is influenced by ethylene responsiveness. Similar results were obtained with ethylene-insensitive tobacco plants (McDonald and Visser, 2003). Very recently, Steffens et al. (2006) highlighted a complex interaction between ethylene, gibberellic acid (GA) and abscisic acid (ABA) in the control of AR formation in deep-water rice. GA was ineffective on its own, but had a synergistic effect together with ethylene to promote ARs and ABA acted as a competitive inhibitor of GA activity. 5.3.3.3 Cytokinins The inhibitory effect of kinetin on rooting of dwarf bean (P. vulgaris) petioles and hypocotyls has been observed before the existence of cytokinins in plants had been confirmed by Humphries (1960). Later, several groups showed that cytokinins (CKs) negatively regulate the formation of ARs in hypocotyls or stems of P. radiata (Smith and Thorpe, 1975), rose (rosae) (De Vries and Dubois, 1988), or pea (Pisum sativum) (Bollmark and Eliasson, 1986). More recently, a trans-zeatin riboside (ZR) present in root xylem sap was shown to negatively regulate AR formation on cucumber (Cucumis sativus) hypocotyls Kuroha et al. (2002). When the roots are cut off, a rapid decrease in the ZR level would occur together with auxin accumulation and ethylene synthesis in the basal part of the cut hypocotyls or stems, resulting in the formation of ARs (Kuroha et al., 2002). Recently, in transgenic tobacco and Arabidopsis plants overexpressing cytokinin oxidases and therefore having a reduced cytokinin content, increased adventitious and lateral root formation were recorded (Werner

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Root Development et al., 2001, 2003). Finally, transgenic tobacco plants overexpressing a zeatin O-glucosyltransferase (ZOG1) gene showed increased ARs on their lower stems, suggesting that the reduction of active cytokinin by O-glucosylation leads to a lower cytokinin/auxin ratio (Martin et al., 2001). All these experiments revealed the inhibitory action of cytokinins on adventitious rooting. The wooden leg-3 (wol-3) Arabidopsis mutant, an allele of the ahk4 mutant, affected in the ARABIDOPSIS HISTIDINE KINASE 4 (AHK4) gene that encodes a cytokinin receptor, is impaired in the development of primary and lateral roots but not that of ARs (Kuroha et al., 2006). Interestingly, the wol-3 and ahk2 ahk3 ahk4 triple mutants developed a normal vascular system in ARs, albeit aborted vascular tissues in the hypocotyl and primary root. The enhanced formation of ARs in wol-3 mutant could be due to the accumulation of auxin in the hypocotyl, because of the defect in vascular tissue development in the hypocotyl and the root. The complete loss of the cytokinin signal in wol-3 and ahk2 ahk3 ahk4 triple mutant seedlings would lead to an inhibition of the negative effect of cytokinin on AR formation. Cytokinin receptors seem to be required for normal development of auxin transporting vascular tissues in the hypocotyl but not in ARs.

5.4

5.4.1

New insights into genetics and molecular mechanisms involved in adventitious rooting Competence to form AR is a heritable quantitative genetic trait

The large array of physiological and biochemical processes involved in the control of AR formation implies an underlying genetic complexity. Recent genetic studies indicate that the competence to form ARs is a quantitative genetic trait. In woody species, competence to form ARs is quantified as percentage of rooted cuttings or root number per rooted cuttings. Economically important genotypes of apple, eucalyptus, pine, or other woody species are classified as easy or difficult to root. Although this variation is better characterized among woody species for which clonal propagation is commonly used in breeding programs, it also exists in herbaceous species as reported in Arabidopsis (King and Stimart, 1998), rapeseed (Brassica napus) (Oldacres et al., 2005), maize (Z. mays) (Mano et al., 2005) or rice (O. sativa) (Zheng et al., 2003; Ikeda et al., 2007). Already in 1968, Wilcox and Farmer reported the existence of quantitative trait loci (QTLs) associated with the root number per rooted cuttings in cottonwood (P. deltoides) showing a broad-sense heritability. A narrowsense heritability has been calculated for the percentage of rooted cuttings in loblolly pine (P. taeda L.). (Foster, 1990). Greenwood and Weir (1994) analyzed the aptitude to form AR in response to IBA in half-sib and full-sib

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families of easy-to-root or difficult-to-root loblolly pine and show that the variation in rooting was highly significant. Grattapaglia et al. (1995) identified four QTLs for the percentage of rooted cuttings in a full-sib cross between E. grandis (difficult to root) and E. urophylla (easy to root). Most of the inherited phenotypic variation (28.5%) was attributed to E. urophylla. E. globulus Labill. is a major plantation species with excellent properties for cellulose pulp production but retains a very irregular adventitious rooting behavior. On the contrary, E. tereticornis has modest pulping qualities but good potential for rooting and therefore is frequently used in hybridization programs to improve adventitious rooting of other eucalyptus species (reviewed in Marques et al., 1999). The genetic transmission of vegetative propagation traits in these two species has been studied and nine QTLs were associated with adventitious rooting (Marques et al., 1999). On average, more adventitious rooting phenotypic variation was contributed by E. tereticornis (16.3%) relative to E. globulus (9.7%). More recently, QTLs that are associated with vegetative propagation traits in pedunculate oak (Q. robur) have been reported (Scotti-Saintagne et al., 2005). An F1 full-sib family of 278 offsprings was used as mapping population. Assessment of rooting ability over 3 years led to the detection of ten QTLs explaining 4.4–13.8% of phenotypic variance, of which the two strongest were stable across years. The ability to dissect the genetic components of vegetative propagation traits is important to clarify the manner they should be incorporated into selection strategies. The identification of molecular markers linked to the genetic traits, such as AFLP markers, microsatellites, or gene sequences, becomes crucial for assisting in the selection of valuable genotypes. Recently, Marques et al. (2002) reported conservation and synteny of SSR loci and vegetative propagation traits in Eucalyptus species. These microsatellite markers should help integrate eucalyptus genetic linkage maps from various sources, leading to more efficient use of genetic information for molecular breeding. A valuable approach to identify new molecular markers associated with AR formation for markerassisted selection would also be the use of global approaches such as transcriptomic or proteomic analysis of the adventitious rooting process and/or to exploit model systems, such as Arabidopsis or rice (O. sativa) for which cloning of genes involved in the regulation of AR formation is accessible (see below). 5.4.2

Transcriptomic and proteomic approaches to identify candidate genes associated with AR formation

In the past few years, several groups performed transcriptomic or proteomic analysis during the process of AR formation in apple (M. domestica), Monterey pine (P. radiata), poplar (P. trichocarpa × deltoides), lodgepole pine (P. contorta) and Arabidopsis, in an attempt to identify candidate genes associated with adventitious rooting (Butler and Gallagher, 1998; Li and Leung, 2001; Kohler et al., 2003; Brinker et al., 2004; Sorin et al., 2006).

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Root Development Differential messenger RNA display and mRNA representational difference analysis have been used to analyze transcripts that are modulated during AR formation in IBA-treated apple stem discs (Butler and Gallagher, 1998) and both upregulated and downregulated transcripts were identified. Clones sharing sequence homology with polygalacturonase and MAP kinases were found among the upregulated transcripts. A full-length cDNA for the most abundant upregulated mRNA, a 2-oxoacid-dependent dioxygenase, designated Adventitious Rooting Related Enzyme-1 (ARRO-1) was further characterized and shown to be strongly induced by IBA and IAA, but not 2,4D during adventitious rooting and also in the primary roots of apple seedlings (Butler and Gallagher, 2000). The transcript abundance of genes involved in water and nutrient absorption during adventitious rooting in the hybrid cottonwood (P. trichocarpa × deltoides) was analyzed with microarrays (Kohler et al., 2003). Hierarchical cluster analysis of aquaporins and transporters identified transcripts that were co-regulated according to the developmental stage. Aquaporins belonging to the PIP family were highly expressed in the bark of dormant cuttings and in root primordia, whereas their expression was downregulated in the adventitious and lateral roots. In contrast, transcripts encoding tonoplastic aquaporins were mostly expressed in root callus and emerging roots or primary and lateral roots. The various transporters analyzed also showed a differential expression during the time course of root development. Sulfate, hexose and oligopeptide transporters were upregulated during callus formation, whereas nitrate and peptide transporters were mainly expressed in primary and lateral roots. The gene expression pattern during rooting of P. contorta hypocotyl cuttings was investigated (Brinker et al., 2004) and a significant change in the transcript level of 220 genes could be observed over the period of root development. During the root initiation phase, genes involved in cell replication and cell wall weakening and a transcript encoding a PINHEAD/ZWILLE-like protein were upregulated, while genes related to auxin transport, photosynthesis and cell wall synthesis were downregulated. Similarly, changes in transcript abundance of genes related to water stress were observed. During the root meristem formation phase, the transcript abundance of genes involved in auxin transport, auxinresponsive transcription and cell wall synthesis increased, while those encoding proteins involved in cell wall loosening decreased. More recently, genetic and physiological analyses were coupled with protein profiling of Arabidopsis mutants altered in the formation of ARs (Sorin et al., 2006), namely the auxin overproducers superroot mutants (sur1 and sur2) (Boerjan et al., 1995; Delarue et al., 1998) that spontaneously develop ARs and the argonaute1 mutant (Bohmert et al., 1998) that poorly make ARs in response to auxin (Sorin et al., 2005). Comparison of two-dimensional gel electrophoresis protein profiles resulted in the identification of 11 proteins whose abundance was either positively or negatively correlated with the endogenous auxin content, the number of AR primordia and/or the number of mature ARs. Interestingly, three auxin-inducible GH3-like proteins were positively correlated with the

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number of mature ARs. The 11 proteins were predicted to be involved in different biological processes, including the regulation of auxin homeostasis and light-associated metabolic pathways. Some overlap was reported among the proteins identified and the genes described to be potentially associated with AR formation in P. contorta (Brinker et al., 2004), suggesting that the function of these genes is at least partially conserved among different plant species. 5.4.3

Genes involved in the control of AR formation

Several mutants affected in different phytohormone signaling or homeostasis are also affected in AR formation. The ethylene-insensitive never ripe tomato mutant developed fewer ARs than the wild type (Clark et al., 1999). In contrast, the ABA-deficient tomato mutants flacca and notabilis produce an excess of ARs on the stems (Tal, 1966). The AR phenotype was attributed to an increased ethylene production in the mutants; however, this could never be demonstrated. Recently, Thompson et al. (2004) showed that the AR phenotype can be restored to that of wild type in the notabilis tomato mutant by expressing the 9-cis-epoxycarotenoid dioxygenase 1 (LeNCED1) gene involved in ABA biosynthesis. These results suggest that ABA can be a negative regulator of AR formation. In Arabidopsis, mutants overproducing auxin such as the superroot mutants sur1 and sur2 (Boerjan et al., 1995; Delarue et al., 1998) or yucca (Zhao et al., 2001) spontaneously produce AR on the hypocotyls of light grown seedlings. SUR1 (SUPERROOT 1) and SUR2 (SUPERROOT 2) genes encode a C-S-lyase protein and the cytochrome P450 Cyp83B1 respectively, both involved in the indole glucosinolate pathway (Barlier et al., 2000; Bak et al., 2001; Mikkelsen et al., 2004). The YUCCA1 gene encodes a flavin mono-oxygenase suitable for converting tryptamine in N-hydroxyl tryptamine in vitro (Zhao et al., 2001; 2002) and belongs to a family of 11 predicted YUC flavin mono-oxigenases from which 4 play essential roles in auxin biosynthesis (Cheng et al., 2006). Although these mutants clearly confirm the importance of hormone homeostasis and of the interactions between different hormones in the control of AR formation, they do not tell much about more specific molecular mechanisms. Three temperature-sensitive Arabidopsis mutants rrd1, rrd2 and rrd4 affected in root redifferentiation from hypocotyl explants cultured in vitro, were identified (Sugiyama, 2003) and their characterization suggested a role for the genes ROOT REDIFFERENTIATION 1 and 2 (RRD1 and RRD2) in some fundamental processes required for active cell proliferation and for RRD4 an involvement in the acquisition step of competence for cell proliferation during callus initiation in hypocotyl explants. Nine other temperature-sensitive mutants defective in various stages in the formation of ARs were isolated (Konishi and Sugiyama, 2003). Whereas root initiation defective mutants (rid1 to rid5) failed to initiate root primordia, it is the development of root primordia, which was arrested in the root primordium defective1 (rpd1) mutant. The root growth defective (rgd1, rgd2 and rgd3) mutants were defective in root growth after the establishment of the root apical

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Root Development meristem. The ROOT INITIATION DEFECTIVE 5 (RID5) gene was identified as the Microtubule ORganisation 1/GEMini pollen 1 (MOR1/GEM1) gene encoding a microtubule-associated protein (Konishi and Sugiyama, 2003) and the RPD1 gene as a member of a novel protein family specific to the plant kingdom whose function is still unknown. A knockout of the RPD1 gene caused embryo lethality, a phenotype that is consistent with the hypothesized function of RPD1 in the maintenance of active cell proliferation (Konishi and Sugiyama, 2006). Recently, the cloning of the rice genes CROWN ROOTLESS 1 (CRL1) and ADVENTITIOUS ROOTLESS 1 (ARL1), which encode the same member of the plant-specific ASYMMETRIC LEAVES2/LATERAL ORGAN BOUNDARIES protein family or LOB family was reported (Inukai et al., 2005; Liu et al., 2005). CRL1/ARL1 is an auxin-responsive gene, which contains two putative auxin response elements (AuxREs) in its promoter region. Its induction by auxin requires the degradation of IAA/Aux proteins and it was shown that the proximal AuxRE specifically interacted with an AUXIN RESPONSE FACTOR (ARF) and acted as a cis-motif for CRL1 expression (Inukai et al., 2005). CRL1/ARL1 can be considered as a positive regulator for crown root formation in rice. Interestingly, an Arabidopsis transgenic line overexpressing the ARF17 gene developed fewer ARs than wild-type plants confirming the potential role of ARF genes in the regulation of AR development by auxin (Sorin et al., 2005). Expression of the auxin-inducible GH3-like genes that is positively correlated with the number of ARs (Sorin et al., 2006) was repressed in the ARF17 overexpressing line. Therefore, ARF17- and GH3like genes might be involved in the control of AR formation. Because several GH3-like genes act in the crosstalk between light and auxin signaling pathways (reviewed in Sorin et al., 2005), ARF17 might be a major regulator of AR formation by repressing GH3-like genes and, therefore, modulating auxin homeostasis in a light-dependent manner. Expression of SCARECROW-LIKE (SCR-LIKE) genes was significantly increased in response to exogenously applied auxin, within 24 hours of the rooting process, in rooting-competent cuttings of two distantly related forest species (P. radiata D. Don and Castanea sativa Mill.). This corresponds to the phase of cell reorganization before the resumption of cell division and the appearance of AR primordia. These results suggest that SCARECROW-LIKE genes play a role during the earliest stages of AR formation (S´anchez et al., 2007). SCARECROW (SCR) is a putative transcription factor, expressed in cortical and endodermal initials, and it is required for the asymmetric cell division that gives rise to the cortex and endodermis, and to other tissues in aerial organs of Arabidopsis thaliana (Di Laurenzio et al., 1996; Wysocka-Diller et al. 2000; Heidstra et al. 2004). Arabidospis SCR (AtSCR) is also involved in the establishment of quiescent center identity and in the maintenance of the stem cell status of the surrounding initial cells during embryonic pattern formation and postembryonal development (Sabatini et al., 2003), and its expression is associated with auxin distribution in the root apical meristem (Di Laurenzio et al., 1996; Sabatini et al., 1999).

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5.5 Conclusion and perspectives AR formation is an essential step in the vegetative propagation of plants. It is a complex process that is affected by multiple endogenous (homeostasis of growth regulators, level of phenolic compounds, enzymatic activities) and exogenous factors, including nutritional status, associated stress responses, biotic interactions and genetic characteristics. The mechanisms by which ARs are formed are still unknown; nevertheless, recent results obtained in Arabidopsis and rice started to shed light on the underlying molecular mechanisms. The further characterization of the isolated AR mutants and the identification of the mutated genes will be instrumental for a better understanding of the regulation of the adventitious rooting mechanisms. In the future, the sequences of the Arabidopsis genes might also represent the basis for a ‘candidate gene’ approach in trees, and lead to the identification of the genes involved in the control of rooting ability in trees; mapping of these genes will possibly help to unveil the role and molecular function of the detected QTLs.

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Fogac¸a, C.M. and Fett-Neto, A.G. (2005). Role of auxin and its modulators in the adventitious rooting of Eucalyptus species differing in recalcitrance. Plant Growth Regul. 45, 1–10. Martin, R.C., Mok, D.W.S., Smets, R., Van Onckelen, H.A. and Mok, M.C. (2001). Development of transgenic tobacco harboring a zeatin O-glucosyltransferase gene from Phaseolus. In Vitro Cell. Dev. Biol. Plant 37, 354–360. Martinez-Pastur, G., Zappacosta, D., Arena, M. and Curvetto, N. (2001). Changes in isoperoxidase patterns during the in vitro rooting of Nothofagus antarctica. Bulg. J. Plant Physiol. 27, 43–53. Maurel, C., Barbier-Brygoo, H., Spena, A., Temp´e, J. and Guern, J. (1991). Single rol genes from the Agrobacterium rhizogenes TL -DNA alter some of the cellular responses to auxin in Nicotiana tabacum. Plant Physiol. 97, 212–216. McDonald, M.P. and Visser, E.J.W. (2003). A study of the interaction between auxin and ethylene in wild type and transgenic ethylene-insensitive tobacco during adventitious root formation induced by stagnant root zone conditions. Plant Biol. 5, 550–556. Metaxas, D.J., Syros, T.D., Yupsanis, T. and Economou, A.S. (2004). Peroxidases during adventitious rooting in cuttings of Arbutus unedo and Taxus baccata as affected by plant genotype and growth regulator treatment. Plant Growth Regul. 44, 257– 266. Mikkelsen, M.D., Naur, P. and Halkier, B.A. (2004). Arabidopsis mutants in the C-S lyase of glucosinolate biosynthesis establish a critical role for indole-3-acetaldoxime in auxin homeostasis. Plant J. 37, 770–777. Milborrow, B.V. (1984). Inhibitors. In Advanced Plant Physiology, M.B. Wilkins (ed.). London, Pitman Publishing, pp. 76–110. Miller, C.R., Ochoa, I., Nielsen, K.L., Beck, D. and Lynch, J.P. (2003). Genetic variation for adventitious rooting in response to low phosphorus availability: potential utility for phosphorus acquisition from stratified soils. Funct. Plant Biol. 30, 973–985. Moriuchi, H., Okamoto, C., Nishihama, R., Yamashita, I., Machida, Y. and Tanaka, N. (2004). Nuclear localization and interaction of RolB with plant 14-3-3 proteins correlates with induction of adventitious roots by the oncogene rolB. Plant J. 38, 260–275. Nag, S., Saha, K. and Choudhuri, M.A. (2001). Role of auxin and polyamines in adventitious root formation in relation to changes in compounds involved in rooting. J. Plant Growth Regul. 20, 182–194. Niemi, K., H¨aggman, H. and Sarjala, T. (2002a). Effects of exogenous diamines on the interaction between ectomycorrhizal fungi and adventitious root formation in Scots pine in vitro. Tree Physiol. 22, 373–381. Niemi, K., Julkunen-Tiitto, R., Tegelberg, R. and H¨aggman, H. (2005). Light sources with different spectra affect root and mycorrhiza formation in Scots pine in vitro. Tree Physiol. 25, 123–128. Niemi, K., Salonen, M., Ernstsen, A., Heinonen-Tanski, H. and H¨aggman, H. (2000). Application of ectomycorrhizal fungi in rooting of Scots pine fascicular shoots. Can. J. For. Res. 30, 1221–1230. Niemi, K., Scagel, C. and H¨aggman, H. (2004). Application of ectomycorrhizal fungi in vegetative propagation of conifers. Plant Cell Tissue Organ Cult. 78, 83–91. Niemi, K., Vuorinen, T., Ernstsen, A. and H¨aggman, H. (2002b). Ectomycorrhizal fungi and exogenous auxins influence root and mycorrhiza formation of Scots pine hypocotyl cuttings in vitro. Tree Physiol. 22, 1231–1239.

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Takahashi, F., Sato-Nara, K., Kobayashi, K., Suzuki, M. and Suzuki, H. (2003). Sugar-induced adventitious roots in Arabidopsis seedlings. J. Plant Res. 116, 83– 91. Tal, M. (1966). Abnormal stomatal behavior in wilty mutants of tomato. Plant Physiol. 41, 1387–1391. Tanaka, N., Fujikawa, Y., Aly, M.A.M., Saneoka, H., Fujita, K. and Yamashita, I. (2001). Proliferation and rol gene expression in hairy root lines of Egyptian clover. Plant Cell Tissue Organ Cult. 66, 175–182. Tang, W. and Newton, R.J. (2005). Polyamines promote root elongation and growth by increasing root cell division in regenerated Virginia pine (Pinus virginiana Mill.) plantlets. Plant Cell Rep. 24, 581–589. Thompson, A.J., Thorne, E.T., Burbidge, A., Jackson, A.C., Sharp, R.E. and Taylor, I.B. (2004). Complementation of notabilis, an abscisic acid-deficient mutant of tomato: importance of sequence context and utility of partial complementation. Plant Cell Environ. 27, 459–471. Tiburcio, A.F., Gendy, C.A. and Tran Thanh Van, K. (1989). Morphogenesis in tobacco subepidermal cells: putrescine as marker of root differentiation. Plant Cell Tissue Organ Cult. 19, 43–54. Tonon, G., Kevers, C. and Gaspar, T. (2001). Changes in polyamines, auxins and peroxidase activity during in vitro rooting of Fraxinus angustifolia shoots: an auxinindependent rooting model. Tree Physiol. 21, 655–663. Ueda, J. (1989). Promotive effect of capillarol and related compounds on root growth. Physiol. Plant. 76, 42–46. Van Der Krieken, W.M., Breteler, H. and Visser, M.H.M. (1992). Uptake and metabolism of indolebutyric acid during root formation on Malus microcuttings. Acta Bot. Neerl. 41, 435–442. Veierskov, B. (1988). Relations between carbohydrates and adventitious root formation. In Adventitious Root Formation in Cuttings, T.D. Davis, B.E. Haissig and N. Sankhla (eds). Portland, Dioscorides Press, pp. 70–78. Vera, P., Lamb, C. and Doerner, P.W. (1994). Cell-cycle regulation of hydroxyprolinerich glycoprotein HRGPnt3 gene expression during the initiation of lateral root meristems. Plant J. 6, 717–727. Vidal, N., Arellano, G., San-Jos´e, M.C., Vieitez, A.M. and Ballester, A. (2003). Developmental stages during the rooting of in-vitro-cultured Quercus robur shoots from material of juvenile and mature origin. Tree Physiol. 23, 1247–1254. Visser, E.J.W., Blom, C.W.P.M. and Voesenek, L.A.C.J. (1996). Flooding-induced adventitious rooting in Rumex: morphology and development in an ecological perspective. Acta Bot. Neerl. 45, 17–28. ¨ Werner, T., Motyka, V., Laucou, V., Smets, R., Van Onckelen, H. and Schmulling, T. (2003). Cytokinin-deficient transgenic Arabidopsis plants show multiple developmental alterations indicating opposite functions of cytokinins in the regulation of shoot and root meristem activity. Plant Cell 15, 2532–2550. ¨ Werner, T., Motyka, V., Strnad, M. and Schmulling, T. (2001). Regulation of plant growth by cytokinin. Proc. Natl. Acad. Sci. U. S. A. 98, 10487–10492. White, F.F., Taylor, B.H., Huffman, G.A., Gordon, M.P. and Nester, E.W. (1985). Molecular and genetic analysis of the transferred DNA regions of the root-inducing plasmid of Agrobacterium rhizogenes. J. Bacteriol. 164, 33–44. Wilcox, J.R. and Farmer, R.E., Jr. (1968). Heritability and C effects in early growth of eastern cottonwood cuttings. Heredity 23, 239–245.

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Annual Plant Reviews (2009) 37, 157–174 doi: 10.1002/9781444310023.ch6

www.interscience.wiley.com

Chapter 6

ROOT GRAVITROPISM Ranjan Swarup and Malcolm J. Bennett Centre for Plant Integrative Biology and Plant Sciences Division, School of Biosciences, University of Nottingham, Sutton Bonington Campus, Loughborough LE12 5RD, UK

Abstract: Gravity represents a critical environmental signal for land plants that profoundly influences their growth and development. Plants correct changes in their orientation employing a differential growth response, termed gravitropism. Root gravitropism can be divided into several successive phases: gravity perception, gravitropic signal transmission, gravitropic response and a fourth phase that we term gravitropic signal attenuation which serves to prevent further root bending. Gravity perception induces the formation of a lateral auxin gradient. A large number of the auxin-related components that make up the gravitropic signalling and response machinery have been identified in the model plant Arabidopsis thaliana highlighting the central importance of this signal. The past 5 years have also seen several important breakthroughs which have led to the identification of the root cells and tissues that transport, and respond to, the lateral auxin gradient. The review will focus on these recent advances in our understanding about the molecular and cellular mechanisms that regulate root gravitropism. Keywords: root gravitropism; gravity perception; auxin transport; Arabidopsis; lateral auxin gradient

6.1 Introduction Primary roots develop the ability to respond to gravity prior to emergence from the seed coat (Ma and Hasenstein, 2006), aiding seedling establishment. When secondary roots emerge from the primary root, they initially grow out horizontally (Mullen and Hangarter, 2003), whilst higher orders of lateral roots often exhibit randomized directions. These diagravitropic and plagiotropic patterns of root growth greatly facilitate the acquisition of water and nutrients (Basu et al., 2007), particularly in the topsoil where phosphorus availability is the greatest (Lynch and Brown, 2001). This chapter will primarily focus on recent advances in our knowledge about the molecular mechanisms that regulate gravitropism in the primary root of the model plant Arabidopsis thaliana. The past 5 years have seen several 157

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Root Development important breakthroughs in our understanding about the molecular components, signals and tissues that perform key functions during root gravitropism. Our review will concentrate on these latest developments. Readers are directed to a number of excellent reviews for details about earlier root gravitropic research (Konings, 1995; Muday, 2001; Tasaka et al., 2001; Boonsirichai et al., 2002; Blancaflor and Masson, 2003; Perbal and Driss-Ecole, 2003; Morita and Tasaka, 2004) or related root tropisms (Esmon et al., 2005). Root gravitropism has traditionally been divided into three phases: gravity perception, gravitropic signal transmission and gravitropic response. Recent research suggests that a later phase exists, which we term gravitropic signal attenuation that serves to prevent further root bending. Each of these phases will be reviewed in sequence, describing the latest understanding of the molecular, cellular and tissue scale mechanisms that underpin root gravitropism.

6.2 6.2.1

Gravity perception The columella represents the primary site of gravity perception

The primary site of gravity perception in the Arabidopsis root resides in the root cap (Blancaflor and Masson, 2003; Morita and Tasaka, 2004). Experiments involving laser ablation (Blancaflor et al., 1998) and heavy-ion microbeam irradiation (Tanaka et al., 2002) have mapped the site of graviperception in the Arabidopsis root cap in great detail. The Arabidopsis root cap is consisted of lateral root cap (LRC) cells flanking several storeys of columella cells termed S1, S2, S3, etc. (Figs. 6.1a and 6.1b). The effect of laser ablating individual columella cells or storeys has been assessed on several gravitropic parameters (Blancaflor et al., 1998). Roots in which S1 and S2 columella cells had been ablated showed the greatest reduction in curvature after gravistimulation, had the longest presentation times (i.e. minimum exposure time to a 1g field to induce a response; Sack, 1991) and deviated the most from vertical growth. Although ablating S3 cells caused a reduction in root gravitropism, it did not affect the presentation time, suggesting an impaired transmission of the gravity signal, rather than a major reduction in sensitivity to the gravity stimulus. These results imply that S1 and S2 columella layers represent the primary site of gravity sensing in the Arabidopsis root. 6.2.2

Statoliths function as gravity sensors

Ultrastructural studies have revealed that columella cells lack a central vacuole and contain freely sedimenting starch-filled amyloplasts, termed statoliths (Sack, 1991). According to the ‘starch-statolith hypothesis’, gravity is perceived by sedimenting statoliths within columella cells (Sack, 1997;

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Fig. 6.1c). The sedimentation of statoliths would activate an, as yet, unknown receptor triggering a signal transduction pathway to promote the development of a gravitropic curvature. This hypothesis has gained significant support from genetic studies that revealed that Arabidopsis starch-deficient mutants were less sensitive to gravity than wild-type roots (Kiss et al., 1996; MacCleery and Kiss, 1999; Fitzelle and Kiss, 2001). For example, starch content and root gravitropic response have been found to correlate directly (Kiss et al., 1996). Three starch-deficient mutants with variable starch contents were selected and four parameters used to assay gravitropism. The starchless mutants had a reduced gravitropic response when compared to the wild-type roots, whereas mutants with intermediate starch contents (between 50 and 60% of the wild-type level) had an intermediate gravitropic response. These results suggest that 51–60% of the wild-type level of starch is near the threshold amount needed for full gravitropic sensitivity. Through hypergravity experiments, the gravitropic defect in starchdeficient mutants could be corrected to wild-type level at 5g and this was correlated with the sedimentation of statoliths (Fitzelle and Kiss, 2001). High gradient magnetic fields that were used to mimic a gravitational field, exploiting the diamagnetic property of starch, Arabidopsis revealed that Arabidopsis wild-type roots could respond to high magnetic fields but under the same conditions the starchless TC7 mutant failed to do so (Kuznetsov and Hasenstein, 1996). In view of the large body of experimental evidence supporting the starch statolith hypothesis, the general consensus is that it represents the main mechanism of gravity perception in roots. However, other mechanisms might also operate in parallel (Wolverton et al., 2002a, 2002b; LaMotte and Pickard, 2004). 6.2.3

Translating statolith sedimentation into (a)gravitropic signal(s)

How does sedimentation of statoliths trigger a gravitropic response? Which molecular components are required to translate statolith sedimentation into (a)gravitropic signal(s)? What is the nature of this gravitropic signal? Whereas answers to the first question remain elusive, significant progress has recently been made addressing the last two questions. Friml et al. (2002) have shown that targeting of the auxin efflux carrier PIN3 is rapidly altered in columella cells following a gravity stimulus: 2 minutes after root reorientation, the originally symmetric distribution of PIN3 partially relocalized to the lower face of columella cells and within 5 minutes PIN3 relocalization to the lower face of columella cells was complete (Fig. 6.1c). This rapid repositioning of PIN3 to the lower face of columella cells provides a novel molecular mechanism for the observed establishment of a lateral auxin response gradient following a gravity stimulus (Rashotte et al., 2001; Ottenschl¨ager et al., 2003; Fig. 6.3). Surprisingly, the pin3 mutant exhibits a weak agravitropic root phenotype that might be due to redundancy in

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Root Development expression among members of the PIN family since PIN3, PIN4 and PIN7 have all been reported to be expressed in columella cells (Blilou et al., 2005; Paponov et al., 2005; Vieten et al., 2005). In addition, interpretation of mutant phenotypes of single pin mutants can be difficult due to redundant ectopic expression by other PIN genes (Blilou et al., 2005). Trafficking of PIN proteins is regulated by the ARF-GEF protein, GNOM (Geldner et al., 2003). The GNOM gene is strongly expressed in columella cells (Geldner et al., 2004). Partial loss-of-function alleles of GNOM disrupt root gravitropism. This phenotype probably results from impaired relocalization of multiple PIN proteins, such as PIN3, following a gravity stimulus. Rapid changes in the pH of root caps have also been observed to occur during the early stages of gravity perception (Scott and Allen, 1999; Fasano et al., 2001; Hou et al., 2004). Cytoplasmic pH in columella cells increased from 7.2 to 7.6 within the first minute and was followed by the sustained acidification of the apoplast (Fasano et al., 2001). These pH changes appear to be linked with statolith sedimentation since they were not observed in the starchless mutant pgm. Significantly, disrupting the pH shift in columella cells by acidifying or alkalinizing agents altered the gravitropic responses, suggesting that these pH changes are of functional importance. How changes in columella pH would initiate a gravitropic response is not yet clear, but a key role for the Dna J-like proteins ALTERED RESPONSE TO GRAVITY (ARG1) and ARG1-like 2 (ARL2) has been proposed (Sedbrook et al., 1999; Boonsirichai et al., 2003; Guan et al., 2003). ARG1 and the closely related gene ARL2 encode Dna J-like peripheral membrane proteins. Mutations in ARG1 confer an agravitropic root phenotype and block cytosolic alkalinization of columella cells (Boonsirichai et al., 2003). Expressing the ARG1 gene in columella cells only is sufficient to rescue the arg1 agravitropic defect, suggesting that ARG1 plays a key role in early stages of gravity sensing. ARG1 is associated with microsomal membranes and the cytoskeleton and has been proposed to be involved in regulating the trafficking or function of membrane proteins needed for the early phases of gravity signal transduction such as PIN3 (Boonsirichai et al., 2003). Consistent with this model, the expression of the auxin-responsive reporter DR5::GUS was altered in the arg1 mutant background.

6.3 6.3.1

Root gravitropic signal transmission Auxin represents the primary gravitropic signal

The root columella and elongation zone tissues are spatially distinct (Fig. 6.2a), necessitating the transmission of (a)gravitropic signal(s). Auxin represents a strong candidate for the gravitropic signal given the large number of auxin transport and response mutants that exhibit root gravitropic defects (Table 6.1). In addition, auxin-responsive reporters (Rashotte et al., 2001;

ABC transporter family ARF-GEF

ABC transporter family Novel protein PP2A Chalcone synthase Potassium transporter

PGP1 GNOM

PGP4 AXR4 RCN1 TT4 TRH1

Phospholipase D

GRAVI-ATTENUATION PGP19 ABC transporter family

PLD Zeta2

GRAVI-RESPONSE AXR1 Ubiquitin-activating enzyme AXR2 AUX/IAA protein AXR3 AUX/IAA protein IAA14/SLR1 AUX/IAA protein IAA3/SHY2 AUX/IAA protein ARF7,19 Auxin response factors TIR1 (AFB2-4) F-box protein (auxin receptor)

Agravitropic roots

Auxin influx carrier

AUX1

Reduced root gravitropism

Reduced root gravitropism

Agravitropic roots Agravitropic roots (GoF) Agravitropic roots (GoF) Agravitropic roots (GoF) Agravitropic roots (GoF) Reduced root gravitropism Reduced root gravitropism

Reduced root gravitropism Reduced root gravitropism in weak alleles Reduced root gravitropism Agravitropic roots Reduced root gravitropism Altered root gravitropism Altered root gravitropism

Reduced root gravitropism Agravitropic roots

Reduced root and shoot gravitropism Reduced root and shoot gravitropism Reduced root and shoot gravitropism Root cap morphogenesis and altered gravisensitivity

Mutant Phenotype

GRAVI-SIGNAL TRANSMISSION PIN3 Auxin efflux carrier PIN2/EIR1/AGR1 Auxin efflux carrier

GRAVI-PERCEPTION PGM Phosphoglucomutase ARG1 Dna J protein ARL2 Dna J protein ADK1 Adenosine kinase

Gene/Protein

Summary of the key genes that regulate root gravitropism in Arabidopsis

Gene Name

Table 6.1

Lin and Wang (2005)

Leyser et al. (1993) Timpte et al. (1994), Nagpal et al. (2000) Leyser et al. (1996), Rouse et al. (1998) Fukaki et al. (2002, 2005) Weijers et al. (2005) Okushima et al. (2005) Ruegger et al. (1998), Dharmasiri et al. (2005a, 2005b), Kepinski and Leyser (2005) Li and Xue (2007)

Terasaka et al. (2005) Hobbie and Estelle (1995), Dharmasiri et al. (2006) Garbers et al. (1996), Deru`ere et al. (1999) Buer et al. (2006) Vicente-Agullo et al. (2004)

Friml et al. (2002) M¨uller et al. (1998), Paciorek et al. (2005), Abas et al. (2006) Bennett et al. (1996), Swarup et al. (2001, 2004, 2005), Yang et al. (2006) Lin and Wang (2005) Geldner et al. (2003, 2004)

Caspar and Pickard (1989) Sedbrook et al. (1999), Boonsirichai et al. (2003) Guan et al. (2003) Young et al. (2006)

References

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Root Development Boonsirichai et al., 2003; Ottenschl¨ager et al., 2003; Fig. 6.3) exhibit asymmetric changes in their patterns of expression in columella and LRC cells following a gravity stimulus, consistent with the transmission of a lateral auxin gradient to the lower side of the root. However, many other plant signals have also been linked with gravitropism (reviewed by Philosoph-Hadas et al., 2005) including brassinosteroids (Li et al., 2005) and ethylene (Buer et al., 2006). Cytokinin has recently been reported to be asymmetrically redistributed (Aloni et al., 2004) during a root gravitropic response. Nevertheless, biosynthetic or response mutants for these hormone signals remain gravitropic, questioning their primary functional importance. Determining whether auxin acts as the gravitropic signal has been hampered by the inability to visualize dynamic expression changes of auxinresponsive reporters in graviresponsive elongation zone tissues (Rashotte et al., 2001; Boonsirichai et al., 2003; Ottenschl¨ager et al., 2003; Fig. 6.3). Hence, it is possible that auxin does not act directly on elongation zone cells but involves other signalling intermediates (Fasano et al., 2002; Monshausen and Sievers, 2002; Plieth and Trewavas, 2002; Hu et al., 2005). This question prompted us to precisely map those root tissues required to transport auxin during a gravitropic response (Swarup et al., 2005). Targeted expression of the auxin influx carrier AUX1 (Yang et al., 2006) to specific tissues in an aux1 mutant background helped us to demonstrate that root gravitropism is dependent on auxin being transported via the LRC and elongating epidermal cells. Our results provide conclusive evidence that auxin functions as the primary gravitropic signal linking gravisensing and graviresponsive tissues (Swarup et al., 2005).

6.3.2

Channelling the lateral auxin gradient via the LRC

The gravity-induced auxin gradient is initially transported from the columella cells to elongation zone tissues via the LRC (Fig. 6.2a). The auxin influx and efflux carriers AUX1 and PIN2 expressed in the LRC, appear critical for auxin transport during a root gravitropic response. Root gravitropism is disrupted when mutations in either of these genes or treatment with inhibitors of each class of auxin carrier block this basipetal transport pathway (Rashotte et al., 2000; Swarup et al., 2001, 2005; Ottenschl¨ager et al., 2003). The asymmetric localization of PIN2 at the apical plasma membrane face of LRC cells directs the lateral auxin gradient toward the elongation zone tissues (Mullen et al., 1998a, 1998b (see references); Figs. 6.2b and 6.2d), while the non-polarlocalized AUX1 is essential for the efficient uptake of auxin by LRC cells (Swarup et al., 2001; Figs. 6.2e and 6.2g). The auxin influx carrier AtPGP4 (that belongs to the P-glycoprotein [PGP] class of transporters) is also expressed in LRC cells (Santelia et al., 2005; Terasaka et al., 2005). However, mutations in AtPGP4 do not confer a strong agravitropic phenotype (Santelia et al., 2005; Terasaka et al., 2005) suggesting that this auxin influx carrier plays

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a relatively minor role in the redistribution of the gravity-induced lateral auxin gradient compared to AUX1. 6.3.3

Delivering auxin to elongation zone tissues

Auxin influx and efflux carrier activities are also required to transport the lateral auxin gradient in elongation zone tissues. Simulations of auxin fluxes through elongation zone tissues suggest that epidermal expressed AUX1 and PIN2 minimize the effect of radial diffusion while facilitating basipetal auxin transport (Swarup et al., 2005). The asymmetric localization of PIN2 at the apical PM face of epidermal cells (Fig. 6.2c) provides directionality to the lateral auxin gradient (Wisniewska et al., 2006), while the non-polar-localized AUX1 (Fig. 6.2f) appears essential for the efficient auxin uptake by expanding epidermal cells. Mutations in the AXR4 gene that disrupt plasma membrane targeting of AUX1 in elongating epidermal cells result in an agravitropic root phenotype (Dharmasiri et al., 2006). The Arabidopsis primary root elongation zone contains approximately 16 rapidly expanding cells in each longitudinal file of cells (Beemster and Baskin, 2000). Since expressing AUX1 in up to the first six of these rapidly expanding epidermal cells was still not sufficient to rescue the aux1 agravitropic phenotype (Swarup et al., 2005), we concluded that the root gravitropic response requires AUX1 to be expressed in every expanding epidermal cell, presumably to ensure that the lateral auxin gradient persists while it migrates through the elongation zone and drives asymmetric root growth. There is a common misconception that because protonated IAA is membrane permeable, influx carriers play only a supplemental role in auxin distribution. However, cells expressing AUX1 are estimated to have a carrier-mediated IAA influx 15 times greater than the diffusive contribution (Kramer and Bennett, 2006). The rate of IAA membrane diffusion is too slow for root gravitropism to occur based on the aux1 mutant phenotype. However, this biophysical constraint can be overcome through the addition of the membrane-permeable auxin 1-NAA that rescues the aux1 agravitropic defect (Yamamoto and Yamamoto, 1998; Marchant et al., 1999). Auxin has been demonstrated to effect endocytosis of PIN2 (Paciorek et al., 2005). This post-translational mechanism of PIN2 regulation has important functional consequences during root gravitropism (Abas et al., 2006). Following a gravity stimulus, inhibition of endocytosis by the lateral auxin gradient results in PIN2 being retained at the plasma membranes of expanding epidermal cells on the lower side of the root. In contrast, PIN2 is rapidly internalized and degraded in equivalent cells on the upper side of the root with a higher rate of auxin transport on the lower versus upper side of the root as a consequence, further maintaining the lateral auxin gradient established by PIN3. The functional importance of this mechanism for root gravitropic curvature is underlined by the phenotype of pin2 missense allele wav6-52. Unlike the wildtype PIN2 protein, PIN2wav6-52 is no longer degraded. PIN2wav6-52 appears to interfere with the redistribution of auxin following a gravity stimulus,

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Root Development resulting in a reduced lateral auxin gradient and causing an agravitropic root phenotype. Disruption of PIN2 cycling by mutations in the phospholipase Dæ2 gene or the PLD-specific inhibitor 1-butanol also reduces root gravitropism (Li and Xue, 2007). Mutations in PIN2 only reduce basipetal auxin transport by approximately 30% (Rashotte et al., 2000; Shin et al., 2005) and do not cause as severe an agravitropic phenotype as aux1, suggesting that other efflux carriers must function in parallel. The auxin efflux carrier AtPGP1 is also expressed in epidermal cells (Blilou et al., 2005; Blakeslee et al., 2007). Nevertheless, mutations in atpgp1 confer an agravitropic root phenotype (Lin and Wang, 2005). However, the pgp1 pgp19 pin2 triple mutant exhibited an agravitropic root phenotype as severe as that of aux1 (Blakeslee et al., 2007). This additive phenotype suggests that both PIN and PGP classes of auxin efflux carriers transport the lateral auxin gradient during a root gravitropic response.

6.4

The root gravitropic response

Detailed growth measurements of gravitropic root growth in wild-type Arabidopsis reveal that initial root curvature results from differential inhibition of elongation in the distal elongation zone (DEZ), followed by an acceleration of elongation along the top side of the DEZ (Ishikawa and Evans, 1997; Mullen et al., 1998a, 1998b). This pattern of differential growth is consistent with the Cholodny-Went hypothesis that predicts that differential growth patterns are induced by a gravity-induced auxin gradient. 6.4.1

The epidermal auxin response is essential for gravitropic curvature

The lateral auxin gradient is presumed to directly act on expanding cells in all elongation zone tissues. However, simulations using a root elongation zone model suggest that the lateral auxin gradient accumulates 10- to 20fold more in the epidermis than in the underlying cortical and endodermal tissues (Swarup et al., 2005). Partitioning the lateral auxin gradient between root tissues in such a manner is predicted to result in root gravitropic bending being driven primarily by differential cell elongation in the epidermis. We directly tested the predicted functional importance of the epidermis to root gravitropism by selectively disrupting the auxin response in this tissue using the stabilized auxin repressor protein axr3-1 (Rouse et al., 1998). Results from our axr3-1 targeted expression studies demonstrate that the auxin response in expanding epidermal cells (but not the underlying tissues) is essential for the root gravitropism (Swarup et al., 2005). In contrast, the basal rate of root growth is only slightly reduced when the auxin response is disrupted in the epidermis. These contrasting growth effects could be explained if the epidermis primarily regulated differential root growth, whereas basal root

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growth was driven by the expansion of the inner tissues. In such a model, the epidermis would restrain basal root growth in response to a lateral auxin gradient. As the outermost root tissue, the epidermis is ideally positioned to regulate organ bending (Kutschera, 1989). 6.4.2

Molecular targets for the lateral auxin gradient

The lateral auxin gradient causes rapid asymmetric changes in root growth that are detectable within 10 minutes after a gravity stimulus (Ishikawa and Evans, 1997). This initial phase of the gravitropic response is likely to be caused by non-genomic effects of auxin. For example, auxin causes the rapid acidification of the apoplastic space which alters the activities of cell wall enzymes (Cosgrove, 1998). On the other hand, Paciorek et al. (2005) recently described how the gravity-induced auxin gradient can differentially inhibit endocytosis in elongating epidermal cells on the lower (versus upper) side of the root. The later phase of the root gravitropic response is dependent on auxinregulated transcription (Abel and Theologis, 1996) that is controlled by two related families of proteins, designated auxin response factors (ARFs) and auxin/indole-3-acetic acids (AUX/IAA) (reviewed by Leyser, 2006). ARFs regulate transcription by binding to a motif in promoters of auxin-regulated genes, the auxin response element (AuxREs; reviewed by Guilfoyle and Hagen, 2001). Two closely related members of the ARF gene family, ARF7 and ARF19, have been linked with root gravitropism (Okushima et al., 2005). Reverse genetic studies have revealed that root gravitropism is disrupted in the arf7 arf19 double mutant. ARF7 and ARF19 appear to be functionally redundant since loss-of-function mutation in either gene does not confer an agravitropic root phenotype. ARF7 and ARF19 encode transcriptional activators (Ulmasov et al., 1999; Tiwari et al., 2003; Wang et al., 2005). ARF-controlled transcription is inhibited by AUX/IAA proteins that function as repressor proteins that heterodimerize with ARFs (Tiwari et al., 2001), and have been demonstrated to interact with several AUX/IAA proteins including IAA14/SLR1 and IAA3/SHY2 (Fukaki et al., 2005; Weijers et al., 2005). Auxin causes the targeted degradation of AUX/IAA proteins by promoting their interaction with the SCFTIR1/AFB ubiquitin–ligase complex (Dharmasiri et al., 2005a, 2005b; Kepinski and Leyser, 2005). The proteolytic degradation of AUX/IAA proteins derepresses ARF-dependent transcription. Mutations that stabilize AUX/IAA proteins that interact with ARF7 and ARF19 such as iaa14/slr1 and iaa3/shy2-2 disrupt root gravitropism (Fukaki et al., 2002; Weijers et al., 2005). Global gene expression analysis has revealed that auxin-induced gene expression is severely impaired in the arf7 single and arf7 arf19 double mutants (Okushima et al., 2005). The data suggest that the ARF7 and ARF19 proteins play essential roles in auxin-mediated root gravitropism by regulating both unique and partially overlapping sets of target genes. Bioinformatics

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Root Development analysis of ARF7 and ARF19 regulated genes expressed in expanding root epidermal cells has identified a large number of genes with cell wall-related functions (Wilson and Bennett, unpublished results). These include genes such as AtXTH18 and AtXTH19 that encode cell wall enzymes that crosslink cell wall components causing inhibition of cell expansion (Vissenberg et al., 2005). Hence, auxin is likely to cause differential inhibition of cell expansion through the upregulation of such cell wall enzymes. 6.4.3

Auxin also regulates the gravitropic response via other signals

Recent evidence suggests that auxin also regulates root gravitropism via additional signals, such as nitric oxide (NO), cGMP, reactive oxygen species (ROS) and IP3 (Joo et al., 2001, 2005; Hu et al., 2005; Perera et al., 2006). Recently, NO has been reported to accumulate in the tissues on the lower side of gravity-stimulated roots (Hu et al., 2005). This pattern of NO accumulation can be blocked by addition of the auxin transport inhibitor NPA and restored by asymmetric application of auxin. Importantly, inhibition of NO production inhibits root curvature, underlining its physiological importance toward the gravitropic bending response. NO can either signal by directly modifying proteins via nitrosylation (Durner and Klessig, 1999) or elevating cGMP levels (Durner et al., 1998). During root gravitropism, auxin-induced NO appears to signal via cGMP because unilateral application of a membrane permeable analogue 8-Br-cGMP can restore curvature of NPA-treated roots (Hu et al., 2005). Nevertheless, treatment with NO or cGMP inhibitors did not completely block root gravitropism, suggesting that additional signals are required. Production of ROS also appears to be required for root gravitropism (Joo et al., 2001). Like NO, ROS accumulation in tissues is found on the lower side of gravity-stimulated roots and can be mimicked by asymmetric application of auxin. Moreover, unilateral addition of hydrogen peroxide (H2 O2 ) can induce curvature even in NPA-treated roots. Hence, ROS appears to act downstream of auxin. Joo et al. (2005) have reported that auxin induces ROS production during root gravitropism by activating phosphatidylinositol 3-kinase (PtdIns 3 kinase) and pre-treatment with an inhibitor of PtdIns 3-kinase activity blocked auxin-mediated ROS generation and reduced root gravitropic curvature.

6.5

Attenuating the root gravitropic response

Recent evidence suggests that an additional phase, which we term gravitropic signal attenuation, functions to reduce the persistence of the root gravitropic signal, auxin. Several distinct mechanisms appear to operate in columella and elongation zone tissues.

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Treatment of Arabidopsis roots with the actin cytoskeleton inhibitor latrunculin B (Lat B) has been observed to result in enhanced gravitropic curvature (Hou et al., 2004). It was concluded that the actin cytoskeleton is unnecessary for the initial perception of gravity, but functions later to downregulate the root gravitropic response. Consistent with this model, Lat B-treated DR5:GUS roots exhibited a more persistent asymmetric pattern of reporter activity than the controls. For a root to resume vertical growth, the actin cytoskeleton may be required to re-localize auxin efflux carriers (such as PIN3) to their original basal position in the columella plasma membrane (Hou et al., 2004). Following a gravity stimulus PIN2 is retained preferentially on the plasma membranes of expanding epidermal cells on the lower (but not the upper) side of the root (Abas et al., 2006). Auxin treatments for periods of 2 hours and longer enriched intracellular PIN2 and reduced protein levels, which might reflect enhanced turnover and represent a homeostatic mechanism by root cells to control PIN2 levels at later stages of a gravitropic response. Auxininduced PIN2 degradation would reduce auxin transport in cells on the lower side of the root, thereby preventing further root bending. Mutations in the auxin efflux carrier gene AtPGP19 exhibit enhanced root gravitropic curvature (Lin and Wang, 2005). Consistent with its mutant phenotype, the atpgp19 mutant exhibited a more persistent asymmetric pattern of DR5:GUS reporter activity than the wild-type control. Recently, AtPGP19 protein has been localized in cortical and other inner root tissues (Blakeslee et al., 2007) and is ideally positioned to facilitate auxin reflux, which involves the recycling of auxin from epidermal to stele tissues (Blilou et al., 2005). Therefore, AtPGP19 appears to reduce the persistence of the gravitropic signal auxin in elongation zone tissues.

6.6 Future directions The past 5 years have witnessed important breakthroughs in our understanding about the molecular components, signals and tissues that perform key functions during root gravitropism. Nevertheless, many important questions remain to be answered. For example, how does statolith sedimentation result in relocalization of PIN proteins to create a lateral auxin gradient? What roles do signals, such as pH and IP3, play during root gravitropism? At later stages of root gravitropism, how does auxin bring about a differential growth response? Which cell wall enzymes and/or components are targets for regulation? To answer these and other future questions, new and existing tools will have to be applied. Mutant screens will continue to underpin studies of root gravitropism (Table 6.1), but approaches, such as quantitative genetics, should also be embraced. Proteomic approaches promise to uncover components undergoing rapid changes taking place during the initial stages of a gravitropic response (Young et al., 2006), while transcriptomics is ideally

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Root Development placed to pinpoint expression changes in downstream components (Moseyko et al., 2002; Kimbrough et al., 2004). Chemical genetic approaches promise to identify new inhibitors for root gravitropism that will uncover novel molecular targets. Recently, Yamazoe et al. (2005) have identified a new inhibitor of root gravitropism, terfestatin A (TrfA), that blocks the auxin-mediated degradation of Aux/IAA proteins without affecting the interaction between Aux/IAAs and TIR1. After defining the key components and intercellular pathways, researchers must begin to develop a quantitative, rather than just a qualitative, understanding of root gravitropic signalling processes in the Arabidopsis root. Simulations of auxin fluxes within and between root tissues using realistic transport parameters, carrier distributions and three-dimensional cellular geometries will help to realize this important goal (Swarup et al., 2005). Such models provide a valuable tool for researchers by generating new predictions that can be tested experimentally. For example, simulations of a gravityinduced auxin gradient in root elongation zone tissues have predicted that the hormone accumulates preferentially in expanding epidermal (versus cortical and endodermal) cells, prompting experiments that have later revealed the primary importance of the epidermis for gravitropic curvature (Swarup et al., 2005). This example illustrates how new insights can be obtained if biologists embrace a systems biology approach and interact with researchers from other disciplines.

Acknowledgements The authors acknowledge research support from the BBSRC and European Space Agency.

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Root Development Mullen, J.L., Turk, E., Johnson, K., Wolverton, C., Ishikawa, H., Simmons, C. et al. (1998b). Root-growth behavior of the Arabidopsis mutant rgr1. Plant Physiol. 118, 1139–1145. ¨ Muller, A., Guan, C., G¨alweiler, L., T¨anzler, P., Huijser, P., Marchant, A. et al. (1998). AtPIN2 defines a locus of Arabidopsis for root gravitropism control. EMBO J. 17, 6903–6911. Nagpal, P., Walker, L.M., Young, J.C., Sonawala, A., Timpte, C., Estelle, M. et al. (2000). AXR2 encodes a member of the Aux/IAA protein family. Plant Physiol. 123, 563– 573. Okushima, Y., Overvoorde, P.J., Arima, K., Alonso, J.M., Chan, A., Chang, C. et al. (2005). Functional genomic analysis of the AUXIN RESPONSE FACTOR gene family members in Arabidopsis thaliana: unique and overlapping functions of ARF7 and ARF19. Plant Cell 17, 444–463. Ottenschl¨ager, I., Wolff, P., Wolverton, C., Bhalerao, R.P., Sandberg, G., Ishikawa, H. et al. (2003). Gravity-regulated differential auxin transport from columella to lateral root cap cells. Proc. Natl. Acad. Sci. U. S. A. 100, 2987–2991. Paciorek, T., Zaz´ımalov´a, E., Ruthardt, N., Petr´asˇ ek, J., Stierhof, Y.-D., Kleine-Vehn, J. et al. (2005). Auxin inhibits endocytosis and promotes its own efflux from cells. Nature 435, 1251–1256. Paponov, I.A., Teale, W.D., Trebar, M., Blilou, I. and Palme, K. (2005). The PIN auxin efflux facilitators: evolutionary and functional perspectives. Trends Plant Sci. 10, 170–177. Perbal, G. and Driss-Ecole, D. (2003). Mechanotransduction in gravisensing cells. Trends Plant Sci. 8, 498–504. Perera, I.Y., Hung, C.-Y., Brady, S., Muday, G.K. and Boss, W.F. (2006). A universal role for inositol 1,4,5-trisphosphate-mediated signaling in plant gravitropism. Plant Physiol. 140, 746–760. Philosoph-Hadas, S., Friedman, H. and Meir, S. (2005). Gravitropic bending and plant hormones. Vitam. Horm. 72, 31–78. Plieth, C. and Trewavas, A.J. (2002). Reorientation of seedlings in the earth’s gravitational field induces cytosolic calcium transients. Plant Physiol. 129, 786–796. Rashotte, A.M., Brady, S.R., Reed, R.C., Ante, S.J. and Muday, G.K. (2000). Basipetal auxin transport is required for gravitropism in roots of Arabidopsis. Plant Physiol. 122, 481–490. Rashotte, A.M., DeLong, A. and Muday, G.K. (2001). Genetic and chemical reductions in protein phosphatase activity alter auxin transport, gravity response, and lateral root growth. Plant Cell 13, 1683–1697. Rouse, D., Mackay, P., Stirnberg, P., Estelle, M. and Leyser, O. (1998). Changes in auxin response from mutations in an AUX/IAA gene. Science 279, 1371–1373. Ruegger, M., Dewey, E., Gray, W.M., Hobbie, L., Turner, J. and Estelle, M. (1998). The TIR1 protein of Arabidopsis functions in auxin response and is related to human SKP2 and yeast Grr1p. Genes Dev. 12, 198–207. Sack, F.D. (1991). Plant gravity sensing. Int. Rev. Cytol. 127, 193–252. Sack, F.D. (1997). Plastids and gravitropic sensing. Planta 203, S63–S68. ¨ Santelia, D., Vincenzetti, V., Azzarello, E., Bovet, L., Fukao, Y., Duchtig, P. et al. (2005). MDR-like ABC transporter AtPGP4 is involved in auxin-mediated lateral root and root hair development. FEBS Lett. 579, 5399–5406. Scott, A.C. and Allen, N.S. (1999). Changes in cytosolic pH within Arabidopsis root columella cells play a key role in the early signaling pathway for root gravitropism. Plant Physiol. 121, 1291–1298.

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Sedbrook, J.C., Chen, R. and Masson, P.H. (1999). ARG1 (Altered Response to Gravity) encodes a DnaJ-like protein that potentially interacts with the cytoskeleton. Proc. Natl. Acad. Sci. U. S. A. 96, 1140–1145. Shin, H., Shin, H.-S., Guo, Z., Blancaflor, E.B., Masson, P.H. and Chen, R. (2005). Complex regulation of Arabidopsis AGR1/PIN2-mediated root gravitropic response and basipetal auxin transport by cantharidin-sensitive protein phosphatases. Plant J. 42, 188–200. Swarup, R., Friml, J., Marchant, A., Ljung, K., Sandberg, G., Palme, K. et al. (2001). Localization of the auxin permease AUX1 suggests two functionally distinct hormone transport pathways operate in the Arabidopsis root apex. Genes Dev. 15, 2648–2653. Swarup, R., Kargul, J., Marchant, A., Zadik, D., Rahman, A., Mills, R. et al. (2004). Structure-function analysis of the presumptive Arabidopsis auxin permease AUX1. Plant Cell 16, 3069–3083. Swarup, R., Kramer, E.M., Perry, P., Knox, K., Leyser, H.M.O., Haseloff, J. et al. (2005). Root gravitropism requires lateral root cap and epidermal cells for transport and response to a mobile auxin signal. Nat. Cell Biol. 7, 1057–1065. Tanaka, A., Kobayashi, Y., Hase, Y. and Watanabe, H. (2002). Positional effect of cell inactivation on root gravitropism using heavy-ion microbeams. J. Exp. Bot. 53, 683–687. Tasaka, M., Kato, T. and Fukaki, H. (2001). Genetic regulation of gravitropism in higher plants. Int. Rev. Cytol. 206, 135–154. Terasaka, K., Blakeslee, J.J., Titapiwatanakun, B., Peer, W.A., Bandyopadhyay, A., Makam, S.N. et al. (2005). PGP4, an ATP binding cassette P-glycoprotein, catalyzes auxin transport in Arabidopsis thaliana roots. Plant Cell 17, 2922–2939. Timpte, C., Wilson, A.K. and Estelle, M. (1994). The axr2-1 mutation of Arabidopsis thaliana is a gain-of-function mutation that disrupts an early step in auxin response. Genetics 138, 1239–1249. Tiwari, S.B., Hagen, G. and Guilfoyle, T. (2003). The roles of auxin response factor domains in auxin-responsive transcription. Plant Cell 15, 533–543. Tiwari, S.B., Wang, X.-J., Hagen, G. and Guilfoyle, T.J. (2001). AUX/IAA proteins are active repressors, and their stability and activity are modulated by auxin. Plant Cell 13, 2809–2822. Ulmasov, T., Hagen, G. and Guilfoyle, T.J. (1999). Dimerization and DNA binding of auxin response factors. Plant J. 19, 309–319. Vicente-Agullo, F., Rigas, S., Desbrosses, G., Dolan, L., Hatzopoulos, P. and Grabov, A. (2004). Potassium carrier TRH1 is required for auxin transport in Arabidopsis roots. Plant J. 40, 523–535. Vieten, A., Vanneste, S., Wisniewska, J., Benkov´a, E., Benjamins, R., Beeckman, T. et al. (2005). Functional redundancy of PIN proteins is accompanied by auxin-dependent cross-regulation of PIN expression. Development 132, 4521–4531. Vissenberg, K., Oyama, M., Osato, Y., Yokoyama, R., Verbelen, J.-P. and Nishitani, K. (2005). Differential expression of AtXTH17, AtXTH19 and AtXTH20 genes in Arabidopsis roots. Physiological roles in specification in cell wall construction. Plant Cell Physiol. 46, 192–200. Wang, S., Tiwari, S.B., Hagen, G. and Guilfoyle, T.J. (2005). AUXIN RESPONSE FACTOR7 restores the expression of auxin-responsive genes in mutant Arabidopsis leaf mesophyll protoplasts. Plant Cell 17, 1979–1993. Weijers, D., Benkova, E., J¨ager, K.E., Schlereth, A., Hamann, T., Kientz, M. et al. (2005). Developmental specificity of auxin response by pairs of ARF and Aux/IAA transcriptional regulators. EMBO J. 24, 1874–1885.

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Root Development Wisniewska, J., Xu, J., Seifertov´a, D., Brewer, P.B., Ruzicka, K., Blilou, I. et al. (2006). Polar PIN localization directs auxin flow in plants. Science 312, 883. Wolverton, C., Ishikawa, H. and Evans, M.L. (2002a). The kinetics of root gravitropism: dual motors and sensors. J. Plant Growth Regul. 21, 102–112. Wolverton, C., Mullen, J.L., Ishikawa, H. and Evans, M.L. (2002b). Root gravitropism in response to a signal originating outside of the cap. Planta 215, 153–157. Yamamoto, M. and Yamamoto, K. (1998). Differential effects of 1-naphthalenic acid, indole-3-acetic acid and 2,4-dichlorophenoxyacetic acid on the gravitropic response of roots in an auxin resistant mutant of Arabidopsis, aux1. Plant Cell Physiol. 39, 660–664. Yamazoe, A., Hayashi, K.-I., Kepinski, S., Leyser, O. and Nozaki, H. (2005). Characterization of terfestatin A, a new specific inhibitor for auxin signaling. Plant Physiol. 139, 779–789. Yang, Y., Hammes, U.Z., Taylor, C.G., Schachtman, D.P. and Nielsen, E. (2006). Highaffinity auxin transport by the AUX1 influx carrier protein. Curr. Biol. 16, 1123–1127. Young, L.-S., Harrison, B.R., Narayana Murthy U.M., Moffatt, B.A., Gilroy, S. et al. (2006). Adenosine kinase modulates root gravitropism and cap morphogenesis in Arabidopsis. Plant Physiol. 142, 564–573.

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Figure 1.1 Root development. (a) Schematic representation of the Arabidopsis root. (b) Root specification during early embryogenesis. Focus is on the instructive elements in root specification. Colored serrated lines at the border of cells indicate respective PIN transport facilitators. (c) Hypophysis specification involving auxin (IAA) and its signal transducers BDL and MP. Abbreviations: Col, columella; Cor, cortex; En, endodermis; Epi, epidermis; Lrc, lateral root cap; Vasc, vascular tissue. Yellow indicates high levels of auxin accumulation. For detailed description, see text.

Stem cells

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Figure 1.2 Genes controlling different aspects of root development. (a) Combinatorial code of root stem cell niche specification. Peak levels of PLT expression are indicated in blue and SHR and SCR protein expression in green. WUS and WOX5 represent conserved gene activities for stem cell maintenance. (b) Asymmetric ground tissue division. (c) Interpretation of PLT proteins by the PIN protein-mediated auxin gradient, determining the zones of cell division, expansion and differentiation. Abbreviations: Cor, cortex; DIFF, cell differentiation; DIV, zone of cell division (meristem); En, endodermis; EXP, cell expansion; QC, quiescent center; SCN, stem cell niche; Vasc, vascular tissue; For detailed description, see text.

Epidermal initial (a)

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Figure 3.2 Formation of the root meristem and epidermal initials in the heart stage embryo. (a) A schematic of a longitudinal section through a heart stage embryo. It is hypothesized that signals (pale blue arrow) from the QC (orange) induce initial status on the adjacent ring of protoderm cells (shaded dark grey). (b) A schematic of a transverse section through the quiescent centre (orange) and the protoderm in grey. (c) A schematic of a longitudinal view of the epidermal initial at late heart stage. The epidermal initial divides to form the lateral root cap (light grey) and a new initial (dark grey).

Figure 3.3 Establishment of cell pattern in the Arabidopsis root epidermis. (a) Transverse section in the meristematic region of an Arabidopsis root stained with toluidine blue. Cells in the hair (H) position (adjacent to two cortical cells) are stained more intensely than epidermal cells in the non-hair (N) position (adjacent to one cortical cell). Scale bar, 25 µm. (b) Schematic representation of the laser ablation experiment that shows that epidermal cell identity is determined by position relative to underlying cortical cells. When a cell in the H position is ablated (1), a cell in the N position divides and takes the position of the ablated cell (2). The new cell in the H position develops H fate despite its N lineage (3). Cortical cells are white, N cells are yellow and H cells are grey. (c) Schematic representation of the clonal analysis that shows that epidermal cell identity is determined by position relative to underlying cortical cells. Occasionally, a cell in the H position (1) undergoes a longitudinal anticlinal division (2). The daughter cell that is adjacent to two cortical cells develops H identity, but the daughter cell that is adjacent to only one cortical cell develops N identity (3). (d) Schematic representation of the pathway that controls cell specification in the root epidermis. Green arrows indicate an unknown positional signal from cortical cells. Black arrows indicate interaction at the level of gene expression. Red arrows indicate cell-to-cell protein movement. See text for details.

Figure 4.1 Confocal images and schematic view of lateral root primordium development in Arabidopsis. (a) Pericycle founder cells undergo asymmetric anticlinal divisions to form a Stage I LRP. Cells then divide periclinally and anticlinally to form a two-layered Stage II LRP. OL, outer layer; IL, inner layer. The outer and inner layers divide periclinally to form first a Stage III and then a Stage IV LRP. OL1, outer layer 1; OL2, outer layer 2; IL1, inner layer 1; IL2, inner layer 2. Stage V is characterized by subdivision of OL1 to form a root cap and epidermis. In Stage VI, OL2 divides to form the endodermal and cortex layers, surrounding the future QC and endodermal/cortex initials. At this stage, a characteristic pattern of four central cells surrounded by four peripheral cells on each side can be clearly seen in median section. Another round of cell divisions creates a pattern of eight central cells surrounded by eight peripheral cells on each side (Stage VII). The LRP is now ready for emergence from the parent root and meristem activation. IL1 and IL2 divide to form the stele and stele initials during Stages V–VII; these divisions are not shown in this schematic model. For more details, see Malamy and Benfey (1997a). (b) Confocal images of LRP development using EGFP-LTI6b plasma membrane marker transgenic plants (Kurup et al., 2005). A primary root tip is also shown to highlight the anatomical similarities. Top row: Stage I, Stage II, Stage IV; bottom row: Stage VI/VII, Stage VII, primary root tip.

Figure 4.2 Comparison of lateral root primordium development and embryonic root development. (a) The scarecrow promoter was fused to GFP and stably transformed into Arabidopsis (Wysocka-Diller et al., 2000), and LRP was visualized at various stages. The left panels are DIC images, the middle panels are GFP in the same tissue and the right panels are tracings of the LRP with scarecrow expression indicated. (b) In situ analysis confirms the authenticity of the GFP patterns observed in A. (c) A schematic model of the expression of scarecrow in the developing embryo and LRP. The divisions involved in the formation of the endodermis and cortex layers appear to be quite similar in these two structures. This similarity is reinforced by the parallel expression patterns of the scarecrow gene. Embryonic stages of development are indicated at the left, and LRP developmental stages are indicated on the right. (Schematic model by Y. Helariutta (unpublished).) Glob, globular stage; triang, triangular stage. For confocal images of scarecrow expression in the embryo, see Wysocka-Diller et al. (2000).

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Figure 6.1 Gravity perception in roots mediated by statolith sedimentation in columella cells. (a) Schematic representation of an Arabidopsis root with grey-marked columella cells. (b) Arabidopsis root apex stained for starch. S1 and S2 layers are denoted by the red and blue arrows, respectively. (c) Upon change in gravity vector, sedimentation of statoliths in the columella cells results in repositioning of auxin efflux facilitator PIN3 (in red) to the new bottom of the cell by a yet unknown mechanism. (a)

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Figure 6.2 Transport of the lateral auxin gradient from the root apex to elongation zone tissues by the auxin influx and efflux carriers AUX1 and PIN2. (a) Schematic representation of an Arabidopsis root marking graviperception (magenta), gravity signal transmission (blue) and graviresponse (green) tissues. (b) Confocal image showing PIN2 (green) localization in the Arabidopsis root apex. (c) Close-up of PIN2 localization in epidermal cells. (d) Close-up of PIN2 localization in LRC cells (indicated by arrows). (e) Confocal image showing AUX1 (red) localization in the Arabidopsis root apex. (f) Close-up of AUX1 localization in epidermal cells. (g) Close-up of AUX1 localization in LRC cells.

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Figure 6.3 Formation of an auxin response gradient prior to a root gravitropic response. (a) Time lapse images of a root curvature following a 90◦ gravity stimulus (at time 0). (b) IAA2:GUS auxin-responsive expression exhibits asymmetric reporter staining following a 90◦ gravity stimulus (at time 0).

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Figure 7.1 Morphology of the cereal root system. (a) Root types of embryonic origin in 5-day-old maize and a rice seedlings. Maize forms a primary root (pr) and seminal roots (sr) during embryogenesis, while in rice only a primary root is formed. Brace roots (br) emerge from aboveground stem nodes late in development in maize (b) and rice (c) and constitute together with the belowground formed crown roots the shoot-borne root system. ZmSCR

ZmNAC5

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Figure 7.2 Comparison of the ZmSCR and ZmNAC5 gene expression patterns on adjacent median longitudinal sections through maize embryos at successive developmental stages. (a, b) Proembryo: ZmSCR is expressed in a central domain of the embryo proper while transcription for ZmNAC5 is not detectable. At the transition stage (c, d) and the coleoptilar stage (e, f), the shape and orientation of the ZmNAC5 domain in the center of the embryo mimics that of the overlying ZmSCR transcription pattern. At its basal end, the ZmNAC5 domain marks the transition toward the suspensor. col, coleoptile.

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QC LTR Columella Figure 12.1 Organization of the Arabidopsis root. The Arabidopsis root has apparent radial and longitudinal organization. In the center is a false-colored confocal image of the root tip. Cortex cells are labeled in green. Note that this cell lineage can be traced back to the root apex where the QC resides. Along the longitudinal axis, the root can be divided into three developmental zones. The meristematic zone includes cells that are actively engaged in the cell cycle. Cells in the elongation zone are rapidly expanding along the longitudinal axis. Cells in the maturation zone take on their final differentiated state. Not shown in the figure is root hair initiation as a key morphological marker for the beginning of the maturation zone. The zonation depicted to the right indicates the divisions used in Birnbaum et al. (2003) and Brady et al. (2007) for their transcriptional profiling experiments.

Figure 12.2 Using DNA microarrays to study global transcriptional events. Two flow charts depicting the overall steps involved in generating probes for transcriptional profiling with RNA as the starting material (left) and ChIP-chip using chromatin as the starting material (right), respectively.

Figure 12.3 GFP marker lines and FACS methodology. (a) Confocal images of three GFP reporter lines that mark distinct populations of cells in the root: the SCR::GFP, the QC and endodermis, S32, the phloem and PET111, the columella root cap cells. Roots are counter-stained with propidium iodide. (b) Cartoon depicting the cell-sorting methodology. Hundreds of roots from a particular GFP reporter line (SCR::GFP) are harvested and treated with an enzyme cocktail that digests the cell walls. Subsequently, cells dissociate from each other and are fed into an FACS device that collects the GFP-fluorescent cells into an extraction buffer. RNA is isolated and the probe is synthesized for hybridization to microarrays.

Figure 12.4 Environmentally regulated and stable biological functions in the root. Analysis of cell-type-specific gene expression profiles in the root after treatment with environmental stimuli allowed the identification of biological functions regulated in each cell layer. GO categories regulated by nitrogen provided by K. D. B. and M. L. G. (unpublished results). Comparison of the salt-stress, iron-deprivation and standard condition data sets led to the identification of environmentally stable biological functions that define each cell layer. Environmental regulation indicated by color: red, salt stress, green, iron deprivation, blue, nitrogen, purple, core biological function.

Annual Plant Reviews (2009) 37, 175–191 doi: 10.1002/9781444310023.ch7

www.interscience.wiley.com

Chapter 7

MOLECULAR AND GENETIC DISSECTION OF CEREAL ROOT SYSTEM DEVELOPMENT Frank Hochholdinger and Roman Zimmermann Center for Plant Molecular Biology (ZMBP), Department of General Genetics, University of T¨ubingen, 72076 T¨ubingen, Germany

Abstract: Monocotyledonous cereal root systems have a complex organization and are composed of primary and seminal roots that are preformed during embryogenesis and roots that emerge postembryonically either from other roots (lateral roots) or from nodal structures of the shoot (shoot-borne roots). In recent years, a number of specific mutants affected in cereal root system development, mainly in maize, rice and barley, have been identified. These mutants are impaired in root formation during embryogenesis, in primary root elongation, in root hair formation, in shoot-borne root initiation and in lateral root initiation and elongation. Recently, for some of these mutants the corresponding genes have been cloned and functionally characterized. In addition, these mutants have also been deployed for comparative proteome and transcriptome analyses to dissect the molecular networks involved in cereal root formation. Keywords: mutants; embryonic roots; postembryonic roots; maize; rice; barley

7.1 Introduction Cereal crops are of paramount importance for human nourishment and their starch provides approximately 70% of human food energy (Chandler and Brendel, 2002). In some developing countries, the diet of almost the entire population depends on cereals. The outstanding role of cereals for human nutrition is reflected in distinct morphological features that allow the optimal acquisition of natural resources and their conversion into valuable components of the human diet. The high productivity of cereals requires an elaborate root system architecture that secures efficient uptake of water and nutrients from the soil and provides anchorage (Lynch, 1995; Aiken and 175

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Root Development Smucker, 1996). The formation of the cereal root system is governed by an endogenous genetic program, but is also regulated by the plasticity of the root system to react to exogenous biotic and abiotic stimuli of the surrounding rhizosphere (McCully, 1999) and allows for the flexible adaptation to changing environmental cues. This chapter will focus on the current status of the genetic and molecular dissection of root development in cereals with special emphasis on maize (Zea mays L.) and rice (Oryza sativa L.) which are the premier model organisms in cereal genetics. The interaction of root systems with the rhizosphere has been recently reviewed elsewhere (Bais et al., 2006; Watt et al., 2006).

7.2

Morphology of cereal root systems

Maize and rice root development can be divided into an embryonic and a postembryonic phase (Feldman, 1994; Feix et al., 2002; Hochholdinger et al., 2004a, 2004b, 2005a). During the embryonic phase in maize, primary and seminal roots are formed that emerge in young seedlings a few days after germination (Fig. 7.1a). While the primary root is formed at the basal pole of the embryo, seminal roots develop in variable numbers at the scutellar node. In contrast to maize, rice does not form seminal roots at the scutellar node (Fig. 7.1a). In literature, there has been some confusion concerning the nomenclature of the different embryonic root types of maize and rice. In some publications, the primary root of rice is designated seminal root (for instance, Matsuo and Hoshikawa, 1993) which is unfortunate because this term is also used for the roots formed at the scutellar node of maize. Therefore, we suggest to use exclusively the term primary root for the root emerging from the basal pole of the embryo. The postembryonic root systems of maize and rice are determined by extensive shoot-borne roots that are formed at consecutive underground and aboveground nodes of the shoot (Feldman, 1994; Moriata and Nemoto, 1995; Hochholdinger et al., 2004a, 2004b). The nodal roots that are formed below the soil level are designated crown roots while the respective shoot-borne roots formed at aboveground nodes are called brace roots (Figs. 7.1b and 7.1c). A common feature of all embryonic and postembryonic root types is their ability to form lateral roots. Lateral roots are defined as roots that are initiated from pericycle cells of other roots. In addition to the roots that are determined by the normal developmental program, maize and rice can also form roots at unusual sites of the plant body, for instance at the mesocotyl or under exceptional circumstances, such as after wounding, hormone application or other exogenous stimuli. These roots are commonly referred to as adventitious roots. Unfortunately, also the term adventitious is not always consistently used in literature. In older references, all roots that are not derived from another root (except the primary root) are described as adventitious. Hence, in this old nomenclature seminal, crown and brace roots were classified as adventitious roots, which is not very

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helpful because molecular and genetic data suggest distinct functions for the different root types (Hochholdinger et al., 2004a, 2004b, 2005a).

7.3 Morphological and anatomical comparison of Arabidopsis and cereal root systems In recent years, Arabidopsis thaliana has become the prime model organism of plant molecular biology and genetics. Impressive progress has been made in the genetic analysis of various aspects of Arabidopsis root development, including pattern formation in the primary root (see Chapter 1), root hair formation (see Chapter 3) or lateral root formation (see Chapter 4). Nevertheless, there are a number of significant morphological and anatomical differences between Arabidopsis and cereal root systems that make it worthwhile to study the molecular and genetic basis of cereal root formation in more detail. To begin with, the architecture of cereal embryos is more complex compared to the simple body plan characteristic for the Arabidopsis embryo. For instance, the root apical meristem (RAM) is enclosed by a structure referred to as the coleorhiza, not present in Arabidopsis, that subtends the RAM to unsheathe and protect it during embryogenesis and upon germination (Tillich, 1977). Therefore, the primary root in cereals is established endogenously (Yamashita, 1991), whereas the primary root in dicots develops exogenously ¨ to constitute the basal pole of the embryo (Jurgens, 1992). Embryo development in maize gives rise to an additional root type, the seminal roots, which are also formed endogenously. Depending on the genetic background, a variable number of seminal roots is initiated between 22 and 40 days after pollination in the region of the scutellar node (Sass, 1977; Erdelska and Vidovencova, 1993; Feldman, 1994). Seminal roots are neither formed in rice (Fig. 7.1a) nor in Arabidopsis. On the cellular level, various differences between Arabidopsis and cereal roots can be observed after germination. The cortex of the cereal root is composed of 10–15 concentric layers of parenchyma cells and one endodermal cell layer (Feldman, 1994; Moriata and Nemoto, 1995), whereas during early development at least the Arabidopsis root is composed of one endodermal and only one cortical cell layer. Moreover, the radial number of cortical cells in maize and rice is variable, but the single Arabidopsis cortex cell layer always contains eight cells circumferentially. The quiescent center (QC) is a region of reduced meristematic activity in the center of the root tip. In cereals, the QC consists of 800–1200 cells (Jiang et al., 2003), whereas in Arabidopsis it is composed of exactly 4 cells (Schiefelbein et al., 1997). The QC is embedded in the proximal and distal meristems. While the distal meristem which is also known as calyptrogen gives rise to the root cap the arc shaped proximal meristem provides the different cell types of the emerging root. The RAMs consist of several hundreds of cells in cereals (Feldman, 1994) and only of a

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Root Development small number of cells surrounding the QC in Arabidopsis (Schiefelbein et al., 1997). In Arabidopsis, lateral roots are formed from pericycle cells (Dubrovsky et al., 2000; Beeckman et al., 2001) but in cereals, endodermal cells give rise to the epidermis and columella and the pericycle contributes to the remaining tissues of the newly formed lateral roots (Fahn, 1990). Detailed accounts of the morphological and histological features of maize and rice root systems are given in Hochholdinger et al. (2004b) and Matsuo and Hoshikawa (1993).

7.4

Molecular and genetic analysis of cereal root formation

Forward genetics to identify specific mutants is a valuable approach to identify genes involved in root development. However, the identification of such mutants in cereals is complicated by several factors. First, roots display an underground phenotype that is not directly accessible for phenotypic analysis. Second, the cereal root system is of considerable size and complexity (Kiesselbach, 1949). Third, root system development is very plastic and largely influenced by mutual interactions of roots and the biotic and abiotic soil environment (Drew and Saker, 1975, 1978; Hawes et al., 1998; McCully, 1999). These difficulties require a standardized procedure for the screening of novel mutant phenotypes, such as the germination of seeds in paper rolls (Wen and Schnable, 1994; Hetz et al., 1996). The fact that maize and rice seedlings can completely live on the nutrients stored in their endosperm in the first two weeks after germination allows mutagenized seeds to germinate in distilled water under very stringently controlled temperature, humidity and light conditions. Moreover, all root types, except the brace roots, can already be observed at this early developmental stage, enabling screening for a wide variety of phenotypic aberrations in the different root types.

7.4.1

Root development during embryogenesis

7.4.1.1 Radial pattern formation of the primary root Cell-type specification processes underlying embryonic root development have been well studied in Arabidopsis, but are still poorly understood in cereals. While in Arabidopsis individual structures of the mature embryo can be traced back to single cells or cell tiers during early developmental stages ¨ (Mansfield and Briarty, 1991; Jurgens and Mayer, 1994), a detailed analysis of radial patterning in cereals is complicated by the variable and unpredictable cell division pattern during embryogenesis (Randolph, 1936; Abbe and Stein, 1954; Poethig et al., 1986). An essential prerequisite for the concentric and regular organization of the different tissue layers in the root is the establishment of radial pattern information in the embryo. In Arabidopsis, the SCARECROW

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(AtSCR) gene that encodes a member of the GRAS gene family of putative transcription factors plays a pivotal role in this developmental process (Scheres et al., 1995; di Laurenzio et al., 1996) and is necessary for the asymmetric division of the descendants of the cortex/endodermis initials (WysockaDiller et al., 2000; Heidstra et al., 2004). Accordingly, Atscr mutants exhibit an incomplete radial organization with only one ground tissue layer sharing both cortex and endodermis characteristics. In longitudinal sections through the primary root, AtSCR gene expression marks a sickle-shaped domain connecting both endodermal cell files throughout the endodermis/cortex initials and the QC (di Laurenzio et al., 1996; Wysocka-Diller et al., 2000). In spite of the morphological differences of Arabidopsis and cereal roots, several indications imply that SCR functions as part of a conserved mechanism for the establishment of the root radial organization in monocots. Until now, genes closely related to AtSCR have been identified in maize and rice, ZmSCR and OsSCR, respectively, that have a similar exon–intron structure and are classified into the same clade as AtSCR in phylogenetic reconstructions (Lim et al., 2000; Kamiya et al., 2003). The ZmSCR and OsSCR gene expression patterns in primary roots are reminiscent of that of AtSCR (Lim et al., 2000; Kamiya et al., 2003). ZmSCR gene expression has been shown to be a primary determinant in the regeneration of radial pattern information in the maize root tip in surgical experiments (Lim et al., 2000) and, finally, ZmSCR expressed under the control of the native AtSCR promoter can rescue the patterning defects observed in Arabidopsis scr mutants (Lim et al., 2005). Despite some differences, both AtSCR and ZmSCR are expressed from the early proembryo stage onwards (Wysocka-Diller et al., 2000; Lim et al., 2005). This indicates that even in evolutionary distant species the establishment of radial pattern information during embryogenesis is a basic requirement for subsequent patterning processes. This notion is supported by the globular embryo4 (gle4) mutant of rice (see Section 7.4.1.3) that is defective in embryonic radial patterning and, as a consequence, does not develop any embryonic organs. 7.4.1.2 Specification of the QC and root cap during embryogenesis Besides ZmSCR and OsSCR, several other informative root-specific molecular markers have been identified in maize and rice in recent years. The QUIESCENT-CENTER-SPECIFIC HOMEOBOX gene (QHB; Kamiya et al., 2003), a member of the WUSCHEL-type homeobox gene family in rice, is most closely related to the WUSCHEL-RELATED HOMEOBOX 5 (AtWOX5) gene from Arabidopsis (Haecker et al., 2004). Both genes are early and selective markers for the acquisition of QC identity in the root. QHB expression in the embryo precedes histological differentiation of the radicle. Comparison to the transcription pattern during crown root formation suggests that cell-type specification of the QC might be controlled by different mechanisms in these root types (Kamiya et al., 2003). In maize, a member of the NAC gene family (named according to its founding members NAM, ATAF1/2 and CUC2), ZmNAC5, has been shown to be specifically associated with the establishment

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Root Development of the coleorhiza during embryogenesis and its expression persists postembryonically in the primary root where it is detected in the 4–5 uppermost layers of the root cap subtending the RAM. Comparative expression studies on adjacent longitudinal sections through the transition stage or coleoptilar embryo reveal that the ZmNAC5 domain in its shape and orientation mimics that of the overlying ZmSCR domain (Fig. 7.2; Zimmermann and Werr, 2005). The close spatiotemporal relation of the ZmNAC5 and ZmSCR transcription patterns suggests that the root cap in maize is established inside the embryo as part of the coleorhiza, dependent on ZmSCR-mediated radial positional information. Gene expression of the ZmNAC5 ortholog from rice (OsNAC7) in embryos (Kikuchi et al., 2000) might be indicative for a similar mechanism in rice. Tissue-specific molecular markers can serve as valuable tools to classify embryo-specific mutants along the course of patterning processes that occur during cereal root formation (Elster et al., 2000; Kamiya et al., 2003). Recently, a number of GFP marker lines associated with individual cell types in the primary root have been established in rice (Johnson et al., 2005). Future expression studies will reveal whether some of these lines might be applicable for the analysis of embryonic root development. Such lines might also be helpful to identify other genes expressed in a cell-type-specific manner in the embryonic root based on fluorescence-activated cell sorting (FACS) and microarray analyses, as already demonstrated for the Arabidopsis primary root (Birnbaum et al., 2003; Nawy et al., 2005, Brady et al., 2007). 7.4.1.3 Mutants of embryonic root formation Only a few mutants impaired in root development during embryogenesis in cereals have been identified so far (Table 7.1). One explanation might be that, in contrast to the diploid rice genome (Goff et al., 2002), many other cereal species, such as maize, are characterized by a high degree of ploidy reflected by an increased genetic redundancy. As a consequence, mutations in individual genes often do not become obvious phenotypically because their loss of function is compensated by other genes acting in the same or a similar developmental process. In the following paragraph, mutants impaired in embryo root development in cereal crops will be described briefly. The globular embryo4 (gle4) mutant of rice does not develop any embryonic organs. Results from marker-assisted studies of the gle4 mutant phenotype demonstrate that the GLE4 gene is involved in radial pattern formation of the L2 and L3 layer during embryogenesis, but does not contribute to the establishment of the L1 layer (Kamiya et al., 2003). Embryos of the rice radicleless1 (ral1) mutant develop normal apical structures, but are impaired in radicle differentiation. The observed phenotype is due to vascular patterning defects in the procambium and point at a role of the RAL1 gene in cell axialization in procambial cells (Scarpella et al., 2003). The maize mutants rootless with undetectable meristems1 (rum1; Woll et al., 2005) and rootless concerning crown and seminal roots (rtcs; Hetz et al., 1996) are both affected in seminal root initiation. Since these mutants are also affected either in shoot-borne (rtcs)

Species

Mutant

Lateral root initiation Lateral root and seminal root initiation Lateral root elongation Lateral root elongation Lateral root initiation

Z. mays Z. mays Z. mays Z. mays O. sativa

O. sativa O. sativa O. sativa O. sativa O. sativa O. sativa

Z. mays Z. mays Z. mays O. sativa H. vulgare H. vulgare

Lateral roots lrt1 rum1 slr1 slr2 rm109

Primary root rm1 rm2 rrl1 rrl2 srt5 srt6

Root hairs rth1 rth2 rth3 rh2 brb rhl1.a

n.d., not determined.

Shoot-borne- and seminal root initiation Shoot-borne root formation Shoot-borne root initiation Shoot-borne root initiation

Shoot-borne roots rtcs Z. mays rt1 Z. mays crl1/arl1 O. sativa crl2 O. sativa

Root hair elongation Root hair elongation Root hair elongation Root hair initiation Root hair initiation Root hair initiation

Primary root elongation Primary root elongation Primary root elongation Primary root elongation Elongation of all root types Primary root elongation

Radial patterning of radicle Radicle differentiation

Embryogenesis gle4 O. sativa ral1 O. sativa

Defect

Cereal mutants affected in root system development

Table 7.1

sec3 homolog n.d. n.d. n.d. n.d. β-expansin

n.d. n.d. n.d. n.d. n.d. n.d.

n.d. n.d. n.d. n.d. n.d.

LOB domain gene n.d. LOB domain gene n.d.

n.d. n.d.

Gene Function

Wen and Schnable (1994), Wen et al. (2005) Wen and Schnable (1994) Wen and Schnable (1994) Suzuki et al. (2003) Gahoonia et al. (2001) Kwasniewski and Szarejko (2006)

Ichii and Ishikawa (1997) Ichii and Ishikawa (1997) Inukai et al. (2001b) Inukai et al. (2001b) Yao et al. (2002) Yao et al. (2003)

Hochholdinger and Feix (1998) Woll et al. (2005) Hochholdinger et al. (2001) Hochholdinger et al. (2001) Hao and Ichii (1999)

Hetz et al. (1996); Taramino et al. (2007) Jenkins (1930) Inukai et al. (2005); Liu et al. (2005) Inukai et al. (2001a, 2001b)

Kamiya et al. (2003) Scarpella et al. (2003)

References

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Root Development or lateral root initiation (rum1), they will be discussed in more detail in the corresponding sections. 7.4.2

Initiation of shoot-borne root formation

As the major portion of the adult root system, shoot-borne roots are the basis for lodging resistance of cereals and are responsible for the major part of water and nutrient uptake via their lateral roots (McCully and Canny, 1988). Crown and brace roots are formed endogenously, and their primordia are initiated in the outermost cell layers of the cortex opposite to collateral vascular bundles in the nodal structures of the shoot (Martin and Harris, 1976). A maize plant develops approximately 70 shoot-borne roots during its life cycle in approximately six whorls of underground crown roots and two to three whorls of aboveground brace roots (Fig. 7.1b; Hoppe et al., 1986). The first four underground internodes are very short (Hoppe et al., 1986), and their crown roots form a dense root system. Not all aboveground brace roots grow into the soil, and only those brace roots that penetrate the soil level form lateral roots. 7.4.2.1 Mutants of shoot-borne root formation In maize, two mutants affected in shoot-borne root formation have been isolated. The mutant rootless1 (rt1) does not form shoot-borne roots at the higher nodes (nodes 7 and 8) while the number of crown roots at the first two nodes is only slightly reduced (Jenkins, 1930), indicating that rt1 is primarily affected in the formation of the aboveground brace roots. The rt1 mutation is inherited as a monogenic recessive trait and maps on chromosome 3 (Emerson et al., 1935). Whereas rt1 is exclusively affected in postembryonic shoot-borne root formation, the rtcs mutant (Hetz et al., 1996) is affected in shoot-borne and seminal root formation and lacks all seminal, crown and brace roots. The only root that remains in rtcs is the primary root. However, the primary root and its lateral roots are sufficient to generate a mature plant in the greenhouse when properly supported. Histological studies revealed that the mutation inhibits the initiation of the affected root types (Hetz et al., 1996). The distinct map positions of the rt1 and rtcs genes of maize indicate that these genes are not allelic. In rice, two mutants crownrootless1 (crl1) (Inukai et al., 2005) and adventitiousrootless1 (arl1) (Liu et al., 2005) have been described that later turned out to be different alleles of the same gene. The mutants crl1 and arl1 are both defective in shoot-borne root initiation (Inukai et al., 2005; Liu et al., 2005). While arl1 does not form any shoot-borne roots at higher shoot nodes, the mutant crl1 occasionally does. Moreover, arl1 is not affected in lateral root formation (Liu et al., 2005) whereas the number of lateral roots of the primary root of the mutant crl1 is reduced to 70% of that of the wild type (Inukai et al., 2005). In addition, gravitropic response of the primary root in crl1 is reduced when compared to wild type, an aspect that was not analyzed in arl1. Both mutants do not display any abnormalities in the aboveground tissues of the

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seedlings. Still, the arl1 mutant dies at the seedling stage, indicating that it might represent a stronger allele than crl1. Another rice mutant, crownrootless2 (crl2; Inukai et al., 2001a, 2001b), forms a reduced number of shoot-borne roots whose growth ceases shortly after germination. The remaining primary root displays an increased thickness and length as well as a reduced number of lateral roots. 7.4.2.2 LOB domain proteins are involved in shoot-borne root formation Recently, the rice gene affected in the allelic mutants crl1 and arl1 was cloned (Inukai et al., 2005; Liu et al., 2005) and shown to map on chromosome 3 and to encode a 259-amino-acid protein with a LATERAL ORGAN BOUNDARIES (LOB) domain (Inukai et al., 2005; Liu et al., 2005). Similarly, the rtcs gene of maize was cloned via a map-based approach and shown to encode a 244amino-acid LOB-domain protein located on chromosome 1S (Taramino et al., 2007). Since RTCS along with 14 other genes in the RTCS region maps in the corresponding synthenic region of CRL1/ARL1, we concluded that RTCS is the ortholog of rice CRL1/ARL1. In contrast to CRL1/ARL1, the RTCS gene has been duplicated during evolution. The paralogous RTCL (RTCSLIKE) gene maps on chromosome 9 and displays 72% sequence identity with the RTCS gene on the protein level (Taramino et al., 2007). Both genes are preferentially expressed in roots. In the coleoptilar node, expression of RTCS is confined to emerging shoot-borne root primordia. The CRL1/ARL1 and RTCS genes share auxin response elements (AuxRE) in their promotors and have indeed been shown to be auxin inducible (Inukai et al., 2005). The observation that a recombinant auxin response factor (ARF) can bind to the promoter of CRL1 (Inukai et al., 2005) also supports the idea that the CRL1/ARL1 and RTCS genes might play a pivotal role in auxin-mediated initiation of shootborne root development. While CRL1/ARL1 is exclusively involved in the initiation of postembryonically formed shoot-borne roots, the RTCS gene also contributes to the establishment of embryonic seminal roots. 7.4.2.3 Proteomic dissection of shoot-borne root formation The most abundant soluble proteins of the first node (coleoptilar node) in the wild type and the rtcs mutant of maize were compared at two early stages of crown root formation via Coomassie blue-stained two-dimensional gels. Five and 14 differentially accumulated proteins were identified in 5-day-old and in 10-day-old coleoptilar nodes, respectively (Sauer et al., 2006). The sequence of all differentially expressed proteins was identified via liquid chromatography electrospray ionization tandem mass spectrometry (LC-ESI MS/MS). RNA gel blot experiments performed for genes encoding five of the identified proteins confirmed differential gene expression and revealed subtle changes in regulation during early coleoptilar node development. Remarkably, several of the differentially expressed proteins represented components involved in auxin signal transduction (Sauer et al., 2006) and might thus give cues on the molecular context of RTCS during shoot-borne root initiation. An

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Root Development auxin-binding protein 1 (ABP1), which is believed to act as a high-affinity auxin receptor (Chen, 2001), was highly expressed in wild-type coleoptilar nodes five days after germination. Expression was downregulated to low levels between five and ten days after germination, suggesting a feedback inhibition of this gene during early coleoptilar node development. In rtcs, the putative feedback inhibition of ABP1, which might act in the earliest stages of auxin signal transduction, was not detected but instead the gene and protein maintained its high expression level (Sauer et al., 2006). Furthermore, calmodulin, that binds to the SAUR class of early auxin-responsive genes (Yang and Poovaiah, 2000), is downregulated in the rtcs mutant (Sauer et al., 2006). Finally, a subunit of a heterotrimeric G-protein was downregulated in 10-day-old wild-type coleoptilar nodes (Sauer et al., 2006). Heterotrimeric G-proteins coupled to a low-affinity auxin receptor are assumed to stimulate cell division in response to high auxin concentrations via a second, yet unidentified, auxin pathway (Chen, 2001). 7.4.3

Formation of lateral roots

Lateral roots are defined as roots that emerge from pericycle cells of other roots and are initiated via dedifferentiation of pericycle cells in the differentiation zone some distance from the root apex (Esau, 1965). The first division of lateral root founder cells is induced by endogenous signals of the maize plant, such as auxin (Dubrovsky et al., 2000). In contrast to Arabidopsis, it is less evident to predict the sites of lateral root initiation in cereal roots (Bell and McCully, 1970). 7.4.3.1 Mutants of lateral root formation In maize, the identification of root type-specific lateral root mutants suggest that there are at least two pathways of lateral root formation or at least two different sensitivities to signals involved in lateral root formation (Hochholdinger and Feix, 1998; Hochholdinger et al., 2001; Woll et al., 2005). This notion is supported by mutants specifically affecting lateral root initiation and elongation on the embryonic, but not the postembryonic, roots. The lateralrootless1 (lrt1) mutant is affected in early postembryonic root development (Hochholdinger and Feix, 1998). This developmental phase is confined to lateral roots initiated in embryonic roots and the formation of crown roots at the first node, i.e. the coleoptilar node. Crown root formation from the second node onwards and lateral root formation in crown roots emerging from these higher nodes are normal in lrt1. Similarly, the mutant rootless with undetectable meristems (rum1) is affected in lateral root initiation in the primary root while lateral root formation is normal in the shoot-borne root system (Woll et al., 2005). The rum1 mutant is also affected in the initiation of the embryonic seminal roots. The mutants short lateral roots1 (slr1) (Hochholdinger et al., 2001) and short lateral roots2 (slr2) (Hochholdinger et al., 2001) also show reduced lateral root elongation of the embryonic roots but form normal

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lateral roots in the postembryonic shoot-borne root system. In rice, the mutant rm109 (2,4-Dichlorophenoxyacetic acid resistant mutant 109) is thus far the only mutant that is defective in lateral root initiation. The pericycle cell number in the primary root of the rm109 mutant was 70–80% of that of the wild type, indicating that the deficient lateral root production might mainly be based on irregular cell division (Hao and Ichii, 1999). 7.4.3.2 Proteome analyses of lateral root formation Three proteome analyses highlighting different aspects of lateral root formation in maize have been published recently (Hochholdinger et al., 2006). First, in a comparative study of 9-day-old wild-type versus lrt1 mutant primary roots that do not initiate lateral roots, 67 proteins were identified via MALDI-ToF (matrix-assisted laser desorption ionization time-of-flight mass spectrometry) mass spectrometry. Interestingly, 10% of the detected proteins were preferentially expressed in the mutant (Hochholdinger et al., 2004c). For the first time, the influence of lateral roots on the proteome composition of the primary root was demonstrated. In a second analysis, the abundant soluble proteins of 2.5-day-old primary roots of wild-type seedlings and the lateral root initiation mutant rum1 were compared before the initiation of lateral roots (Liu et al., 2006). Among 350 proteins detected in Coomassie blue-stained twodimensional gels, 14 proteins encoded by 12 different genes were identified via LC-ESI MS/MS as differentially accumulated. Functionally, these proteins are involved in lignin biosynthesis, defense, and the citrate cycle. This study represented the first comparative proteomic analysis of maize primary roots prior to lateral root initiation. Third, in a reference data set of early root formation, the 81 most abundant soluble proteins were identified via MALDI-ToF from 5-day-old primary roots of maize (Hochholdinger et al., 2005b). This study demonstrated the remarkable changes in the accumulation of abundant soluble proteins between 5 and 9 days after germination, hence, during the early stages of lateral root formation on the primary root of maize. 7.4.3.3

Cell-type-specific transcriptome analyses of lateral root formation Plant organs are characterized by their composite structure. Roots, for example, are made up of epidermal, cortical, endodermal, and various cell types in the central cylinder and RAM. Each of these cell types expresses a unique transcriptome. Therefore, microarray analyses of a complete root only provide average gene expression levels integrated over all cell types bearing the potential to mask genes of interest that are expressed in a particular cell type. Laser capture microdissection (LCM), that allows for the isolation of a particular cell type that can be unambiguously distinguished at the histological level, can overcome this problem (Schnable et al., 2004). This technique is of particular value for the study of processes leading to lateral root initiation in the single cell layer of the pericycle. The lateral root initiation defective mutant of maize, rum1, was used to compare the cell-type-specific transcriptome

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Root Development of the pericycle via the combination of LCM and microarray experiments 64 hours after germination, thus before lateral root initiation. This study revealed 90 and 73 genes preferentially expressed in the wild-type and in the rum1 pericycle, respectively. Among the 51 annotated genes predominately expressed in the wild-type pericycle, 19 genes are involved in signal transduction, transcription, and the cell cycle, thus defining an array of genes that is active in the pericycle before lateral root initiation (Woll et al., 2005). 7.4.4

Mutants of primary root formation

In cereals, the primary root is the first root type that emerges 2–3 days after germination. Several rice mutants are specifically impaired in primary root elongation [root mutant 1 (rm1; Ichii and Ishikawa, 1997), root mutant 2 (rm2; Ichii and Ishikawa, 1997), reduced root length 1 (rrl1; Inukai et al., 2001b), reduced root length 2 (rrl2; Inukai et al., 2001b), short root 6 (srt6; Yao et al., 2003)]. In contrast to these mutants, the mutant short root 5 (srt5; Yao et al., 2002) has reduced root elongation also during crown and lateral root development. As a common feature, all these mutants display reduced cortical cell elongation in the affected roots. Until now, none of the corresponding genes has been cloned. 7.4.5

Mutants of root hair formation

In maize, the mutants roothairless1 to roothairless3 (rth1, rth2 and rth3; Wen and Schnable, 1994) have been identified to be defective in root hair elongation. Interestingly, rth2 and rth3 grow normally, implicating that root hairs might be dispensable under specific environmental conditions. Furthermore, the rice root hair mutant roothairless2 (rh2) is able to develop very short root hairs after auxin (NAA) treatment (Suzuki et al., 2003). Finally, in barley (Hordeum vulgare), a spontaneously induced monogenic recessive roothairless mutant, bald root barley (brb; Gahoonia et al., 2001) and the roothairless1.a (rhl1.a) mutant (Kwasniewski and Szarejko, 2006), have been isolated. 7.4.5.1 A SEC3 homolog is involved in root hair elongation in maize Unicellular root hairs elongate via localized tip growth, a process that is mediated by polar exocytosis of secretory vesicles. The RTH1 gene encodes a SEC3 homolog (Wen et al., 2005). In yeast and mammals, SEC3 is a subunit of the exocyst complex that tethers exocytotic vesicles prior to their fusion. For the first time, the cloning of the RTH1 gene links a component of the postulated exocyst complex to root hair elongation and supports the notion that the exocyst complex might be involved in plant exocytosis. Comparative proteomic analyses identified four proteins that accumulate to different levels in wild type and rth1 primary roots (Wen et al., 2005). The preferential accumulation of a prohibitin, that is a negative regulator of the cell cycle in the rth1 mutant proteome, might at least partially explain the delayed development and flowering of the rth1 mutant.

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7.4.5.2 A ␤-expansin is tightly linked to root hair initiation in barley By means of subtractive hybridization with RNA from wild type and the rhl1.a mutant, a member of the EXPB family of ␤-expansins (HvEXPB1) was demonstrated to be exclusively expressed in roots of wild-type plants (Kwasniewski and Szarejko, 2006). HvEXPB1 expression strictly cosegregated with the root hair phenotype in the F2 generation of crosses between rhl1.a and wild type. Expression of the gene in the root hair mutants rhp1.a, which only develops root hair primordia, and in rhs1.a, which forms very short root hairs, and a complete lack of HvEXPB1 expression in the roothairless mutant brb suggest a role of this gene in root hair formation in barley.

7.5 Prospects Currently, a growing number of cereal mutants specifically affected in different aspects of root development paves the way for a better understanding of the molecular basis of cereal root system architecture. Isolation of genes has already and will further help to determine root development-specific checkpoints unique to monocot plants. Moreover, identification of up- and downstream interaction partners will reveal the molecular context of the cloned genes. Furthermore, comparative transcriptome- and proteome-wide analyses of specific mutants, in particular on the cell-type-specific level, will reveal components involved in the complex network of cereal root formation. Finally, reverse genetic dissection of root-specific genes and the analysis of root traits that are inherited in a quantitative way will increase our understanding of the functional networks involved in root formation in monocotyledonous plants.

Acknowledgments We thank Caroline Gutjahr (University of Lausanne, Switzerland) for a photo of the aboveground rice root system. This research is supported by the German Research Council (DFG) grants HO2249/4, HO2249/6, and SFB446 project B16. R.Z. is supported by a DuPont research grant.

References Abbe, E.C. and Stein, O.L. (1954). The growth of the shoot apex in maize: embryogeny. Am. J. Bot. 41, 285–293. Aiken, R.M. and Smucker, A.J.M. (1996). Root system regulation of whole plant growth. Annu. Rev. Phytopathol. 34, 325–346. Bais, H.P., Weir, T.L., Perry, L.G., Gilroy, S. and Vivanco, J.M. (2006). The role of root exudates in rhizosphere interactions with plants and other organisms. Annu. Rev. Plant Biol. 57, 233–266.

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Hao, Z. and Ichii, M. (1999). A mutant RM109 of rice (Oryza sativa L.) exhibiting altered lateral root initiation and gravitropism. Jpn. J. Crop. Sci. 68, 245–252. Hawes, M.C., Brigham, L.A., Wen, F., Woo, H.H. and Zhu, Y. (1998). Function of root border cells in plant health: pioneers in the rhizosphere. Annu. Rev. Phytopathol. 36, 311–327. Heidstra, R., Welch, D. and Scheres, B. (2004). Mosaic analyses using marked activation and deletion clones dissect Arabidopsis SCARECROW action in asymmetric cell division. Genes Dev. 18, 1964–1969. Hetz, W., Hochholdinger, F., Schwall, M. and Feix, G. (1996). Isolation and characterization of rtcs, a maize mutant deficient in the formation of nodal roots. Plant J. 10, 845–857. Hochholdinger, F. and Feix, G. (1998). Early post-embryonic root formation is specifically affected in the maize mutant lrt1. Plant J. 16, 247–255. Hochholdinger, F., Guo, L. and Schnable, P.S. (2004c). Lateral roots affect the proteome of the primary root of maize (Zea mays L.). Plant Mol. Biol. 56, 397–412. Hochholdinger, F., Park, W.J. and Feix, G.H. (2001). Cooperative action of SLR1 and SLR2 is required for lateral root-specific cell elongation in maize. Plant Physiol. 125, 1529–1539. Hochholdinger, F., Park, W.J., Sauer, M. and Woll, K. (2004a). From weeds to crops: genetic analysis of root development in cereals. Trends Plant Sci. 9, 42–48. Hochholdinger, F., Sauer, M., Dembinsky, D., Hoecker, N., Muthreich, N., Saleem, M. et al. (2006). Proteomic dissection of plant development. Proteomics 6, 4076–4083. Hochholdinger, F., Woll, K., Guo, L. and Schnable, P.S. (2005b). The accumulation of abundant soluble proteins changes early in the development of the primary roots of maize (Zea mays L.). Proteomics 5, 4885–4893. Hochholdinger, F., Woll, K., Sauer, M. and Dembinsky, D. (2004b). Genetic dissection of root formation in maize (Zea mays) reveals root-type specific developmental programmes. Ann. Bot. 93, 359–368. Hochholdinger, F., Woll, K., Sauer, M. and Feix, G. (2005a). Functional genomic tools in support of the genetic analysis of root development in maize (Zea mays L.). Maydica 50, 437–442. Hoppe, D.C., McCully, M.E. and Wenzel, C.L. (1986). The nodal roots of Zea: their development in relation to structural features of the stem. Can. J. Bot. 64, 2524– 2537. Ichii, M. and Ishikawa, M. (1997). Genetic analysis of newly induced short-root mutants in rice (Oryza sativa L.). Breed. Sci. 47, 121–125. Inukai, Y., Miwa, M., Nagato, Y., Kitano, H. and Yamauchi, A. (2001a). Characterization of rice mutants deficient in the formation of crown roots. Breed. Sci. 51, 123–129. Inukai, Y., Miwa, M., Nagato, Y., Kitano, H. and Yamauchi, A. (2001b). RRL1, RRL2 and CRL2 loci regulating root elongation in rice. Breed. Sci. 51, 231–239. Inukai, Y., Sakamoto, T., Ueguchi-Tanaka, M., Shibata, Y., Gomi, K., Umemura, I. et al. (2005). Crown rootless1, which is essential for crown root formation in rice, is a target of an AUXIN RESPONSE FACTOR in auxin signaling. Plant Cell 17, 1387–1396. Jenkins, M.T. (1930). Heritable characters of maize XXXIV-rootless. J. Hered. 21, 79–80. Jiang, K., Meng, Y.L. and Feldman, L.J. (2003). Quiescent center formation in maize roots is associated with an auxin-regulated oxidizing environment. Development 130, 1429–1438. Johnson, A.A.T., Hibberd, J.M., Gay, C., Essah, P.A., Haseloff, J., Tester, M. et al. (2005). Spatial control of transgene expression in rice (Oryza sativa L.) using the GAL4 enhancer trapping system. Plant J. 41, 779–789.

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Sauer, M., Jakob, A., Nordheim, A. and Hochholdinger, F. (2006). Proteomic analysis of shoot-borne root initiation in maize (Zea mays L.). Proteomics 6, 2530–2541. Scarpella, E., Rueb, S. and Meijer, A.H. (2003). The RADICLELESS1 gene is required for vascular pattern formation in rice. Development 130, 645–658. Scheres, B., Di Laurenzio, L., Willemsen, V., Hauser, M.-T., Janmaat, K., Weisbeek, P. et al. (1995). Mutations affecting the radial organisation of the Arabidopsis root display specific defects throughout the embryonic axis. Development 121, 53–62. Schiefelbein, J.W., Masucci, J.D. and Wang, H. (1997). Building a root: the control of patterning and morphogenesis during root development. Plant Cell 9, 1089–1098. Schnable, P.S., Hochholdinger, F. and Nakazono, M. (2004). Global expression profiling applied to plant development. Curr. Opin. Plant Biol. 7, 50–56. Suzuki, N., Taketa, S. and Ichii, M. (2003). Morphological and physiological characteristics of a root-hairless mutant in rice (Oryza sativa L.). Plant Soil 255, 9–17. Taramino, G., Sauer, M., Stauffer, J.L., Jr., Multani, D., Niu, X., Sakai, H. et al. (2007). The maize (Zea mays L.) RTCS gene encodes a LOB domain protein that is a key regulator of embryonic seminal and post-embryonic shoot-borne root initiation. Plant J. 50, 649–659. Tillich, H.J. (1977). Vergleichend morphologische Untersuchungen zur Identit¨at der Gramineen-Prim¨arwurzel. Flora 166, 415–421. Watt, M., Silk, W.K. and Passioura, J.B. (2006). Rates of root and organism growth, soil conditions, and temporal and spatial development of the rhizosphere. Ann. Bot. 97, 839–855. Wen, T.-J., Hochholdinger, F., Sauer, M., Bruce, W. and Schnable, P.S. (2005). The roothairless1 gene of maize (Zea mays) encodes a homolog of sec3, which is involved in polar exocytosis. Plant Physiol. 138, 1637–1643. Wen, T.-J. and Schnable, P.S. (1994). Analyses of mutants of three genes that influence root hair development in Zea mays (Gramineae) suggest that root hairs are dispensable. Am. J. Bot. 81, 833–843. Woll, K., Borsuk, L.A., Stransky, H., Nettleton, D., Schnable, P.S. and Hochholdinger, F. (2005). Isolation, characterization and pericycle-specific transcriptome analyses of the novel maize lateral and seminal root initiation mutant rum1. Plant Physiol. 139, 1255–1267. Wysocka-Diller, J.W., Helariutta, Y., Fukaki, H., Malamy, J.E. and Benfey, P.N. (2000). Molecular analysis of SCARECROW function reveals a radial patterning mechanism common to root and shoot. Development 127, 595–603. Yamashita, T. (1991). Ist die Prim¨arwurzel bei Samenpflanzen exogen oder endogen? Beitr. Biol. Pflanzen 66, 371–391. Yang, T. and Poovaiah, B.W. (2000). Molecular and biochemical evidence for the involvement of calcium/calmodulin in auxin action. J. Biol. Chem. 275, 3137–3143. Yao, S.-G., Taketa, S. and Ichii, M. (2002). A novel short-root gene that affects specifically early root development in rice (Oryza sativa L.). Plant Sci. 163, 207–215. Yao, S.-G., Taketa, S. and Ichii, M. (2003). Isolation and characterization of an abscisic acid-insensitive mutation that affects specifically primary root elongation in rice (Oryza sativa L.). Plant Sci. 164, 971–978. Zimmermann, R. and Werr, W. (2005). Pattern formation in the monocot embryo as revealed by NAM and CUC3 orthologues from Zea mays L. Plant Mol. Biol. 58, 669–685.

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Chapter 8

FERN ROOT DEVELOPMENT Guichuan Hou1 and Elison B. Blancaflor2 1 Dewel Microscopy Facility and Department of Biology, Appalachian State University, 112 Rankin Science South, Boone, NC 28608-2027, USA 2 Plant Biology Division, The Samuel Roberts Noble Foundation, 2510 Sam Noble Parkway, Ardmore, OK 73401, USA

Abstract: Ferns have a special niche among land vascular plants since they have a different mode of root system construction in comparison with that of seed plants. Ferns have nonbipolar embryos and their ‘primary roots’ are lateral in embryonic origin. There are no distinct embryonic and postembryonic phases of root development in ferns because nonseed plant embryos do not undergo dormancy and germination. A recent study in the model fern Ceratopteris richardii has identified two shoot-borne root populations: stem-born and leaf-born, which exhibit different temporal and spatial developmental patterns. In contrast to seed plants, roots of most ferns possess a single apical cell that is the ultimate source of all cells in the root. The single root apical cell divides in a predictable manner producing daughter merophytes and each merophyte in turn divides and differentiates in a predictable pattern. Lateral root development in ferns also involves initiation of a single lateral root apical cell from the endodermal layer of the parent root. Previous studies show that exogenous auxins could not induce additional lateral root formation in ferns but inhibited both parent and lateral root growth. Although this indicates that the physiology of fern roots is similar in some respects to that of flowering plant roots, the developmental program for lateral root initiation in ferns might be different compared to flowering plants. Molecular and genetic studies of fern roots have lagged behind that of flowering plant roots. By drawing attention to the unique anatomical features of fern roots, we hope to generate more interest in ferns as a model for root developmental studies. Keywords: fern root development; Ceratopteris richardii; root apical cell; merophyte; endodermis

8.1

Introduction

As we have already learned from several chapters in this volume, studies on root development have positively impacted our understanding of basic plant 192

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developmental processes. The available genomic resources for model plants, such as Arabidopsis, have led to an enhanced appreciation of the molecular mechanisms underlying root biology, from transcriptional networks that specify epidermal cell fate (Bernhardt et al., 2005; Kwak and Schiefelbein, 2007; see Chapters 3 and 12, this volume) to signaling and hormonal regulation of root growth responses (Kramer and Bennet, 2006; see Chapters 5 and 6, this volume). Although a considerable amount of new information has been uncovered from work using flowering plant models, a renewed awareness of root anatomical and developmental processes in other plant systems is needed, especially if we are to achieve a better grasp on understanding how plant roots evolved. This chapter diverts from the common theme of angiosperm root development in that it focuses on ferns, a group of land plants that, although a major component of modern land flora, has lagged behind flowering plants in terms of understanding their biology. Ferns are classically organized within the Division Filicophyta (or Pteridophyta) sensu (Gifford and Foster, 1989). Filicophyta and its (fern) allies, including Psilophyta (such as Psilotum), Lycophyta (such as Lycopodium and Selaginella) and Sphenophyta (such as Equisetum), are seedless sporeproducing vascular plants that are collectively called pteridophytes sensu lato. Several anatomical features set pteridophyte roots apart from flowering plant roots. For instance, Selaginella roots are capless early in development whereas some roots of lycophytes exhibit apical meristem bifurcation. Furthermore, meristems in roots of other species within the Lycophyta develop either as a shoot or as a root depending on growth conditions (Webster and Steeves, 1967). Within the pteridophytes, there are cases where root structure and development differ between species. For example, although each root of Equisetum possesses a prominent pyramidal apical cell (see below), early cell division patterns in daughter merophytes and lateral root (LR) initiation are distinct from those of most ferns (Gifford, 1993). Ferns, as a systematic group, share many important features that would make them excellent models to study novel aspects of root development. The first (embryonic) root in ferns is initiated in a lateral position on the young shoot, i.e. ferns have unipolar (nonbipolar) embryos (Gifford and Foster, 1989; Barlow and Palma, 1997) with subsequent root formation closely associated with the shoot in contrast to flowering plants. The latter have bipolar embryos where in many cases the root system shows minimal structural association with the shoot system, even though shoot-borne roots do occur in some monocotyledonous plants (Hou and Hill, 2002; see Chapter 7, this volume). The formation of shoot-borne roots in ferns is called primary homorhizy and the postembryonic development of shoot-borne roots in flowering plants secondary homorhizy (see Groff and Kaplan, 1988). Furthermore, variations in root development during early sporophyte ontogeny are correlated with shoot aging (Hou and Hill, 2002), a process comparable to heteroblastic development in seed plants where leaf shape

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Root Development assumes different morphologies between juvenile and adult phases (Jones, 1999). Perhaps the most common anatomical feature of fern roots is the occurrence of a single root apical cell (RAC) that is the ultimate source of all cells for that particular root and divides in a regular sequence, resulting in predictable cellular patterns (Gunning et al., 1978; Gifford, 1991; Hou and Hill, 2004). LR development in ferns also involves the initiation of a single lateral RAC. However, unlike roots of flowering plants in which LR initials emerge from the pericycle (De Smet et al., 2006; see Chapter 4, this volume), the lateral RAC and as a result the emerging fern LR, originates from the endodermal layer of parent roots (Lin and Raghavan, 1991; Hou et al., 2004). The aforementioned examples simply highlight some unique aspects of fern roots that will allow researchers to ask fundamental questions about root development that cannot be addressed in flowering plant models. At the same time, questions as to whether ferns utilize mechanisms similar to those of flowering plants in specifying root developmental programs should lead to a better understanding of plant root evolutionary processes. Most of our current botanical textbooks treat plant roots as a single entity; however, it is not known if all land plant roots (as an organ per se) share a common ancestor. Paleobotanical evidence indicates that Sphenophytes, Lycopodophytes and Pteridophytes may have been the first root-bearing plants and that this group of land plants evolved during the mid-Devonian (Brundrett, 2002). Some of their descendants such as Lycopodium, Selaginella and Isoetes are still extant. One hypothetical scenario is that plant roots evolved from a common ancestor, beginning with a dichotomous branched root type (e.g. Selaginella) and progressing to roots with a single apical cell (Equisetum and ferns). This eventually gave rise to gymnosperm roots that are characterized by indistinct cell layers, and finally to angiosperm roots with the most highly organized cell layers (Foster and Gifford, 1974). Alternatively, true roots could have evolved independently in several plant lineages that arose in mid-Devonian times during a period of rapid plant diversification and increasing complexity (Kenrick and Crane, 1997; Gensel et al., 2001). Due to the paucity of root fossils, many questions regarding the origin of plant roots remain unanswered. Since recent phylogenetic analysis indicate that ferns are the closest relatives of seed plants (Pryer et al., 2001), detailed studies on the anatomy, development and molecular genetic control mechanisms of fern roots promise to shed more light on our understanding of plant root evolution. As the focus of this chapter is not plant root evolution, the reader is referred to a recent review article (Friedman et al., 2004). In this chapter, we will limit the coverage of fern root development to anatomical studies with Ceratopteris richardii focusing on the formation of shoot-borne roots, LR development and the intricacies of the RAC. By providing this broad overview of fern root anatomy, we hope to generate increased interest in this important group of land plants for addressing important issues in plant developmental biology.

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8.2 Overview of the fern root system – shoot-borne roots As already mentioned, our general concept about root systems is derived mainly from studies of flowering plants. To describe plant root systems, most botany/biology textbooks use the terms taproot (or primary root) for the first root to develop from the embryo and fibrous roots for adventitious roots arising from the shoot (see also Chapter 5, this volume). To emphasize ‘the continuity of origin from like organs’, Groff and Kaplan (1988) defined the root system as the total of all roots whose origin can be traced back to other roots or to the same parent root. In accordance with this definition, the taproot is a single root system, whereas the fibrous root system refers to multiple root systems. At the same time, the concept of primary homorhizy in fern root development, in which all roots originate from a single shoot system during normal development (Hou and Hill, 2002; Fig. 8.1), draws attention to the need to reconsider the traditional description of plant roots. Groff and Kaplan (1988) rightfully suggested that the term ‘shoot-borne roots’ should be used rather than ‘adventitious’, because of the predictable pattern in which fern roots arise from the shoots. In the homosporous fern C. richardii, two shoot-borne root populations exist, even though all roots arise from the shoot system via (de-)differentiation of an expanding root apical mother cell (RAMC) in the hypodermal cell layer (Hou and Hill, 2002). The first root population is categorized as stem-borne roots, where roots are produced by a developing shoot with one root per node (Figs. 8.1a and 8.1b). The second root population, referred to as leaf-borne

Figure 8.1 Young sporophytes with roots formed during early ontogeny in the fern Ceratopteris richardii. (a) Roots numbered sequentially according to their position along the shoot. One root is produced in association with one leaf per node. An LR begins to emerge on root 4 (arrow). (b) Emergence of many more LRs from roots at position 4–6 (arrows) in an older sporeling. Roots at position 1–3 generally do not produce LRs and show determinate growth. (c) Portion of the third sporophyll of C. richardii with a stem-borne root (arrow) and four leaf-borne roots (arrowheads). Bar = 5 mm (a, b) and 1 mm (c).

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Root Development roots, originates from the base of the leaves (Fig. 8.1c), with leaf 7 as the typical first node that bears such roots. In leaf-borne roots, the root number per leaf increases with increasing leaf position in young sporophytes of C. richardii. The leaf-borne root population probably represents a heteroblastic variation among the leaves from which these roots are initiated (Hou and Hill, 2002). 8.2.1

Stem-borne roots

In C. richardii, stem-borne roots are distinguished from leaf-borne roots by two salient features. First, only one root initiates per node for the stem-borne root along the shoot, whereas in leaf-borne roots, the number of initiating roots increases at each node, usually beginning with leaf 7. Second, successive stem-borne roots gradually change in root size, structural complexity, growth and LR production capabilities as evidenced in C. richardii (Figs. 8.1a and 8.1b; Hou and Hill, 2002). On the other hand, leaf-borne roots possibly lack such predictable patterns of individual root growth changes. For instance, the root diameter varies even among leaf-borne roots produced at the same node (G. Hou, unpublished data). The distinct root changes during ontogeny in stem-borne roots are comparable to the heteroblastic development well known for leaves of many plant species. In C. richardii, differentiation of the RAMC of a stem-borne root and the corresponding leaf apical cell occurs in the same merophyte, which is originally derived from a single shoot apical cell division (Hou and Hill, 2002). However, the nature and significance of root heteroblastic development have not been widely studied and the fern system provides an excellent model to begin and address this issue. Plant heteroblastic development has been shown to depend on age and reflects ontogenetic phase changes (Poethig, 1990). Several mutants that affect leaf and shoot heteroblasty in maize (Poethig, 1990) and Arabidopsis (Tsukaya et al., 2000) have been identified clearly indicating that heteroblastic traits are under tight molecular genetic control. In leaf studies, however, although the basic pattern of ontogenetic change in leaf shape is intrinsic and therefore genetically determined, the transition can be slowed or reversed (Allsopp, 1965), and the sequence of leaf shapes can be modified by environmental conditions (Jones, 1995). Similar mechanisms appear to operate in root heteroblastic development. In the first detailed study of root heteroblasty in C. richardii, roots at positions 1–3 generally lack LRs (Fig. 8.1b; Hou and Hill, 2002). However, changing the growth conditions, such as medium composition and illumination period, can induce roots at position 3 (position 2 in some individual plants) to produce LRs (G. Hou and E.B. Blancaflor, unpublished data), implying that similar to leaf heteroblastic development, the sequence of LR production along the stem-borne roots is not predetermined. Outstanding questions that remain include the identification of genes that regulate fern root heteroblastic development and how shoot aging guides root system formation. Future molecular studies in ferns should bring some answers to these important questions.

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8.2.2

Leaf-borne roots

As stated above, leaf-borne roots originate from the leaf base right above the stem-borne root in C. richardii (Fig. 8.1c; Hou and Hill, 2002), beginning from position (node) 7. The number of the leaf-borne roots increases with increasing leaf position as successive leaves become bigger and more complex. The leafborne roots originate from individual hypodermal cells at the abaxial side of the leaf base (Hou and Hill, 2002), but anatomically stem-borne and leaf-borne roots do not substantially differ except for root size (G. Hou, unpublished data). The distinction between stem-borne and leaf-borne roots emphasizes the existence of two developmental programs in root system formation in C. richardii. Each of these developmental programs is mirrored in certain monocotyledonous plants. For instance, several Philodendrons produce stem-borne-like roots only. In this case, root formation typically follows a pattern of one root produced per leaf/stem node, although roots at many nodes may not emerge or die prematurely because of environmental conditions. Leaf-borne roots are generally not formed in several Philodendron species observed (G. Hou, unpublished data). At the same time, the leaf-borne root population in ferns (Hou and Hill, 2002) is reminiscent of the postembryonic shoot-borne root formation in Zea mays and some other monocotyledonous species. However, the shoot-rooting pattern in corn conforms to secondary homorhizy to distinguish it from that of primary homorhizy in ferns. Mutations in C. richardii that affect stem-borne but not the leaf-borne root formation, or vice versa, will shed more light on our understanding of the molecular genetic control of the shoot-borne root system formation (Hou and Hill, 2002).

8.3 Anatomical and structural aspects of fern root development It is well established that most fern roots possess a tetrahedral/pyramidal RAC (Fig. 8.2) that is the ultimate source of all cells in the root (Gunning et al., 1978; Gifford, 1991; Hou and Hill, 2004). The RAC always divides asymmetrically and sequentially and each division produces a daughter merophyte and a self-perpetuating RAC. Merophytes are cell packets that were originally referred to as ‘equivalent parts of the plant’ (Gunning et al., 1978). Merophyte boundaries can be clearly identified within developing roots thereby facilitating detailed studies on the cellular basis of root development. For example, merophytes derived from the RAC distal division face contribute to the root cap, while merophytes from the three RAC proximal division planes give rise to the root proper (Figs. 8.2 and 8.3). A consequence of sequential RAC divisions, especially along the three proximal faces, is the generation of a strict merophyte helix along the root longitudinal axis during development. Each merophyte seems to acquire its unique temporal and special identity

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Figure 8.2 Representative median longitudinal sections of a stem-borne root at position 4 (a), and an arbitrary leaf-borne root (b) of Ceratopteris richardii. All roots, regardless of origin, possess a single RAC that is the ultimate source of all cells in the root. (c) Indirect immunofluorescence labeling of microtubules in a stem-borne root with an RAC in mitosis. Bar = 10 µm (c).

at the onset of RAC division and this unique identity is maintained in coordination with other merophytes throughout root ontogeny (Gunning, 1982; Fig. 8.2). 8.3.1

RAC formation and division

The first sign of shoot-borne root initiation in young sporophytes of C. richardii is cell expansion to form an RAMC at the base of each leaf primordium and beneath the protoderm (Hou and Hill, 2002). Upon reaching a certain size, the RAMC divides four times sequentially, producing four daughter merophytes resulting in the formation of an RAC in the middle. Thereafter, the RAC divides along the three proximal division planes (Fig. 8.3b) and contributes to the root proper formation. Therefore, root formation in C. richardii can be divided into three developmental stages based on anatomical criteria: cell expansion to form an RAMC; RAMC division to produce four daughter merophytes and a tetrahedral RAC and RAC divisions continuously adding merophytes to the root. The general sequence of the three stages in the formation of roots is essentially the same as that for LR formation in C. richardii (Hou et al., 2004; also see below). The RAC divisions are predictable because subsequent division planes are always positioned parallel to the oldest of the three immediate daughter merophytes (Fig. 8.3b). Each of the merophytes occupies a sector of approximately 120◦ in transverse view. Successive merophytes overlap with one another along an imaginable helix around the root longitudinal axis. A merophyte that is closer to the RAC is always younger (Fig. 8.2) and a root can also be viewed as consisting of three columns (or orthostichies) of merophytes. How the RAC division sequence is determined and maintained throughout fern root ontogeny is still a matter of speculation. Positional signaling

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(a)

(b)

(c)

Arabidopsis

Ceratopteris

Figure 8.3 Morphology and anatomy of roots at position 1 of Ceratopteris richardii. (a) Scanning electron micrograph of the root tip region showing the root cap (arrow). (b) A single RAC is the ultimate source of all cells in the root. The RAC divides following a rigid sequence specified during the RAC formation. The daughter merophytes are labeled with numerals and the RAC division is clockwise in the root. The next RAC division along the three proximal division planes will be parallel to merophyte 3. RC, root cap. (c) Representative micrographs showing the root cap of Arabidopsis thaliana and the fifth root in C. richardii. Merophytes derived from the distal division plane of the RAC contribute to the cap formation. Note that the central region of the fern root cap resembles the columella region (cc) in the root cap of Arabidopsis. Bars = 50 µm (c).

probably plays an important role in the formation of the RAC because the shoot-borne roots always form at predictable sites on the shoot in C. richardii (see above; Hou and Hill, 2002) and the RAC divides clockwise (Fig. 8.3b) or counter-clockwise depending on its position on the shoot, as is the case in Azolla (Gunning et al., 1978). The nature and the origin of a positional signal that allows the fern root to maintain this highly predictable pattern of division that begins with the formation of the RAC and derivative merophytes remain unknown. Physical and genetic ablation studies that have been useful in defining positional relationships and individual cell function in Arabidopsis should shed light into these important processes (van den Berg et al., 1995; Blancaflor et al., 1998; Tsugeki and Fedoroff, 1999).

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Root Development Unlike studies with flowering plants (Hawes et al., 2003), root cap development in ferns has not attracted much attention. As noted above, the RAC divides in its distal face and produces merophytes for root cap development (Fig. 8.3c), but not with equal frequency along its distal division plane when compared with the division frequency along the three proximal division planes in C. thalictroides (Chiang and Gifford, 1971). The RAC produces one merophyte for root cap development in approximately every ten RAC divisions (Chiang and Gifford, 1971). Anatomically, the root cap of C. richardii appears to have some features similar to those of flowering plants. For example, a columella-like region, an important root cap structure required for gravitropism that is often described in flowering plants, is apparent in longitudinal sections of the root cap of C. richardii (Fig. 8.3c; see also Chapter 6, this volume). A closer examination of cap cells in fern roots shows that like flowering plant roots, the putative columella region in fern roots caps contains dense plastids (Fig. 8.3c). It will be interesting to determine whether fern root plastids function in gravity sensing similar to angiosperm roots. 8.3.2

Formative and proliferative cell divisions

In a growing fern root, an older merophyte is representative for the future of the next younger merophyte within the same orthostichy (Gunning et al., 1978; Hou and Hill, 2004). Therefore, the sequence of cell division within a merophyte can be inferred from the analysis of successive merophytes by serial transverse sections. Two general types of cell divisions have been recognized during merophyte ontogeny in C. richardii roots: formative and proliferative. Formative divisions occur in the zone where different cell file initials are formed (Gunning et al., 1978) and are either periclinal or radial anticlinal, but the resulting two daughter cells usually have different developmental fates. Formative divisions establish cell initials for all tissue types in a merophyte and, in turn, make up the various tissues of the root. On the other hand, proliferative divisions occur so that two daughter cells have the same developmental fates. The function of proliferative cell divisions is to increase the cell numbers within the same file/tissue type. Eight or nine characteristic formative divisions have been identified within each merophyte in the fifth stem-borne roots of C. richardii (Fig. 8.4; Hou and Hill, 2004). The first four formative divisions (D1–D4) are further classified as precursor divisions because daughter cells do not directly form specific cell or tissue types. D5–D7 are founder divisions because new cells with specified developmental fates are generated (Fig. 8.4; Hou and Hill, 2004). D8 is considered an early proliferative division, because the resulting two daughter cells usually develop into vascular cells (i.e. xylem or phloem; Hou and Hill, 2004). Among the formative divisions in the fifth root, D9 appears to be the most variable in terms of differentiation patterns leading to either two pericycle cells or a pericycle cell and a sieve cell. Thus, it is difficult to classify

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Figure 8.4 Cell lineage diagram of formative divisions in a root merophyte (derived from Hou and Hill, 2004). Seven formative divisions (D1 to D7) occur in every merophyte of the fifth stem-borne root in C. richardii. Division 8 (D8) occurs in one of two inner cells formed by D6 (∗∗ ) and is classified as an early proliferative division and D9 in one of two outer cells formed by D6 (∗ ). In approximately half of the D9s examined, each produces a pericycle initial and an outer sieve element. D9 takes place in two of three successive merophytes.

D9 as exclusively formative or proliferative. Subsequent cell divisions after D9 in a merophyte are strictly proliferative (Hou and Hill, 2004). In contrast to the homosporous fern Ceratopteris, which has eight to nine formative divisions, 11 formative divisions have been identified in developing roots of the heterosporous fern Azolla (Gunning et al., 1978). Like Ceratopteris roots, the temporal regulation of formative division sequence in Azolla is rigid and predictable. The root radial structural pattern generated by D1–D9 in each merophyte of Azolla is essentially the same as that described for the fifth root of C. richardii, although the early division sequence/order (i.e. precursor divisions) differs between the two species (Gunning et al., 1978; Hou and Hill, 2004). In Azolla roots, D10 delineates the protoxylem from metaxylem and D11 partitions the trichoblast from atrichoblast. Meanwhile, all Azolla roots show determinate growth and bear no LRs (Gunning et al., 1978). Although the role of proliferative cell divisions is to increase the number of cells within a tissue, the pattern of such divisions often varies depending on the tissue type, among different roots of the same species, and among different species (Gunning et al., 1978; Hou and Hill, 2004). For example, in A. pinnata the metaxylem initial differentiates without proliferative division, whereas each endodermal initial undergoes four complete rounds of

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Root Development proliferative divisions (Gunning et al., 1978). Roots of Asplenium vary considerably in diameter/size; the formative divisions in early merophyte ontogeny are essentially the same among different roots. In Asplenium, root size is believed to depend primarily on the proliferative divisions of cortex initials (Gifford, 1991). However, root size variations in the root heteroblasty of young C. richardii sporophytes involve proliferative divisions in all cell files, including the central stele, of the roots (Hou and Hill, 2002). It is interesting to note that roots of the above three genera are all diarch with twofold anatomical symmetry characteristic of mature roots superimposing on a threefold radial symmetry that originates behind the RAC. This observation implies that root radial structural pattern formation is specified by the orchestration of successive merophytes and signaling for this process must transcend merophyte boundaries (see below). 8.3.3

Differentiation and structural pattern formation

The above discussion has so far focused on differentiation into specific root cell types based on lineage. However, like in flowering plant roots there is also evidence that cells in fern roots can differentiate according to their position rather than their lineages (Gunning et al., 1978; Hou and Hill, 2004). In C. richardii, cells can differentiate into LR mother cells (see below) or typical endodermal cells, depending on their position within the root. A similar argument can be made for cells whose lineage can be traced to the pericycle since cells within this lineage can continue to produce pericycle cells or outer sieve elements (Hou and Hill, 2004). The latter case is the same in Azolla and Asplenium (Gunning et al., 1978; Gifford, 1991). Considering that positions of cell differentiation progress toward the root apex as the individual root ages (Gunning et al., 1978) and most differentiation processes transcend the merophyte boundaries, for example twofold radial structural pattern arising from threefold merophyte arrangement (Gunning et al., 1978; Hou and Hill, 2004), the positional signals specifying cell developmental fate must be interactive and cooperative. The nature of the positional signal, where the signal originates, and how the signal is propagated among merophytes are outstanding questions that remain to be answered. Emphasizing the role of global positional information does not necessarily discount the contributions of cell division per se in root morphogenesis. In fact, it is the ordered (rather than random) cell divisions that delineate daughter cells to a specific position relevant to the other cells in the root as a whole. Gunning (1982) hypothesized that microtubules might provide a means of receiving and translating positional information to guide cell division, expansion and other aspects of differentiation. It is interesting to note that when the formative division 7 of Azolla roots that forms inner cortex and endodermis lineages is disrupted, subsequent division patterns in the two cell files are also changed (Gunning, 1982). This is reminiscent of the defects observed in the Arabidopsis loss-of-function mutants scarecrow (scr) and

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short root (shr), that lack asymmetric cell division in the cortex/endodermal initials resulting in a single ground tissue layer rather than a normal cortex and endodermal layer (Helariutta et al., 2000). Both SHR and SCR encode putative transcription factors and SHR functions in a noncell-autonomous way upstream of SCR (Helariutta et al., 2000; Nakajima and Benfey, 2002). The link between SHR/SCR and cell cycle control that guarantees the cortex/endodermal initial cell division is still unknown.

8.4 LR formation in ferns LRs can be defined as roots that are endogenously initiated from existing (parent) roots. Studies of LRs open another window for understanding plant morphogenesis since LR formation is a de novo process and exhibits regular developmental patterns (Lin and Raghavan, 1991; Hou et al., 2004). In contrast to flowering plants, LRs in ferns are initiated in the endodermal layer and emerge in an acropetal sequence in rows or ranks (Fig. 8.5a). Each of the LRs also possesses a single tetrahedral/pyramidal apical cell (LRAC), which is the ultimate source of all cells in the LR. The initiation of the lateral root apical mother cell (LRMC) occurs very close to the parent root apex (Figs. 8.5b and 8.5c). One LRMC is produced per merophyte and only two in every three successive merophytes are involved in the LR formation, i.e. three parent RAC divisions along the proximal division planes will form two LRs in the

Figure 8.5 LR developmental pattern on representative sixth stem-borne roots in Ceratopteris richardii. (a) Emergence of LRs alternatively in two ranks along the parent root longitudinal axis. (b) LR initiation (black arrows) very close to the parent root apex and formation of several LR primordia before emergence. (c) LR formed in the endodermal layer with the lateral RAC (white arrow) representing the ultimate source of all cells in the LR. Bar = 5 mm (a).

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Root Development opposite position. Interestingly, the two ranks of LR patterning are correlated with the parent root stele structure, i.e. diarch organization (Chiang and Gifford, 1971; Lin and Raghavan, 1991; Hou et al., 2004). LR development can be divided into three stages in C. richardii (Hou et al., 2004). The first indication of LR formation is cell expansion prior to any cell division of a particular cell in the endodermal layer to form an LRMC adjacent to the xylem pole, which is designated as stage 1 of LR development. When LRMC reaches a threshold size, it divides four times asymmetrically and sequentially to form a pyramidal LRAC in the middle and four daughter merophytes (stage 2; Fig. 8.5c). The division sequence along the three proximal planes of the LRAC established during this second stage perpetuates throughout LR ontogeny. Stage 3 is defined as LR growth and emergence via the continued activity of the LRAC and its daughter merophytes (Hou et al., 2004). LR development in Marsilea quadrifolia is similar to that described for C. richardii and the formative division patterns in merophyte ontogeny in LRs are identical to those observed in the parent roots (Lin and Raghavan, 1991). It is well established that LR initiation in flowering plant roots is under tight regulation by the plant hormone auxin (De Smet et al., 2006; see Chapter 4, this volume). Interestingly, fern LR initiation generally does not respond to exogenous addition of auxins, but strong growth inhibitory effects on root elongation are still apparent (Hou et al., 2004). It is well known that exogenous auxin can increase the number of LRs formed on parent roots in Raphanus sativus and Arabidopsis (Laskowski et al., 1995). Blocking endogenous auxin transport also affects LR development in Arabidopsis (Casimiro et al., 2003). However, exogenous indole acetic acid (IAA) or indole butyric acid (IBA) does not induce additional LRs in C. richardii. LRs continue to emerge in two ranks along the parent root with the pattern of alternative long and short intervals between successive LRs, even after the addition of exogenous auxins (Hou et al., 2004). Also, the auxin transport inhibitor 1-N-naphthylphtalamic acid (NPA) does not change the LR developmental patterns in C. richardii. Nevertheless, like exogenous auxin application, NPA significantly inhibited parent and LR root growth (Hou et al., 2004). In some flowering plants, the existence of two LR populations has been described: acropetal and adventitious (reviewed by Charlton, 1996), in the sense that the acropetal LR population is initiated in the parent meristematic zone and the LR initials do not undergo a de-differentiation process (Beeckman et al., 2001), LR formation on the fifth stem-borne roots of C. richardii resembles the acropetal LR population described in flowering plants and the fifth stemborne root might simply lack the ‘adventitious’ LR population (Hou et al., 2004). However, studies of solitary-root (slr) mutant indicate that acropetal LR formation in Arabidopsis is also regulated by hormone auxin (Fukaki et al., 2002, 2005). The dominant slr-1 mutant completely lacks LRs and this phenotype cannot be rescued by the application of exogenous auxin. SLR encodes IAA14, a member of the Aux/IAA protein family (Fukaki et al., 2002, 2005). Therefore, further molecular and genetic investigations of mutants

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with any disruption in LR developmental pattern in C. richardii promise to shed light on our understanding of the control mechanism of LR formation in the fern specifically and of the differences between the fern and the seed plant generally.

8.5 Concluding remarks and future prospects Our coverage of fern root development has focused exclusively on anatomical aspects because most of the work to date on fern sporophytic development lacks a significant molecular component and stands in stark contrast to studies in fern gametophytes where tools for gene silencing are continuously being refined (Klink and Wolniak, 2001; Stout et al., 2003; Rutherford et al., 2004) and mutants that affect various stages of gametophyte development can be readily scored and generated (Banks, 1999). More recently, an expressed sequence tag library constructed from C. richardii spores and used for subsequent microarray analysis during early stages of spore germination points to parallel regulatory mechanisms with Arabidopsis pollen germination (Salmi et al., 2005; Bushart and Roux, 2007). A high-resolution genetic linkage map in C. richardii has also been generated that should pave the way for more in-depth genetic studies in this model fern (Nakazato et al., 2006). Although research on fern sporophytic development has yet to exploit the above genetic resources, anatomical studies of fern roots have revealed visible and predictable patterns of development that enable great opportunities for further experimental manipulations. For example, it is obvious that positional signaling plays a critical role in cell fate acquisition in fern roots. Cell division, especially the founder divisions, is a likely component of this positional signaling network since cell division partitions a cell to a specific position relevant to other cells within the organ. Studies on the RAC and its derivative merophytes using currently available cell biological and genomic tools such as specific cell laser ablation and laser capture microdissection (Blancaflor et al. 1998; Woll et al., 2005) should help unravel aspects of the molecular genetic and cellular networks that govern the formation of the RAC and how this single cell eventually defines root structural pattern formation. Furthermore, comparative analysis of gene expression changes in fern sporophytes and flowering plants provide a good entry point (Sano et al., 2005) for dissecting the molecular basis of fern root development and complement the rich fern anatomical data that are currently available.

Acknowledgments We thank the Noble Foundation and the National Science Foundation (grant number DBI- 0400580) for funding the research related to this work.

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Gifford, E.M. (1993). The root apical meristem of Equisetum diffusum: structure and development. Am. J. Bot. 80, 468–473. Gifford, E.M. and Foster, A.S. (1989). Morphology and Evolution of Vascular Plants. New York, W.H. Freeman, p. 625. Groff, P.A. and Kaplan, D.R. (1988). The relation of root systems to shoot systems in vascular plants. Bot. Rev. 54, 387–422. Gunning, B.E.S. (1982). The root of water fern Azolla: cellular basis of development and multiple roles for cortical microtubules. In Developmental Order: Its Origin and Regulation, S. Subtelny and P.B. Green (eds). New York, Alan Liss, pp. 379– 421. Gunning, B.E.S., Hughes, J.E. and Hardham, A.R. (1978). Formative and proliferative cell divisions, cell differentiation, and developmental changes in the meristem of Azolla roots. Planta 143, 121–144. Hawes, M.C., Bengough, G.A., Cassab, G. and Ponce, G. (2003). Root caps and rhizosphere. J. Plant Growth Regul. 21, 352–367. Helariutta, Y., Fukaki, H., Wysocka-Diller, J., Nakajima, K., Jung, J., Sena, G. et al. (2000). The SHORT-ROOT gene controls radial patterning of the Arabidopsis root through radial signaling. Cell 101, 555–567. Hou, G., Hill, J.P. and Blancaflor, E.B. (2004). Developmental anatomy and auxin response of lateral root formation in Ceratopteris richardii. J. Exp. Bot. 55, 685– 693. Hou, G.-C. and Hill, J.P. (2002). Heteroblastic root development in Ceratopteris richardii (Parkeriaceae). Int. J. Plant Sci. 163, 341–351. Hou, G.-C. and Hill, J.P. (2004). Developmental anatomy of the fifth shoot-borne root in young sporophytes of Ceratopteris richardii. Planta 219, 212–220. Jones, C.S. (1995). Does shape prolong juvenile development? A morphological analysis of leaf shape changes in Cucurbita argyrosperma subsp. sororia (Cucurbitaceae). Am. J. Bot. 82, 346–359. Jones, C.S. (1999). An essay on juvenility, phase change, and heteroblasty in seed plants. Int. J. Plant Sci. 160, S105–S111. Kenrick, P. and Crane, P.R. (1997). The origin and early evolution of plants on land. Nature 389, 33–39. Klink, V.P. and Wolniak, S.M. (2001). Centrin is necessary for the formation of the motile apparatus in spermatids of Marsilea. Mol. Biol. Cell 12, 761–776. Kramer, E.M. and Bennett, M.J. (2006). Auxin transport: a field in flux. Trends Plant Sci. 11, 382–386. Kwak, S.-H. and Schiefelbein, J. (2007). The role of the SCRAMBLED receptor-like kinase in patterning the Arabidopsis root epidermis. Dev. Biol. 302, 118–131. Laskowski, M.J., Williams, M.E., Nusbaum, H.C. and Sussex, I.M. (1995). Formation of lateral root meristems is a two-stage process. Development 121, 3303–3310. Lin, B.-L. and Raghavan, V. (1991). Lateral root initiation in Marsilea quadrifolia. I. Origin and histogenesis of lateral roots. Can. J. Bot. 69, 123–135. Nakajima, K. and Benfey, P.N. (2002). Signaling in and out: control of cell division and differentiation in the shoot and root. Plant Cell 14 (Suppl), S265–S276. Nakazato, T., Jung, M.-K., Housworth, E.A., Rieseberg, L.H. and Gastony, G.J. (2006). Genetic map-based analysis of genome structure in the homosporous fern Ceratopteris richardii. Genetics 173, 1585–1597. Poethig, R.S. (1990). Phase change and the regulation of shoot morphogenesis in plants. Science 250, 923–930.

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Annual Plant Reviews (2009) 37, 209–238 doi: 10.1002/9781444310023.ch9

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Chapter 9

WHEN PLANTS SOCIALIZE: SYMBIOSES AND ROOT DEVELOPMENT Benjamin P´eret, Sergio Svistoonoff and Laurent Laplaze Institut de Recherche pour le D´eveloppement (IRD), UMR Diversit´e et Adaptation des Plantes Cultiv´ees (Agro.M/INRA/IRD/UM2), Equipe rhizogen`ese, 911 Avenue Agropolis, F-34394 Montpellier Cedex 5, France

Abstract: Plant roots encounter a large variety of living organisms in the soil. Some soil microorganisms associate with plant roots to form symbiotic interactions. In this chapter, we describe the establishment of two widespread and important symbiotic associations: mycorrhizae and actinorhizae; and the developmental changes that they induce both at the site of interaction and on the whole root system. Keywords: arbuscular mycorrhizae; ectomycorrhizae; fungi; actinorhizal symbioses; nitrogen fixation; Frankia

9.1 Introduction The soil is a heterogeneous and changing environment where plants need to find the water and nutrients necessary for their growth (Hodge, 2006). It is also a living environment containing a diverse and complex community of microorganisms and animals as well as plant root systems. These living organisms influence plant growth and nutrition and, in return, plant root growth and physiology have marked effects on this community. In some cases, plant roots enter mutualistic associations with soil microorganisms leading to intimate relationships that can even result in the development of specialized symbiotic root organs. In this chapter, we describe the establishment of two widespread and important symbiotic associations: mycorrhizae and actinorhizae. We focus on the developmental changes that occur both at the site of interaction and on the whole root system. For legume nodulation, see Chapter 10. 209

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Root Development Mycorrhizae are symbioses between plant roots and fungi that increase plant nutrients and water uptake. For a long time, mineral nutrient deficiency, especially phosphorus and nitrogen, was known to be a prerequisite for mycorrhiza formation (Hatch, 1937; Smith and Read, 1997). In their natural environment, most plants associate with mycorrhizal fungi and therefore this symbiosis is a rule rather than an exception. There are two main types of mycorrhizal interactions: arbuscular mycorrhizae and ectomycorrhizae. Arbuscular mycorrhizae (AM) are a very ancient symbiosis that evolved during plant colonization of land and occurs in more than 80% of land plants. On the other hand, ectomycorrhizal associations (EC) evolved more recently and are the predominant symbiotic interactions for most temperate trees. Other more specific types of mycorrhizae, such as orchid mycorrhizae or ericoid mycorrhizae, exist (Smith and Read, 1997), but will not be discussed here. The amount of available nitrogen in the soil is an important limiting factor for plant growth. Although dinitrogen is plentiful in the air, only a few prokaryotes possess the biochemical machinery to use this source of N for their nutrition. During the course of evolution, certain plant species developed an intimate relationship with nitrogen-fixing bacteria leading to the formation of novel root symbiotic organs. Two groups of plants can develop nitrogen-fixing root nodules: legumes that associate symbiotically with rhizobia (see Chapter 10) and so-called actinorhizal plants that belong to eight angiosperm families associated with the actinomycete Frankia. Interestingly, although actinorhizal and legume nodules differ in their development and structure (Pawlowski and Bisseling, 1996), molecular phylogenetic studies indicate that legume and actinorhizal plants group in the same clade (rosid I) suggesting a single origin for nodulation capacity (Soltis et al., 1995). Therefore, a comparison of the molecular mechanisms involved in establishing these two plant–microbe interactions should shed light on how these important nitrogen-fixing symbioses have evolved. Moreover, some results indicate that a subset of genes involved in nitrogen-fixing root nodule symbioses were recruited from the ancient AM symbiosis (Paszkowski, 2006). Future work should help to understand to what extent molecular mechanisms are shared and to identify core symbiotic genes.

9.2

Arbuscular mycorrhizae

Arbuscular mycorrhizae (AM) are widespread associations formed between approximately 200 fungal species from the order of Glomeromycota, named arbuscular mycorrhizal fungi (AMF), and more than 80% of all land plants, including angiosperms, gymnosperms, pteridophytes and bryophytes (Trappe, 1987; Wang and Qiu, 2006). AM symbioses appear to be at least as old as terrestrial colonization by plants and mycorrhization probably played a central role in this evolutionary development by assisting plants with primitive

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root systems to deal with adverse conditions, such as limited nutrients and water availability (Pirozynski and Malloch, 1975; Simon et al., 1993; Remy et al., 1994; Redecker et al., 2000). In most cases, AMF are beneficial to the plant host, thanks to (1) improved nutrient uptake mediated by fungal hyphae that are able to explore a larger volume of soil and mobilize nutrients more efficiently than roots, and (2) protection against root fungal pathogens, mainly because their niche is already occupied by AMF (Smith and Read, 1997). The association with the plant partner is absolutely essential for AMF to complete their life cycle. AMF are obligate biotrophs that rely entirely on the plant partner for their carbohydrate supply (Smith and Read, 1997). AMF produce multinucleate spores asexually, and sexual reproduction has almost never been observed (Pawlowska, 2005). The mechanisms by which AMF maintain their genome remain to be elucidated – AMF might be clonal organisms lacking any form of recombination, or genetic exchange might still occur through conventional cryptic sexual exchanges or illegitimate hyphal fusions (Pawlowska, 2005). Surprisingly, despite its enormous ecological importance, little information is available concerning the molecular mechanisms involved in this symbiosis, mainly due to AMF biotrophy and recalcitrance to genetic transformation (Paszkowski, 2006). 9.2.1

Preinfection events

Three successive stages have been proposed to occur during the plant interaction with AMF (Fig. 9.1). During the first or asymbiotic stage, germination of dormant AMF spores and limited hyphal growth is induced by a variety of stimuli, including water, CO2 and root exudates (B´ecard et al., 2004). In the absence of host plant roots in the vicinity of the developing hyphae, growth stops before the spore reserves are depleted and the fungus reenters the dormant state (Fig. 9.1a). Depending on the size of the spore, up to ten germinations can take place (B´ecard et al., 2004). When the hyphae come close to a host root, the fungal cells undergo strong metabolic changes, including increased respiration, mitochondria biogenesis, changes in cytosolic pH and membrane polarization (Tamasloukht et al., 2003; B´ecard et al., 2004). Shortly afterwards, extensive hyphal branching takes place, thus increasing the chances of contact with the host root. At this point, hyphal growth continues until all the reserves are depleted, the fungus enters the so-called presymbiotic phase and return to the dormant stage is no longer possible (Fig. 9.1b) (B´ecard et al., 2004). AMF detect the presence of a host root by sensing so-called ‘branching factors’, i.e. signaling compounds emitted by the root that stimulate hyphal branching (Buee et al., 2000). Recently, strigolactones, a group of sesquiterpenes, have been identified as branching factors (Akiyama et al., 2005). These molecules were identified as stimulators of seed germination in parasitic weeds Striga and Orobanche.

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(b)

(c)

Figure 9.1 Formation of Arum-type arbuscular mycorrhizal symbiosis. (a) Symbiotic stage: spore germination is followed by limited hyphal growth. (b) Presymbiotic stage: strigolactones secreted by the plant root induce metabolic changes and extensive hyphal branching. (c) Symbiotic stage: upon contact with the host root a hyphophodium is formed enabling hyphal penetration through an epidermal cell. Cortical cells are then colonized by intercellular hyphae that develop vesicles and intracellular arbuscules.

In return, diffusible signals produced by AMF are presumably perceived by the host plant prior to infection (Vierheilig, 2004). Hence, by homology to Nod factors, ‘Myc’ factors might activate a signaling pathway analogous to that elicited by Nod factors in legumes (Albrecht et al., 1998; Blilou et al., 1999; Ol´ah et al., 2005; see also Chapter 10). Interestingly, diffusible fungal factors have been shown to increase the level of cytosolic calcium in soybean cell cultures (Navazio et al., 2007) and to activate the promoter of MtEnod11 (Kosuta et al., 2003), a gene previously shown to be expressed during early stages of the AM interaction (Chabaud et al., 2002). This activation can be observed even when both partners are separated by a cellophane membrane. The chemical nature of these ‘Myc’ factors is still unknown, and the analogy with Nod factor’s signaling is unclear, since the activation of MtEnod11 does not depend on functional Does not Make Infection (DMI) genes (see below). Genetic analyses in legumes suggest that Nod factors and putative Myc factor responses share part of a preinfection signal transduction cascade. Many legume nodulation mutants have also been shown to be defective in

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AM symbiosis (Marsh and Schultze, 2001; Oldroyd and Downie, 2004; see Chapter 10). Common genes include (1) receptor-like kinases with extracellular leucine-rich-repeat (LRR) domains encoded by Medicago truncatula Mt-DMI2 and the corresponding genes from M. sativa (Ms-NORK), pea (PsSYM19) and Lotus japonicus (Lj-SYMRK) (Endre et al., 2002; Stracke et al., 2002), (2) ion channel-like proteins encoded by Mt-DMI1 and its counterparts in L. japonicus (Lj-POLLUX and Lj-CASTOR) (An´e et al., 2004; Imaizumi-Anraku et al., 2005), (3) LjNUP133 encoding a protein similar to the mammalian nucleoporin 133 (Kanamori et al., 2006), (4) Ca2+ /calmodulin (CaM)-dependent protein kinases that act downstream of the Nod factor-induced calcium spiking encoded by Mt-DMI3 from M. truncatula and Ps-SYM9 from pea (L´evy et al., 2004; Mitra et al., 2004). Mutants affected in steps upstream of DMI1/DMI2 or downstream of DMI3 are not affected in AM interactions, suggesting that the hypothetical common signaling pathway diverges before DMI1/DMI2 and after DMI3. Myc factors are probably chemically different from Nod factors and require different receptors (Paszkowski, 2006). The evolutionarily recent legume–Rhizobium symbiosis (less than 60 million years) probably evolved by recycling parts of the much older AM signaling pathway. If so, orthologs of DMI genes would be expected to exist in nonnodulating plants, which is indeed the case as they have been found in rice, tobacco, wheat and even in the nonmycorrhizal plant Arabidopsis (Zhu et al., 2006). Recently, the rice ortholog of DMI3 was shown to play a crucial role in AM symbiosis, thus suggesting the widespread role of the DMI signaling pathway among plants that form AM (Chen et al., 2007). Several other mutants affected in both the AM symbiosis and nodulation have been described (Marsh and Schultze, 2001) and new screens to isolate mutants affected in the AM symbiosis but not in nodulation have been conducted on the model legumes L. japonicus and M. truncatula. The identification of the corresponding genes will certainly provide new insights into the molecular mechanisms of the AM symbiosis, including Myc factor perception and transduction. 9.2.2

Root infection and colonization

The third stage of the interaction is the symbiotic phase. This phase starts when fungal hyphae contact the host root, triggering the differentiation of a hyphophodium that will enable hyphal penetration into the epidermal cells and colonization of the root cortex (Figs. 9.1c and 9.3d; Smith and Read, 1997). Hyphal infection occurs through invagination of the plasma membrane that separates the hyphae from the plant cell cytoplasm. In the space between the fungal cell wall and the plasma membrane, extracellular matrix material is deposited whose composition resembles that of primary cell wall (Smith and Read, 1997). Confocal microscopy has revealed that, just before infection, the cytoskeleton/ER of the epidermal cell undergoes major reorganization, involving nuclear movement and the assembly of a tubular structure termed the

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Root Development prepenetration apparatus (PPA). This transient assembly is thought to play a key role in constructing the transcellular invagination that subsequently guides hyphal growth through the cell (Genre et al., 2005). Importantly, mutants in at least two M. truncatula DMI genes are blocked in PPA formation, suggesting that the perception of a ‘Myc’ signal might be essential for initiation of this specialized cytoskeletal assembly (Genre et al., 2005). After crossing the epidermis, infection proceeds into the root cortex. Depending on the infection patterns two main types of AM can be distinguished. On the one hand, the Arum-type AM (named after the species in which it was first described) is characterized by extensive intercellular development of fungal hyphae and associated formation of arbuscules, intracellular structures that result from dichotomous ramification of hyphae (Figs. 9.1c, 9.3d and 9.3e). On the other hand, the Paris-type AM is typified by mainly intracellular hyphae that often develop intracellular coils and very rarely arbuscules. While the Paris-type AM are more frequent in nature, most research has been performed on the Arum-type AM (Smith and Smith, 1997). Arbuscules and coiled hyphae are believed to be the primary sites for nutrient exchange in the two types of AM (Harrison, 2005). Most AMF also differentiate intra- or intercellular storage organs, called vesicles that contain mainly lipids (Smith and Read, 1997). Mycorrhization and rhizobia–legume interactions also appear to share certain mechanisms that control the development of the symbioses. Split-root experiments have shown that mycorrhization systemically suppresses further nodulation and vice versa (Vierheilig, 2004), and furthermore some supernodulating mutants, such as LjSym78 also show increased mycorrhization (Kawaguchi et al., 2002). Very few mutants that are defective in AM symbiosis have been studied in nonlegume species. The tomato rmc mutant is resistant to colonization by most AMF species (Barker et al., 1998). Maize mutants affected in appressoria formation or cortex invasion and mutants showing enhanced and more rapid fungal invasion have been described (Paszkowski et al., 2006). Unfortunately, the identities of the corresponding genes are still unknown but reverse-genetics approaches based on transcriptomics and proteomics are currently increasing our understanding of the molecular events that regulate AM symbiosis (Paszkowski, 2006). Such an approach has recently been used with LjPT3 and MtPT4, two genes from L. japonicus and M. truncatula respectively that encode phosphate (Pi) transporters highly expressed in infected cells. Silencing or mutation of these genes resulted in reduced Pi uptake via the AMF, growth retardation and a decreased number of arbuscules, suggesting that Pi supply through mycorrhizae is a key factor that controls mycorrhization itself (Maeda et al., 2006; Javot et al., 2007). Recently, lysophosphatidylcholine was identified as a signaling molecule mediating the transcription of the StPT3 Pi transporter in arbuscule-containing cells of potato (Drissner et al., 2007).

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9.2.3

Effects of AM on root development

Originally AM was thought to have little effect on root development (Atkinson et al., 1994) because changes in root development upon AM colonization are subtle and difficult to assess using global approaches, such as measurement of total root weight or length. Furthermore, the wide range of taxonomically distant species that exhibit AM symbiosis often respond in different or even opposing ways to AM colonization (Atkinson et al., 1994; Berta et al., 2002). The effects of AMF colonization on root development depend on the characteristics of each plant species but also on ecological conditions, which affect the cost/benefit balance of each strategy. Topological models to describe root systems have provided tools to analyze and quantify differences between root architectures and have been especially useful in comparing branching patterns between two root systems (Fitter, 1985). By careful comparison of individual root lengths, mycorrhization has been shown to result in most cases in increased root branching. In Allium porrum, branching increased by up to 145% after infection by AMF (Berta et al., 1990). Increased branching has been observed in many species such as garlic (Allium sativum L.) (Regvar et al., 1996), maize (Zea mays) (Kaldorf and ¨ Ludwig-Muller, 2000), barrel medic (Medicago truncatula) (Ol´ah et al., 2005), grape vine (Vitis vinifera) (Schellenbaum et al., 1991), cotton (Gossypium sp.) (Torrisi et al., 1999) and poplar (Populus sp.), the latter producing up to sixfold as many lateral roots when colonized by Glomus caledonium (Hooker et al., 1992). However, in a few plant species, mycorrhization had no effect or even decreased root branching (Hetrick et al., 1988; Price et al., 1989; Trotta et al., 1991; Forbes et al., 1996; Vigo et al., 2000). AMF colonization can also affect other parameters of root architecture, such as increase of root diameter in A. porrum (Fusconi et al., 1994), S. esculentum (Fusconi et al., 1999), A. gerardii (Hetrick et al., 1988) and Prunus cerasifera (Berta et al., 1995), or change in root topology, albeit with contrasting responses. For instance, mycorrhized A. reinhardii plants develop a more elongated exploratory root system (Hetrick et al., 1988) whereas V. vinifera reacts with a more random pattern of root development. In some cases, both responses can be observed at different stages of the symbiotic association: mycorrhization of Platanus acerifolia results in a herringbone pattern at the beginning that changes over time and becomes more dichotomous in older mycorrhized plants compared to uninfected plants (Tisserant et al., 1996). Since mycorrhization substantially improves nutrient and water uptake, two key factors regulate root architecture (see Chapter 11), it is difficult to separate indirect nutritional effects from those directly caused by AMF colonization. In order to discriminate between architectural changes due to improved nutrition and those arising specifically from AMF colonization, increased amounts of nutrients and water were provided to nonmycorrhized plants, thus allowing comparisons between plants of the same size and with

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Root Development similar nutritional levels. Using this experimental design, Trotta et al. (1991) confirmed the results of Berta et al. (1990) on A. porrum that AMF increased branching and decreased the length of lateral roots. The effect on first- and second-order lateral roots was observed even when Pi was supplied, suggesting that at least part of the response was not due to improved nutrition, but to colonization per se. Similar results were obtained on poplar (Hooker et al., 1992), P. cerasifera (Berta et al., 1995), A. reinhardii (Hetrick et al., 1988) and O. europeaea (Citernesi et al., 1998). Another important factor to be considered when studying the impact of AMF on root development is the temporal dimension. In particular for the root systems of perennial plants, the duration of root growth and branching and the time during which the root will stay alive are crucial parameters. Only a small fraction of roots survive over time and the probability of survival decreases with the rank of the root. Primary and secondary roots are likely to be lignified, whereas higher order roots rapidly die and are replaced. Since in most cases AMF colonization increases root branching mycorrhization is to be expected to have a negative impact on root survival in woody plants. This assumption has been experimentally confirmed for at least two species: in poplar, colonization by G. mosseae increased root death up to fivefold (Hooker et al., 1995) while in the herbaceous species A. porrum, mycorrhization delayed senescence of cortical cells (Lingua et al., 1999). In woody species, in which effective AM are restricted to fine roots, enhanced root turnover would result in a higher proportion of fine roots that are more suitable for mycorrhization. In contrast, in A. porrum, most roots are functional as long as the whole plant survives and the delayed senescence observed upon mycorrhization would consequently benefit the plant by increasing the period during which each root is functional (Berta et al., 2002). In conclusion, different species can respond in very different (even opposite) ways to AMF colonization. This variability probably reflects the diversity of strategies used by plants to improve mycorrhization (Berta et al., 2002), but also the plasticity of the root system that is able to adapt to changing environmental conditions. For instance, different topologies might be adopted (even by the same plant) to improve mycorrhization: a branched root system could be very useful at the start to increase the chances of infection while later the development of a herringbone-like root system would be advantageous because the extraradical hyphae, being very efficient in nutrient uptake could play the role of branched lateral roots. Root survival and branching depend on the activity of meristems. In A. porrum, AM was found to slow down the mitotic cycle of root apical meristems, correlated with increased root apex size, a larger meristem and quiescent center (Fusconi et al., 1994). A longer cell cycle was also observed upon increased Pi availability, but without complete blockage of the cell cycle for AMF (Fusconi et al., 2000). In P. cerasifera, mycorrhized roots showed an increased percentage of cells in metaphase, also suggesting blockage of the cell cycle (Berta et al., 1995). However, some species appear to respond in a

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different way: for instance colonization of S. esculentum by G. mossae was found to increase mitotic activity in the apex (Fusconi et al., 1999). Although currently few data are available on the mechanisms that mediate the morphological changes observed upon mycorrhization, several studies have concluded that at least some of the observed morphological changes are due to increased nutrient supply (Hetrick et al., 1988; Hooker et al., 1992; Berta et al., 1995; Fusconi et al., 2000). Most probably the same signaling pathways are involved in the nutrient-dependent morphological changes in Arabidopsis and other species (see Chapter 11). For instance, both Pi supply and mycorrhization are able to rescue the lrt1 mutant that lacks lateral roots when grown without mycorrhizae at low Pi, indicating that similar mechanisms are involved in the formation of lateral roots in both cases (Paszkowski and Boller, 2002). The involvement of plant hormones in developmental changes induced by AMF, in particular, auxins and cytokinins, was suspected early on (Atkinson et al., 1994). The first measurements of indole-3-acetic acid (IAA) concentrations in maize revealed that colonized and noncolonized roots did not differ ¨ (Ludwig-Muller et al., 1997), and similar results were obtained in tobacco and leek (Torelli et al., 2000; Shaul-Keinan et al., 2002). Nevertheless, the levels and synthesis of another auxin, indole-3-butyric acid (IBA), were found to increase ¨ ¨ (Ludwig-Muller et al., 1997; Kaldorf and Ludwig-Muller, 2000). In maize, levels of free IBA, IAA- and IBA-amide conjugates increased upon mycorrhization, whereas free IAA, IAA- and IBA-esters were unchanged (Fitze et al., 2005). However, the role of auxins remains unclear. Concerning cytokinins it was shown that germinated spores of G. mosseae produce gibberellin- and cytokinin-like compounds (Barea and Azc´an-Aguilar, 1982) and that levels of cytokinins increase in mycorrhizal plants (Baas and Kuiper, 1989; van Rhijn et al., 1997; Torelli et al., 2000). Cytokinins might play a role similar to that in Arabidopsis in which they have been shown to be key regulators of lateral root development (Werner et al., 2003; To et al., 2004; Riefler et al., 2006). Another plant hormone seems to have an important role is jasmonic acid (JA). Both JA application and AMF colonization increased root branching in garlic, and JA application promoted the development of arbuscules and vesicles (Regvar et al., 1996). In barley (Hordeum vulgarum), colonization by G. intraradices resulted in elevated levels of JA and JA–isoleucine conjugates (Hause et al., 2002). In M. truncatula, low JA levels resulting from a JA biosynthesis gene knockdown correlated with delayed colonization and reduced arbuscule formation, suggesting an important role of JA in mycorrhiza development (Isayenkov et al., 2005). Still, the mode of action of JA remains to be elucidated. Recently, Ol´ah et al. (2005) have found that AMF produce a diffusible factor that is able to induce branching on M. truncatula roots. In contrast to the morphological changes induced by auxin, this factor induces branching without a reduction in growth suggesting that this factor is not an auxin. The observed lateral root formation requires the symbiotic genes DMI1 and

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Root Development DMI2 but not DMI3 and NFR. The plant ‘branching factor’ produced by the fungus was suggested to be perceived by a receptor which differs from the Nod factor receptor, leading to the steps in which DMI1/DMI2 are involved and to subsequent divergence of the pathway after DMI2. Over the past decade, increasing evidence has emerged favoring links between molecular mechanisms controlling lateral root formation, mycorrhization and nodulation. For instance, in L. japonicus the LjSym78 mutant possesses a hypernodulating phenotype as well as enhanced mycorrhization and root branching (Solaiman et al., 2000; Kawaguchi et al., 2002). LjSym78 is allelic to hypernodulation aberrant root 1 (har1), a gene encoding an LRR receptor-like kinase (Kinkema et al., 2006). Furthermore, a recent screen for suppressors of hypernodulation in L. japonicus har1-1 led to the identification of mutants with altered interactions with AMF (Murray et al., 2006). Recently, Bright et al. (2005) identified the lateral root organ-defective (latd) mutant in M. truncatula affected in both lateral root formation and nodulation, again suggesting common mechanisms for lateral root formation and nodulation (see Chapter 10). The percentage of mycorrhization in latd was comparable to that in wild-type plants, but further investigation is needed to determine whether latd also affects root branching induced by mycorrhization factors as described by Ol´ah et al. (2005).

9.3

Ectomycorrhizae

Ectomycorrhizae (EC) are symbiotic associations between 3 and 5% of the seed plants (angiosperms and gymnosperms) and more than 5000 fungal species belonging mostly to the Basidiomycetes and Ascomycetes. EC symbiosis evolved recently with the oldest evidence of EC found in 50-millionyear-old fossils recovered in British Columbia (LePage et al., 1997). In spite of their limited number, ectomycorrhizal plants are of great ecological and economic importance (timber, edible fungi such as truffles, etc.). Like AM symbioses, EC increase water and nutrient acquisition of the plant partner in exchange for photosynthetates (10–20% of the plant’s assimilates; Smith and Read, 1997). The Pi inflow in EC and AM roots is of the same order of magnitude (Smith and Read, 1997). Three distinct structures define EC: the extraradical mycelium, the sheath and the Hartig net, and, of course, the absence of intracellular penetration of plant cells by the fungus. In the case of intracellular penetration by hyphae from the Hartig net or the sheath, the structure is called an ectendomycorrhiza (Smith and Read, 1997). Diverse forms of EC appear to be controlled by the plant partner, as shown by the fact that the same fungus can make different types of EC in different plants. Some plants are exclusively ectomycorrhizal while others can also enter endomycorrhizal symbiosis. Some EC fungi have a broad host range, while others only interact with a limited number of host plant species.

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EC formation has mostly been studied with simplified laboratory models, involving seedlings in sterile or semi-sterile cultures and a single selected fungus. Until now, progress has been slow in this field mainly because of the nature of the plant partner (woody perennials), but the recent completion of the sequencing of the poplar genome (Tuskan et al., 2006) should speed things up. 9.3.1

Preinfection events

Little is known about the molecular dialogue between the two symbiotic partners forming EC. Data suggest that specific chemicals secreted by the plant root are perceived by and trigger changes in compatible ectomycorrhizal fungi, such as spore germination, increased hyphal growth and branching, and chemotropic growth. For instance, Horan and Chilvers (1990) showed that specific root exudates attracted compatible strains of Paxillus involutus or Pisolithus tinctorius but had no effect on incompatible strains. As for other root symbioses, secreted root phenylpropanoids are likely to play a role. Phenylpropanoids accumulate during larch mycorrhization (Weiss et al., 1997) whereas rutin, a phenolic compound produced by eucalyptus roots, stimulates Pisolithus tinctorius hyphal growth at picomolar concentrations (Lagrange et al., 2001). Finally, some plant hormones, such as cytokinins, have been shown to change hyphal growth and branching (Gogala, 1991), even if they are unlikely to serve as specific signals. However, the molecular mechanisms of perception of these specific signals are still unknown. On the other hand, the fungal partner produces some compounds that are perceived by and have phenotypic effects on the host plant. Contact with eucalyptus roots increases the production of an indolic compound, hypaphorine, in P. tinctorius, that inhibits root hair elongation (B´eguiristain and Lapeyrie, 1997) and that could act as an antagonist of auxin (for review, see Jambois et al., 2005). Nevertheless, hypaphorine is not a signal common to all EC fungi and its symbiotic role remains to be unravelled. In summary, diffusible signals are exchanged during the first stage of EC formation (Fig. 9.2a) but the nature of the symbiotic signals and their mechanisms of perception are still unidentified. Even if the establishment of the symbiosis leads to changes in gene expression in both partners, no EC-specific genes have been identified so far (for review, see Wiemken and Boller, 2002; Duplessis et al., 2005). The absence of molecular markers of the symbiosis impedes further progress in the identification of the various symbiotic signals. 9.3.2

Root colonization

Colonization occurs mostly in the elongation zone, just above the root cap of first-order lateral roots whose formation is increased by auxin produced by fungal hyphae. The entire root surface is compatible for fungal adhesion. Compatible strains attach to the root surface and fungal cells enlarge and

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(b)

(c)

Figure 9.2 Different stages of Ectomycorrhizal symbiosis. (a) The symbiotic interaction starts with the exchange of signals between the two partners. The chemical nature of these signals is still unknown. (b) Fungal hyphae attach to the root surface and start to swell and branch until they completely colonize the root surface. Hyphae penetrate between epidermal cells to form a dense intercellular hyphal network, the Hartig net. (c) Mature ectomycorrhizae show three typical structures: the intercellular Hartig net (1) involved in the exchanges of nutrients between the two partners, a dense mantle (2) that covers the root surface and (3) an extraradical mycelium that explores the soil.

branch (Fig. 9.2b). Attachment is associated with changes in the properties of the fungal cell wall and production of adhesin-like polypeptides and hydrophobins (Tagu and Martin, 1996). Binding sites for Lactarius deterrimus cell wall lectins were detected on the root surface of spruce, its host plant (Giollant et al., 1993), suggesting that lectin–carbohydrate recognition could also be involved in attachment of compatible strains to the plant root. In contrast, incompatible strains induce defense processes, including the deposition of phenolic material at the point of contact (Malajczuk et al., 1984; Horan et al., 1988). The mechanisms that enable the discrimination of compatible and incompatible strains remain unknown. 9.3.3

Morphogenesis

Hyphae completely colonize the growing root surface and form a dense mantle. The fungus then penetrates between epidermal cells just behind the

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Figure 9.3 Examples of Arbuscular and ectomycorrhizal roots. (a–c) Ectomycorrhizae; (d and e) Arbuscular mycorrhizae. (a) Allocasuarina verticillata roots colonized by Pisolitus tinctorius. (b and c) Transversal sections of a Leptolaena multiflora root colonized by an unknown EC fungus. (d and e) Petunia hybrida colonized by Glomus mossae. HN, Hartig network; M, mantle; H, hyphophodium; Arb, arbuscule; Ch, coiled hyphae. (Pictures (b and c) and (d and e) were kindly provided by Dr. R. Duponnois (IRD, Dakar, S´en´egal) and Dr. S. Wegm¨uller (ETH, Zurich, Switzerland).)

root cap to form a dense network of intercellular hyphae, called the Hartig net (Figs. 9.2c, 9.3b and 9.3c) that leads to swelling of root tip. The Hartig net is the physiological equivalent of the arbuscule involved in the exchange of nutrients with the plant cell. In some cases, ingrowths have been observed in the plant cell walls presumably increasing the exchange surfaces. In the majority of angiosperms, Hartig net penetration is limited to the epidermis. In this case, infection induces radial expansion of epidermal cells, leading to increased surface contact. In ectomycorrhizal gymnosperms and a few genera of angiosperms, the Hartig net also extends to several layers of cortical cells and no radial enlargement of epidermal cells is observed. In both cases, root hair development is abolished and meristem activity reduced. Ramification occurs close to the EC root tip. Newly formed lateral roots are colonized as they emerge from the parent root. This gives rise to short, thick and ramified EC roots (Fig. 9.3a).

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Root Development The role of auxin in EC development is still unclear (for review, see Barker and Tagu, 2000). The phenotype of short ramified EC roots suggests that auxin could be a key factor in EC development. EC fungi are known to produce auxins in vitro (Ek et al., 1983; Frankenberger and Poth, 1987) and to increase lateral root formation in inoculated plants (Karabaghli-Degron ´ et al., 2003). However, there are conet al., 1998; Niemi et al., 2002b; Rincon tradictory reports on the effect of EC formation on IAA concentration in the roots. Surprisingly, an auxin-induced GH3 homolog from Pinus pinaster was downregulated following inoculation with compatible, but not after inoculation with incompatible EC fungi (Reddy et al., 2006). An IAA-overproducing mutant strain of the EC fungus Hebeloma cylindrosporum was used to study the role of auxin during EC formation in Pinus pinaster (Gay et al., 1994). The mutant strain produced threefold to sixfold more colonized roots than the wild-type fungal strain and the Hartig net was more developed and thicker. Moreover, whereas the Hartig net of wild-type strains was restricted to the outer half of the cortex, that of the auxin overproducing strain extended to the endodermis (Tranvan et al., 2000). This increased infection did not lead to pathogenesis or to increased plant growth, suggesting that auxin is involved in the infection process and in the formation of the Hartig net. This hypothesis is supported by the inhibition of EC development by auxin transport inhibitors, such as naphthylphthalamic acid (NPA) or tri-iodobenzoic ´ acid (TIBA) (Karabaghli-Degron et al., 1998; Niemi et al., 2002b; Rincon et al., 2003). On the other hand, the EC phenotype of aspen trees overexpressing Agrobacterium IAA-biosynthetic genes resembled that of wild-type plants (Hampp et al., 1996). In conclusion, clear experimental data point to a role for auxin in EC development and Hartig net formation. However, the precise effect of fungal colonization on auxin homeostasis (synthesis, transport, degradation and conjugation) in infected roots remains to be analyzed. Polyamines (PAs) also seem to be involved in EC formation. Exogenous PAs can promote EC formation in vitro (Niemi et al., 2002a). Fungal ornithine decarboxylase (ODC) transcripts are expressed in the developing hyphal mantle and Hartig net (Morel et al., 2005; Niemi et al., 2006). ODC is a key enzyme in PA biosynthesis. Different EC fungi have been shown to produce PAs, such as putrescin, in vitro and the concentration of PA increases in the roots of EC seedlings (Niemi et al., 2006). Taken together these data point to a positive role of PAs during the establishment of the EC symbiosis. 9.3.4

Effect of ectomycorrhizal symbioses on root system architecture

Very few studies address the effects of EC symbiosis on root system architecture. Plants bearing EC show increased plant growth with both root system and aerial parts of the plants being more developed than in noninoculated control plants (reviewed in Smith and Read, 1997). Frequently, there is an initial phase of EC formation where growth is similar or even slower in

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inoculated plants than in control plants due to the carbon resources consumed by the fungal partner and the establishment of the symbiosis. A similar phenomenon has been described for AM. Concerning the effects of EC on the root:shoot ratio, EC formation seems to reduce the ratio in most cases. Nevertheless, the interaction with EC fungi clearly changes root architecture with an increase in number of short roots (Alexander, 1981) and the appearance of short, thick, ramified EC. EC fungi also increase lateral root formation in inoculated plants (Karabaghli-Degron et al., 1998; Niemi et al., ´ et al., 2003). 2002b; Rincon

9.4 Actinorhizal symbioses Actinorhizal symbioses result from the interaction between a soil actinomycete, Frankia and plants distributed in 8 angiosperm families and 24 genera collectively called actinorhizal plants including Betulaceae, Casuarinaceae, Elaeagnaceae and Myricaceae, mostly trees or shrubs that can grow in poor and damaged soils under a range of environmental stresses, such as high salinity, the presence of heavy metal and extreme pH (Dawson, 1990). As a consequence, actinorhizal plants are important elements in plant communities worldwide, in particular several species of Casuarinaceae that are used in agroforestry, land reclamation and fuelwood production (Diem and Dommergues, 1990). Frankia is a filamentous Gram-positive bacterium (reviewed by Benson and Silvester, 1993) that by interacting with actinorhizal plants leads to the formation of a new root organ called the actinorhizal nodule, in which the bacterium is hosted and provides reduced nitrogen to the plant. While the use of two model legume species, Medicago truncatula and Lotus japonicus, has accelerated the genetic/molecular characterization of the Rhizobium–legume symbiosis (see Chapter 10), not a single model actinorhizal plant has emerged so far. All actinorhizal species, except one (Datisca glomerata), are woody perennials and are therefore not amenable to genetics. Moreover, since it is not possible to transform Frankia, genetic analysis of the bacterial symbiotic genes is very limited. For all these reasons, our knowledge of actinorhizal nodule formation is still in its infancy. Plant genes specifically expressed in response to Frankia infection, defined as actinorhizal nodulin genes (Mullin and Dobritsa, 1996) by homology with legume nodulin genes, have been identified in several actinorhizal species. Early nodulin genes are expressed before nitrogen fixation occurs and late nodulin genes are involved in the functioning of the nodule. The recent sequencing of three Frankia genomes (Normand et al., 2007), together with the development of RNAi strategies for actinorhizal plants, such as Allocasuarina verticillata (unpublished data) open new avenues to identify plant and bacterial genes involved in the symbiotic interaction.

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Preinfection events

Actinorhizal nodule development only occurs under conditions of nitrogen deprivation and plant roots are assumed to emit signals of unknown nature that are perceived by Frankia. Root phenylpropanoids are prime candidates for these signaling molecules. For instance, the nodulation of Alnus sp. is enhanced by the addition of flavonone (Benoit and Berry, 1997) or flavonol compounds (Hughes et al., 1999). In return, Frankia produces symbiotic signals that are perceived by the plant root (Fig. 9.4a). In intracellularly infected plants, this leads to root hair

(a)

(e) (b)

(d) (c)

Figure 9.4 Formation of actinorhizal symbioses. (a) Signal exchanges between the actinorhizal plant and Frankia leads to root hair deformation. (b) Frankia penetrates a deformed root hair triggering cortical cell divisions. (c) Dividing cortical cells are infected by Frankia hyphae and hypertrophy thus leading to the formation of the prenodule. At the same time, pericycle cell divisions occur in front of a xylem pole to form a nodule primordium. (d) Frankia hyphae coming from the prenodule invade the cortex of the nodule primordium. (e) In the mature nodule, four zones are observed: the meristematic zone of the nodule which is always free of Frankia, the infection zone containing newly formed cortical cells that are progressively infected and hypertrophy, the fixation zone where infected cells fix atmospheric nitrogen and a senescence zone where Frankia and plant cells degenerate (only in old nodules). The nodule presents a root-like structure with a central vasculature.

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deformation. The chemical nature of the actinorhizal factors is still unknown, although data suggest that they are biochemically different from Rhizobium Nod factors (see Chapter 10). Nod factor application has no effect on Alnus root hairs (C´er´emonie et al., 1999), but a bacterial fraction containing a root hair deforming factor has been characterized (C´er´emonie et al., 1999) that had biochemical properties that differed from those of Nod factors. Moreover, because Rhizobium nod mutants cannot be complemented with Frankia DNA, different genes might be involved in the production of Frankia actinorhizal factors (C´er´emonie et al., 1998). Identification of a Frankia actinorhizal factor(s) is hampered by the lack of a genetic analysis system for Frankia. The recent sequencing of the Frankia genome will hopefully shed new light on the chemical nature of the actinorhizal factors. Unfortunately, no plant gene specifically induced in response to a diffusible Frankia symbiotic signal has been identified so far. Thus, a sensitive and reproducible biotest is needed that could be used to purify Frankia actinorhizal factor(s). Frankia also produces phytohormones that could play a role in the symbiotic interaction. Natural auxins, such as IAA or phenylacetic acid (PAA) can be found in Frankia cultures at relatively high concentrations (10−5 to 10−6 M; Wheeler et al., 1979; Hammad et al., 2003). The cytokinin isopentenyl adenosine (iPA) has also been detected at a concentration of 10−6 M (Gordons et al., 1988). However, the role of these hormones in the nodulation process is still unknown. 9.4.2

Infection process

There are two modes of infection of actinorhizal plants by Frankia: intracellularly (also called root hair infection; Fig. 9.4b) and intercellularly (Berry and Sunnel, 1990). The mode of infection is under the control of the host plant. Intracellular infection (e.g. Myrica, Comptonia, Alnus and Casuarina) starts with root hair deformation 24–48 hours after inoculation. Depending on the species, either all (e.g. Casuarina; Torrey, 1976) or only a few root hairs (e.g. Comptonia; Callaham et al., 1979) deform in response to inoculation. Only growing root hairs that are not fully differentiated are competent for infection (Callaham and Torrey, 1977; Callaham et al., 1979). Frankia hyphae are trapped by plant cell polysaccharides at the tip of some deforming root hairs that initiate an intracellular infection thread in a folded part of the plant cell (Berry et al., 1986; Berg, 1999a). Local disorganization of the primary cell wall is observed at the site of Frankia penetration (Berry et al., 1986). During infection, hyphae are always surrounded by the plant cell membrane and plant cell wall-derived material. Cg12, a Casuarina glauca gene encoding a subtilisin-like protease, is specifically expressed during intracellular infection of plant cells including root hairs (Laplaze et al., 2000b; Svistoonoff et al., 2003). Cg12 is secreted and could be involved in cell wall remodeling during Frankia infection (Svistoonoff et al., 2003). Infected root hairs display a high metabolic activity while noninfected hairs start to degenerate (Berry et al., 1986; Berry and

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Root Development Sunnel, 1990). Root hair infection triggers a limited number of cortical cell divisions close to the infection site, leading to the formation of a protuberance on the root, called the prenodule (Fig. 9.4c) (Callaham and Torrey, 1977). The prenodule is an obligatory step in intracellular infection, but is not involved in nodule formation (Laplaze et al., 2000a). Frankia progresses intracellularly from the infected root hair to prenodule cells through cytoplasmic bridges, called preinfection threads (Berg, 1999b) that are induced before Frankia infection by an unknown signal. Infected prenodule cells fill with Frankia and subsequently hypertrophy. In the prenodule, Frankia and infected plant cells differentiate to fix nitrogen (Angulo Carmona, 1974; Laplaze et al., 2000a). Intercellular infection (e.g. Eleagnus, Ceanothus and Cercocarpus) begins with Frankia entering the root tissue between epidermal cells. This mode of infection is not associated with root hair deformation. Dense intercellular material is secreted by plant cells at the site of infection (Miller and Baker; 1985). Frankia progresses through the middle lamella in the cortical tissues. 9.4.3

Nodule formation

In both the intracellular and intercellular infection modes, Frankia infection induces cell divisions in the pericycle in front of a xylem pole that give rise to a nodule lobe primordium (Fig. 9.4d). In Comptonia, some cortical cells divide and participate in the formation of the nodule lobe primordium (Callaham and Torrey, 1977). Each nodule lobe presents a meristem at its apex, has a central vascular bundle, and is surrounded by a periderm. Frankia hyphae coming either from the prenodule (intracellular infection) or progressing intercellularly (intercellular infection) invade some cortical cells at the base of the young nodule lobe. During intercellular infection, bacterial hyphae become intracellular when they invade the young nodule primordium. Frankia hyphae then start to grow toward the apical meristem creating a gradient of development of both Frankia and infected plant cells (Fig. 9.4e). Four zones are commonly described: (1) a meristematic zone at the apex that is responsible for the indeterminate growth of the nodule and is always devoid of Frankia; (2) an infection zone adjacent to the apical meristem where Frankia hyphae infect some newly produced cortical cells in which the bacterium starts to proliferate; (3) a fixation zone that contains both infected and uninfected cells, the first being hypertrophied and full of Frankia hyphae that differentiate to fix nitrogen; and (4) a senescence zone found at the base of old nodules in which both the host cells and Frankia degenerate and Frankia sporangia can be seen. In some species, such as Casuarina or Myrica, a so-called ‘nodule root’ devoid of Frankia infection is found at the apex of each nodule lobe. This modified root lacks root hairs and shows negative geotropism (i.e. grows toward the surface). The presence of a developed aerenchyma suggests that nodule roots facilitate gas exchanges (Silvester et al., 1990). In this case, the nodule lobe meristem changes fate at some stage and starts to form a

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nodule root. Nothing is known about the mechanisms leading to this switch in behavior of the nodule lobe meristem. Growth of the nodule is maintained by ramification of the existing nodule lobes. Cell divisions in the nodule lobe pericycle lead to the formation of new lobes that become infected by Frankia hyphae coming from the nodule giving rise to a coraloid structure. Because they have the same origin as lateral roots (cell divisions in the pericycle in front of xylem poles) and a root-like structure, actinorhizal nodule lobes have been considered as modified lateral roots (Pawlowski and Bisseling, 1996). The formation of an apical nodule root at the apex of some actinorhizal nodule lobes is consistent with this theory. However, a careful study in Alnus glutinosa suggests that nodules do not form from preexisting lateral root primordia (Angulo Carmona, 1974). Moreover, in some species, such as Comptonia, cortical cells are involved in the formation of the nodule lobe primordium while lateral root initiation is restricted to the pericycle (Callaham and Torrey, 1977). Consequently, even if actinorhizal nodules have root-like features, it is not clear to what extent nodule and lateral roots share common developmental pathways. Unfortunately, few studies have addressed this important issue. The Nicotiana tabacum gene HRGPnt3 encodes a plant cell-wall protein expressed at early stages of lateral root development (Keller and Lamb, 1989) and is therefore a good marker for lateral root initiation. In our laboratory, we introduced the HRGPnt3 promoter fused to the ␤-glucuronidase gene into transgenic Allocasuarina verticillata plants. Unfortunately, no HRGPnt3 was expressed during lateral root or nodule development in A. verticillata, suggesting that tissue-specific expression of this marker is not conserved in a heterologous environment (unpublished results). Similarly, despite its central role during lateral root development (Casimiro et al., 2003), little is known about the role of auxins during actinorhizal nodule development. Application of auxin efflux transport inhibitors, such as 1- NPA or TIBA, on C. glauca roots leads to the formation of nodule-like structures, called ‘pseudonodules’ (Duhoux et al., 1996). On the other hand, inhibition of auxin influx using 1-naphtoxyacetic acid (1-NOA) delays nodule formation in C. glauca (P´eret et al., 2007). Moreover, CgAUX1, a C. glauca gene encoding a functional auxin influx carrier, is expressed in plant cells infected by Frankia throughout the course of actinorhizal nodule formation (P´eret et al., 2007). This together with the higher auxin content of nodulated roots compared to noninoculated roots and auxin production by Frankia (Wheeler et al., 1979; Hammad et al., 2003) suggests that auxin might play an important role during plant cell infection (P´eret et al., 2008). Unfortunately, molecular markers of auxin response, such as soybean GH3 gene or the synthetic DR5 promoter (Ulmasov et al., 1995) are not functional in transgenic actinorhizal plants of the Casuarinaceae family (unpublished results). Interestingly, CgAUX1 is expressed in lateral root primordia but not in actinorhizal nodule primordia (P´eret et al., 2007). This suggests that these two organs have, at least in part, different developmental programs. Clearly, further work is needed to

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Root Development understand how actinorhizal nodules develop and to what extent this developmental program is derived from lateral root development. 9.4.4

Effect of nodulation on root system architecture

Very little is known about the effect of actinorhizal symbioses on root architecture. Plants nodulated by Frankia grow better than noninoculated plants with increased growth of the root system. Since actinorhizal nodules are developmentally related to lateral roots, it was questioned whether their development might change root architecture. A detailed study in Alnus glutinosa showed that actinorhizal nodule formation has no impact on the number or the distribution of lateral roots (Angulo Carmona, 1974). Actinorhizal nodules are therefore additional lateral organs and do not form from preexisting lateral root primordia (Angulo Carmona, 1974). Moreover, the production of lateral roots is not increased as a result of Frankia infection, suggesting that the formation of these two types of organs is regulated independently. In conclusion, actinorhizal symbioses lead to the formation of new supplementary lateral organs on the root system without impact on its overall architecture.

9.5

Concluding remarks

Under natural conditions, plant roots are always associated with a diverse population of microorganisms that can culminate in intimate, mutualistic symbiotic associations. Actinorhizal and mycorrhizal symbioses together with the Rhizobium–legume associations are important adaptative responses allowing the root system to improve acquisition of nutrients and water. For instance, under natural conditions, most plant species are associated with mycorrhizal fungi and such mycorrhizal symbioses are therefore the main adaptative strategy for these plants to cope with poor nutrient (phosphorus) availability. The importance of root hair development or root architecture responses to nutrient deprivation would be mainly limited to those plants that do not form this symbiotic association (such as Brassicaceae or Proteaceae) or to soils deprived of mycorrhizal fungi. These different symbioses thus play an important ecological role by enabling plant growth on poor soils and stabilizing plant communities. Comparison of these different symbiotic systems reveals some common themes. First, the establishment of all these symbiotic interactions is controlled by the physiological status of the plant. Since plants have to invest a large proportion of their photosynthetate in establishing and maintaining these associations, they only enter symbioses when key nutrients (nitrogen or phosphorus) are limiting. Root exudates and particularly phenylpropanoids are key signals for symbiotic microorganisms to recognize their host plant and to change their development accordingly. This might explain why the secretion of such molecules, which are also signals for root pathogens or

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parasitic weeds, has been conserved during evolution. These root-derived organic compounds have a large impact on the rhizospheric microbial community in general. Our understanding of the molecular mechanisms of symbiotic recognition in actinorhizal and mycorrhizal symbiosis is still in its infancy. However, data suggest that part of the signaling pathway involved in the recognition of different symbiotic microorganisms by their plant host is conserved. Recently, we have isolated a DMI2/SymRK homolog from the actinorhizal tree C. glauca. Previous work in legumes showed that this gene is involved in both Rhizobium and AMF recognition (Endre et al., 2002; Stracke et al., 2002). Ongoing studies suggest that CgSymRK is also involved in the signaling pathway leading to actinorhizal symbioses (unpublished results), consistent with the recycling of molecular mechanisms from the ancient AM symbiosis. However, even if the central core of the signal transduction cascade is conserved, there are differences in signal perception and of course also in the downstream signal for each symbiosis. So far, only Nod factors and their putative receptors have been identified. Identification of the Myc and actinorhizal factors and the corresponding perception mechanisms will allow insights into the evolution of these different symbiotic signaling pathways. Interestingly, some pathogenic organisms are able to mimic symbiotic signaling in order to infect some plants. For example, Penicillium nodositatum can form actinorhizal nodule-like structures, called myconodules, in some Alnus species (Capellano et al., 1987). The infection process is identical to that of Frankia but myconodules remain small and unilobed and their morphology resembles that of incompatible Frankia nodules (Sequerra et al., 1994). The effect of symbiotic associations on root architecture has been poorly studied, although it is clear that they have an impact on the development of the root system. Some of these effects are due to changes in the acquisition of nutrients by the plant. The association itself can occur on an existing root with either limited (arbuscular mycorrhizae) or major (ectomycorrhizae) developmental effects or lead to the formation of a new symbiotic organ that can derive from a lateral root developmental program (actinorhizae or legume nodules: see also Chapter 10).

Acknowledgments The authors thank Dr. R. Duponnois (IRD, Dakar, S´en´egal) and Dr. S. ¨ Wegmuller for providing pictures. We are also grateful to Dr. D. Barker, Dr. D. Bogusz, Dr. P. Doumas, Dr. E. Duhoux, Dr. R. Duponnois and Dr. S. ¨ Wegmuller for comments on the manuscript. Research in our laboratory is supported by the Institut de Recherche pour le D´eveloppement (IRD), the Agence Nationale pour la Recherche (ANR; Grant No. 06-3-134048) and Genoscope. B.P. is supported by a grant from the Minist`ere de la Recherche et de la Technologie.

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cortical cells during Casuarina glauca and Allocasuarina verticillata actinorhizal nodule development. Mol. Plant Microbe Interact. 16, 600–607. Tagu, D. and Martin, F. (1996). Molecular analysis of cell wall proteins expressed during the early steps of ectomycorrhiza development. New Phytol. 133, 73–85. Tamasloukht, M., S´ejalon-Delmas, N., Kluever, A., Jauneau, A., Roux, C., B´ecard, G. et al. (2003). Root factors induce mitochondrial-related gene expression and fungal respiration during the developmental switch from asymbiosis to presymbiosis in the arbuscular mycorrhizal fungus Gigaspora rosea. Plant Physiol. 131, 1468–1478. Tisserant, B., Gianinazzi, S. and Gianinazzi-Pearson, V. (1996). Relationships between lateral root order, arbuscular mycorrhiza development, and the physiological state of the symbiotic fungus in Platanus acerifolia. Can. J. Bot. 74, 1947–1955. To, J.P.C., Haberer, G., Ferreira, F.J., Deru`ere, J., Mason, M.G., Schaller, G.E. et al. (2004). Type-A Arabidopsis response regulators are partially redundant negative regulators of cytokinin signaling. Plant Cell 16, 658–671. Torelli, A., Trotta, A., Acerbi, L., Arcidiacono, G., Berta, G. and Branca, C. (2000). IAA and ZR content in leek (Allium porrum L.), as influenced by P nutrition and arbuscular mycorrhizae, in relation to plant development. Plant Soil 226, 29–35. Torrey, J.G. (1976). Initiation and development of root nodules of Casuarina (Casuarinaceae). Am. J. Bot. 63, 335–344. Torrisi, V., Pattinson, G.S. and McGee, P.A. (1999). Localized elongation of roots of cotton follows establishment of arbuscular mycorrhizas. New Phytol. 142, 103– 112. Tranvan, H., Habricot, Y., Jeannette, E., Gay, G. and Sotta, B. (2000). Dynamics of symbiotic establishment between an IAA-overproducing mutant of the ectomycorrhizal fungus Hebeloma cylindrosporum and Pinus pinaster. Tree Physiol. 20, 123–129. Trappe, J.M. (1987). Phylogenetic and ecologic aspects of mycotrophy in the angiosperms from an evolutionary standpoint. In Ecophysiology of VA Mycorrhizal Plants, G.R. Safir (ed.). Boca Raton, CRC Press, pp. 5–25. Trotta, A., Carminati, C., Schellenbaum, L., Scannerini, S., Fusconi, A. and Berta, G. (1991). Correlation between root morphogenesis, VA mycorrhizal infection and phosphorus nutrition. In Plant Roots and Their Environment. Vol. 24: Developments in Agricultural and Managed Forest Ecology, B.L. McMichael and H. Persson (eds). Amsterdam, Elsevier, pp. 333–339. Tuskan, G.A., DiFazio, S.P., Jansson, S., Bohlmann, J., Grigoriev, I., Hellsten, U. et al. (2006). The genome of black cottonwood, Populus trichocarpa (Torr. and Gray ex Brayshaw). Science 313, 1596–1604. Ulmasov, T., Liu, Z.-B., Hagen, G. and Guilfoyle, T.J. (1995). Composite structure of auxin response elements. Plant Cell 7, 1611–1623. van Rhijn, P., Fang, Y., Galili, S., Shaul, O., Atzmon, N., Wininger, S. et al. (1997). Expression of early nodulin genes in alfalfa mycorrhizae indicates that signal transduction pathways used in forming arbuscular mycorrhizae and Rhizobium-induced nodules may be conserved. Proc. Natl. Acad. Sci. U. S. A. 94, 5467–5472. Vierheilig, H. (2004). Regulatory mechanisms during the plant–arbuscular mycorrhizal fungus interaction. Can. J. Bot. 82, 1166–1176. Vigo, C., Norman, J.R. and Hooker, J.E. (2000). Biocontrol of the pathogen Phytophthora parasitica by arbuscular mycorrhizal fungi is a consequence of effects on infection loci. Plant Pathol. 49, 509–514. Wang, B. and Qiu, Y.-L. (2006). Phylogenetic distribution and evolution of mycorrhizas in land plants. Mycorrhiza 16, 299–363.

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Annual Plant Reviews (2009) 37, 239–287 doi: 10.1002/9781444310023.ch10

www.interscience.wiley.com

Chapter 10

LEGUME ROOT ARCHITECTURE: A PECULIAR ROOT SYSTEM Silvina Gonzalez-Rizzo, Philippe Laporte, Martin Crespi and Florian Frugier Institut des Sciences du V´eg´etal (ISV), Centre National de la Recherche Scientifique, 91198 Gif-sur-Yvette, France

Abstract: The soil environmental conditions and symbiotic interactions are the major determinants of legume root architecture. Investigating cues that affect root developmental adaptations to the environment as well as understanding the mechanisms underlying the control of the root architecture are crucial to improve agronomical traits, notably in the legume family. Indeed, legumes are one of the most widespread crops, in terms of number and diversity of cultivated species. Legume roots can develop two types of secondary root organs: lateral roots and nitrogen-fixing nodules. Lateral root formation is common to all higher plants; however, nodules are present only on legume roots as a result of the symbiotic interaction with nitrogen-fixing soil bacteria, collectively known as rhizobia. The only nonlegume plants described to be able to interact with rhizobia to form nitrogen-fixing nodules are Parasponia spp. What makes the legume root system so peculiar? The aim of this chapter is to give an overview of the current knowledge of the development of secondary root organs in legumes. A comparative analysis of their structure and ontogeny will be presented, and the specific and common regulatory mechanisms involved will be described. Keywords: nitrogen-fixing nodule; lateral root; Rhizobium; symbiosis; abiotic stress; phytohormones

239

240
Root Development

10.1

10.1.1

Comparison of legume lateral roots and nitrogen-fixing nodules Lateral root development in legumes

In general, legume roots are composed by a single layer of epidermal cells at the surface, with no specific root hair patterning, followed by three to five irregularly shaped cortical cell layers, usually defined as outer, middle and inner cortex (Gage, 2004) (Fig. 10.1) and a single layer of endodermal cells that encircles a pericycle layer. Within the pericycle the stele region encloses the vascular bundles.

Figure 10.1 Lateral root (LR) formation in legume roots. (1) The initial periclinal divisions in the pericycle (P). (2) The following periclinal and anticlinal divisions give rise to the LR primordium, with contribution of endodermis (En) and cortex layers (C). (3) After emergence, distinct tissues start to differentiate in the LR, following the parental root pattern. E, epidermis.

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Two types of lateral root (LR) development have been distinguished in flowering plants (Berthon, 1943): one in which LR primordia arise only from pericycle cells (such as Arabidopsis thaliana; see Chapter 4) and a second in which pericycle, endodermis and cortex layers from the parental root contribute to LR primordia formation. This latter type corresponds to LRs in legumes (Fig. 10.1). The pericycle, however, is the main layer involved in LR initiation and first divisions take place opposite to a protoxylem pole (Torrey, 1986). Thereafter, an LR primordium is formed with simultaneous divisions of endodermal and cortical cells (von Guttenberg, 1968). A cavity is then formed in the cortex adjacent to the elongating primordia, allowing LR emergence. At this stage, distinct tissues start to differentiate, in accordance with the parental root pattern (Peterson and Peterson, 1986; Malamy and Benfey, 1997; see Chapter 4). 10.1.2

Nitrogen-fixing nodule development in legumes

The rhizobia–legume interaction is a highly specific process that leads to the development of a new root-derived organ, the symbiotic nitrogen-fixing nodule. First, rhizobia colonize the root surface, attach to root hairs and induce their deformation and curling (Callaham and Torrey, 1981). The rhizobial invasion process occurs preferentially in the zone of emerging root hairs (Bhuvaneswari et al., 1980). Rhizobia induce a series of rapid changes in root hair cells, such a calcium spiking, pH variation and depolarization of the plasma membrane (Oldroyd and Downie, 2004). Recently, a root-hair-independent mode of rhizobial infection has been described based on the identification of a Lotus (Lotus japonicus) mutant defective in root hair formation, root hairless 1 (rhl1) (Karas et al., 2005). This mutant was identified as a suppressor of the hypernodulation and aberrant root 1 (har1) hypernodulation phenotype (see Section 10.3.4), indicating that root hairs are not an absolute requirement for the initiation of nodule organogenesis and rhizobial infection. Concomitantly to rhizobial infection, the pericycle is generally the first cell layer to be stimulated for division, which then proceeds in the cortex layer usually in front of a protoxylem pole close to the infection point (Timmers et al., 1999). In alfalfa (Medicago sativa), pericycle and cortical cells are activated and divide simultaneously (Thornton and Rudorf, 1936), while pericycle division has not been described in pea (Pisum sativum) (Bond, 1948). Despite these differences, actively dividing cortical cells form most of the nodule primordium in all cases, wherein large amounts of amyloplasts accumulate (Mylona et al., 1995; Bauer et al., 1996). At the root surface, rhizobia locally degrade the plant cell wall and penetrate the root hair cell by means of an infection thread. Infection threads are plant-derived structures originating from plasma membrane invaginations, followed by external deposition of cell wall material (van Spronsen et al., 1994). Infection threads progress intracellularly through the outer cortex, ramify, and finally reach the nodule primordium cells. In most cases, however, progression of infection threads aborts. Only very few infection

242
Root Development threads reach the nodule primordium cells (Vasse et al., 1993) and finally penetrate into plant cell walls, initiating a differentiation process that is heralded by cell enlargement. Bacteria differentiate into specific nitrogen-fixing forms, called bacteroids, that are released from infection threads into the cytoplasm of the plant cells surrounded by a peribacteroid membrane to form the symbiosome compartment (Roth and Stacey, 1989; Mylona et al., 1995). In parallel to bacteroid differentiation, the nodule primordium develops into a mature nodule (Brewin, 1991). Two major types of legume nodules are distinguished: determinate and indeterminate (Fig. 10.2). In the first type, cell divisions are initiated in the outer cortex of the root and nodule meristematic activity is blocked early after initiation, yielding spherical nodules. In contrast, cell divisions of indeterminate nodule primordia arise in the inner cortex and a persistent apical meristem is formed to allow continuous growth of the nodule. These nodules are elongated and contain all developmental stages of plant and bacterial cells. A differentiation gradient is found from the distal (apical) meristem to the proximal region attached to the main root. Several zones have been defined (Vasse et al., 1990): the meristematic region or zone I; the invasion region (a) I

Meristem

II

Outer cortex

II–III

Endodermis Pericycle

III

IV

(b)

Vascular bundle Inner cortex

III

Root hair Endodermis Pericycle Phloem Xylem

Figure 10.2 Schematic representation of longitudinal sections through an indeterminate (a) or a determinate (b) legume nodule. Several regions have been defined, which coexist in the same organ in the case of the indeterminate nodule: a persistent apical meristem (I); a rhizobial invasion zone (II); an interzone (II and III); a nitrogen-fixing zone (III); and a zone of senescence in older nodules (IV). In the case of determinate nodulation, a single differentiation zone is present at each developmental stage.

Legume Root Architecture
243

or zone II, where bacteria are released from infection threads into plant cells; a region characterized by an accumulation of amyloplasts, the interzone II-III; a nitrogen-fixing region or zone III, where symbiosomes are differentiated, nitrogen is fixed by the bacteroids, and ammonium is assimilated by plant cells; and a senescing region, or zone IV, present in older nodules, where both bacteroids and plant cells degenerate. Peripheral cell layers are found, including a parenchyma, an outer and an inner cortex, an endodermis, as well as vascular bundles connecting the nodule to the parental root. The type of nodule formed depends on the host plant or in particular cases on environmental conditions: in Sesbania rostrata, determinate nodules develop under aquatic conditions while roots grown on aerated conditions ´ form indeterminate nodules (Fern´andez-Lopez et al., 1998). 10.1.3

Interactions between LR formation and rhizobial infection

Rhizobial infection via root hair curling (RHC) is the best-known entry mechanism and occurs in most members of the three subfamilies of the Leguminosae. However, other modes for rhizobial invasion exist, linked to the development of LRs (Fig. 10.3). Rhizobia can directly colonize the cortex via a crack in the epidermis layer (‘crack entry’) that originates from lateral or adventitious roots protrusion (James et al., 1992; Boogerd and Van Rossum, 1997; Goormachtig et al., 2004). This lateral root base (LRB) nodulation observed in tropical legumes is however highly dependent on environmental conditions: bacterial infection via LRB invasion only occurs under hydroponic conditions (Goormachtig et al., 2004). Moreover, a recent transcriptomic study comparing LRB and RHC nodulation in S. rostrata has highlighted that a large number of genes were specific for crack-entry epidermal invasion (Capoen et al., 2007). RHC invasion occurs generally in the zone of emerging root hairs (Bhuvaneswari et al., 1980); however, in white clover (Trifolium repens), rhizobia can also enter the root in the mature region at the site of LR emergence (Mathesius et al., 2000). Indeed, cortical cells activated during LR development recapitulate some of the early responses associated with nodule formation. These cortical cells can be ‘hijacked’ by the rhizobia to form a lateral root associated nodule (LRAN). The ability of rhizobia to induce nodules in mature regions of roots is however reduced. Hence, rhizobial infection modes linked to LR formation exist among different legume–rhizobia interactions, notably when root hair development is impaired. 10.1.4

Nodulation can be induced in the absence of rhizobia or by using invasion-defective rhizobial strains

In alfalfa plants, a phenotype of nodulation in the absence of Rhizobium (NAR) has been reported (Truchet et al., 1989). These spontaneous nodules

244
Root Development

Figure 10.3 Schematic representation of the diversity of infection modes in legume-rhizobia symbioses. (a, c): Root Hair Curling (RHC). This is the main rhizobial infection mode. Rhizobia enter shepherd’s crook root hairs and progress through an intracellular infection thread towards the nodule primordia. Cortical cell divisions occurs in outer cortex in the case of determinate nodulation (a) and in the inner cortex in the case of indeterminate nodulation (c). (b): Lateral Root Associated Nodulation. In some legumes, rhizobia can also enter the root in the mature region at the site of lateral root emergence. Dividing cortical cells associated to lateral root development are “hijacked” by rhizobia to form a nodule primordium. (d): “Empty nodules” formed following the interaction with exopolysaccharide (exo-) mutants. Infection threads are aborted in the cortex. LR: Lateral Root. VT: Vascular Tissue. NP: Nodule primordia.

possess all the histological features characteristic of indeterminate nodule: an apical meristem, peripheral vascular bundles connected to the plant vascular system, endodermis and cortex layers surrounding the nodule. Like nitrogen-fixing nodules, these structures are inhibited by the presence of combined nitrogen (see Section 10.3.5) and express early nodulin genes (e.g. MsEnod2 and MsEnod40; see Section 10.2.1). Similarly, spontaneous nodules have been recently identified in Lotus through a genetic screen (Tirichine

Legume Root Architecture
245

et al., 2006a). As in alfalfa, these spontaneous nodules formed (snf ) mutants show an ontogeny and histology characteristic of Mesorhizobium loti-induced nodules, including induction of early nodulin gene expression, and spontaneous nodule formation is inhibited by nitrate and ethylene. Studies with Rhizobium mutants impaired in the synthesis of surface components, such as certain acidic exopolysaccharides (exo− mutants), ␤-glucans (␤-glu− mutants), or lipopolysaccharides (lps− mutants), have revealed that these mutant bacteria cannot be released from infection threads or differentiate into bacteroids, and therefore lead to the formation of the so-called ‘empty’ non-nitrogen-fixing nodules (Truchet et al., 1980; Finan et al., 1985; Gray et al., 1992; Fraysse et al., 2003). Recently, the Sinorhizobium meliloti msbA2 (multicopy suppressor of htrB mutations A) gene encoding a putative ATPbinding cassette exporter has been characterized in a screening for symbiotic mutants (Griffitts and Long, 2008). MsbA2 might export a polysaccharide specifically produced during symbiotic interactions. In contrast to spontaneous nodules, these structures induced by defective rhizobial strains do not form a persistent meristem (Leigh et al., 1987). Altogether these data demonstrate that rhizobia can be dispensed for nodule initiation: plant genetic determinants seem sufficient to allow nodulation in the absence of rhizobia.

10.2

Recent advances in genetics and genomics of nitrogen-fixing nodule development in legumes

The organogenesis of legume nodules requires a precise spatiotemporal expression of specific genes during the different stages of the symbiotic interaction. Analysis of plant signaling pathways involved in the early stages of this developmental process have been carried out, mainly based on genetic approaches and high-throughput gene expression (transcriptomics) studies.

10.2.1

Signaling pathways involved in early stages of the symbiotic interaction

Plant and bacteria symbiotic partners exchange signals at all steps of the interaction. The plant roots exude flavonoids able to activate the expression of rhizobial nodulation (nod) genes. These genes lead to the synthesis of lipochitooligosaccharides signal molecules, designated Nod factors, which induce plant host responses corresponding to the early steps of nodule formation. Nod factors are strictly necessary for the infection process and subsequent nodule formation (Crespi and G´alvez, 2000). The perception of Nod factors by the host plant triggers a series of morphological and physiological changes in the root, such as root hair deformations and curling, cortical cell

246
Root Development divisions, changes in actin filaments near root hair tips, preinfection threads formation and up to the formation of nodule-like structures in some legumes (Stacey et al., 2006a). In root hairs, Nod factors additionally induce a rapid influx of calcium followed by a KCl efflux leading to a transient depolarization of the plasma membrane (Felle et al., 1995); and, few minutes later, calcium oscillations (Ca2+ spiking) (Ehrhardt et al., 1996; C´ardenas et al., 1999). In addition to Nod factors, other bacterial surface components are required for the accurate development of infection threads and further stages of nodulation, such as exopolysaccharides or lipopolysaccharides (see Section 10.1.4). These studies indicate that besides the key role of Nod factors, other bacterial signals are required for a successful symbiotic interaction. Furthermore, it was recently reported that nodulation arises in some legumes in the absence of nodABC genes, required for Nod factor synthesis. Indeed, two photosynthetic Bradyrhizobia sp. use an alternative pathway (potentially involving purine derivatives) to establish nodules on stems of sensitive jointvetch (Aeschynomene sensitiva) (Giraud et al., 2007).

10.2.2

Perception and transduction of the Nod factor signal: a genetic approach

The Nod factor signaling pathway has been recently dissected through the identification of mutants showing early defects in nodulation (nod− ; Table 10.1). Most of these genetic studies have been carried out in the model legumes Lotus and Medicago truncatula (Barker et al., 1990; Handberg and Stougaard, 1992). The nfr1 and nfr5 mutants of Lotus (Nod factor receptor), sym2 of pea (symbiotic locus) and nfp mutants of Medicago (Nod factor perception) are affected in all the earliest Nod factor responses, such as root hair deformation, curling and calcium spiking (Ben Amor et al., 2003; Madsen et al., 2003; Radutoiu et al., 2003) (Table 10.1). The corresponding genes have been cloned and encode transmembrane LysM-type serine/threonine receptor kinases (LYKs). The pea sym2 mutation, which shows a symbiotic phenotype dependent on the specific recognition of an acetylated Nod factor, links PsSYM2 directly to Nod signal recognition (Lie, 1984; Geurts et al., 1997) and a Medicago orthologous region contains a cluster of LysM RLK genes in Medicago (Limpens et al., 2003). The LysM domain corresponds to a peptidoglycan-binding motif recently shown to bind chitin, a well known pathogenic elicitor (Bateman and Bycroft, 2000; Kaku et al., 2006; Mulder et al., 2006). Therefore, LysM RLKs might be good candidates for Nod factor receptors that contain N-acetylglucosamine backbones. In M. truncatula, two of these genes, MtLYK3 and MtLYK4, are closely related to LjNFR1. RNA interference (RNAi) constructs for each of these genes reduces infection thread formation (Limpens et al., 2003). More recently, study of a weak allele for the Medicago hcl (hair curling) mutant led to the proposition that HCL, which corresponds to MtLYK3, could be

Legume Root Architecture
247

a component of a Nod factor entry receptor (Smit et al., 2007). MtNFP is likely an ortholog of LjNFR5 (Arrighi et al., 2006), as these LysM kinases both lack the kinase activation loop and may not be able to autophosphorylate. LjNFR5/LjNFR1 and MtNFP/MtLYK3 or MtNFP/MtLYK4 proteins might act as heteromeric Nod factor receptors. Recently, the LysM2 domain of Lotus NFR5 has been implicated in the determination of Nod factor host range specificity (Radutoiu et al., 2007). Even though at least three different Nod factor-binding sites have been biochemically identified (Hogg et al., 2006), a direct biochemical interaction between Nod factors and LYK proteins remains however to be demonstrated. These various LYK complexes may be required at different steps of the interaction (Radutoiu et al., 2003; Arrighi et al., 2006), as suggested by the various expression patterns of LYK/NFR genes: LjNFR1 seems root specific, and both LjNFR1 and LjNFR5 expressions are not affected by Mesorhizobium loti inoculation. In M. truncatula, MtLYK3 is expressed in root tissues, but not in nodules, whereas MtLYK4 was not detected using Northern analysis. Four other LYK or LYK-related genes (LYR2, LYR3, LYR6 and LYK10) were expressed in roots and nodules, two mainly in roots (LYR1 and LYK8), and three in all organs tested (Arrighi et al., 2006). More detailed studies revealed that MtNFP is expressed only in growing root hairs susceptible to Rhizobium and that, upon rhizobial inoculation, its expression follows nodule primordia formation and is found later in the infection zone of mature nodules. These results indicate that several heterodimeric Nod factor receptors might be involved all throughout the rhizobia infection process, for infection thread development and eventually bacterial release. Other components of the Nod factor signaling pathway, acting downstream of the Nod factor perception have been identified in Medicago by characterizing does not make infection mutants (dmi1, dmi2, dmi3) that are affected in nodulation as well as in mycorrhiza symbiotic interactions (Weidmann et al., 2004; Ol´ah et al., 2005; and orthologous mutants in Lotus: Kistner et al., 2005); see Chapter 9). dmi1 and dmi2 mutants show wild-type root hair deformations, but no calcium spiking, in response to rhizobia. The MtDMI1 and MtDMI2 genes code for a putative cation channel and a leucine-rich-repeats receptor-like kinase (referred to NORK in alfalfa, for NOdulation Receptor Kinase), respectively, and might act downstream of the putative MtNFP receptor (Endre et al., 2002; An´e et al., 2004). Recently, MtDMI1 was shown to transport monovalent cations such as Na+ /Li+ but not Ca2+ , using heterologous expression in yeast (Peiter et al., 2007). This channel may then be indirectly linked to Ca2+ oscillation induced in response to Nod (and Myc) factors. PsSYM8 (for SYMbiosis locus 8) in pea, and CASTOR and POLLUX in Lotus are orthologs of MtDMI1 (ImaizumiAnraku et al., 2005; Edwards et al., 2007). The Lotus proteins contain functional plastid localization signals and were proposed to form a heteromeric channel that might mediate the ion fluxes required for calcium spiking between plastids and cytosol (Imaizumi-Anraku et al., 2005). However, a MtDMI1:GFP

nod −

Lj

Ms Ps Mt

Lj

pollux

Ps Mt

sym2 dmi2

nork sym19 dmi1

Mt

hcl

symrk

Mt

nfp

nod −

Lj Lj

nfr1 nfr5

nod −

Plant Species

Mutant Name

Ion-channel protein

LRR RK LRR RK Ion-channel protein

LRR RK

LysM RK LRR RK

LysM RK (LYK3)

LysM RK*

LysM RK LysM RK*

Mutated Gene

Mutants affected in nodule development

Nodulation Phenotype

Table 10.1

Roots and infection zone of nodules Various organs; stronger in nodules

Roots and infection zone of nodules

Roots and infection zone of nodules Root

Roots

Gene Expression

Enhanced root hair response to touch

myc-

Defective microtubule organization in root hairs

Other Phenotypes

Plastid localization

Nuclear envelope myc-

Plasma membrane, infection thread membrane

Protein Localization

Imaizumi-Anraku et al. (2005)

Endre et al. (2002) Stracke et al. (2002) An´e et al. (2004), Riely et al. (2007)

Stracke et al. (2002)

Esseling et al. (2004)

Limpens et al. (2003) An´e et al. (2002)

Catoira et al. (2001), Smit et al. (2007)

Radutoiu et al. (2003) Madsen et al. (2003), Radutoiu et al. (2007) Ben Amor et al. (2003)

Selected References

−

Lj

sym15

nod −

snf2 (gof) nin nin sym35 bit1/pdl

snf nod −

Lj Mt Ps Mt

Lj

Lj

nsp2

hit1

Mt

Lj

nsp1

nsp2

Mt

nsp1

snf1 (gof)

Lj Lj Mt

Lj

nup85 nup133 dmi3

nod −

nod −

nod

snf

nod −

nod −

castor

GRAS-family TF Cytokinin receptor LHK1 Putative TF Putative TF Putative TF AP2/ERF family TF

GRAS-family TF GRAS-family TF GRAS-family TF

CCaMK

Ion-channel protein Nucleoporin Nucleoporin CCaMK

Induced by Rhizobia

Nodules Nodules

Roots and nodules Roots, nodules, and shoots

Preferentially in roots Roots and nodules Various organs and induced by Rhizobia

Roots and nodules

Ubiquitous Ubiquitous Roots and nodules

Various organs

Nuclear envelope, endoplasmic reticulum

Nucleus

Aberrant or aborted infection threads

(Continued)

Tirichine et al. (2007) Schauser et al. (1999) Marsh et al. (2007) Borisov et al. (2003) Middleton et al. (2007)

Heckmann et al. (2006) Murray et al. (2007),

Heckmann et al. (2006) Kalo´ et al. (2005)

Smit et al. (2005)

Imaizumi-Anraku et al. (2005) myc − Saito et al. (2007) Kanamori et al. (2006) Nuclear envelope myc − Nucleus myc − Pseudomonas L´evy et al. (2004), fluoresens Sanchez et al. (2005) defective perception Tirichine et al. (2006b) Plastid localization

Ps

Gm

lin

latd/nip

lot1

sunn

har1

sym29

nark

nod −

nod −

nod −

nod ++

astray/sym77

Lj

crinkle/sym79

nod −

nod ++

Mt

alb1/sym74

nod −

Lj

Lj

Mt

Mt

Lj

Lj

Mutant Name

Nodulation Phenotype

Plant Species

(Continued)

Table 10.1

CLAVATA1like RK CLAVATA1like RK bZIP TF (HY5)

CLAVATA1like RK CLAVATA1like RK

Mutated Gene

Ubiquitous

Ubiquitous

Gene Expression

Protein Localization

Krusell et al. (2002)

Nishimura et al. (2002)

Schnabel et al. (2005)

Kuppusamy et al. (2004) Bright et al. (2005), Veereshlingam et al. (2004) Ooki et al. (2005)

Imaizumi-Anraku et al. (2000) Tansengco et al. (2003)

Selected References

Nitrate tolerant, Searle et al. (2003) increased density of lateral roots Pleiotropic effects Nishimura et al. on photomorpho- (2002) genesis

Moderate dwarf, distorted trichome, high sensitivity to nitrate, ethylene insensitivity Nitrate tolerant, shortened primary root Nitrate tolerant, increased density of lateral root, shorter primary root Nitrate tolerant

Defect in lateral root formation

Pleiotropically affected in development

Other Phenotypes

Lj

Gm

klavier

nts382

EIN2

Increased in root length, delayed petal and leaf senescence, ethylene insensitivity Dwarf, delayed flowering, short root and low lateral root density Nitrate tolerant nodulation, early lateral root formation

Carroll et al. (1985)

Oka-Kira et al. (2005)

Penmetsa and Cook (1997)

Abbreviations: alb, aberrant localization of bacteria inside the nodule; AP, APetala; bit, branching infection threads; CCaMK, calcium-calmodulin-dependent protein kinase; dmi, does not make infections; ERF, Ethylene Response Factor; ERN, ERF Required for Nodulation; har, hypernodulation and aberrant root; hcl, hair curling; hit, hyperinfected; HY, long HYpocotyl; klv, klavier; latd, lateral root organ defective; LHK, Lotus Histidine Kinase; lin, lumpy infections; lot, low nodulation and trichome distortion; LYK, LysM receptor Kinase; LysM RK∗ , LysM RK without autoactivation loop; nark, nodule autoregulation receptor kinase; nfp, Nod factors perception; nfr, Nod factors receptor kinase; nin, nodule inception; nip, numerous infections and polyphenolics; nork, nodulation receptor kinase; nsp, nodulation signaling pathway; nup, nucleoporin; pdl, poodle; RK, receptor kinase; skl, sickle; snf , spontaneous nodules formed; sunn, super numeric nodules; sym, symbiotic; symrk, symbiosis receptor kinase; TF, transcription factor; gof , gain of function; Gm, Glycine max; Lj, Lotus japonicus; Ms, Medicago sativa; Mt, Medicago truncatula; Ps, Pisum sativum.

Mt

sickle

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Root Development fusion expressed from its own promoter in a Medicago dmi1 background did not lead to a plastid localization (Riely et al., 2007). In fact, these careful experiments located the MtDMI1 channel in the nuclear envelope. Orthologous genes of DMI2 have been identified in Lotus (LjSYMRK for SYMbiosis Receptor-like Kinase; Schauser et al., 1998; Stracke et al., 2002; Kistner et al., 2005), in S. rostrata (SrSYMRK; Capoen et al., 2005), in alfalfa (MsNORK for NOdulation Receptor Kinase; Endre et al., 2002) and in pea (PsSYM19 for SYMbiosis locus 19; Stracke et al., 2002). A recent study revealed that a SYMRK gene from a nonnodulating Brassicales species (Garden nasturtium, Tropaeolum majus) is able to complement the nodulation phenotype of the Lotus symrk mutant in contrast to genes from more distantly related species such as tomato or monocots (Markmann et al., 2008). Nevertheless, all these SYMRK genes are able to complement the defect in mycorrhizal interactions. As SYMRK gene structure is conserved among Eurosids but not in other phylogenic groups, these results suggest that SYMRK has been recruited for nodulation from a preexisting signaling pathway linked to mycorrhiza. In addition, the Medicago MtDMI2 homolog has been associated to root hair sensitivity to touch and pathogenic responses, suggesting broader functions for this receptor in root hair perception of the biotic environment (Esseling et al., 2004). Like many of its orthologs, the MtDMI2 gene is expressed at high levels in roots and in the infection zone of the nodule. MtDMI2 localizes in the plasma membrane and the infection thread, suggesting a late function in nodulation to control infection thread, rhizobial release and symbiosome formation (Bersoult et al., 2005; Capoen et al., 2005; Limpens et al., 2005). Recently, Kevei et al. (2007) have identified a protein interacting with the DMI2 kinase domain, and corresponding to a key enzyme for mevalonate synthesis (3-Hydroxy-3-MethylGlutaryl CoA Reductase 1; MtHMGR1). This general pathway seems to play an essential role for nodule development although which isoprenoid compound(s) may be involved could not yet be determined. The Lotus mutants (nup133 and nup85) affected in nucleoporin genes show thermosensitive mycorrhiza and nodulation phenotypes, calcium spiking and RHC defects (Kanamori et al., 2006; Saito et al., 2007; see Chapter 9). These nucleoporin genes are expressed in all plant organs and NUP133 is localized in the nuclear envelope of root cells. These nucleoporins might be taking part of a nuclear pore subcomplex required for calcium spiking and then implicated in symbiotic interactions. Further downstream components could be identified with the Medicago dmi3 mutant that is able to allow calcium spiking in response to Nod factors, but is defective at later stages of the symbiotic response and also in the mycorrhizal interaction (Catoira et al., 2000). Similarly, sym9 pea mutants are altered at the same nodulation stage, suggesting that PsSYM9 is a putative ortholog of MtDMI3 (Duc et al., 1989; Walker et al., 2000). MtDMI3 encodes a calciumcalmodulin-dependent protein kinase (Ca/CaM-DPK; L´evy et al., 2004; Mitra et al., 2004), located in the nucleus (Kalo´ et al., 2005). This gene is strongly

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expressed in roots and nodules, at lower levels in flowers, and is undetectable in stems and leaves (L´evy et al., 2004). Recently, a modified MtDMI3 protein lacking the predicted autoinhibition domain and a gain-of-function mutant in Lotus (snf1) similarly affected in the orthologous gene have been shown to induce the formation of nodules in the absence of rhizobia (spontaneous nodules, see Section 10.1.4) (Gleason et al., 2006; Tirichine et al., 2006b). Thus, the potential activation of MtDMI3 by both calcium and calmodulin suggests that this Ca/CaM kinase may integrate the oscillatory calcium signal into the Nod factor signaling pathway leading to nodule organogenesis. Mutants specifically affected in nodulation and not in mycorrhiza have also been identified downstream in the pathway through the cloning and characterization of Medicago mutants (nodulation signaling pathway 1 and 2, nsp1 and nsp2). The pea SYM7 gene might be an ortholog of nsp2 based on genetic map location (Kalo´ et al., 2005). Both Medicago mutants have similar phenotypes, showing wild-type calcium spiking, but reduced root hair deformations, blocked infection thread formation, drastically reduced expression of early nodulin genes, and no initiation of nodule primordia (Catoira et al., 2000; Oldroyd and Long, 2003). MtNSP gene products probably operate downstream of the previous genes, but upstream of Nod factor-induced early nodulin genes (such as MtRIP1 (Rhizobium Induced Peroxidase) and MtEnod11) (Catoira et al., 2000; Wais et al., 2000; Oldroyd and Long, 2003). MtNSP1 and MtNSP2 genes encode putative GRAS family transcription factors, respectively, expressed preferentially in roots or in roots, shoots and leaves (Kalo´ et al., 2005; Smit et al., 2005). MtNSP1 and Lotus LjNSP2 are located in the nucleus, whereas MtNSP2 is predominantly localized in the nuclear envelope and endoplasmic reticulum (Kalo´ et al., 2005; Smit et al., 2005; Murakami et al., 2006; Heckmann et al., 2006). Expression of LjNSP1, LjNSP2 and MtNSP2 is induced after rhizobial inoculation or Nod factor treatment, and the MtNSP2 protein is also relocalized into the nucleus after such treatments. Expression pattern, as well as heterologous complementation experiments with a Nicotiana benthamiana homolog, suggests that NSP1 may also be involved in later stages of nodulation, related to infection, bacterial release and bacteroid differentiation. Another gene that acts probably as transcriptional regulator is LjNIN (for Nodule INception): indeed, LjNIN contains membrane-spanning helices and a nuclear localization signal, similar to Notch and SREBEP transcription factors in animals (Schauser et al., 1999). The Lotus nin mutant shows excessive RHC, but aborted infection threads, no cortical cell division nor nodule primordia formation. Nod factor-induced calcium spiking and Enod expression patterns placed NIN downstream of the early signal exchanges between symbionts. LjNIN expression is strongly upregulated during nodulation and transcripts are localized in dividing cells of nodule primordia as well as in mature nodule parenchyma, nodule vascular bundles and nitrogen-fixing zones (Schauser et al., 1999). Orthologs are encoded by the sym35 locus in pea (Borisov et al., 2003) and MtNIN in Medicago (Marsh et al., 2007). Interestingly, MtNIN may

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Root Development negatively regulate the spatial expression domain of MtEnod11 in Rhizobiuminfected roots, acting on the ‘early’ downstream NFP-dependent Nod factor pathway but not on the ‘late’ MtHCL/LYK3 pathway. Recently, Middleton et al. (2007) have identified the bit1/pdl mutants (branching infection threads 1/poodle) as affected in the ERN1 gene (for ERF Required for Nodulation), encoding a transcription factor of the AP2/ERF family (APetala 2/Ethylene Response Element). The bit mutant is blocked in infection thread formation and Nod factor-induced gene expression. In parallel, a small family of ERN proteins (MtERN1 to 3; ERN1 corresponding to the previously mentioned ERN) was characterized as able to bind the Nod factor responsive regulatory unit (the NF-box) present in the MtENOD11 promotor (Andriankaja et al., 2007). In root hairs, Nod factor treatment upregulates MtENOD11 expression via an MtERN1 and 2 dependent pathways, whereas MtERN3 would repress MtERN1 and 2 activities. Antagonistic ERN transcription factor activities may be involved in fine-tuning gene expression in response to Nod factors. Several other mutants affected in different aspects of the symbiosis have been identified in Medicago or Lotus, showing frequently pleiotropic phenotypes (reviewed in Stacey et al., 2006a; Table 10.1). These mutants may be notably useful to analyze the integration of nodulation in whole plant physiology. Altogether, based on the identification through genetic screens of nonmodulating (nod− ) plants in different legume species, several genes have been cloned that affect Nod factor perception and signaling as well as early stages of the symbiotic interaction, allowing a transduction pathway leading to nodule organogenesis to be proposed (Jones et al., 2007). 10.2.3

Transcriptomics of early nodule development

Genes induced during nodule development, the so-called nodulin genes, are coordinately expressed in the different steps of the symbiotic interaction (van Kammen, 1984). It is interesting to note that some of the downstream Nod factor signaling pathway components identified through genetic approaches (see Section 10.2.2) behave as nodulins, such as LjPOLLUX, Mt/LjNIN, LjNSP1, Mt/LjNSP2 or MtERN1. This observation indicates that genetic and transcriptomic approaches can be complementary for the identification of genes involved in Nod factor signaling and nodule organogenesis. Reverse genetic approaches such as RNAi can be of particular interest when functional genetic redundancy might exist. Two classes have been distinguished according to their temporal expression pattern: early nodulin (or Enod) or late nodulin (or Nod) genes, relatively to the initiation of the nitrogen fixation process. Several strategies have been used to identify differentially expressed genes in nodules or infected root hairs: differential display, differential and cold-plaque screenings, as well as systematic sequencing of symbiotic cDNA libraries (Covitz et al., 1998;

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¨ Frugier et al., 1998; Gyorgyey et al., 2000; Medicago EST Navigation System [MENS] and The Gene Index [TGI] EST databases: http://medicago. toulouse.inra.fr/cgi-bin/Mt/MtESTMtCD2003.cgi.pl and http://compbio. dfci.harvard.edu/tgi/cgi-bin/tgi/gimain.pl?gudb=medicago), yielding a large diversity of molecular markers associated with different stages of nodule development in several legumes. A large number of early nodulin codes for proteins putatively associated to cell walls (e.g. proline-rich proteins, glycine-rich proteins and extensins; reviewed in Bladergroen and Spaink, 1998). Other early nodulins are homologous to lectins (Bauchrowitz et al., 1996), enzymes of the phenylpropanoid pathway (Estabrook and Sengupta-Gopalan, 1991; Savour´e et al., 1994), or pathogenesis-related proteins (Gamas et al., 1998; Goormachtig et al., 1998). Late nodulins are mainly related to nitrogen fixation and assimilation (e.g. sucrose synthase, glutamine synthetase, phosphoenolpyruvate [PEP] carboxylase and leghemoglobin; Crespi and G´alvez, 2000), or constituents of the peribacteroid membrane, such as the aquaporin Nod26, peptide transporters and a cytochrome P450 (Szczyglowski et al., 1997, 1998). More recently, genomic approaches have been developed in the model legumes Lotus and Medicago such as Serial Analysis of Gene Expression (SAGE; Asamizu et al., 2005), transcriptome profiling (Colebatch et al., 2002; El Yahyaoui et al., 2004; Kouchi et al., 2004; Mitra et al., 2004; Lohar et al., 2006) and substractive hybridization (Godiard et al., 2007). A GeneChip containing the genome of Sinorhizobium meliloti and approximately 10 000 Medicago probes has also been used to study simultaneously gene expression profiles in both symbiotic partners (Barnett et al., 2004). More than 200 Medicago genes were identified of which many had been formerly characterized as nodulins. Deep sequencing of cDNA libraries together with in silico screenings of available EST databases have been used to isolate nodule-specific genes with conserved motifs, such as F-box proteins, proline-rich proteins and cysteine-rich proteins (Fedorova et al., 2002; Mergaert et al., 2003; Graham et al., 2004). Roles for these genes in nodule differentiation have been mainly deduced from their homologies and their spatiotemporal expression patterns. In a few cases, transgenic approaches to modify nodulin gene expression have been utilized to address their function in the symbiotic interaction. One of the first studies concerned the Enod40 gene, a nonprotein coding RNA with highly conserved secondary structures, suggesting a possible role as riboregulator (Crespi et al., 1994). This RNA does not contain any long ORF but can be translated in vitro into two small peptides spanning a conserved 5 region. ¨ These ENOD40 peptides are able to bind sucrose synthase (Rohrig et al., 2002). In addition, the MtEnod40 RNA has been shown to be involved during nodulation in the relocalization of an RNA-binding protein, MtRBP1, from nuclear speckles to the cytoplasm (Campalans et al., 2004). Overexpression of MtEnod40 resulted in rapid rhizobial infection of cortical cells and acceleration of the nodulation process (Charon et al., 1999). Among the transgenic plants obtained, two lines in which this gene was cosuppressed formed few

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Root Development nodules with arrested meristems, suggesting that MtEnod40 function is required for nodule morphogenesis. More recently, a second MtEnod40 gene has been identified in Medicago, and RNAi plants showing reduced expression of both genes formed only few and modified nodule-like structures devoid of nitrogen-fixing bacteroids (Wan et al., 2007). These two genes might be required for nodule initiation and bacteroid development. In contrast, RNAi analysis of LjEnod40 in Lotus revealed a strong inhibition of nodule primordia initiation, but normal infection thread formation (Kumagai et al., 2006). Antisense Medicago plants in which an activator component of the Anaphase-Promoting Complex/Cyclosome (APC/C), MtCcs52A (cell cycle switch 52-kDa protein) was downregulated, showed defects in nodule development, correlated with cells having lower ploidy levels and reduced size (Vinardell et al., 2003). Moreover, ineffective invasion as well as early bacterial and nodule plant cell senescence observed in these plants revealed that this limiting factor for the regulation of endoreduplication is crucial for the differentiation of nitrogen-fixing symbiotic cells. A specific soybean apyrase gene, GS52 (for Glycine soja 52-kDa protein), was constitutively expressed in Lotus. The resulting transgenic plants showed increased infection thread formation and enhanced length of the infection zone (McAlvin and Stacey, 2005). More recently, functional analysis of the flavonoid pathway using RNAi directed against different biosynthetic enzymes confirmed its crucial involvement in the early stages of the rhizobial interaction (Subramanian et al., 2006; Wasson et al., 2006). Few putative regulatory genes induced during nodulation have been functionally studied. Genes encoding small G-proteins of the Rab type, GmRab1 and GmRab7, induced in mature nodules were characterized in soybean using their antisense expression driven by a leghemoglobin promoter (Cheon et al., 1993). For both genes, this resulted in the blocking of vesicles transport into the peribacteroid membrane. For GmRab1, a decrease in nodule size, bacteroid number per cell, and nitrogen fixation activity was also observed. Another study using an antisense strategy in Medicago revealed the role of a vascular-associated C2 H2 zinc finger putative transcription factor, MtZpt2-1 (Frugier et al., 2000): antisense plants grew normally but developed non-nitrogen-fixing nodules, in which differentiation of the bacteroids was impaired. Hence, this transcription factor seems to be required for the differentiation of the nitrogen-fixing zone of the root nodule. More recently, a transcription factor of the CCAAT-box binding factor family (CBF/NF-Y/ HAP), MtHAP2-1, whose transcript was posttranscriptionally regulated by microRNA 169, was associated to nodule organogenesis and the balance between nodule meristematic activity versus differentiation (Combier et al., 2006). Functional studies have also been attempted with late nodulin genes showing their involvement in different aspects of the carbon/nitrogen metabolism, e.g. glutamine synthetase (Carvalho et al., 2003; Ortega et al., 2006), sucrose synthase (Baier et al., 2007; Horst et al., 2007), PEP carboxylase (Nomura et al.,

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2006) and leghemoglobin (Ott et al., 2005). In addition, a Lotus non-nitrogenfixing mutant (fix− ), ign1 (for ineffective greenish nodules) showing premature senescence and abnormal symbiosome formation was characterized as affected in a novel ankyrin-repeat protein localized at the plasma membrane (Kumagai et al., 2007).

10.3

10.3.1

Evidences for a crosstalk between symbiotic nodule and LR developmental pathways Differences and similarities between LR and nodule: ontology and histology

The evolutionary origin of nodules has been the subject of many speculations. No fossil nodules have been described, making it difficult to determine whether ancestral legumes were even nodulated (Hirsch and LaRue, 1997). Several hypotheses have been proposed: nodules could be highly modified preexisting organs, such as stems (Sprent and Raven, 1992), LRs (Nutman, 1948), or carbon storage organs (Caetano-Anoll´es et al., 1993). Alternatively, nodules could be novel organs (Libbenga et al., 1973) that might have evolved from another symbiotic interaction (e.g. endomycorrhiza) or from a plant–bacteria pathogenic interaction (Hirsch and LaRue, 1997). Nodules and roots share many aspects of their development, consistent with nodulation evolving from preexisting LR organs (Hirsch and LaRue, 1997; Mathesius et al., 2000). Homology between these organs has been inferred in nonlegume nitrogen-fixing nodules, especially those formed on actinorhizal plants (see Chapter 9) or on Parasponia roots. These nodules are clearly modified LRs in terms of their structure and development. In contrast, legume nodules diverge from LRs in their developmental origin, anatomy and patterns of gene expression (Hirsch and LaRue, 1997). The orientation of the initial division leading to LR or nodule formation in legumes seems controversial in the literature, likely because studies were based on electron microscopy of fixed sections (Torrey, 1986). The statement that the first pericycle cell division occurring at the onset of nodule formation is anticlinal whereas it is periclinal during LR initiation may need to be reconsidered. In Arabidopsis, where LR initiation can be observed maintaining whole root integrity, the initial division occurring in the pericycle is anticlinal (Malamy and Benfey, 1997), and this model may be extrapolated to legumes. Hence, both legume root-derived organs are likely to initiate through an initial anticlinal division of the pericyle in front of a protoxylem pole. Nevertheless, LRs and nodule primordia are formed primarily from different tissues, pericycle and cortex, respectively (Brewin, 1991; Hirsch, 1992). Thus, even though the same root tissue layers are involved, they have different relative contributions. Nodules of peanut (Arachis hypogaea) originate

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Root Development predominantly from the pericycle (Allen and Allen, 1940). Additional major differences between root and nodule histology are that nodules lack an apical cap and have a peripheral vasculature. Intermediate lateral organs have been identified and the existence of such ‘root-nodule hybrids’ supports the theory that nodule formation evolved from developmental pathways activated during LR formation. Root-nodule hybrids have been described in alfalfa (Dudley and Long, 1989), white clover (McIver et al., 1993) and bean (Phaseolus vulgaris) (Vandenbosch et al., 1985; Ferraioli et al., 2004) following inoculation with specific Rhizobium strains. In the case of bean, the different R. etli mutants collectively called ‘root inducer’ (or RIND) provoked the development of ectopic roots from abortive nodule primordia (Ferraioli et al., 2004). These mutants initiate a wild-type early sequence of nodulation events in the root, including root hair deformation and development of nodule primordia. Later on, the primordia adopt an elongated shape and in the distal region one or more ectopic roots can emerge. The homeotic mutant cochleata (coch) of pea, having stipules replaced by alternative leaf components (Wellensiek, 1959) has been also recently shown to be affected in nodule development (Ferguson and Reid, 2005). These nodules are dichotomously branched and multiple callus and root structures emerge from their apical meristem, creating root-nodule hybrid structures. These peculiar roots, generally protruding from the side of the nodule lobes, display agravitropism. The nodules formed are functional and the hybrid root structures have an anatomy similar to wild-type LRs. High temperatures have also been reported to convert nodule apex into roots and calli structures in alfalfa and various Trifolium species (Dart, 1977). Moreover, treating roots with auxin transport inhibitors lead to the formation of nodule-like structures with some histological traits typical of LRs in alfalfa and pea (see Section 10.4.2.1) (Hirsch et al., 1989; Scheres et al., 1992). 10.3.2

Common marker genes between LR and nodules

As mentioned above, many plant genes, known as nodulin genes, are induced during different stages of nodule development but several are also expressed in nonsymbiotic tissues, and closely related genes exist in nonlegume plants (Arredondo-Peter et al., 1998; Kouchi et al., 1999). These observations suggest that several functions required for nodulation have been recruited from preexisting genes that are present in both leguminous and nonleguminous plants (Bladergroen and Spaink, 1998). Indeed, several nodulins are also expressed in LRs, such as the early nodulins Enod2, Enod11, Enod12, Enod40, MtAnn1 (coding for an Annexin), MtLAX1/MtLAX2 (coding for AUX1-like genes; see Section 10.4.2.1) (Hirsch et al., 1989; Yang et al., 1993; Crespi et al., 1994; Journet et al., 1994; Papadopoulou et al., 1996; de Carvalho Niebel et al., 1998; de Billy et al., 2001; Journet et al., 2001), or even late nodulins linked to carbon and nitrogen metabolisms (Govers et al., 1985). In addition, nodule and LR also show similar patterns of ProGH3 :GUS expression during

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comparable developmental stages, suggesting that changes in auxin concentration or transport are required in both organs (Mathesius et al., 1998) (see Section 10.4.2.1). 10.3.3

Cell-to-cell communication in LR and nodule formation

Nodule and LR initiation are controlled by a number of hormonal factors (see Section 10.4.2) that emanate from the stele and shoot-dependent autoregulation mechanisms (Caetano-Anoll´es and Gresshoff, 1991) (see Section 10.3.4), suggesting the involvement of long-distance and local cell-to-cell communication mechanisms. The cytoplasms of neighboring plant cells can be connected by channels spanning the cell walls called plasmodesmata (PD) (Complainville and Crespi, 2004). Regulation of PD-mediated communication seems to play a role in inter-organ communication at the whole plant level; therefore, plants can be divided into symplasmic domains, e.g. groups of cells interconnected by functional PDs that ensure the coordination of development by allowing diffusion of signal molecules, such as transcription factors, only within the cells of a symplasmic domain. As a tracer of macromolecular phloem unloading, GFP has been expressed under the control of the companion cell-specific AtSUC2 (Sucrose transporter 2) promoter in Arabidopsis (Imlau et al., 1999). In Medicago roots, the use of the AtSUC2-GFP reporter has revealed PD-mediated communication between the root phloem and LR primordia as well as root apical meristems (Complainville et al., 2003). After Rhizobium inoculation, GFP could be detected in the nodule primordia, as soon as the first cortical cell divisions occurred, as well as in the meristems, the invasion zones, and the vascular tissue of mature nodules. This movement of the GFP protein precedes cortical cell division and was concomitant with a rearrangement of the PD network in this initial cell as shown by electron microscopy. Perturbation of this process with viral proteins further indicated that cell-to-cell communication between nodule initials and the phloem is crucial for this de novo organogenesis (Complainville et al., 2003). The phenotype of several supernodulating mutants depends on the shoot genotype, as shown by grafting experiments, suggesting that a shootderived mobile signal is responsible for nodule number autoregulation and further addressing the role of long distance cell-to-cell communication in nodule organogenesis. 10.3.4

Mutants affected in both root lateral organogeneses

Several mutants affected in genes with a dual function in nodule formation and root development have been identified, such as lateral root organdefective (latd/nip) (Veereshlingam et al., 2004; Bright et al., 2005) and several hypernodulating mutants (har1, super numerous nodulese [sunn], nitrate tolerant symbiosis 382 [nts382] and sickle [skl]) (Day et al., 1986; Penmetsa and Cook, 1997; Wopereis et al., 2000; Penmetsa et al., 2003; Table 10.1), suggesting the

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Root Development existence of common regulatory pathways between these two root-derived organogeneses. Other mutants such as crinkle, klavier or astray are not only affected in root and nodule formation, but also in other plant organs development (Nishimura et al., 2002b; Tansengco et al., 2003; Oka-Kira et al., 2005; Table 10.1). More recently, an RNAi strategy used in Medicago to downregulate specifically the MtCRE1 (Cytokinin Response 1) gene encoding a cytokinin receptor revealed a positive and negative role of this pathway in regulating nodule or LR formation, respectively (Gonzalez-Rizzo et al., 2006; see Section 10.4.2.2). In addition, overexpression of microRNA 166 downregulates class III HD-ZIP transcription factors, and leads to an abnormal patterning of root vascular bundles correlated with defective LR and nodule formation (Boualem et al., 2008). The Medicago latd mutant is defective in both nodule formation and root development (Bright et al., 2005). Even though the latd main root grows normally a few days after germination, it stops later on with a strong inhibition of LR formation. The latd LRs are disorganized and lack a visible root cap. In addition, latd nodulation is also blocked, and nodule primordia remain small, white and undifferentiated. Therefore, the MtLATD gene seems required for the function of three root-derived meristems (e.g. primary root, LR and symbiotic nodule). Interestingly, the shoot meristem is not modified in these mutants. The crinkle mutant (Ljsym79) is perturbed in root growth and root hairs, but also in trichome and seed development (Tansengco et al., 2003). The abnormal nodulation of crinkle, characterized by arrested infection thread growth in the root epidermis, might be associated with an enlarged root hair base phenotype independent from the symbiotic interaction. Blocked infection thread development leads to the formation two types of nodules on crinkle roots: small white nodules, which are frequently found, and enlarged, irregularly shaped nodules. After inoculation with rhizobia, shoot growth and LR formation are significantly decreased when compared to the wild type. Another class of mutants (hypernodulating or supernodulating; nod++ ) are affected in autoregulation, a systemic feedback mechanism negatively controlling the final number of nodules formed in legume root systems (CaetanoAnoll´es and Gresshoff, 1991). Grafting experiments have revealed that nodule autoregulation is controlled by the shoot and the root, suggesting the existence of crosstalks involving the whole plant to limit the number of nodule primordia and of rhizobial infections within the root nodulation zone, respectively. These negative autoregulatory mechanisms may also affect the regulation of other root meristems, and, consequently, the whole root architecture of legumes (Jiang and Gresshoff, 1997). Shoot-dependent hypernodulating mutants have been identified in several legumes: har1 in Lotus (Nishimura et al., 2002a), nodule autoregulation receptor kinase (nark) in soybean (Searle et al., 2003), sym29 in pea (Krusell et al., 2002), and sunn in Medicago (Penmetsa et al., 2003; Schnabel et al., 2005). These mutants display similar nodulation phenotypes, with infection events

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(root hair deformation and infection thread formation) similar to those of wild type, but with increased nodule formation all along the root. They are all affected in LRR-type receptor kinases similar to Arabidopsis CLAVATA1 (CLV1), a gene involved in the restriction of shoot meristem proliferation through a short-distance negative feedback loop (Clark et al., 1997). In Lotus, the har1 mutant also displays a drastically altered root development controlled by the shoot (Wopereis et al., 2000; Krusell et al., 2002): a short main root, with an increased LR density in uninoculated plants compared to wild type. In contrast, the sunn Medicago mutant displays a shortened primary root growth in presence or absence of rhizobia, but no significant LR phenotype (Penmetsa et al., 2003; Schnabel et al., 2005). Changes in root morphology have been associated also to other hypernodulating mutants. In the case of the soybean nts382 mutant showing a shootdependent hypernodulation phenotype, early LR formation is increased independently of the application of rhizobia and/or nitrate (Day et al., 1986). The pea nod3 mutant displays a root-dependent hypernodulation phenotype (Gremaud and Harper, 1989) and might be affected in the production of a signal molecule by roots or nodule primordia that would move to the shoot to activate systemic negative feedback mechanisms. This mutant also displays an altered root morphology phenotype: grown on nitrate conditions main root growth is reduced and LR formation is increased (Jacobsen and Feenstra, 1984). Other nod++ mutants are affected more pleiotropically. In the Medicago skl mutant, nodulation enhancement is linked to an increased persistence of successful infections which therefore lead to a local increase in nodule number within the initial infection zone. In addition, these nodules are not always formed in front of protoxylem poles (Penmetsa and Cook, 1997). An increased root length as well as delayed petal and leaf senescence are also observed together with ethylene insensitivity (Penmetsa and Cook, 1997; Penmetsa et al., 2003; Prayitno et al., 2006). Indeed, root or hypocotyl growth as well as nodulation are not inhibited in response to this phytohormone (Penmetsa and Cook, 1997). Grafting experiments indicated that MtSKL is involved in a root-dependent autoregulatory mechanism (Prayitno et al., 2006). The klavier mutant of Lotus is dwarf and exhibits aberrant leaf veins, markedly delayed flowering, and short roots with a decreased number of LRs (Oka-Kira et al., 2005). Grafting klv shoots on wild-type roots caused an increase in the number of nodules and the width of the nodulation zone. The astray mutant in Lotus (Ljsym77) shows enhanced nodulation and has pleiotropic effects on root, shoot and hypocotyl elongation under different light conditions (Nishimura et al., 2002b). The mutant name astray is derived from the agravitropic LR phenotype (going ‘astray’ against gravity). The gene responsible for these phenotypes, LjBZF encodes a homolog of the Arabidopsis bZIP transcription factor HY5 (long HYpocotyl 5) involved in photomorphogenesis (Hardtke et al., 2000). Unlike the hy5 mutant, astray did not show an enhancement of LR formation but increased nodulation.

262
Root Development Collectively, these data suggest that a single gene can regulate several aspects of legume root architecture (rhizobial infection, nodule development, root growth and LR formation) as well as other plant developmental programs (in shoots, hypocotyls, leaves, or seeds). Certain genes can have either a positive role on the formation of both types of root lateral organs (such as latd) or inhibitory effects on these organogeneses (such as har1). 10.3.5

Environmental control of legume root architecture

Several environmental factors have been reported to influence the development of root-derived organs in legumes, among which availability of nutrients such as nitrate or phosphate, and growth under abiotic stress conditions, such as salt, water deficit, temperature or pH. Besides LR development, the ability of legume roots to interact with symbiotic microorganisms, such as rhizobia or mycorrhizal fungi (see Chapter 9), constitutes an adaptation to specific nutrient starvation conditions. These interactions are themselves influenced by environmental conditions of the soil (such as abiotic stresses). Nitrate (NO3 − ) is particularly relevant for legume root architecture, because its availability modulates plant growth and exerts complex effects on root growth, LR formation and the symbiotic interaction (Dazzo and Brill, 1978; Gresshoff, 1993). In fact, nitrate has been shown to locally stimulate LR formation but also to systemically inhibit the formation of nodules at early developmental stages (Carroll et al., 1985; Zhang and Forde, 1998). Indeed, nitrate deprivation represents the major environmental factor that regulates nodulation (Streeter, 1985a, 1985b) and most hypernodulating mutants, such as har1 in Lotus, or several nitrate tolerant symbiosis (nts) mutants in soybean are also affected in their nitrate regulation, suggesting that these two pathways are tightly interconnected (Carroll et al., 1985; Wopereis et al., 2000). In addition, inhibition of nodulation by nitrate can be overcome by the ethylene inhibitor aminoethoxyvinylglycine, suggesting that this phytohormone mediate response to this environmental signal (Ligero et al., 1991). Phosphate (PO4 3− ) availability is, together with nitrogen, a major yield-limiting factor in many regions of the world. This compound affects root development and nodulation (Pereira and Bliss, 1989). In common bean, phosphorus deficiency increases root growth and stimulates root hair development and this response might be mediated by ethylene (Lynch and Brown, 1997). Phosphorus appears to be also essential for proper nodule development and nitrogen fixation (Ssali and Keya, 1983; Pereira and Bliss, 1989). Among the abiotic stresses, studies involving physiological, molecular and functional data have been carried out mainly on salt stress in legumes. Salinity in the arid and semiarid regions of the world is a serious threat to agriculture. Increasing salt concentrations in soils lead to marked changes in the root growth pattern of legumes and also affect the symbiotic nitrogen fixation process. Legumes are very sensitive to salt levels in soils (Arrese-Igor et al.,

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1999), whereas rhizobia are generally much more tolerant (up to 700 mM NaCl) than their respective host legumes (Lauter et al., 1981; Singleton and Bohlool, 1984). Different steps of the symbiotic interaction and nodule development are affected by salt stress, reducing nodule number and limiting nitrogen fixation (Singleton and Bohlool, 1984). Reduced colonization and early rhizobial infection events (such as production of signaling molecules, RHC, infection thread formation and nodule initiation) are particularly sensitive to salt stress (Tu, 1981; McKay and Djordjevic, 1993). For example, salt stress (100 mM NaCl) decreases significantly the soybean root hair responses to Nod factors and at higher concentrations (such as 250 mM NaCl) causes shrinkage of root hairs, affecting consequently root hair deformation and inhibiting the infection process (Duzan et al., 2004). The later stages of nodule organogenesis have been shown to be more resistant to salinity than the infection process. However, degradation of peribacteroid membranes or decrease in bacteroid number have been reported in pea, leading to a reduction in ˜ et al., 2003). nitrogen fixation (Bolanos Molecular studies have been used to identify candidate genes as regulators of osmotolerant responses, particularly putative regulatory genes such as transcription factors (Hasegawa et al., 2000). In alfalfa, MsAlfin1 has been identified as a salt-inducible transcript that encodes a zinc-finger protein predominantly expressed in roots (Winicov, 1993). Overexpression of this putative transcription factor enhances root growth under control and saline conditions (Winicov, 2000). The lack of root formation of alfalfa transgenic lines expressing MsAlfin1 in an antisense orientation also supports a crucial role for this gene in root development. Another C2 H2 zinc-finger transcription factor (ZPT2-1) has been identified in alfalfa and characterized in Medicago (Frugier et al., 1998, 2000). This gene, expressed in vascular tissues of roots and nodules, is also induced by salt stress (Merchan et al., 2003; de Lorenzo et al., 2007; Merchan et al., 2007). Medicago MtZPT2-1 antisense lines develop nodules unable to fix nitrogen because of a block in bacteroid differentiation (Frugier et al., 2000), and are also less able to recover from a salt stress than the wild-type plant (Merchan et al., 2003). This suggests that the transcription factor might be involved in nodule and root-adaptive responses to osmotic and salt stresses.

10.4

Common signals in nodulation and LR development

One of the first to analyze the global regulation between LR and nodule formation was Nutman (1948) who found that roots of red clover (Trifolium pratense) with many LRs were able to develop more nodules after inoculation by rhizobia. Moreover, nodule formation in red clover and in pea is stimulated by the excision of either mature nodules or root tips (Nutman, 1952; Caetano-Anoll´es and Gresshoff, 1991). These findings indicate that similarly to differentiated nodules, main and LR tips can affect nodule formation, based

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Root Development on a negative feedback autoregulatory mechanism that might involve similar signal molecules. 10.4.1

Effects of Nod factors on root development

Recently, Nod factors of Bradyrhizobium japonicus have been reported to stimulate development of soybean root systems by increasing their total lengths and dry weights, indicating that these bacterial molecular signals also affect root development (Souleimanov et al., 2002). Nod factors of S. meliloti stimulate LR formation in M. truncatula, and this response depends on the specific structure of bioactive Nod factors (Ol´ah et al., 2005): indeed no effect could be detected using Nod factors from rhizobial species unable to nodulate M. truncatula. Moreover, no significant Nod factor-dependent stimulation of LR formation has been observed in nod− mutants (nfp, dmi1, dmi2, dmi3 and nsp1; see Section 10.2.2). Another effect of Nod factors on root development is the induction of root hair formation that has been described in vetch (Vicia sativa), Sesbania rostrata and Lotus, including in the Ljrhl1 mutant background defective in root hair formation in uninoculated roots (Roche et al., 1991; Mergaert et al., 1993; van Spronsen et al., 2001; Karas et al., 2005). In this last case, only inoculation with a Mesorhizobium loti strain able to synthesize Nod factors was able to induce de novo formation of root hairs in this mutant, mainly in association with nodules or nodule primordia. Nod factor signaling might thus be shared by legume root and nodule developmental pathways. Therefore, this stimulation of LR and root hair formation might be considered as part of the symbiotic responses. 10.4.2

Hormonal controls involved in both root lateral organogeneses

Endogenous plant hormones are involved in the regulation of many physiological processes during plant growth and development and also during plant adaptation to environmental conditions, such as nutrient starvation or stress (Martin et al., 2000; Mok and Mok, 2001). In the context of legume root development, Libbenga et al. (1973) showed that the joint addition of auxins and cytokinins on root explants of pea induced cortical cell divisions similar to those induced during nodulation, whereas auxin alone induced only pericycle cell divisions. These results revealed the role of various phytohormonal balances to allow cell division in the different layers of the root. Data suggesting a phytohormonal control of legume root architecture came initially from pharmacological approaches and, more recently, from genetic studies and transgenic approaches. 10.4.2.1 Auxin The phytohormone auxin and its polar transport have been shown to be crucial for the initiation and development of LRs in many plant species,

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including legumes (Casimiro et al., 2003). Involvement of auxin during symbiotic nodule formation was initially shown with auxin transport inhibitors (ATIs) such as 2,3,5-triiodobenzoic acid (TIBA) and N-(1-naphthyl) phthalamic acid (NPA) that induce the formation of nodule-like structures on alfalfa roots (Hirsch et al., 1989). These pseudonodules arise from pericycle cells, have central vascular bundles, similarly to LRs, and can be formed in the presence of nitrogen. However, they lack a root cap and exhibit a determinate growth even in species with indeterminate nodulation. These nodule-like structures express certain early nodulin genes, such as MsEnod2 and MsEnod12 (Hirsch et al., 1989; Scheres et al., 1992), although these genes are also expressed in LRs (see Section 10.3.2). Exogenous application of NPA activates cortical cell divisions and formation of pseudonodules also in nonnodulating mutants of white sweetclover (Melilotus alba) (Wu et al., 1996) or nonlegumes (Hirsch and LaRue, 1997). Moreover, NPA and purified Nod factors induce different early cellular and molecular responses in legume roots and their action is differentially controlled by environmental factors (light and nitrate) (Bauer et al., 1996). Altogether, these data suggest that these root-derived structures might result from an inhibition of LR elongation and subsequent fusion of closely spaced root primordia rather than from a nodulation-related process. Direct measurement of polar auxin transport (PAT) using radiolabelled auxin revealed that rhizobia or bio-active Nod factors locally inhibit acropetal auxin transport in vetch roots (Boot et al., 1999). Accordingly, the use of an auxin-responsive promoter fused to a reporter gene (ProGH3 :GUS) revealed a localized accumulation of auxin in white clover roots rapidly after rhizobia infection or Nod factor application (Mathesius et al., 1998). More recently, specific auxin efflux facilitators of the PIN family, such as MtPIN2, have been shown to be induced during nodulation and MtPIN2 silencing reduce the number of nodules formed (Schnabel and Frugoli, 2004; Huo et al., 2006). During Lotus determinate nodulation, an increase in PAT could be detected in response to Nod factors using a similar ProGH3 -reporter system or radiolabelled auxin (Pacios-Bras et al., 2003). Interestingly, modification of the expression pattern of this auxin-sensitive promoter during early stages of nodule and LR development are very similar, suggesting that similar auxin requirements might be shared in both types of root lateral organogeneses (Mathesius et al., 1998). Similarly, AUX1-like genes of Medicago (MtLAX) show an increased expression during LR and nodule formation at two stages, first during primordia formation and then in association with vascular bundles development (de Billy et al., 2001). In the search for a secondary signal that could mediate the action of Nod factors on the redistribution of auxin during nodulation, notably by mediating a block in PAT, plant flavonoids have been suggested as possible candidates (Hirsch et al., 1992). The strong temporal and spatial correlation between accumulation of specific flavonoids and the ProGH3 :GUS reporter construct indeed suggested that flavonoids could specifically regulate local auxin levels (Mathesius et al., 1998). Recently, the use of RNA interference to silence genes

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Root Development involved in the flavonoid pathway, a chalcone synthase gene in Medicago or an isoflavone synthase gene in soybean, has led to roots deficient in flavonoid content that exhibited reduced nodulation and increased auxin transport compared to control roots (Subramanian et al., 2006; Wasson et al., 2006). However, in soybean, the ability of isoflavones to modulate auxin transport does not seem essential for nodulation (Subramanian et al., 2006). In addition, long-distance auxin transport from shoot to root was shown to be increased in the sunn mutant (van Noorden et al., 2006), possibly leading to higher auxin accumulation in roots and hypernodulation, even though surprisingly no effect on LR development has been observed in this mutant (Schnabel et al., 2005). Finally, certain ethylene effects on nodulation or phenotypes of ethylene-insensitive mutants might involve the local regulation of PAT in roots (Prayitno et al., 2006). During the different stages of indeterminate legume nodule organogenesis, auxin accumulation might be the consequence of two synergistic, but independent, regulations of auxin transport: (1) a local inhibition of auxin transport during nodule initiation, which may be related to an induction of flavonoids synthesis and ethylene action (see Section 10.4.2.3) and (2) a reduction of long-distance PAT from the aerial part that would control auxin loading in roots and be part of a negative autoregulation mechanism (van Noorden et al., 2006; Wasson et al., 2006). A model can be proposed where activation of the SUNN/NARK/SYMRK receptor kinase (see Section 10.2.2) by an unknown ligand might produce a shoot-derived mobile signal linked to regulation of PAT and eventually to nodule autoregulation (Prayitno et al., 2006; van Noorden et al., 2006). Even though mechanisms involved remain to be precised, it is clear that both root lateral organogeneses involve a role for auxin local accumulation and/or signaling pathways. 10.4.2.2 Cytokinin Physiological studies based on exogenous application of cytokinins in legumes have shown that these phytohormones have an inhibitory effect on root growth and stimulate root hair formation (Lorteau et al., 2001). In etiolated pea seedlings, an inhibition of LR initiation has also been observed, as well as an additional thickening of the root tips (Bertell and Eliasson, 1992). Cytokinin also appears to stimulate root growth in the har1 mutant of Lotus (see Section 10.3.4) in an ethylene-independent manner (Wopereis et al., 2000). Effects of cytokinins on legume symbiotic nodule development have initially been analyzed using a Rhizobium nod− mutant strain (unable to synthesize Nod factors) that had been genetically modified to secrete the trans-zeatin cytokinin. This specific strain induced nodule-like structures on alfalfa roots with initiation of peripheral vascular bundles and expression of some early nodulins (Cooper and Long, 1994). Additionally, cytokinins elicit similar responses in alfalfa roots to Nod factors such as amyloplast deposition, cortical cell divisions and early nodulin gene expression (Bauer et al., 1996; Hirsch and LaRue, 1997; Jim´enez-Zurdo et al., 2000). Concomitant application of

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cytokinins and rhizobia increases the number of nodules formed in burr medic (Medicago polymorpha) (Yahalom et al., 1990). Cytokinins also affect nodulation in soybean and pea, with a stimulatory or an inhibitory effect depending on their concentrations (Taller and Sturtevant, 1991; Lorteau et al., 2001). Finally, accumulation of cytokinins in the supernodulating soybean mutant nts382 was higher after infection by rhizobia, compared to that of wild-type plants (Caba et al., 2000). More recently, promoter–GUS fusion studies in transgenic Lotus roots using an Arabidopsis cytokinin primary response gene (the Arabidopsis Response Regulator ARR5) have revealed that this promoter is rapidly activated after rhizobial infection in curled root hairs, dividing cortical cells, and at the base of LR primordia (Lohar et al., 2004). These data suggest a local accumulation of cytokinins and/or activation of cytokinin signaling during LR formation and upon rhizobial infection. Moreover, reduction of endogenous cytokinin content in Lotus transgenic roots overexpressing a heterologous cytokinin oxydase gene leads to reduced nodulation and enhanced LR formation. Several Medicago homologs of Arabidopsis cytokinin signaling genes are significantly upregulated a few hours after rhizobial infection, including a receptor kinase similar to AHK4/CRE1 (Arabidopsis Histidine Kinase 4/Cytokinin Response 1) and a response regulator similar to ARR4 (Gonzalez-Rizzo et al., 2006; Lohar et al., 2006). Use of the promoter region of this CRE1-homologous gene fused to GUS revealed that this putative cytokinin receptor is expressed in tips of uninoculated roots as well as in LR primordia (Lohar et al., 2006). In roots inoculated by rhizobia, expression is detected in the infection zone and in some cortical cells of the mature root. Cytokinin signaling genes have recently been characterized in Medicago and RNAi of a specific cytokinin receptor (MtCRE1) leads to cytokinin-insensitive roots, increases LR formation and inhibits nodulation (Gonzalez-Rizzo et al., 2006). Early stages of the symbiotic interaction such as growth of infection threads in the epidermis and induction of cortical cell divisions are blocked. In addition, the hit1 (hyperinfected 1) Lotus mutant corresponding to a loss of function mutation in the MtCRE1 orthologous gene (Lotus Histidine Kinase 1, LHK1) shows a drastically inhibited nodulation associated with a proliferation of infection threads (Murray et al., 2007). A gain of function mutation in the same LHK1 cytokinin receptor yield plants developing spontaneous nodules in the absence of rhizobia (snf2; Tirichine et al., 2007), indicating that cytokinins are necessary and sufficient to induce cortical cell divisions and nodule organogenesis. A crosstalk between Nod factors and cytokinin signaling pathways has been identified: indeed, the cytokinin regulation of the early nodulin MtNIN expression depends on MtCRE1; conversely, rhizobial induction of certain cytokinin signaling Response Regulator (MtRR) genes is dependent on Nod factor signaling (Gonzalez-Rizzo et al., 2006). Collectively, these data suggest a role for local accumulation of cytokinins and/or regulation of cytokinin sensitivity in relation with root lateral organs

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Root Development development, with a negative role in LR formation, and a positive one during the early stages of the symbiotic nodule organogenesis (Frugier et al., 2008). 10.4.2.3 Ethylene Ethylene is involved in the control and coordination of a diverse range of plant growth and developmental processes. In legume roots, ethylene inhibits primary root elongation in pea and negatively affects lateral and adventitious roots formation (Clark et al., 1999). In addition, several aspects of legume symbiotic nodule development have been well documented to be controlled by this hormone. Ethylene production significantly increases in roots after infection with symbiotic rhizobia (Ligero et al., 1986), whereas exogenous applications of 1-aminocyclopropane-1-carboxylic acid (ACC), the ethylene precursor, decreases the effectiveness of the symbiotic interaction, resulting in decreased number of infection events and nodule primordia (Oldroyd et al., 2001a). Ethylene also negatively modulates Nod factor-triggered calcium signaling in root hairs and the induction of early nodulin genes. These results indicate that exogenous ethylene can downregulate early steps in the infection process at, or upstream, of calcium spiking. In addition, inhibition of endogenous ethylene synthesis or perception in indeterminate nodules revealed a role of this phytohormone in the determination of the spatial window for rhizobial infection in the root differentiation zone (Heidstra et al., 1997; Oldroyd et al., 2001a). Another role for ethylene in positioning nodule formation has been proposed in pea based on the expression pattern of the ethylene biosynthetic enzyme, ACC oxidase, which is expressed in pericycle cell layers in between protoxylem poles (Heidstra et al., 1997). This local production of ethylene could restrict spatially the induction of cortical cell divisions by Nod factors, therefore determining the position where nodule primordia can be formed. However, ethylene does not seem to affect nodulation in soybean (Lee and LaRue, 1992; Schmidt et al., 1999). In S. rostrata, ethylene has a positive effect on LRB nodulation under hydroponic growth conditions, playing a crucial role in Nod factor-induced cell death during infection pocket formation (D’Haeze et al., 2003). Genetic data further indicate that nodulation can be affected by perturbation of ethylene responses: sym5 in pea and skl in M. truncatula, the latter corresponding to an ortholog of the Arabidopsis ethylene INsensitive (EIN2) gene, are insensitive to exogenous application of ACC and are hyperinfected by rhizobia (supernodulators) (Ligero et al., 1991; Fearn et al., 1992; Penmetsa and Cook, 1997; Prayitno et al., 2006). Moreover, nodules are not always formed in front of xylem poles. Recently, it was shown that, in contrast to wild-type plants, rhizobia or ACC could not inhibit long-distance PAT from shoot to root in skl mutants (Prayitno et al., 2006). In the infection zone, auxin transport locally increases in this mutant, as well as the expression of MtPIN1 and MtPIN2 auxin efflux carrier encoding genes, which could lead to an increased nodulation. These results suggest that ethylene signaling modulates

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both long-distance and local auxin transport regulation, and, therefore affects the number of nodules formed. However, ethylene-insensitive mutants (ethylene response 1, etr1) of soybean show a wild-type nodulation (Schmidt et al., 1999) and in Lotus, heterologous expression of a dominant-negative mutated ethylene receptor leads to ethylene insensitive plants which show increased rhizobial infection and nodule primordia, changes in their spatial localization, but not to a hypernodulation phenotype (Nukui et al., 2004). This suggests that determinate nodule primordia may be controlled by an ethylene-independent autoregulation mechanism. Altogether, ethylene probably plays different roles in root growth and nodulation control. Positive or negative effects on nodulation are observed, mostly dependent on different types of nodulation. These various controls might involve directly ethylene signaling or the regulation of auxin transport as intermediate. 10.4.2.4 Abscisic acid Even though in some species (such as Arabidopsis), abscisic acid (ABA) acts as an inhibitor of LR formation (De Smet et al., 2003), it increases LR development in several nodulated legumes species, suggesting that the negative role of ABA on root branching is correlated with the ability to nodulate (Liang and Harris, 2005). Thus, ABA might represent one of the mechanisms involved in maintaining the balance between LRs and nodules in legume roots. Several studies have also shown that exogenous ABA application inhibits nodule formation in various legumes, such as pea (Phillips, 1971), soybean (Bano et al., 2002), white clover and Lotus (Suzuki et al., 2004). Observation of root hair infection events in white clover revealed that ABA blocked early steps of infection such as root hair deformation (Suzuki et al., 2004). Moreover, decreasing ABA levels by using specific inhibitors led to an increase in nodule number in white clover and Lotus (Asami et al., 2003). In addition, Medicago latd mutants (see Section 10.3.4) have a lower sensitivity to ABA, and this hormone could rescue the short-root phenotype but not the nodulation phenotype of these mutants (Liang et al., 2007). Altogether, these results suggest a possible negative role for ABA in the control of nodule number, and a positive role in LR formation in nodulated legumes. 10.4.2.5 Gibberellins Little is known about the role of gibberellins (GAs) in the development of lateral organs in root legumes. Inhibition of LR formation by GAs has been observed in pea roots (Goodwin and Morris, 1979), whereas induction of nodule-like structures on the roots of Lotus has been reported (Kawaguchi et al., 1996). GAs are also involved in infection pocket and infection thread formation during S. rostrata LRB nodulation in hydroponic growth conditions (Lievens et al., 2005). In contrast, RHC nodulation is inhibited in the presence of GAs in the same semiaquatic plant. Furthermore, pea mutants deficient in GAs exhibit a reduction in LR and nodule formation that could be

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Root Development complemented by exogenous application of GAs (Reid et al., 2004; Ferguson et al., 2005). These data suggest a role for GAs in both types of legume rootderived organogeneses. 10.4.2.6 Salicylic acid Salicylic acid (SA) is involved in diverse plant–pathogen interactions (Vernooij et al., 1994; Schaller, 2001) and can block rhizobia infection threads in alfalfa by mediating related hypersensitive responses and local negative autoregulation of effective infections (Vasse et al., 1993; Mart´ınez-Abarca et al., 1998). Moreover, SA can completely block indeterminate nodule development at the level of primordia formation in legume species such as pea, alfalfa and white clover (van Spronsen et al., 2003). In the case of determinate nodulation, results are contradictory, maybe because SA has a major inhibitory effect on rhizobial growth (Stacey et al., 2006b). A strategy to reduce endogenous levels of SA by expressing a salicylate hydoxylase (NahG) gene either in Lotus or Medicago led to an increase in the number of infections, the size of the infection zone, and nodulation. Interestingly, only transgenic lines of Lotus also showed an increased root growth. Altogether, both determinate and indeterminate nodulation involve SA to control infections, probably via the regulation of localized defense response pathways. 10.4.2.7 Brassinosteroids and jasmonic acid Brassinosteroids (BRs) have no direct effect on nodule number, but might influence shoot mechanisms involved in autoregulation of nodule numbers (Ferguson et al., 2005). Indeed, mutants deficient in BRs showed a reduction in nodulation, and grafting experiments suggested that these phytohormones influence a shoot mechanism that controls nodulation. Interestingly, a correlation between nodules and LR numbers has been observed in the lines affected in BRs metabolism. Application of methyl jasmonate (JA) on shoots strongly suppressed nodulation in Lotus and the har1 mutant (see Section 10.3.4; Nakagawa and Kawaguchi, 2006). JA inhibits early stages of nodulation, including infection thread formation and LjNIN expression, and also suppresses LR formation. In Medicago also, JA inhibits early stages of nodulation as well as root growth (Sun et al., 2006): in contrast to ethylene, a negative effect of JA on calcium spiking and on the frequency of calcium oscillations has been identified. This effect is amplified in the ethylene-insensitive mutant skl, indicating an antagonistic interaction between these two hormones for the regulation of early Nod factor signaling. These findings suggest that JA and/or its related compounds might as well participate in nodule autoregulation signaling and the control of legume root architecture. In conclusion, many hormones might be involved in rhizobial infection, nodule development, local or long-distance autoregulation mechanisms and/or root growth and LR formation. As plant hormones interact at various levels to form a signaling network, many points of crosstalk should exist

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between their metabolism, transport and signaling pathways, and also with the Nod factor pathway. In other words, crosstalks between phytohormonal pathways seem crucial to rapidly integrate environmental, developmental and biotic stimuli to adapt legume root architecture.

Acknowledgments We thank D. Meyer for artwork. S.G.-R. was the recipient of a grant from the Consejo Nacional de Ciencia y Tecnologia (CONACyT), Mexico. P.L. was the recipient of a grant from the Minist`ere de l’Education Nationale, de la Recherche et de la Technologie (MENRT), France.

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Root Development DMI3-dependent activation of genes related to a signal transduction pathway in roots of Medicago truncatula. Plant Physiol. 139, 1065–1077. Savour´e, A., Magyar, Z., Pierre, M., Brown, S., Schultze, M., Dudits, D. et al. (1994). Activation of the cell cycle machinery and the isoflavonoid biosynthesis pathway by active Rhizobium meliloti Nod signal molecules in Medicago microcallus suspensions. EMBO J. 13, 1093–1102. Schaller, F. (2001). Enzymes of the biosynthesis of octadecanoid-derived signalling molecules. J. Exp. Bot. 52, 11–23. Schauser, L., Handberg, K., Sandal, N., Stiller, J., Thykjær, T., Pajuelo, E. et al. (1998). Symbiotic mutants deficient in nodule establishment identified after T-DNA transformation of Lotus japonicus. Mol. Gen. Genet. 259, 414–423. Schauser, L., Roussis, A., Stiller, J. and Stougaard, J. (1999). A plant regulator controlling development of symbiotic root nodules. Nature 402, 191–195. ¨ Scheres, B., McKhann, H.I., Zalensky, A., Lobler, M., Bisseling, T. and Hirsch, A.M. (1992). The PsENOD12 gene is expressed at two different sites in Afghanistan pea pseudonodules induced by auxin transport inhibitors. Plant Physiol. 100, 1649– 1655. Schmidt, J.S., Harper, J.E., Hoffman, T.K. and Bent, A.F. (1999). Regulation of soybean nodulation independent of ethylene signaling. Plant Physiol. 119, 951–959. Schnabel, E., Journet, E.-P., de Carvalho-Niebel, F., Duc, G. and Frugoli, J. (2005). The Medicago truncatula SUNN gene encodes a CLV1-like leucine-rich repeat receptor kinase that regulates nodule number and root length. Plant Mol. Biol. 58, 809– 822. Schnabel, E.L. and Frugoli, J. (2004). The PIN and LAX families of auxin transport genes in Medicago truncatula. Mol. Genet. Genomics 272, 420–432. Searle, I.R., Men, A.E., Laniya, T.S., Buzas, D.M., Iturbe-Ormaetxe, I., Carroll, B.J. et al. (2003). Long-distance signaling in nodulation directed by a CLAVATA1-like receptor kinase. Science 299, 109–112. Singleton, P.W. and Bohlool, B.B. (1984). Effect of salinity on nodule formation by soybean. Plant Physiol. 74, 72–76. Smit, P., Limpens, E., Geurts, R., Fedorova, E., Dolgikh, E., Gough, C. et al. (2007). Medicago LYK3, an entry receptor in rhizobial nodulation factor signaling. Plant Physiol. 145, 183–191. Smit, P., Raedts, J., Portyanko, V., Debell´e, F., Gough, C., Bisseling, T. et al. (2005). NSP1 of the GRAS protein family is essential for rhizobial Nod factor-induced transcription. Science 308, 1789–1791. Souleimanov, A., Prithiviraj, B. and Smith, D.L. (2002). The major Nod factor of Bradyrhizobium japonicum promotes early growth of soybean and corn. J. Exp. Bot. 53, 1929–1934. Sprent, J.I. and Raven, J.A. (1992). Evolution of nitrogen fixing symbioses. In Biological Nitrogen Fixation, G. Stacey, R.H. Burris and H.J. Evans (eds). New York, Chapman and Hall, pp. 461–496. Ssali, H. and Keya, S.O. (1983). The effect of phosphorus on nodulation, growth and dinitrogen fixation by beans. Biol. Agric. Hortic. 1, 135–144. Stacey, G., Libault, M., Brechenmacher, L., Wan, J. and May, G.D. (2006a). Genetics and functional genomics of legume nodulation. Curr. Opin. Plant Biol. 9, 110–121. Stacey, G., McAlvin, C.B., Kim, S.-Y., Olivares, J. and Soto, M.J. (2006b). Effects of endogenous salicylic acid on nodulation in the model legumes Lotus japonicus and Medicago truncatula. Plant Physiol. 141, 1473–1481.

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Annual Plant Reviews (2009) 37, 288–324 doi: 10.1002/9781444310023.ch11

www.interscience.wiley.com

Chapter 11

EFFECT OF NUTRIENT AVAILABILITY ON ROOT SYSTEM DEVELOPMENT ´ Alfredo Cruz-Ram´ırez, Carlos Calderon-V´ azquez and Luis Herrera-Estrella Nacional Laboratory of Genomics for Biodiversity, CINVESTAV Campus Guanajuato, Km. 9.6 Carretera Irapuato-Le´on Z.C. 36500, Irapuato, Guanajuato, M´exico

Abstract: Modern-day plant root systems have been modeled from distinct ancestors and by different evolutionary pathways influenced by environmental cues, leading to varied root system architectures. Availability of mineral nutrients has played a major role in evolution and development of the diverse root system architectures. In this chapter, we review the importance of the study of mineral nutrition on root development and describe the physiological, morphological and molecular responses of plant roots to diverse nutrient deficiencies and how the different hormone signaling pathways participate in these responses. Keywords: plasticity

11.1 11.1.1

adaptation;

nutrient starvation;

root architecture;

phenotypic

Introduction Evolution and diversity of root systems

Evolutionary adaptation is an intrinsic response to environmental change. In the course of geological time, the environments in which plants grow have changed continuously. Under such unstable conditions, plants have evolved and developed phenotypic plasticity as a valuable capability that allowed them to adapt to the diverse physical and chemical environments they have faced during the last 400 million years. Fossil findings point to lycopods as the earliest vascular plants with a root-like organ; however, they did not succeed and became extinct in the Devonian period (Gensel and Edwards, 2001). Thus, 288

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adaptation of ancient plants to a terrestrial existence involved overcoming several challenges through a myriad of physiological and morphological changes during thousands of years. Early land plants, such as bryophytes and embryophytes, were faced with limitations in their photosynthetic capacity and many other physiological and morphological disadvantages to ensure the colonization of the land. We know that this struggle was successful: modern plants have achieved permanence and diversity (Hemsley and Poole, 2004). Root system evolution is a notable process that has led to a progressive transformation from the very simple root systems of early land plants to the diverse and complex root systems of the modern plants. Ancient plants did not have to face obstacles to acquire water, obtain nutrients, or efficiently anchor to the substrate, as long as they kept growing in moist environments. However, as plants began to be confronted with drier environments in their conquest of land, availability of nutrients represented an important challenge and, particularly, for their root systems, that had to secure nutrient and water uptake and to become a vigorous anchor for establishment in soils with diverse physical and chemical characteristics. To overcome such challenges, it is easy to speculate that two of the very early events in the evolution of root systems were root branching and root hair development, processes that involve the main phenomena studied by developmental biology: cell proliferation, and cell growth and differentiation. The influence of the environment on the genetic program regulating these cellular processes led to the origin of the elements of a root system and established the necessary arrangement in a spatial configuration, determining the root system architecture (Raven and Edwards, 2001). Modern-day plant root systems have been modeled from distinct ancestors and by different evolutionary pathways influenced by environmental cues, leading to varied root system architectures. For example, the root system of most gymnosperms and dicotyledonous plants is markedly regulated by mineral nutrient and water availability. These plants have developed a taproot system in which the embryonic root matures and becomes a primary root. From this primary root, or taproot, lateral roots emerge and the whole root system is formed. Under optimal environmental conditions, when nutrients and water are nonlimiting, the primary root grows continuously downward reaching a depth greater than that of the lateral roots. This growth pattern is called indeterminate and can be drastically altered by the scarcity of water and nutrients (Veit, 2004). In contrast to dicotyledonous plants, the primary root of monocotyledonous plants has a short life span and the root system develops from postembryonic stem-borne roots that, growing above or below ground, give rise to branches that build a root system, called fibrous, in which no specific root grows preferentially longer than the others. In fibrous root systems most stem-borne roots develop underground, but in some species, such as maize (Zea mays), roots are produced from above ground structures, generally named aerial or prop roots. Prop roots stabilize the main stem and are also

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Root Development capable of branching and absorbing nutrients and water (Hochholdinger et al., 2004; see also Chapter 7). A remarkable example of how environmental cues influence root-structure modeling is the root system of desert plants. Agave and cactus plants grow in arid and semiarid regions where water uptake is a crucial challenge for root systems. Under such environmental conditions, exploration of the upper soil layers must be efficient not only to seek for and obtain nutrients, but also to acquire the scarce and only occasionally available water. There are evident differences among the root systems of cacti and the taproot and fibrous root systems. In the taproot system, the primary root continuously grows because its meristem follows an indeterminate growth program, whereas agaves and cacti have a short-lived primary root that is due to a particular growth pattern in which root meristematic cells divide for only a few days after germination and then differentiate (Dubrovsky, 1997). In other words, the primary root meristem enters a determinate pathway of root growth that has a remarkable ecological significance (reviewed in Shishkova et al., 2008). Upon arrest of the primary root, pericycle cells are activated and become founder cells from which lateral roots develop rapidly forming lateral roots that grow in shallow depths to make water uptake more efficient and increase the survival potential of plants that experience drought for prolonged periods. Although the root systems of cacti are characterized by short-lived primary and lateral roots, they do not develop a fibrous root system. Cacti and agaves develop a multiple shallow root system that grows vertically and gives rise to usually horizontal lateral roots that branch in the same direction, producing a root system that extends in a radial pattern. Although diverse root types can initiate from different tissues during embryonic and postembryonic development, their stem or initial cells have numerous common characteristics, among which are the location of the apical stem cells within the meristem, their position-dependent identity, and their infrequent and polarized division (Nakajima and Benfey, 2002). Fossil registers of fungi associated with plants demonstrate that the occurrence of fungal–plant symbioses are as old as early land plants, indicating that this type of association played a fundamental role not only in the conquest of land by plants but also in root system evolution. These symbiotic interactions with microorganisms present in the soil facilitate nutrient uptake and are proposed to have played an important role in the adaptation of early land plants (Barker and Tagu, 2000; see also Chapter 9). The vegetative mycelium of mycorrhizal fungi can establish a mutualistic symbiosis with root tips, allowing plants to efficiently grow in soils with suboptimal nutrient availability. Three types of associations are found in most annual and perennial plants: arbuscular endomycorrhiza (AM), ectomycorrhiza (ECM) and ericoid associations. The impact of the association on root development depends on the type of fungi. Ectomycorrhizal fungi surround the root cells, whereas endomycorrhiza penetrate them to form the so-called ‘vesicular–arbuscular’ structures in the symbiotic root (Barker and Tagu, 2000). Vesicular–arbuscular

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structures consist of fungal hyphae surrounded by host plant plasma membranes and profoundly influence root development and architecture. The so-called extramatrical and intraradicular hyphal networks are metabolically active and provide essential nutrient resources to the host plant in exchange for photosynthates that serve as a carbon source to support fungal growth. Mycorrhiza-generated structures evolved as nutrient and water-absorbing organs in 90% of vascular plants and are not just peripheral to the architecture of root systems. Nevertheless, the fact that diverse vascular plants do not establish symbiotic associations with mycorrhiza and that most plant species are capable of responding to nutrient stress conditions even in the absence of mycorrhizal fungi, suggests a parallel evolution of adaptive strategies to cope with nutrient deficiencies through their own biochemical, physiological and morphological resources. In this chapter, we will focus on the developmental responses of plant root systems to low nutrient availability. 11.1.2

Historical perspective

The naturalists of Ancient Greece believed that plants drew their nutrients from the soil but never proved their conclusion by any experimental evidence. This state of ignorance persisted during the Middle Ages and The Renaissance (Saltini, 1984). It was not until the seventeenth and eighteenth centuries that European naturalists reported the first experimental findings to support what had been empirically known for two millennia: the essential role of mineral nutrition in plant development. The work of Stephen Hales, considered the first plant physiologist, is of absolute importance to our understanding of the fundamental processes, such as water and solute transport. In his book Vegetable Staticks (1727), Hales considered transpiration as a function that regulates the transport of water and the substances dissolved in it from roots to leaves and he presumed the existence of a transport mechanism for certain ‘substances’. It was not until the first half of the nineteenth century that Karl Sprengel and Justus von Liebig, who focused their research on uncovering some ‘indispensable mineral salts’, demonstrated that certain elements are absolutely required for plant growth and development. They influenced the concept known as ‘essentiality’ of an element. Liebig, considered one of the pioneers of agricultural chemistry, was also the promoter of the so-called ‘law of the minima’ (reviewed in Penazzio, 2005). In 1860, the botanist Julius von Sachs carried out an aquatic procedure similar to modern hydroponic cultures and introduced the first standard formula of liquid culture medium that included all the mineral salts believed to be necessary for plant growth. After Sachs and until 1900, other scientists documented the essentiality of carbon, hydrogen, oxygen, nitrogen, calcium, iron, magnesium, sulfur, phosphorus and potassium. In 1939, Arnon and Stout refined essentiality as a concept in plant biology and proposed to consider an element as ‘essential’ when it accomplishes three basic principles: (i) in its

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Root Development absence, plants must be incapable of continuing normal growth; (ii) its biological functions cannot be carried out by any other element and (iii) it must be directly involved in plant metabolism. These basic rules have been successfully used to document the disorders produced by nutrient deficiency and established mineral nutrition of plants (reviewed in Penazzio, 2005). The essentiality of macronutrients for plant metabolism and growth was discovered early in the discipline’s history but the concern of plant biologists in unraveling the biochemical, morphological and physiological phenomena related to plant mineral nutrition was only modestly explored until approximately 30 years ago. It was only after 1950 that research on plant mineral nutrition considerably developed when Epstein and Hagen (1952) explored ion uptake mechanisms in excised roots of barley (Hordeum vulgare) using radioisotopes. In addition, remarkable studies have also been reported for potassium (Cooper et al., 1967; Satter and Galston, 1971; Hawker et al., 1974), nitrogen (Goldbach et al., 1975), phosphorus (Heldt et al., 1977) and sulfur (Vange et al., 1974; Rennenberg et al., 1979). Reports of diverse research groups in the early 1970s illustrate the interest in understanding the effects of nutrient availability on root system development and its impact on plant productivity (Bar-Yosef, 1971; Riley and Barber, 1971; Drew et al., 1973). Since then plant biologists study intensively the effects caused by macronutrient and micronutrient availability, metabolism and transport. Currently, this research field has generated several powerful tools that can be employed to unravel in detail the molecular and developmental mechanisms involved in the diverse responses under study.

11.2

11.2.1

Regulation of the root system architecture by nutrient availability Phosphate

11.2.1.1 A general response Phosphate (Pi) has a fundamental role in most developmental and biochemical processes in plants. It is not only a constituent of key cell molecules, such as ATP, nucleic acids and phospholipids, but it is also an essential metabolic regulator of diverse processes, such as protein activation, energy transfer, and carbon and nitrogen metabolism. Moreover, Pi availability represents one of the major constraints for growth and development of terrestrial plants in both natural and agricultural ecosystems, due to its low mobility and high absorption capacity in the soil. The environmental challenge posed by Pi availability in the soil was a major selective pressure for plants to evolve a range of developmental, biochemical and symbiotic strategies to adapt to Pi deprivation (Lynch, 1995; Raghothama, 1999). In both monocotyledonous and dicotyledonous plants, a general strategy to cope with low Pi availability

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293 (a)

(b)

(c)

(d)

(e)

Figure 11.1 Root architecture of Col-0 Arabidopsis plants in response to optimal nutrient conditions and diverse nutrient deficiencies: (a) optimal nutrient conditions, (b) phosphate, (c) nitrogen, (d) sulfate and (e) potassium deficiencies.

has been described that involves three fundamental mechanisms: (i) release and uptake of Pi from external organic and inorganic sources (Bariola et al., 1994; Rubio et al., 2001), (ii) optimization of Pi utilization by a wide range of metabolic alterations and mobilization of internal Pi and (iii) increase in the exploratory capacity of the root and the absorptive surface area by alteration ´ ´ et of the root system architecture (Lopez-Bucio et al., 2003; S´anchez-Calderon al., 2005; see Fig. 11.1). The first mechanism involves biochemical responses directed to augment soil Pi availability by increasing Pi-uptake capacity through the induction of high-affinity Pi transporters and Pi recycling and mobilization through the induction of endogenous and secreted acid phosphatases and RNases and the increased excretion of organic acids (Raghothama, 1999). The second mechanism includes the utilization of alternative glycolytic pathways that involve Pi-independent enzymes (Duff et al., 1989), changes in carbohydrate metabolism, and the hydrolysis of phospholipids to release Pi for other metabolic processes and their replacement for nonphospholipids, such as sulfolipids and galactolipids (Essigmann et al., 1998; Cruz-Ram´ırez et al., 2006). Recently, microarray analyses in Arabidopsis (Arabidopsis thaliana) demonstrated or confirmed that genes involved in several of the previously described processes are transcriptionally upregulated by Pi starvation. From 732 differentially regulated genes, 501 genes, including both novel and previously reported Pi starvation-induced genes, are upregulated and 231 are downregulated. Genes involved in lipid biochemical pathways, such as

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Root Development phospholipid hydrolysis and galacto- and sulfolipid synthesis, are upregulated by Pi deprivation (Misson et al., 2005). 11.2.1.2 Main adaptations of root system architecture To face Pi stress, diverse wild and cultivated plant species adapt their root developmental programs toward the formation of shallow, highly branched root systems that increase the soil exploratory capacity of the plant (Forde and Lorenzo, 2001). Pi-dependent root architecture alterations have been studied in diverse crop species, such as maize (Zea mays), rice (Oriza sativa) and common bean (Phaseolus vulgaris). Despite divergence, most of these root systems experience an increase in adventitious root production and lateral root density under Pilimiting conditions. Under natural conditions, such alterations are thought to be directed to maximize Pi acquisition as the nutrient becomes more limiting in the soil (Lynch and Brown, 2001). Several of the changes caused by Pi starvation on the root architecture have been well characterized in the model plant Arabidopsis. This plant develops a taproot system that experiences three fundamental alterations upon Pi deficiency: a reduction in primary root growth, an increase in lateral root development and more and longer root hairs (see Fig. 11.1). 11.2.1.3 Postembryonic effects of Pi availability on the root meristem Determinate root development has been reported as a stable program of certain desert cacti (Dubrovsky, 1997; Rodr´ıguez-Rodr´ıguez et al., 2003). In most species, this pattern of root development appears to be triggered by specific biotic and abiotic factors, as in the case of Grevillea robusta, in which the formation of clusters of short determinate roots or proteoid roots is induced by Pi deprivation (Skene et al., 1996). In the case of Arabidopsis, the reduction in primary root growth suggests that Pi stress has an effect on individual cell length and in controlling the proliferation of initial cells at the primary ´ root meristem (Williamson et al., 2001; Lopez-Bucio et al., 2002). Recently, Pi deprivation has been shown to alter cell division and elongation of stem cells and their derivatives and to induce an irreversible shift from indeterminate to determinate growth program in which the quiescent center (QC) ´ et al., 2005). By using the QC identity plays a central role (S´anchez-Calderon marker QC46, the earliest alteration observed in Pi-deprived seedlings was the periclinal cell divisions of QC cells, which occurred at day 2 of growth in Pi-deficient medium and preceded changes in cell elongation and meristem size. A reduced mitotic activity was also observed using the CycB1,1::uidA reporter gene. In conclusion, roots of stressed plants progressively lose stem cells in the meristem as a consequence of decreased cell division events and premature cell differentiation. Also, in crop species, such as rice and maize, Pi starvation significantly affects the total root length of both primary and lateral roots, although detailed analyses are needed to establish whether determinate growth is induced by Pi starvation (He et al., 2003).

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Figure 11.2 Proliferative capacity of primary and lateral root meristems altered by Pi starvation. PLETHORA 1 (PLT1) and PLETHORA 2 (PLT2) are essential transcription factors for QC identity and stem cell niche maintenance. Both genes are positively regulated by auxins via auxin responsive factors (ARFs), such as ARF19 and ARF7, during embryonic and postembryonic developmental stages (Aida et al., 2004). A hypothetical model for lateral roots is that low Pi availability may alter auxin perception and induce ARFs by affecting AUX/IAA proteins, such as IAA14, which are negative regulators of ARFs. In the primary root Pi starvation may modulate ARFs in a negative way through effects on auxin concentration, transport or redistribution. In both cases the expression of ARF-regulated genes such as PLT1 and PLT2, may be altered and in consequence the proliferative capacity of the root meristems modified.

In Arabidopsis, the maintenance and development of the primary root meristem apparently requires an optimal auxin level that correlates with an auxin response maximum in the meristematic zone. Sabatini et al. (1999) demonstrated that alterations in this maximum lead to abnormalities in cell fate and meristem organization. In Arabidopsis seedlings subjected to Pi starvation, the auxin maximum decreases and this alteration correlates with the loss of ´ et al., 2005). Nevertheless, further studies are QC identity (S´anchez-Calderon needed to determine whether this phenomenon is caused by alterations in auxin synthesis, transport, or sensitivity of root meristem cells to this hormone (Fig. 11.2). One possible pathway through which Pi and auxins might regulate QC maintenance and root meristem activity could be through transcription

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Root Development factors required for the integrity and maintenance of the QC, such as those encoded by the PLETHORA (PLT) genes. The transcriptional factors PLETHORA 1 (PLT1) and PLETHORA 2 (PLT2) of the AP2 class, which are essential for determining QC identity and maintenance of the stem cell niche, are positively regulated by auxin via auxin responsive factors (ARFs) during embryonic and postembryonic developmental stages (Aida et al., 2004). The phenotype of the primary root meristem of seedlings from the double mutant plt1-4 plt2-2 closely resembles the meristem exhaustion caused by Pi starvation in wild-type seedlings. It is possible that low Pi availability might alter root meristem development by modulating the expression of ARFs and PLT genes. Nevertheless, it cannot be ruled out that other specific transcription factors, whose expression is regulated by ARFs, might be involved in ´ ´ the meristem exhaustion process caused by Pi starvation (A. Chacon-L opez, B. Scheres and L. Herrera-Estrella, unpublished data; see Fig. 11.2). 11.2.1.4 Low Pi availability and lateral root development Root branching extends the capacity of root systems to explore the soil and interact with beneficial soil microorganisms. Diverse plant species adapt their growth in soils with low amounts of available Pi through changes in root branching patterns. The root system of Lupinus albus, a legume that is well adapted to infertile soils, forms clusters of short lateral roots (LRs) that arise from the pericycle. Such cluster roots, also called proteoid roots, are specialized in Pi uptake. After a few days of growth, proteoid roots enter a determinate developmental program in which proliferative cells differentiate and numerous root hairs are formed (Johnson et al., 1996). The increased Pi uptake capacity of proteoid roots compared to roots with indeterminate growth is provided by the increased absorptive surface and exudation of organic acids and phosphatases (Neumann and Martinoia, 2002). A similar phenomenon was reported for Fraxinus mandshurica Rupr. seedlings (Wu et al., 2004), in which the elongation rates of primary roots increased as phosphorus availability augmented while the elongation rate of lateral roots decreased. These findings suggest that high Pi availability inhibits the formation and elongation of LRs. In Arabidopsis plants, Pi-limiting conditions result in an increase in LR over ´ primary root growth (Williamson et al., 2001; Lopez-Bucio et al., 2002; Nacry et al., 2005; see Fig. 11.1). The increase in emergence and rate of growth of LR primordia as a response to Pi starvation is a phenomenon that involves a fundamental developmental process: the early establishment and activation of mature LR primordia. One of the major questions is how the existing signaling pathway works between Pi starvation sensing and lateral root development. Diverse studies support the notion that effects of Pi shortage on lateral roots are tightly related to auxin sensitivity and/or local concentration, re´ distribution and signaling (Lopez-Bucio et al., 2002; Malamy, 2005; Nacry et al., 2005). Several aspects of the response of the Arabidopsis root system to low Pi have been shown to be triggered by local changes in auxin concentration

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(Nacry et al., 2005). However, increase in lateral root formation might also be due to an increase in the sensitivity to auxin of the roots of Pi-deprived plants. Whether local changes in auxin distribution or an increase in auxin sensitivity or both are responsible for the increase in lateral root formation under Pi starvation remains to be clarified. Cell cycle reactivation in the pericycle is a stage of LR development that depends on auxin (Casimiro et al., 2001, 2003; Himanen et al., 2002). Therefore, polar transport and local concentration of auxins have a direct effect on cell cycle and stem cell proliferation of initial cells in root meristems. Recent studies have unraveled one of the molecular mechanisms that link auxins to stem cell specification in the root. The PLETHORA genes (PLT1 and PLT2), key factors in root stem cell patterning, are regulated in response to auxin treatment and mutations affecting auxin signaling such as mutations in PIN proteins and ARFs can alter their expression. Thus, it is possible that local changes in auxin distribution or sensitivity to auxins in pericycle cells could alter the expression of PLT genes giving rise to a higher degree of lateral root formation in Pi-deprived Arabidopsis plants (see Fig. 11.2) (Aida et al., 2004; Blilou et al., 2005). Recently, the IAA14 protein has been shown to physically interact specifically with ARF7 and ARF19, suggesting that this binding inhibits ARF7/ARF19 function (Fukaki et al., 2005). Plants of both solitary root-1 (slr-1) (a gain-of-function mutant in IAA14) and the double mutant arf7/arf19 are severely affected in their capacity to form LRs. Therefore, auxin signaling as well as LR primordia formation and cell cycle progression appear to be mediated by ARFs and are probably components of the low Pi signaling pathway that regulates LR formation (Fig. 11.2). After prolonged Pi starvation, the primary and lateral roots of Arabidopsis seedlings experience exhaustion of their root meristem. Expression of the genes ATPT1 and ATPT2 that encode the high-affinity Pi transporters is dramatically increased and acid phosphatases are located at the exhausted root tip, suggesting that the developmental program induced by Pi starvation enhances the Pi uptake capacity of the Arabidopsis root tip by inducing expression of genes involved in Pi uptake. Thus, Pi deficiency exhausts lateral root meristems by triggering a determinate growth program that converts differentiated meristems into organs with a high capacity to express genes involved in Pi scavenging. These phenomena suggest that exhaustion of primary and secondary root meristems is a mechanism through which the Arabidopsis root system induces root branching and develops root architecture elements specialized in excreting enzymes to free Pi from external sources and internalize it by the action of high-affinity transporters. 11.2.1.5 Effects on cell fate and differentiation of epidermal cells Root hairs are important structures of the root system with a major role in water and nutrient acquisition. The limited availability of Pi in the soil makes necessary a strategy to increase the absorptive surface for Pi acquisition.

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Figure 11.3 Root hair patterning and development in response to nutrient stress (a–c). Radial structure of the Arabidopsis root at the differentiation zone under (a) optimal growth conditions, (b) Pi starvation and (c) iron stress. Photographs of the root hair density of Arabidopsis wild-type plants grown under (d) optimal nutrient conditions, (e) phosphate deficiency and (f) iron starvation. H, root hair cells.

Root hairs contribute with up to 80% of the surface area of the root and are the main site of Pi uptake in species that do not establish associations with mycorrhiza (Jungk, 2001). For diverse plant species, one of the most evident effects of low Pi availability on root development is the alteration of these epidermal cell structures (Bates and Lynch, 1996; Lynch and Brown, 2001; Ma ¨ et al, 2001; Zhang et al., 2003; Muller and Schmidt, 2004). In Arabidopsis roots, Pi availability regulates the elongation and density of root hairs, suggesting a phenomenon directed to improve Pi acquisition in response to an uneven ´ distribution of the nutrient in the soil (Fig. 11.3; Lopez-Bucio et al., 2003; ¨ Muller and Schmidt, 2004). Root hair formation takes place at the differentiation zone of the root, which under normal conditions is located 1–2 mm above the root tip whereas under low Pi conditions it is localized very close to the meristematic zone. The molecular basis determining the position of the differentiation zone in response to environmental cues remains to be determined; however, several of the components that regulate the specification of root hair cells and how certain genetic and physiological determinants interact to induce epidermal cells to form a root hair have been characterized in Arabidopsis thaliana. From diverse studies the hair (H)/nonhair (N) cell fate has been determined at early stages of the epidermis cell layer formation. As soon as the division of their initials in the meristem produces epidermal cells, cells destined to become a root hair can be identified by cytological differences, including a denser cytoplasm. This early determination involves cell-to-cell

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communication not only between immature H and N cells but also with the cortical cells adjacent to them; however, little is known about the specific signals that mediate this communication. Fundamental gene products associated with root hair cell determination include the homeodomain protein GLABRA 2 (GL2) (Masucci et al., 1996), the MYB transcriptional factor WEREWOLF (WER) (Schiefelbein, 2003) and TRANSPARENT TESTA GLABRA (TTG), which encodes a WD-repeat protein. Mutants at these loci develop extra root hairs, suggesting that they negatively regulate root hair formation. Two additional transcriptional factors have been determined that determine the N cell type, the bHLH proteins GLABRA 3 (GL3) and ENHANCER OF GLABRA 3 (EGL3) (Bernhardt et al., 2003). Each of these mutants fails to specify the N cell fate and produces ectopic root hairs. Although the wer, gl2, gl3, egl3 and ttg mutants share a common characteristic in their phenotypes (extra root hairs), they differ in the way they affect the H or N cell fates. Results from mutant and reporter analyses suggest that disruptions in WER, GL3, EGL3 and TTG expression affect all aspects of N cell differentiation because plant mutants at these loci form immature epidermal cells. Mutations in GL2 have a later effect on differentiating epidermal cells affecting only the final cell morphology of N cells, which become H cells. In fact, GL2 expression is drastically reduced in ttg and gl3 mutants and completely abolished in the ttg and gl3-egl3 double mutant. Taken together, this evidence indicates that WER, GL3, EGL3 and TTG positively regulate GL2 expression and that all affect the N cell fate. In contrast to GL2, GL3, EGL3, TTG and WER, the small one-repeat MYB proteins CAPRICE (CPC) and TRYPTICHON (TRY) help to determine the H cell fate. Mutants at both CPC and TRY loci show a reduction in root hair formation, implying that CPC and TRY positively regulate H cell differentiation (Wada et al., 2002). Once H cell fate is established, the root hair growth process starts to form a bulge by cell wall expansion. The swelling process implies cytoskeletal rearrangements and the activity of cell wall-modifying enzymes, such as cellulases and expansins. 11.2.1.6 Changes in root hair patterning under Pi-deficient conditions The fact that Pi deprivation increases root hair density suggests that two developmental processes might be affected: cell fate and/or positional signaling (Schiefelbein, 2003). As previously mentioned, H and N epidermal cell fate are fixed in the root meristem before root hairs are formed. However, the position of the epidermal cell in relation with cortical cells also determines their final destiny. Some studies point to the possibility that low Pi induces the formation of ectopic root hairs because of an alteration in the positional arrangement of epidermal cells. Pi starvation affects radial patterning of the root by reducing the size and number of cortical cells, which disorders the position of epidermal cells in contact with cortical cells, generating more epidermal

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Root Development cells located in the H position (Ma et al., 2001; Zhang et al., 2003) (Fig. 11.3). However, it has also been observed that the increase in root hair density in Pi-starved plants includes the formation of extra hairs over both H and N ¨ positions (Muller and Schmidt, 2004). The formation of extra root hairs could be related to auxin efflux, since in response to low Pi availability eir1-1, a mutant impaired in auxin efflux, shows a slight increase in cortical cell number when compared with the wild type. Although there is no conclusive evidence about the effect of Pi starvation on the key transcriptional factors that regulate epidermal cell fate, it is possible that Pi availability regulates the expression or function of the negative or positive molecular determinants of the H cell fate. In the aging primary roots of the wer, ttg and gl2 mutants, all mutated in negative regulators of root hair fate, root hair density increases in a similar way to that of the wild type when grown under Pi deficiency conditions. This is in contrast with the dramatic phenotype of this group of mutants that form ectopic root hairs. Furthermore, this group of genes is more important for the fate of cells in the N position than for adjusting the number of hairs in the ‘normal’ position suggesting that the effects of Pi starvation on root hair development is not only a consequence of ¨ alterations in the positional pattern but also in cell fate (Muller and Schmidt, 2004). In an attempt to elucidate the mechanism by which low Pi availability is perceived by the plant and translated into alterations in root hair number and size, Zhang et al. (2003) tested whether ethylene mediates the responses of root hair growth and development to low Pi availability. In plants grown in high Pi-containing media supplemented with the ethylene precursor 1aminocyclopropane-1-carboxylate (ACC), a response in root hair length similar to that of low Pi availability was observed. On the other hand, elongation of root hairs of Pi-starved plants was reduced when inhibitors of ethylene synthesis were added to the media. In agreement with these findings, the root hair length phenotype of ethylene response mutants depended on Pi availability. This illustrates the existence of a crosstalk between ethylene and Pi in the root hair elongation process induced by Pi starvation and shows the essentiality of ethylene in this phenomenon. Other lines of evidence have revealed the essential function in root hair development of phospholipid-derived molecules, which are intracellular messengers that mediate a myriad of cellular processes, such as cell elongation and membrane trafficking. These signaling lipids include phosphatidic acid (PA) that is the metabolic product of the hydrolization of glycerophospholipids by phospholipases D (PLDs) (Wang, 2004). PA has recently emerged as an important regulator of root hair elongation and epidermal cell integrity. The root hair negative regulator GL2 was found to repress the expression of the PHOSPHOLIPASE D Z1 (PLDZ1) gene of Arabidopsis (Ohashi et al., 2003). Negative regulation of PLDZ1 in transgenic Arabidopsis plants by RNA interference caused formation of globular-shaped root hairs whereas its overexpression led to the ectopic development of root

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hairs. Since PLDZ1 produces PA, using phosphatidylcholine (PtdCho) as a specific substrate, it is possible that GL2 regulates root hair development by modulating PA production. In agreement, the decrease in PtdCho and PA contents causes severe alterations in root development as revealed by the analysis of the xipotl (xpl) Arabidopsis mutant. This is an insertional mutant in a gene encoding an S-adenosyl-L-methionine:phosphoethanolamine Nmethyltransferase (PEAMT), an enzyme that catalyses the triple, sequential N-methylation of phosphoethanolamine (PEA) that are critical steps in phosphocholine (PCho) and phosphatdylcholine (PtdCho) biosynthesis. Consistently with a major role for phospholipids in root hair development, xpl forms globular-shaped root hairs and epidermal cells die at certain developmental stages. Interestingly, normal root hair formation was restored by exogenous addition of PA to the media, suggesting that phenotypical alterations in root hairs are caused by deficient PA production (Cruz-Ram´ırez et al., 2004). Recently, PLDZ1 and PHOSPHOLIPASE D Z2 (PLDZ2) were reported as regulated by Pi starvation (Cruz-Ram´ırez et al., 2006; Li et al., 2006), directly linking phospholipid degradation with Pi starvation responses. However, future studies are needed to elucidate the role of PA as a molecular messenger, rather than as a transient metabolic precursor, in root hair elongation in response to low Pi availability. 11.2.1.7 Novel regulators in the Pi starvation response As previously described, Pi deficiency responses are regulated at the transcriptional level, confirmed by microarray techniques (Misson et al., 2005). Promoters of the Pi-responsive genes encoding Pi transporters, phosphatases and genes involved in lipid recycling such as PLDZ1 and PLDZ2 are significantly enriched with the motif known as phosphate-binding sequence (PBS); it has been experimentally demonstrated that this motif is recognized by the Phosphate Starvation Response 1 transcription factor (PHR1), a member of the MYB coiled–coiled transcription factor family. PHR1 was mapped and isolated through screening of a mutagenized population from a transgenic line in which the GUS gene was under the control of the promoter of a Pi starvation-responsive gene that contains PBS motifs. To date, PHR1 is the only well-defined transcriptional regulator of Pi starvation responses (Rubio et al., 2001). Recently, Miura et al. (2005) have established that PHR1 is a sumoylation target of a small ubiquitin-like modifier, the Arabidopsis SUMO E3 ligase encoded by AtSIZ1, suggesting that AtSIZ1 is a negative regulator of the low Pi-induced responses controlled by PHR1, such as upregulation of highaffinity transporters and acid phosphatases. Nevertheless, AtSIZ1 also alters Pi starvation responses that are not under the control of PHR1, indicating that sumoylation might be a broader process in the control of adaptation to Pi deficiency. In agreement with this hypothesis, allelic mutants in AtSIZ1 (At5g60410) mimic several of the phenotypical responses of the root to Pi starvation, such as alteration of the proliferative capacity of the primary root meristem, lateral root branching increase and alterations in root hair density

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Root Development and length. In other words, AtSIZ1 might be a critical regulator of the Pi starvation response and root system development and sumoylation might be a regulatory process with an important role in root phenotypic plasticity. Recent findings in Arabidopsis (Fujii et al., 2005) indicate that microRNAs (miRNAs) participate in the regulation of the Pi starvation response. The Arabidopsis pho2 mutant shows a two- to fourfold increase of Pi in leaves and unaltered levels in the root, suggesting that the PHO2 participates in Pi homeostasis. The pho2 is caused by a nonsense mutation in a gene coding for an unusual ubiquitin-conjugating E2 enzyme (UBC), which is the target of the miR399. This miRNA is complementary to the 5
UTR of the PHO2 mRNA in five different positions. Transcription of miR399 is strongly induced by Pi starvation, whereas the levels of PHO2 mRNA decrease concomitantly to the increase of mi399 (Aung et al., 2006; Bari et al., 2006). Transgenic Arabidopsis plants with constitutive expression of miR399 mimic the pho2 mutation and show an overaccumulation of Pi in the shoot due to a decrease of PHO2 mRNA when compared to the wild-type control (Aung et al., 2006; Bari et al., 2006). More recently, mi399 was reported to be present in the phloem sap of rapeseed and pumpkin, where it is specifically increased during Pi deprivation, suggesting a potential role of mir399 in long-distance Pi signaling (Pant et al., 2008). Using reciprocal Arabidopsis micrografting between wild-type and mi399 overexpressing plants, two groups were able to demonstrate that mi399 moves from the shoot of overexpressing plants to the root of wildtype plants and that this movement leads to a high accumulation of Pi in the shoot because the level of PHO2 is reduced in the wild-type rootstock (Lin et al., 2008; Pant et al., 2008). These experiments show that mi399 is an important phloem-mobile, systemic signal that participates in Pi homeostasis at the whole plant level. Related to this regulatory network, it has been shown that the Pi starvationinduced gene INDUCED BY PHOSPHATE STARVATION 1 (IPS1) a nonprotein coding gene, contains a conserved 23 nt motif, present in the members of this gene family in different plant species, that is complementary with the sequence of miR399 (Franco-Zorrilla et al., 2007). This study demonstrates that IPS mRNA is not a target but a sequester of miR399 and plays a role in modulating the degree of degradation of PHO2 trascripts displayed by miR399 under Pi starvation conditions. This elegant work shows a very sophisticated regulatory network displayed by Arabidopsis as a response to Pi starvation that regulates Pi homeostasis; however, more detailed analyses on the effects of the deregulation of IPS, miR399 and/or PHO2 on the root system architecture are needed. 11.2.2

Nitrate

11.2.2.1 Main adaptations of root system architecture Nitrogen is one of the most important elements for plant mineral nutrition and is mainly present in soils in the form of nitrate (NO3 − ) and ammonium (NH4 + ). However, the availability of these compounds is frequently

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poor because of microbial competition or because of the low and heterogeneous concentrations of the nutrient in the soil. Nitrogen limitation affects vital metabolic processes related to energy in plants such as photosynthesis and respiration (de Groot et al., 2003). As a consequence of alterations in photosynthetic structures that are rich in nitrogen, such as chlorophyll and the protein ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco), CO2 fixation severely decreases under low nitrogen conditions (Evans and Poorter, 2001). In order to deal with N deficiency, plants establish symbiotic and nonsymbiotic relations with soil microorganisms, such as bacteria or fungi, that provide the nutrient to root systems in the form of amino acids, amides and ureides (Owen and Jones, 2001). Another general response of plant root systems to N deficiency is the increase in root surface and size (total mass, length and area) and rooting depth, both important factors that enable the interception of nitrate leached from soils (Gastal and Lemaire, 2002). Evidence of an involvement of root hairs in N acquisition has also been reported (Forde and Lorenzo, 2001). Although the relation between root systems and nitrogen nutrition has been extensively studied, the molecular, biochemical and developmental pathways that make up the response of plants to N starvation remain to be elucidated. In most soils, NO3 − , in contrast to NH4 + , is more abundant and has a higher diffusion coefficient making it more available for uptake by the plant (Miller et al., 2008). However, the availability of these N sources depends on ecological factors, such as soil composition and environmental conditions. For example, in soils under anaerobic and humid conditions and with low pH and temperature, nitrification is inhibited and the ammonium concentration increases. Although NH4 + might be more readily available in some soils, NO3 − is the main N form taken up by plants (Miller et al., 2008). Nevertheless, concentration of NO3 in the soil rhizosphere is heterogeneous in time and space. NO3 − is localized in randomly distributed patches. Such environmental conditions have been a selective pressure that has led to the evolution of root developmental programs in response to both, nonhomogeneous nitrate supply and fluctuating concentrations in soil. The root systems of plants respond to localized patches of nitrate, but not to those rich in ammonium. When a mature lateral root meets a nitrate-rich patch, an increase in lateral root elongation occurs, suggesting a positive regulation of root meristem proliferation, whereas the rest of the root axis does not change dramatically. In contrast, when plants are grown under a low, uniform, nitrate supply, an increase in lateral root proliferation in the entire root axis is observed (Fig. 11.1). Thus, N availability regulates root system architecture at least via two distinct pathways: one triggered by local sensing of the N status and one regulated by systemic processes (Malamy and Ryan, 2001). The first steps to understand root developmental responses to N availability have involved the study of lateral root growth at the tissue level. It is known that root systems respond not only to the N supply but also to the

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Root Development molecular N source. In tomato and maize, shoot growth does not differ when comparing nitrate or ammonium as N sources, but root growth is enhanced by ammonium (Bloom et al., 1993, 2003). Root density and extension of the root system of maize seedlings are larger in nutrient solutions containing ammonium than in those containing nitrate as the sole nitrogen source, suggesting that cell division in the root apical meristem might be more rapid when NH4 is used as the N source, perhaps because ammonium assimilation is less energy demanding than nitrate assimilation. The responses in root development to differential supplies of N have been proposed to be caused by the altered redox potential or pH during N uptake rather than by a direct inference of either ammonium or nitrate (Bloom et al., 1993, 2003). Although future research in the area is needed, recent reports have contributed to the understanding of the complex and coordinated response of plant root systems to N availability, mainly in relation to regulation of lateral root initiation, meristem activation and root elongation. Strong evidence points toward NO3 − eliciting a systemic signal to regulate root growth. 11.2.2.2 Effects of N availability on root meristem development In Arabidopsis and tobacco, the growth rate of the primary root is almost completely insensitive to a uniform nitrate supply (Zhang and Forde, 1998; Stitt and Feil, 1999; Filleur et al., 2005). However, in Arabidopsis, relative to shoot dry weight, primary root length slightly decreases as uniform nitrate availability increases. In contrast, in roots that encounter a high nitrate patch, primary root growth does not change (Linkohr et al., 2002). Therefore, lateral roots might have a nitrate sensory mechanism that enables them to modulate meristematic activity in response to localized sources of nitrate while the primary root tip might lack one or more components of this regulatory pathway. However, effects of glutamate on root development, including primary and lateral root growth inhibition resulting in a short and highly branched root system (Filleur et al., 2005) (Fig. 11.4). Root tip cells might be able to sense extracellular glutamate that triggers a reduction in the rate of cell production and/or cell expansion and, therefore, promotes rapid colonization of soil patches with high nutrient concentrations. Besides these advances, a detailed analysis of the developmental changes in primary root length as a consequence of modifications in cell size or cell number remains to be carried out. In the case of maize, an increase in dry matter accumulation has been observed when one root axis is supplied with nitrate in split root experiments (Granato and Raper, 1989). This dry matter accumulation was not attributable to growth of the primary axis but to increased lateral root growth. Also, the maize primary seminal root showed a greater extension rate in correlation with both nitrate and ammonium supplies, in contrast with the elongation rate in roots without nitrate (Bloom et al., 2003), suggesting that the elongation is N dependent and that this nutrient is acquired from the growth media and not from internal sources, because in this region of the root, the phloem

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Figure 11.4 Lateral and primary root development regulated by nitrogen availability. LIN1 (NRT2.1), a nitrate high-affinity transporter is a putative sensor which coordinates sucrose levels or related metabolites with nitrate concentration, thus directing LR initiation. LR activation is also modulated by sucrose/nitrate ratios. When plants are grown under low sucrose conditions there is a repression of the meristematic activity (Little et al., 2005; Remans et al., 2006). Similarly, this repression is also observed as a consequence of a systemic signal derived from shoots with high nitrate contents. ANR1 and AXR4 seem to mediate the modulation of LR elongation, directed both by localized high nitrate patches as well as by uniform nitrate deprivation (Zhang and Forde, 2000). Nitrate is considered to be the signal molecule controlling the developmental programs in roots but glutamate also seems to negatively regulate cell division and/or elongation since inhibition in primary root development and later also in LR growth occurs in the presence of glutamate (Filleur et al., 2005).

system is not fully developed to supply the necessary N from mature tissues. Furthermore, Tian et al. (2005) reported a decrease in root length for primary, seminal and crown roots when the N supply increased from 0.05 to 20 mM for two contrasting genotypes. A positive correlation between the quantity of nitrate applied and the internal cytokinin concentration has also been observed, establishing a possible role of cytokinin in the nitrate-mediated root growth found for the most nitrate-responsive genotype (Tian et al., 2005).

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Root Development 11.2.2.3 Nitrogen availability and lateral root development Changes in lateral root growth in nitrate patches has been reported in several plant species, including Arabidopsis, maize, barley, legumes, citrus, rice and tobacco (Samuelson et al., 1992; Sattelmacher et al., 1993; Dunbabin et al., 2001; Forde and Lorenzo, 2001; Linkohr et al., 2002; Hodge, 2004; Huang et al., 2004; Sorgon`a et al., 2005), and some components of the signaling and regulation pathways have been described. Through split-root experiments, nitrate-rich patches and homogeneous nitrogen supplied to one root axis were both found to produce an increase in lateral root formation and elongation (Farley and Fitter, 1999). In barley, this response has also been observed as a response to ammonium (Drew and Saker, 1975). In Arabidopsis, the response has been proposed to be specific to nitrate, because neither ammonium nor glutamine stimulates lateral root growth (Zhang et al., 1999; Tranbarger et al., 2003) (Fig. 11.4). Nitrate might be the signal molecule capable of inducing changes in the developmental program of plants when they are grown in heterogeneous N soils because of the relative mobility of this molecule when compared to the low mobility of ammonium or glutamate. This response might facilitate the uptake of N in soils in which nitrate is produced from immobile organic compounds and/or ammonium as the result of bacterial activity and chemical reactions, directing lateral growth to these patches (Miller et al., 2008). Furthermore, when Arabidopsis seedlings are grown under homogeneous nitrate conditions, particular responses depending on the level of nitrate limitation have been reported. Transfer of Arabidopsis seedlings from high nitrate conditions to very low nitrate concentrations triggers a response in lateral root length, whereas transfer to medium nitrate increases lateral root initiation, reflecting the existence of levels of regulation related to homogeneous low nitrate availability (Remans et al., 2006) (Fig. 11.4). Regarding lateral primordia initiation, the histology of lateral root formation and pericycle cell dedifferentiation has been studied. It is known that high nitrate patches increase lateral root density (Friend et al., 1990; Linkohr et al., 2002) by enhancing LR primordia formation (Drew and Saker, 1975; Granato and Raper, 1989) (Fig. 11.4). However, reports that describe in detail how this process occurs are scarce. The role of nitrate as a signal molecule is supported by the responses of tobacco and Arabidopsis lines with very low nitrate reductase (NR) activity (Scheible et al., 1997; Zhang and Forde, 2000) and, thus, with limited capacity to assimilate NO3 . The higher accumulation of free NO3 in shoots of transgenic lines with low NR activity growing under both low and high nitrate conditions inhibited lateral root growth similar to that observed in wild-type plants growing under high NO3 conditions, thus reflecting the capacity of NO3 as a signal molecule capable of direct developmental responses in roots. Such a role in long-distance signaling has been demonstrated in split root experiments, in which the accumulation of nitrate in shoots, but not in roots, triggered lateral root development inhibition over the whole root system (Stitt and Feil, 1999). Therefore, a dual system has been proposed for controlling

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developmental changes in nitrate availability: one locally induces lateral root elongation by high nitrate patches and one involves a systemic signaling system that mediates repression of the meristematic activation of lateral roots that depends on the nitrate levels in the shoot (Zhang et al., 1999; Zhang and Forde, 2000). To date, the regulation of lateral root development by internal and external nitrate levels has been reported at three stages: (1) lateral root primordia initiation; (2) emergence and meristem activation; and (3) lateral root elongation. The mechanisms that mediate the systemic and local sensing activation or repression of pericycle founder cells to form new lateral root primordia are currently under investigation. Malamy and Ryan (2001) reported that when Arabidopsis seedlings are grown on media with a high sucrose to nitrogen ratio, lateral root initiation is repressed, thus implying a possible role of sugars or sugar–N balance in root responses to nitrate availability. Analyses of lin1, a mutant that overcomes the limitation of lateral root initiation by the sucrose:N ratio, reveal that the gene is involved in the coordination of the root system responses with the nitrogen status, possibly through sensing the levels of sucrose, their metabolites or even the sucrose:N ratio as possible intermediates in the signal transduction pathway (Malamy and Ryan, 2001) (Fig. 11.4). The characterization of lin1 as an allele of NRT2.1 illustrates part of the regulatory pathway for lateral root initiation. NRT2.1 encodes a putative high-affinity nitrate transporter that functions at low external nitrate concentrations and has been proposed to be a sensor in the signal transduction pathway that represses lateral root initiation related to the sucrose:N status. Lateral root initiation seems to be independent of the transporter function because lateral root formation is still increased despite the decrease in nitrate uptake levels (Little et al., 2005). Although further research is needed to demonstrate how the sucrose concentration affects lateral root initiation, by using low sucrose levels in the media, Zhang et al. (1999) positioned the inhibitory effect of high nitrate levels at a late stage of lateral root development; delaying the activation of the meristem and root elongation when lateral roots have already emerged (Zhang and Forde, 2000) (Fig. 11.4). However, Little et al. (2005) suggest that under experimental conditions of high sucrose:low N a stress situation might be stimulated in which it would be advantageous to repress lateral root initiation as a way to regulate carbon usage. High sucrose:low N levels might also simulate conditions in which root proliferation is not necessary, thus repressing lateral root initiation because nitrate levels are low, whereas carbon supply is directed through roots that are actively acquiring N. Apparently, the role of NRT2.1 goes beyond lateral root initiation. Recently, the effect on lateral root elongation in nrt2.1 mutants transferred from homogeneous high-nitrate growth media to low-nitrate conditions has been reported (Remans et al., 2006). These experimental conditions differ from those reported in Little et al. (2005), in which the functionality of NRT2.1 was analyzed with localized nitrate and high sucrose concentrations. Remans

308
Root Development et al. (2006) suggested that NRT2.1 integrates nitrate uptake/nitrate concentration levels with developmental responses because nrt2.1 is unable to respond through the number of lateral roots, but increases the lateral root length, like wild-type plants do when they are transferred from high to very low levels of nitrate (Fig. 11.4). However, lateral root initiation is not explained just by considering the nitrate content/uptake, instead, it is necessary to consider the dual and independent role of NRT2.1 in developmental nitrate responses. In relation to emergence and meristem activation, in Arabidopsis seedlings under low sucrose availability, an indirect pathway for the regulation of lateral root development by nitrate seems to have an effect on mature lateral roots. Repression of lateral root growth by a high and uniform nitrate supply has been reported: lateral root growth was inhibited before lateral root meristem activation and after lateral root primordia had just emerged from the primary root (Zhang et al., 1999; Zhang and Forde, 2000). This repression seems to be reversible and specific when plants are transferred to low and uniform nitrate, but is not observed when a localized nutrient patch is present. Tranbarger et al. (2003) also documented this arrest by using sucrose-free media and described this effect to occur after primordia initiation, but prior to lateral root emergence (Fig. 11.4). One element potentially implicated in this stage of lateral root development under a homogeneous nitrate supply is the AtNRT1 nitrate transporter. AtNRT1 expression coincides with developing organs, such as primary and lateral root tips as well as young leaves and developing flower buds. In the nrt1.1 mutant, both the maturation of lateral root primordia and the emergence of lateral roots are inhibited when compared to those of wild-type seedlings, but only under specific conditions: acid pH, low homogeneous nitrate supply and high ammonium concentrations (Guo et al., 2001). A possible role for ABA in nitrogen-mediated LR formation and growth has been proposed. LR development is less repressed by high nitrate in the ABA-insensitive Arabidopsis mutants abi4 and abi5. This de-repression occurs when plants are grown in concentrations of up to 1 mM nitrate, whereas under low concentrations, the phenotype of both wild-type and mutants is similar (Signora et al., 2001). These results also relate nitrate responses with sugar signaling because ABI4 seems to be involved in sugar signaling (Rook et al., 2006). Also, a role of the hormone auxin in LR development in function of nitrate availability has been demonstrated by showing that AtNRT1.1 possesses a functional auxin response element and is transcriptionally induced by exogenous auxin (Guo et al., 2002). Also, the axr4 mutant of Arabidopsis does not respond to a localized supply of nitrate, suggesting a role for auxin signaling in lateral root development in response to the nitrate supply (Zhang et al., 1999). Additionally, an increase in indole-3-acetic acid (IAA) levels has been observed when roots of Arabidopsis seedlings are transferred from highnitrate to low-nitrate concentrations when compared with those maintained

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in high levels implying auxin is necessary at some checkpoint for lateral root elongation (Walch-Liu et al., 2006). However, these results are in contrast to those obtained by Linkohr et al. (2002), who observed that axr4 seedlings respond to a localized nitrate supply in the same way as wild type. In contrast with the above-mentioned results (Zhang and Forde, 1998, 2000), recent reports about ANR1 (for A NO3 -inducible Arabidopsis gene), expression describe an upregulation by low nitrate and downregulation by high nitrate under uniform hydroponic conditions (Gan et al., 2005). Therefore, it is necessary to evaluate how nutrient conditions affect ANR1 expression. Despite the still unsolved questions about ANR1 expression, this gene plays clearly a direct role in lateral root elongation and mediates the sensing of nitrate availability. A feedback mechanism has been proposed (Gan et al., 2005) in which the overall N status of the plant might participate, thus integrating local and long-distance nitrate sensing and transduction, phenomena that might include other members of the MADS box family, such as AGAMOUS LIKE 16 (AGL16) and AGAMOUS LIKE 21 (AGL21), whose expression is also induced under low nitrate conditions, albeit to a lower extent than that of ANR1 (Gan et al., 2005; Walch-Liu et al., 2006). The last stage of regulation in lateral root growth by N availability lies in the modulation of root elongation. Roots tend to proliferate in high nitrate patches (Drew and Saker, 1975) and this localized stimulation has been observed in the presence of nitrate patches even with very low levels of the nutrient (Zhang and Forde, 2000). The enhanced development has been attributed to an increase in the rate of meristematic cell production (Zhang and Forde, 1998). This level of regulation is also registered under a low (0.05–1 ␮M) uniform nitrate supply, with a de-repression of lateral root growth inhibition in preformed and emerged lateral roots (Remans et al., 2006). ANR1, which codes for a MADS box protein, has been identified as a critical factor in the root elongation response to local nitrate, as revealed by mutant analysis in which lateral root elongation was reduced or eliminated in localized high nitrate patches. In agreement with these results, ANR1 is rapidly upregulated by the nitrate supply and is proposed to control a set of still unknown genes that might be related to the activation of lateral root elongation under localized high nitrate (Zhang and Forde, 1998) (Fig. 11.4). 11.2.2.4 The effect of nitrogen availability on root hair development The isolation of tomato cDNAs encoding an ammonia transporter and two putative low-affinity nitrate transporters in root hair-specific cDNA libraries suggest that root hairs could transport N-compounds such as ammonium and nitrate and that root hair length could influence N-uptake capacity (Lauter et al., 1996). There is evidence that root hair development is negatively affected by a high nitrate supply (Foehse and Jungk, 1983; Robinson and Rorison, 1987; Boot and Mensink, 1990). In agreement with these observations, spinach root hair length is longer under low nitrate conditions than under high nitrate conditions (Foehse and Jungk, 1983). However, the root hair response seems

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Root Development to be heterogeneous between different species, since tomato and some grasses do not show root hair responses to nitrate (Foehse and Jungk, 1983; Robinson and Rorison, 1987; Forde and Lorenzo, 2001). Present evidence suggests that N availability could influence root hair elongation, but not epidermal cell fate since no alterations in root hair number have been reported. 11.2.3

Potassium

Among the mineral nutrients acquired by plants, potassium (K) is essential for plant metabolism, growth and stress adaptation. Plant roots are able to accumulate K to a level exceeding 100 mM, despite that concentrations of this ion in soils may be found in the range of only 0.1–6 mM (Ashley et al., 2006). K is the most abundant inorganic cation in plants, comprising up to 10% of a plant’s dry weight. In the cytosol, K participates as an osmoregulator and plays a role in the neutralization of anionic groups and in the control of cell membrane polarization (V´ery and Sentenac, 2003). Potassium also participates in enzyme activation, stabilization of protein synthesis and translocation of assimilates in the phloem (Maathuis and Sanders, 1997). Transport of K across the vacuolar and plasma membranes drives stomatal movement, light-dependent movements of organs and translocation of assimilates and other ions in the phloem. K also participates directly in cell growth, contributing to turgor regulation and cell extension through K movement into the vacuole (Rigas et al., 2001). Plants can only acquire K from solutions and, although this element is the fourth most abundant mineral, concentrations of K available in soil vary widely. So, conditions of K deficiency are frequent and lead to growth arrest, due to the lack of osmoticum (Leigh and Jones, 1984) and to impaired sugar and nitrogen balance through inhibition of protein synthesis and transport. To face this challenge, plants have acquired mechanisms during evolution to adapt to the shortage of K. An increase in the K transport systems in root hairs and epidermis but also across membranes of various tissues is regulated and activated under low K availability (V´ery and Sentenac, 2003). Nevertheless, and despite the important role of potassium in growth, little is known about the molecular nature of adaptive responses with an impact at the developmental level (Armengaud et al., 2004). The root developmental response to K availability has been studied in barley (Drew and Saker, 1975). K patches affect root system growth and development by decreasing lateral root growth, not only in the roots next to the patch, but also of the entire root system. In contrast to the effect of nitrate or Pi patches, where a compensation in growth in the high nutrient patches was observed, the general response upon K might be due to the high mobility of K in the soil solution. Uniform K deprivation also affects root development because both a decrease in lateral root length and in number of lateral roots are observed in plants grown under low K (Shin and Schachtman, 2004) (Fig. 11.1). Microarray analysis of K-deprived plants revealed a possible role for ethylene in the repression of

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lateral root initiation because at the beginning of the K deprivation, genes related to ethylene biosynthesis are induced. Reactive oxygen species (ROS) seem to play an important role both in the developmental programs and uptake systems induced by K stress because a preferential accumulation of those compounds was found in the region just behind the elongation zone, where K-uptake and translocation to the shoots is more active (Shin and Schachtman, 2004). Additional analyses using ROS localization suggest that root hairs might possess a sensing system for K deprivation (Shin et al., 2005). ´ et al. (2005) compared the responses of the Arabidopsis S´anchez-Calderon root system to starvation conditions for diverse nutrients and showed that under low-K conditions, an overall effect on plant development including a diminishing of primary root length occurs. This result is in contrast to that of Shin and Schachtman (2004), who found that the final length of the primary roots was not altered when plants were germinated for 3 days in complete media and then transferred and incubated on a K-free media for 10 days. However, it is possible that plants acquired enough K during germination to maintain primary root growth. Although K availability has no very marked effects on the developmental processes in the epidermis of Arabidopsis, a link between root hair development and K availability has been evidenced. Both TINY ROOT HAIR 1 (TRH1), a K translocator, as well as Arabidopsis K TRANSPORTER1 (AKT1), a low-affinity K transporter, participate in the regulation of root hair elongation. In addition, the role of RHD2, an NADPH oxidase, on root hair development could be related to at least a subset of K uptake systems. The rhd2 mutant, which initiates root hairs that cease growing almost immediately, is unable to induce two K transporters, WRKY9 transcription factor, two peroxidases, and an unknown protein. These results suggest that potassium uptake systems and root hair elongation might be linked (Rigas et al., 2001; Desbrosses et al., 2003; Shin and Schachtman, 2004). An agravitropic response has also been observed in Arabidopsis primary roots grown under K deprivation: possibly to cope with the heterogeneous distribution of K in soils, enabling the exploration of areas with higher nutrient contents (Vicente-Agullo et al., 2004; Fig. 11.1). An important discovery was that disruption of the TRH1 gene affects root gravitropic responses under high K conditions demonstrating that the reductions in the K concentration might be linked with the root gravitropic response. These responses might be mediated by changes in auxin transport, because TRH1 seems to affect auxin distribution in the root tip when roots are submitted to low K availability (Vicente-Agullo et al., 2004). 11.2.4

Iron

At the biochemical level iron (Fe) is required for vital processes, such as photosynthesis and respiration. Plants have evolved two different strategies to ensure Fe uptake under iron-deficient growth conditions. Dicotyledonous

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Root Development and nongrass monocotyledonous plants develop the so-called Strategy I response that induces three activities under low iron conditions: (1) acidification of the rhizosphere performed by H-ATPase in order to make Fe (III) more soluble; (2) increase in the activity of FRO2, a plasma membrane-bound protein that reduces Fe (III) to Fe (II) (Robinson et al., 1999); and (3) upregulation of IRON TRANSPORTER 1 (IRT1), the major Fe (II) transporter present in Arabidopsis roots (Eide et al., 1996; Vert et al., 2003). In agreement with this phenomenon, FRO2 and IRT1 are transcriptionally activated by Fe-deficiency (Connolly et al., 2002). Strategy II, which is used by grasses, depends on the secretion of phytosiderophores that chelate Fe (III) to be internalized by specific transporters without requiring a previous reduction to Fe (II) (Curie et al., 2001). Little is known about signal transduction pathways and transcriptional networks that function upstream of any of these mechanisms. Recently, two bHLH-type transcription factors involved in the response to iron deficiency in tomato (FER; Ling et al., 2002) and Arabidopsis FeDEFICIENCY INDUCED TRANSCRIPTION FACTOR 1 (FIT1; Colangelo and Guerinot, 2004) have been isolated. The tomato fer mutant was isolated because of its chlorotic phenotype and because it fails to activate the Fe uptake strategy I. FER encodes a protein containing a basic helix-loop-helix domain and fer plants exhibit altered root developmental phenotypes under both low and normal iron conditions, indicating that FER acts irrespective of iron supply. However, in the roots of fer plants, the transcript level of Leirt1, the ortholog of IRT, is lower than that observed in the wild type (Ling et al., 2002). FIT1 encodes a putative transcription factor that modulates iron uptake responses in Arabidopsis. In response to iron starvation, FIT1 transcripts significantly increase in the outer cell layers of the root. fit1 mutant plants are chlorotic and die as seedlings, but can be rescued by the addition of iron suggesting a defect in iron uptake. Also, fit1 plants accumulate less iron than wild-type plants in both root and shoot tissues. Microarray analysis of the fit1 mutant background showed that FIT1 acts upstream of various genes implicated in iron homeostasis, including FRO2 (Colangelo and Guerinot, 2004). At the developmental level, iron deficiency has been found to alter root hair number. In potato and sugar beet not only root hairs, but also the number of transfer cells increase under iron deficiency (Bienfait et al., 1987). A more drastic effect has been described for Casuarina glauca, in which cluster (proteoid) roots develop in response to iron deficiency (Arahou and Diem, 1997). In the model plant Arabidopsis, how Fe deficiency alters root hair number has been remarkably described. When growth under Fe-deficient conditions the primary root of wild-type seedlings shows that almost 50% of the root hairs are branched. In contrast to seedling roots, in which the majority of H cells form root hairs, in the aging primary root and in laterals, only 37% of the epidermal cells in this position differentiate as root hairs. Under Fe starvation the root hair frequency in the H position in the primary root increases to 42% ¨ (Schikora and Schmidt, 2001; Muller and Schmidt, 2004; Perry et al., 2007).

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These two strategies seem to be directed to increase the surface area of the root system in diverse stages of development as a response to deal with Fe deficiency. Microarray analysis of Arabidopsis plants subjected to Fe deficiency demonstrated that the expression of genes involved in ethylene synthesis and signal transduction is transcriptionally induced, thus, suggesting that ethylene is involved in the root hair phenotype triggered by Fe deficiency (Thimm et al., 2001). This phenomenon correlates well with the multiple-tip root hairs observed by Wilson et al. (1990) in the ethylene- and auxin-insensitive mutant axr2 analyzed. The phenotypes of diverse hormone-related mutants under Fe starvation have been analyzed (Schmidt and Schikora, 2001) and ethylene and auxin were found to be essential for the development of extra root hairs in response to Fe deficiency. Beyond the importance of these findings, new approaches are needed to characterize the signaling pathway and molecular networks that connect Fe starvation and ethylene-mediated root hair responses. Recently, new genes involved in root hair morphogenesis that code for transcription factors, GPI-anchored proteins and inositol triphosphate phosphatase have been isolated and characterized (Jones et al., 2006). The detailed analysis and unraveling of the regulatory networks that involve novel ‘root hair genes’ and transcription factors, such as FIT1 and FER, remain a necessary future step in this field of research. 11.2.5

Sulfate

Plants have a constitutive demand for sulfur (S) to synthesize proteins, sulfolipids and other essential S-containing molecules for growth and development. Plants take sulfur from the soil mainly in the form of sulfate, but also from the atmosphere as SO2 or H2 S. Sulfate is directly assimilated into cysteine, the key entry point of the natural sulfur cycle (Buchner et al., 2004). However, sulfate represents only a very small part of the S present in most soils because approximately 95% is bound to organic molecules that cannot be directly assimilated by plants. Thus, symptoms of sulfur deficiency are frequently encountered in crop plants such as retarded and chlorotic growth with pale-green young leaves, while mature leaves remain dark-green. In addition, sulfate starvation also depresses N transport and the hydraulic root conductivity (Karmoker et al., 1991), with significant reduction of crop productivity. The nutritional status of the plant controls both biochemical (high-affinity sulfate uptake, low-affinity vascular transport and vacuolar efflux) and developmental programs. Indirect evidence suggests that sulfate itself has the potential to play a role as a sensor or regulatory signal in the roots (Buchner et al., 2004; Kataoka et al., 2004). Under low-sulfate conditions, some plants rapidly induce the sulfate uptake capacity and assimilation mainly by the root epidermis, root hairs and cortex (Yoshimoto et al., 2002; MaruyamaNakashita et al., 2004b). This increased uptake capacity is accompanied by

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Root Development the transcriptional activation of genes encoding sulfate transporters in the roots and in the rest of the plant (Rae and Smith, 2002), as well as a remobilization of sulfur metabolites (Maruyama-Nakashita et al., 2003). Prolonged sulfate deprivation results in an altered shoot–root partitioning of biomass in favor of the root. In Arabidopsis, a more prolific root system and an increase in lateral root number that develop closer to the tip of the major roots than those of plants growing in the presence of sulfate, have been observed (Kutz et al., 2002) (Fig. 11.1). B. oleracea is able to utilize atmospheric H2 S as S source; however, root proliferation and increased sulfate transporter expression also occur as in other plant species. Upon more prolonged sulfate deprivation, shoot and root biomass partitioning in favor of root production (Buchner et al., 2004). Genetic engineering of root development in response to low S availability has been considered as a target to improve sulfate uptake despite the lack of studies concerning the root developmental responses to sulfate availability, and the direction of most research to the understanding of sulfate uptake capacity and assimilation. A developmental program induced by sulfate starvation has been always accompanied by a biochemical program leading to a better adaptation to low nutrient levels in plants (Kutz et al., 2002; Buchner et al., 2004). Nitrilase 3 (NIT3) has been identified as a direct link between sulfate deprivation and auxin synthesis (Kutz et al., 2002). NIT3 catalyzes the conversion of indole-3acetonitrile into IAA and is specifically induced by low sulfate conditions in conductive tissues and lateral root primordia. The induction of NITRILASE 3 (NIT3) by sulfate deprivation has been proposed to increase auxin production, thus leading to increased root growth and branching. Low sulfate conditions also induce the expression of other genes related to auxin synthesis and auxininduced proteins (Hirai et al., 2003; Nikiforova et al., 2003). These results point to a direct link between sulfate availability and proliferation in root meristems mediated by auxin accumulation. In summary, low sulfate conditions increase local auxin synthesis and, probably, induce the expression of genes involved in the formation, establishment and maintenance of root meristems. Notwithstanding that cellular and molecular modifications related to the alterations in root architecture directed by nutrient status are still unclear and not delimited, novel molecular actors are emerging to help understanding the physiological and morphological responses of roots to sulfate starvation. Although the role of cytokinins in the regulation of root developmental responses to sulfate deprivation has not been unraveled in detail, cytokinins have been demonstrated to mediate the uptake response through the repression of the high-affinity sulfate transporters SULTR1;1 and SULTR1;2 in Arabidopsis roots (Maruyama-Nakashita et al., 2004a). More recently, the role of a microRNA involved in sulfate stress responses has been reported and the expression of the microRNA family MIR395 to be highly upregulated after sulfate starvation. The mRNAs of four members of a gene family that code for ATP sulfurylases (APS) have been predicted to be potential targets of this

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microRNA family (Jones-Rhoades and Bartel, 2004). In fact, the mRNA of Arabidopsis SULFATE TRANSPORTER 68 (AST68) has experimentally been demonstrated to be a target of the miR395 family (Allen et al., 2005). It has been demonstrated that APS1 and AST68 transcripts are more abundant under low-sulfur conditions (Lappartient et al., 1999). At first sight, the fact that both miR395 and target genes are upregulated under stress conditions might be perceived as contradictory, but, we might speculate that the posttranscriptional regulation of APS1 and AST68 is directed to modulate the levels of the transcript amounts of transcripts, in a tissue-specific manner. The existence of diverse members of the MIR395 family supports this idea. From these studies, potential areas of research include the detailed analysis of the expression pattern of each of the MIR395 family members and the mutation and overexpression of mir395 which could determine a possible role of this type of regulation in root development.

11.3

Conclusions

The study of model species, such as Arabidopsis, rice, Medicago and Lupinus albus, has expanded our knowledge on the biochemical and regulatory mechanisms through which nutrient ion signals alter several phenomena, such as root cell length and proliferative capacity, control root growth and alter root system architecture. In angiosperms, most of the responses of root architecture to nutrient deficiencies seem to play a similar role: the increase in the capacity of the root to capture nutrients. The current state of the art in research on the responses of diverse species to various nutrient deficiencies suggests that the stress is sensed by nutrientspecific signal transduction pathways that respond to both external and internal concentrations of nutrients. In some cases, auxins, cytokinins and ethylene appear to play diverse roles in each specific deficiency. Nevertheless, other signaling molecules are also emerging as novel players in these signal transduction pathways. Great advances in the understanding of the physiological, biochemical and cellular responses of higher plants to the availability of different nutrients are being achieved by the use of genetics and novel and powerful tools, such as microarray analysis. Common as well as specific responses to different nutrient deficiencies have been reported, but more experimentation is still needed to fully understand the signaling pathways and their potential interactions. It is a reality that the regulation of genes involved in the adaptation to nutrient deficiencies is more complex than originally thought, because it includes posttranscriptional regulation by microRNAs. This regulation must also be considered by the various research groups in the field to eventually obtain a detailed picture of the molecular and biochemical processes that control the general responses to nutrient stress in plants.

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Acknowledgments The authors thank Dr. June Simpson Williamson for critically reviewing the manuscript. The work from our research group reported in this review was supported in part by the Howard Hughes Medical Institute (Grant No. Nbr55003677) and Consejo Nacional de Ciencia y Tecnolog´ıa (Grant No. SEP2003-C02-43979).

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Chapter 12

STUDYING ROOT DEVELOPMENT USING A GENOMIC APPROACH Jose R. Dinneny1,2 and Philip N. Benfey1 1

Department of Biology and IGSP Center for Systems Biology, Duke University, Durham, NC, USA 2 Temasek Lifesciences Laboratory, National University of Singapore, Singapore

Abstract: The completion of the Arabidopsis thaliana genome sequencing project represented the beginning of a new way to approach biological questions in plants (The Arabidopsis Genome Initiative, 2000). At a basic level, a fully sequenced genome makes it possible to predict the identity and structure of most genes. This information, in conjunction with recent advances in technology to quantitatively detect the expression of thousands of genes simultaneously, has greatly expanded the number of genes a biologist can study. Thus, the postgenome era is characterized by a wealth of data, but also by new challenges. Large data sets require analytical methods such as statistical analysis and computationally intensive methods, with which most biologists are not necessarily familiar. To address these challenges, the new biologist must collaborate with researchers from other fields such as computer science and engineering, to design experiments, interpret data, generate models and build bioinformatics infrastructure to share data. While large data sets facilitate rapid progress using traditional reductionist approaches to answering biological questions, they also enable a new approach termed systems biology that aims to understand the structure of large-scale phenomena not easily studied through the examination of individual genes. In this chapter, we will give an overview of the technologies that allow the root developmental biologist to acquire large data sets and examine how these techniques have been used to study root development. Finally, we will explore the types of questions that remain in root developmental biology for which genomic approaches are likely to provide useful insights. Keywords: root development; genomics; transcriptional networks; microarray; transcription factor; systems biology

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12.1

Introduction: how root development enables a genomic approach

Development is a complicated process. Therefore, developmental biologists have often chosen relatively simple model organisms to study the mechanisms that pattern structures of interest. Subsequently, models of development can be tested in other organisms to determine their generality and, in more complicated organisms, to determine how a simple system can be elaborated upon. In this way, the root of Arabidopsis thaliana has served as one of the simplest developmental systems available in plants (Fig. 12.1). The rotational symmetry that characterizes the arrangement of the approximately 15 cell types in the root simplifies an anatomical analysis. Cells are arranged into files that originate from the root apex, where a population of self-renewing stem cells, termed initials, divide to produce the different cell layers and feed cells into the various files (Scheres et al., 2002). Cells closest to the initials represent the most recently born daughters, while cells further away are more mature descendants. This arrangement along the longitudinal axis allows the observation of cells at every developmental stage in a single root without the need for a time-course analysis. Certain cellular processes, such as cell division and elongation, are limited to particular stages of differentiation in plants, creating zones along the longitudinal axis of the root that roughly demarcate particular developmental transitions in the life of a root cell (Fig. 12.1). Near the root tip is the meristematic zone, which contains cells actively engaged in the cell cycle. Subsequently, cells begin to mature and enter the elongation zone where they start to grow anisotropically along the longitudinal axis. Finally, cells in the maturation zone take on their final differentiated state. This zone is demarcated by the initiation of root hairs from the trichoblast epidermal cell file and also coincides with the maturation of the protoxylem cells in the stele. The structural characteristics of the root allow certain developmental processes to be easily studied using a genomic approach. For example, because cells along the length of the root represent successively later stages of development, transcripts of the longitudinal zones can be profiled to identify genes involved in the maturation of cells in the root (Birnbaum et al., 2003; Brady et al., 2007). As described below, a compendium of expression profiles for many cell/tissue types in the root has been generated using the cellsorting methodology developed by Birnbaum et al. (2005). For almost any gene in the genome, its expression pattern in the root can be inferred with high spatial resolution. Finally, the translucent optical properties of the root allow an unobstructed view of development using confocal microscopy; live imaging is easily accomplished, enabling the study of the function of genes of interest by using fluorescent marker lines. This feature, combined with the small size of the Arabidopsis seedling, potentially makes it possible to visualize many different reporter lines at one time to obtain large-scale expression data sets.

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12.2

12.2.1

Genome-scale technologies for understanding gene function Genomic sequence

Genome-scale strategies are very difficult to employ unless a complete genomic sequence has been determined. Fully sequenced plant genomes include Arabidopsis (The Arabidopsis Genome Initiative, 2000), rice (Oryza sativa) (Goff et al., 2002; Yu et al., 2002) and poplar (Populus trichocarpa) (Tuskan et al., 2006). In addition, 20 other plant genomes are in the process of being sequenced (Jackson et al., 2006). These species cover 13 of the 606 extant plant families and include basal land plants that lack structures homologous to the roots found in seed plants. The next task will be to unravel the gene networks involved in root development in these disparate species to understand how developmental pathways have evolved (Menand et al., 2007).

12.2.2

Genome-wide perturbation of gene activity

Often the immediate result of a genome-scale experiment is a list of genes with a particular expression pattern or property of interest. Identification of a loss-of-function mutant phenotype associated with these genes can help determine the critical roles a gene product has. Furthermore, transcriptional profiling of these mutants can be useful for further characterizing a biological pathway, especially when mapping a transcriptional network. Several collections of insertion alleles have been sequenced and mapped to specific regions of the genome in Arabidopsis and rice and are publicly available from stock centers (Alonso et al., 2003; An et al., 2003; Sallaud et al., 2004). An important goal of one such stock center, the Salk Institute Genomic Analysis Lab (SIGnAL), has been to establish homozygous lines for many of the T-DNA insertions (Alonso and Ecker, 2006). This collection will enable researchers to easily conduct large-scale phenotypic screens of insertion alleles. Thus, it should be possible to characterize the role of all transcription factors (TFs) or signaling components in a particular process of interest in a high-throughput manner, generating what has been termed the mutant ‘phenome’. Recently, synthetic versions of miRNAs (artificial miRNAs, amiRNA) have been designed that have been shown to be effective in silencing gene expression in Arabidopsis, tomato and tobacco and should enable researchers to functionally test the role of presumptive root development genes in other species (Alvarez et al., 2006; Schwab et al., 2006). Viral-induced gene silencing (VIGS) has also been found effective in reducing the activity of genes in the basal eudicot, poppy (Papaver somniferum), indicating that experiments testing gene function do not need to be limited to species in which transgenic organisms are easily generated (Hileman et al., 2005).

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Root Development In addition to transgenic resources for disrupting gene function, nontransgenic methodologies have also been developed for reverse genetics. TILLING (targeting-induced local lesions in genomes) is a method that utilizes a chemical mutagen to induce mutations in the genome of the organism of interest (Henikoff et al., 2004). DNA from several thousand mutant lines is pooled and then screened to identify a pool that contains mutations in the gene of interest. TILLING is particularly useful because it does not require knowledge of the complete genomic sequence and can be performed in species in which generating transgenic plants is not easy. Several TILLING projects are in progress in Arabidopsis, maize and Lotus japonicus and resources are available to enable researchers interested in generating TILLING populations in their species of interest (Henikoff et al., 2004). 12.2.3

DNA microarray technology

Besides sequencing, no other technology has had as large an effect on the amount of data a biologist can analyze as the DNA microarray. While the technology used to generate microarrays can vary, the basics of how they function are the same (Fig. 12.2) (Schaffer et al., 2000; Galbraith, 2006). Fluorescently labeled RNA or DNA is hybridized to a collection of different DNA molecules that are arrayed at high density on a fixed surface. The DNA molecules are complementary to a set of sequences of interest depending on the use. For example, DNA microarrays used to assay gene expression have probes that are complementary to mRNA sequences while arrays used for ChIP-chip (Chromatin Immunoprecipitation followed by hybridization to microarrays, see Section 12.2.4) generally have probes that are complementary to intergenic sequences. Hybridization efficiency is correlated to the concentration of the RNA/DNA molecules in the solution and will determine the amount of hybridized probe. The fluorescent tag is then detected with a laser scanner and the image intensity is converted into arbitrary expression units. The two most widely used microarray platforms are the Affymetrix GeneChip and the spotted cDNA/oligonucleotide microarray. Affymetrix microarrays are generally considered to give more reproducible results, while spotted microarrays are less expensive and can be easily customized to contain probe sets of interest. Spotted cDNA arrays can be generated without knowledge of the genomic sequence while generating a probe set for Affymetrix arrays requires sequence information to design 25-mer probes. 12.2.4

ChIP-chip

The ChIP-chip technique has begun to transform work on transcriptional networks by allowing researchers to identify all the sites in a genome that are bound by TFs of interest. The technique is a combination of two commonly used techniques: Chromatin Immunoprecipitation (ChIP) and DNA microarray hybridization (Fig. 12.2). First, cells are treated with a chemical

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that cross-links proteins to DNA. For example, if a TF is bound to a promoter, it will become chemically bonded to that piece of DNA. The chromatin is then fragmented and an antibody is used to immunoprecipitate the TF/chromatin complex. The cross-linking is reversed and the DNA is purified. In a typical ChIP experiment, sequence-specific primers can be used to amplify regions of interest that are suspected of being bound by the TF in question. The difficulty with ChIP is that prior knowledge of which promoters are bound by the TF and approximately where in the promoter the TF binds is necessary to detect enrichment of a particular sequence in the immunoprecipitated DNA pool. This problem was solved with the advent of the ChIP-chip technique in which a DNA microarray is designed with probes complementary to intergenic sequences of all genes (Ren et al., 2000; Iyer et al., 2001). The immunoprecipitated DNA is labeled with a particular fluorophore and hybridized to the array. Nonimmunoprecipitated chromatin is also hybridized to the same array as a control, but this DNA pool is labeled with a different fluorophore. The hybridization signal is compared between the two-labeled DNA populations and microarray probe sets with an increased hybridization of the immunoprecipitated sample relative to the control are considered regions of the genome bound by the TF of interest. The density of probes on the array and the length of each probe will determine the resolution at which a particular binding site can be defined. The ChIP-chip technique has been used extensively in yeast to determine the binding site for hundreds of TFs (Harbison et al., 2004). These data can be used to determine binding site profiles for TFs and can be compared to expression profiling experiments to determine if the identified TF–promoter interactions make biological sense. ChIP with PCR is now used more extensively in plants to test direct interactions between TFs and promoter sequences. The recent availability of tiling arrays that contain probe sets covering the entire genomic sequence of Arabidopsis (available from Affymetrix, Nimblegen and Agilent) will enable plant biologists to scan the entire genome for TF-binding sites and begin assembling large-scale transcriptional networks of direct interactions (Mockler et al., 2005), which will help facilitate a systems biology approach for understanding developmental processes. 12.2.5

High-throughput sequencing

In addition to hybridization-based techniques, high-throughput sequencing technologies are set to revolutionize studies of gene regulation in plants. The major difficulty with hybridization-based techniques is that they provide indirect measures of RNA or DNA concentration in the cell. Thus, the data primarily provide a relative quantitative estimate. High-throughput sequencing enables the direct quantitation of DNA molecules in a population of interest providing an assessment of absolute concentration. This approach was recently utilized to assay the methylation status of the entire Arabidopsis genome as well as to determine the transcriptional effect of mutants defective in DNA-methylation (Lister et al., 2008).

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Large-scale data analyses

The large amount of data produced in a microarray experiment (thousands to millions of data points) requires the developmental biologist to consider statistics during data analysis. The main reason that standard statistical analyses are often not sufficient relates to the scale of genomic experiments. Some of the earliest microarray-based studies identified genes as differentially expressed between two experiments based solely on a fold-change criterion. Researchers now realize that statistics should be used to measure confidence associated with observed expression differences. While determining the significance of differential expression would be relatively straightforward when only one gene is considered, the examination of thousands of genes at a time requires that the effects of multiple testing be considered and more stringent criteria utilized when classifying a gene as differentially expressed. An indepth description of the methods used for microarray data analysis is beyond the scope of this chapter and the reader is referred to several good resources that discuss available options (Draghici, 2003; Allison et al., 2006).

12.3

Building transcriptional networks: an introduction

The simplicity of the root enables the developmental biologist to focus on a few fundamental processes. Perhaps the most studied process thus far in root development is cellular differentiation. Differentiation is the process by which cell identity is specified and includes the developmental path that establishes a cell’s final form and function. As with most developmental processes, TFs play a crucial role in differentiation processes by regulating the expression of genes that ultimately control changes in cell structure and physiology. Thus, to understand how differentiation occurs in the root, it is necessary to study the network of TFs that regulate changes in gene expression. Pregenomic strategies for understanding differentiation in the root focused on a group of TFs that, when disrupted, caused changes in patterning along the radial axis (Di Laurenzio et al., 1996; Helariutta et al., 2000; Aida et al., 2004). The current goal of postgenomics biology is to characterize the chain of transcriptional events that are controlled by these TFs. To build a detailed transcriptional network, it is important to establish at least three data sets: (1) a high-resolution spatial map of gene expression, including an understanding of how gene expression changes along both the radial and longitudinal axis; (2) the localization of the TFs of interest since mRNA expression is not perfectly correlated with TF activity; and (3) identification of the direct targets for the TFs. In the next section, we will detail several studies that have sought to characterize the transcriptional networks controlling root development (Table 12.1). Most of these studies have focused on transcriptional profiling of root cells/tissues to identify genes that are expressed during a particular process or in a cell type of interest. These data sets provide a foundation of

Studying Root Development Using a Genomic Approach
331 Table 12.1 List of publications describing microarray data sets discussed in this chapter Publication

Technique Used

Tissue/Cell Type Profiled

Himanen et al. (2002)

Pharmacological treatment used to induce synchronous lateral root initiation

Primary roots and lateral root initials

Vanneste et al. (2005)

Same as Himanen et al.

Birnbaum et al. (2003)

FACS using GFP-reporter lines to profile the expression of individual cell/tissue types

Nawy et al. (2005) Lee et al. (2006) Woll et al. (2005)

Same as Birnbaum et al. Same as Birnbaum et al. Laser capture microdissection

Same as Himanen et al., except that slr mutant roots were also profiled Epidermis (atrichoblasts), lateral root cap, endodermis with cortex, endodermis with QC, stele QC and columella root cap Cortex, xylem and phloem Maize wild-type and rum1 mutant pericycle cells

Casson et al. (2005)

Laser capture microdissection

Apical and basal regions of Arabidopsis globular and heart stage embryos

Levesque et al. (2006)

Same as Birnbaum et al.

Pericycle, epidermis (atrichoblasts) with lateral root cap

Mouchel et al. (2006)

Expression assayed in plant carrying a natural variant of the BRX gene from Uk-1 AtGenExpress Consortium set of microarray experiments Same as Birnbaum et al., Microdissection and analysis of tissue segments from individual root Same as Birnbaum et al. except roots treated for 1 hour with salt stress or 24 hours with iron deprivation.

Whole roots

Nemhauser et al. (2006) Brady et al. (2007)

Dinneny et al. (2008)

7-day-old seedlings All tissue layers of the root except the procambium

Epidermis with Lateral Root Cap, Columella Root Cap, Cortex, Endodermis with QC, Stele, Protophloem

techniques and information for characterizing gene expression in a developmental system.

12.4

Building transcriptional networks I: creating high-resolution spatial maps of gene expression

To identify genes involved in a process of interest, it is necessary to compare the transcriptional profiles between at least two different samples. The type of comparison can be as simple as comparing the differences in gene expression between the shoot and the root (Schena et al., 1995), but many processes under consideration often require a much more fine-scale comparison.

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Root Development Pharmacological tricks have been used to synchronously induce the development of specific cells involved in lateral root initiation (LRI) (Himanen et al., 2002, 2004). In addition, several methods have recently been created that allow the developmental biologist to dissect the root into distinct tissues and even cell types for transcriptional profiling (Birnbaum et al., 2003, 2005; Casson et al., 2005; Woll et al., 2005; Brady et al., 2007). 12.4.1

Characterizing the cascade of transcriptional events leading to LRI

The regulation of LRI is an excellent system for studying the interplay between patterning and environmental regulation of development. As a result of both extrinsic and intrinsic signals, cells in the pericycle become competent to initiate lateral roots (Malamy, 2005). First, the cytoplasm of a subset of pericycle cells at the xylem pole becomes optically dense (Malamy and Benfey, 1997). Subsequently, anticlinal divisions occur in the pericycle that reduces cell length, which is followed by periclinal divisions that produce a multitiered lateral root primordium. Further divisions lead to the outgrowth of the root through the outer tissue layers (endodermis, cortex and epidermis) and into the environment. During normal development, lateral root primordia initiate at different times along the length of the primary root, typically with lateral roots initiating farthest away from the root apical meristem first. To perform a time-course microarray experiment to examine the waves of transcription that occur during the different stages of LRI, it would be difficult to obtain enough tissue from any particular stage, especially the earliest stages, due to the irregularity of the process. Work done in the laboratory of Tom Beeckman has circumvented this problem through the use of pharmacological conditions that lead to the synchronous initiation of many lateral root primordia from the primary root (Himanen et al., 2002, 2004). Plants are first germinated on media that contains the polar auxin-transport inhibitor 1-N-naphthylphthalamic acid (NPA), which inhibits the formation of LRI sites. Roots are then transferred to media lacking NPA, but containing the auxin 1-naphthalene acetic acid (NAA). The increase in auxin induces lateral root primordia along the entire length of the primary root in near perfect synchrony. Root segments with the root tip and hypocotyl regions dissected away are collected at several time points after transfer to NAA. In an initial transcriptional profiling experiment with spotted cDNA microarrays, 906 significantly differentially expressed genes were found during the 6-hour time course (Himanen et al., 2002). Clustering of the differentially expressed genes revealed that particular biological processes were selectively up- or downregulated. Of particular interest, genes involved in cell-cycle regulation were activated while cellular differentiation genes were repressed. A second set of transcriptional profiling studies expanded the set of genes analyzed by using the Affymetrix ATH1 array containing approximately 24 000 probe sets (Vanneste et al., 2005).

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To identify more specifically genes downstream of auxin sensing that control LRI, the transcriptional profiles of wild type and solitary root (slr) mutant roots were compared. The slr mutation results in the stabilization of the SLR protein and constitutive suppression of auxin responses in the pericycle (Fukaki et al., 2002). Since auxin perception is necessary to initiate lateral roots, slr mutants lack lateral roots after treatment. A cluster comparison method termed cross-table clustering was used to identify sets of genes induced in wild-type roots after transfer to NAA but which had reduced activation, or which were repressed along the time course in slr mutants. These genes are referred to as LRI genes and depend upon nonstabilized SLR for their activation by auxin. Analysis of the LRI gene set revealed increased enrichment of genes involved in cell proliferation, such as those controlling the DNA synthesis phase of the mitotic cell cycle. Consistent with these findings, DNA metabolism was also more highly enriched in the LRI gene set compared to all significantly differentially expressed genes. A glimpse of the transcriptional network regulating the initiation of lateral roots was also uncovered with the discovery that a large set of AUX/IAA and ARF (auxin response factor) TFs is part of the LRI gene set. Several AUX/IAA genes, which have been previously shown to negatively regulate lateral root formation, were activated indicating the presence of a negative feedback loop downstream of SLR-dependent auxin signaling. In addition, several positive regulators of auxin action were uncovered. For example, ARF19, which shows SLR-dependent auxin activation, directly interacts with the SLR protein and when knocked-out in combination with arf 7, results in similar LRI defects as slr (Fukaki et al., 2005; Okushima et al., 2005; Wilmoth et al., 2005). This auxin-responsive network of TFs may in turn regulate auxin-homeostasis and transport pathways that are included in the LRI gene set. 12.4.2

Cell-type-specific transcriptional profiling using fluorescence-activated cell sorting

Organ-specific transcriptional profiling is useful for gathering information on the general differences between a control and a target population of cells and will identify those genes that are transcriptionally regulated in most cell types of that organ. However, since cell types have specialized functions, it is likely that they express different genes. Thus, pooling these differences will likely dilute many of the most interesting transcriptional changes beyond detection. Recently, a technique that combines the use of green fluorescent protein (GFP) reporter lines marking specific cell/tissue types with fluorescence-activated cell sorting (FACS) has allowed the capture of highly specific transcriptional profiles of nearly every cell type in the root (Fig. 12.3) (Birnbaum et al., 2003, 2005; Brady et al., 2007). To obtain cell-type-specific expression profiles, reporter lines must first be generated that express a fluorophore, such as GFP, in cells of interest. Thousands of roots are grown that contain such a reporter and harvested to

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Root Development obtain enough cells for sorting. Whole roots are placed in a solution containing enzymes that break down cell wall components (protoplasting). After the cell walls are digested, cells dissociate from each other. The protoplasted cells are fed into an FACS device, and the fluorescent GFP-expressing cells are sorted from the nonfluorescent cells. Typically, ∼10 000 fluorescent cells are collected to obtain enough RNA. Because of the limiting quantities of materials used, two rounds of linear amplification are needed to generate sufficient quantities of cRNA probes for hybridization. In the initial studies using FACS (Birnbaum et al., 2003) a set of five characterized marker lines were used covering most of the cell/tissue types of the root. For example, the WOL::GFP reporter marks all cells of the stele including the pericycle, ¨ xylem, phloem and vascular cambium (M¨ahonen et al., 2000). The SCR::GFP reporter marks cells of the endodermal cell lineage, the cortex/endodermal initial (CEI) cell, and the quiescent center (QC) (Sabatini et al., 2003). In combination with the cell-sorting experiments, the root was also divided into three zones along the longitudinal axis, which were distinguished by morphological landmarks (Fig. 12.1) (Birnbaum et al., 2003). The combination of the expression profiles of the three longitudinal zones with the results obtained through cell sorting of tissues encompassing five radial zones resulted in 15 subzones of the root by which differential expression could be characterized. This high-resolution representation of expression levels was designated a ‘digital in situ’ because it could be used to infer spatial information about gene expression similar to that obtained with RNA in situ hybridization. One obvious concern with the transcriptional profiles obtained from sorted cells is that these cells might not retain their transcriptional characteristics long enough after dissociation to provide meaningful profiles. The transcriptional profiles of cells were characterized at several time points after protoplasting and cells were found to retain distinct profiles for up to 1.5 hours, which is within the time frame of a typical sorting experiment (Birnbaum et al., 2003). A small subset of genes consistently affected by cell sorting (3.4%) was removed from any further analysis. In addition, the data set was validated by determining whether published expression patterns of known genes were faithfully represented. Out of 26 published profiles, all but one matched the digital in situ profile. Finally, the expression patterns of two uncharacterized genes were determined by RNA in situ hybridization and found to match the digital in situ profile (Birnbaum et al., 2003). With these expression profiles, Birnbaum et al. (2003) sought groups of genes that were expressed in a similar manner suggesting a shared function. The analysis of differential expression identified between 8 and 10 patterns into which most genes could be classified. As expected, genes involved in cell division and nuclear organization were enriched in a group of genes whose expression encompassed all tissues in zone 1 (root tip). On the other hand, genes expressed in tissues of zone 3 (elongation and maturation regions), were enriched for kinases and TFs. Since hormones have been shown to play important roles in regulating patterning and growth in the root, the question

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was addressed whether hormone regulation and response pathways were regionalized in the root. Some sets of genes involved in auxin, gibberellic acid (GA) and jasmonic acid biosynthesis and response were found to be overrepresented in a few nonoverlapping expression domains, suggesting the presence of hormone response centers in the root (Birnbaum et al., 2003). 12.4.3

Expression profiling of the stem-cell niche

One of the most important populations of cells in the root tip not specifically profiled in Birnbaum et al. is the QC. The QC acts as a stem-cell organizing center, specifying stem cell identity in the immediately surrounding cell population, similar to other stem cell organizing centers in animal systems (Dinneny and Benfey, 2008). To obtain cell-type-specific expression profiles for this cell population, a marker for the QC was identified in an enhancertrap insertion collection (Nawy et al., 2005). An enhancer trap line with a T-DNA inserted into the intron of a gene encoding the AGL42 MADS-box TF was identified that drove expression in the QC and surrounding stem cell niche. Subsequent cloning of genomic fragments from the AGL42 locus determined that elements in the promoter as well as the first intron were required to drive expression in the QC. Using this GFP reporter for cell-sorting experiments, 290 genes were identified that are significantly enriched in the QC compared to other sorted cell profiles. Several of these genes are involved in GA biosynthesis suggesting the existence of a GA point source centered in the vicinity of the QC. In addition, genes involved in inhibiting auxin biosynthesis (SUPERROOT1/SUPERROOT2) (Boerjan et al., 1995; Barlier et al., 2000) were specifically enriched as well as the brassinosteroid receptor, ˜ BRI1 (Li and Chory, 1997; Cano-Delgado et al., 2004), indicating that the QC is a center for hormone-related regulation. 12.4.4

Analysis of spatiotemporal expression patterns in the root

While the Birnbaum et al. data set represented a uniquely high-resolution view of gene expression in a multicellular organ, several cell types remained to be examined and most of the reporter lines utilized for cell sorting marked multiple tissue layers. To add additional detail to the root expression map and generate a bioinformatic framework necessary to analyze a more complex data set, Brady et al. generated additional cell-type-specific expression profiles by FACS (Brady et al., 2007). In total, expression profiles for 14 of the 15 cell types present in the root were analyzed. In addition, the first-ever expression map for a single organ was generated by microdissection of a root into 13 sections along the longitudinal axis (Fig. 12.1). To analyze this data set, two approaches were taken. First, a supervised approach was used in which genes were identified that displayed cell-typespecific expression patterns. Brady et al. found that this method was able to

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Root Development identify unknown biological functions for particular cell types using gene ontology (GO) category enrichment. For every gene in the genome of sequenced model organisms, gene functions are annotated through the use of GO categories, which are cross-species comparable labels based on homology and published descriptions of function (Ashburner et al., 2000). To determine whether a particular biological function is regulated at a frequency significantly higher than expected by chance, the hypergeometric distribution is used to allow a comparison of the distribution of expected values to those observed. One difficulty with taking a supervised approach is that patterns of expression not originally anticipated may be missed. Brady et al., therefore, developed a novel computational pipeline to identify gene expression patterns that are prevalent in the expression data set. These ‘dominant’ expression patterns represented common transcriptional programs in the root. Interestingly, 34 of the 51 dominant patterns identified displayed peak expression that spanned multiple cell types, including several patterns that demonstrated transcriptional similarities amongst disparate cell types. For example, one particular pattern exhibited peak expression in hair and xylem cells and showed enrichment in genes involved in cell wall deposition. These common expression patterns hint at shared regulatory pathways that are active in these tissues. Regulation of the plant hormone, auxin, was also found to exhibit exceptional spatial complexity. Auxin biosynthesis was enriched in the QC, lateral root primordia, pericycle and phloem-pole pericycle, while auxin-conjugating enzymes involved in regulating pools of active auxin showed peak expression in the columella. Conversely, genes associated with auxin transport and transport regulation showed enriched expression in the QC. Brady et al. extended the use of their computational pipeline to identify dominant expression patterns along the longitudinal axis of the root. Previous attempts to analyze gene expression along this axis utilized tissue pooled from multiple roots (Birnbaum et al., 2003). Root development is highly variable, however, and pooling may inadvertently result in tissues of different developmental stages and transcriptional states being combined together. In addition, while certain biological functions such as cell-cycle activity and root hair initiation have historically been used to define specific developmental zones along the longitudinal axis, it is unclear whether these zones define the spatial regulation of other biological functions. In fact, Brady et al. found that many of the 40 dominant expression patterns identified by their computational pipeline showed peak expression in subsets of the previously defined developmental zones. The unsupervised approach taken also led to the finding that 17 of the dominant patterns showed multiple peaks of expression along the longitudinal axis. This result contrasts with the traditional view of root development as a unidirectional process. Instead, it appears that certain transcriptional programs may be required at multiple phases of development, some of which may pattern periodic structures such as lateral root primordia.

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12.4.5

Other methods for tissue-specific expression profiling: laser capture microdissection

The use of the cell-sorting methodology is limited by the availability of fluorescent marker lines and the ability to efficiently dissociate cells by protoplasting. Another technique to obtain cell/tissue-type-specific transcriptional profiles is laser capture microdissection (LCM). In LCM, tissue is lightly fixed, sectioned and annealed to a microscope slide. Cells from the tissue sections are collected by physical attachment to a thermoplastic film with a low-power laser beam. The number of cells obtained using LCM is often limiting, thus requiring 2–3 rounds of linear amplification to prepare enough cRNA probe. LCM has been used in maize to characterize the transcriptional effects in the pericycle of a mutant that lacks lateral roots (rootless with undetectable meristems 1 (rum1)) (Woll et al., 2005) and 163 differentially expressed genes between the mutant and wild type have been detected. Of the highly expressed genes in wild-type pericycle, the most frequent functional classes were metabolism, transcription and cellular transport. Particularly interesting, several genes involved in the cell cycle were significantly repressed in the rum1 mutant pericycle when compared to wild type. LCM has also recently been applied to compare the expression profiles of collected cells from the apical and basal regions of globular- and heartstaged Arabidopsis embryos (Casson et al., 2005). The expression of genes that mark either the apical or basal poles of the embryo was enriched in the proper domains based on the microarray data. Analysis of these data should allow for identification of genes involved in the early patterning steps that distinguish the root from the shoot.

12.4.6

Environmental regulation of cell-type-specific gene expression

Roots are intimately associated with their environment and exhibit substantial developmental plasticity in response to changes in growth conditions. In addition to lateral root development, root growth, cell shape (Burssens et al., 2000) and gravitropism (Sun et al., 2008) can all be influenced by the various environmental stimuli that constantly bombard the root. As described previously, high-resolution transcriptional profiling of roots has clearly shown that each cell type controls a select set of biological functions important for growth under standard laboratory conditions. An important question to be addressed is what the regulatory mechanisms are that control these spatially complex patterns of expression? While developmental pathways are likely to play an important role in determining the transcriptional state of a cell, environmental conditions may also act as important cues. To address the role of environmental stimuli in regulating the transcriptional states of cells, Gifford et al. and Dinneny et al. utilized the cell-sorting methodology to generate transcriptional profiles of the various cell layers

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Root Development before and after treatment with an environmental stimulus (Dinneny et al., 2008; Gifford et al., 2008). Gifford et al. utilized a nitrogen induction system to simulate the presentation of nitrogen, an essential macronutrient. Dinneny et al. characterized the effect of two different abiotic stresses: high salinity, a pervasive agricultural contaminant and deprivation of iron, an important micronutrient. Interestingly, treatment of roots to each stimulus results in a massive transcriptional response that is, to a large degree, undetectable in whole root transcriptional analyses. The majority of transcriptional responses are cell-type specific, while responses detectable by whole root transcriptional analysis tend to occur in multiple cell layers. Thus, cell sorting is able to identify more responsive genes because the response is no longer diluted out. In addition, use of cell sorting was able to uncover genes with mixed regulation (Gifford et al., 2008). For example, the auxin response factor, ARF8, is activated by nitrogen treatment in the pericycle and lateral root cap, but shows repressed expression in the stele. The identification of more stimulus responsive genes revealed novel regulatory modules, which may be important for enabling the root to adapt to the change in environment (Fig. 12.4). Salt stress, for instance, results in upregulated expression of genes involved in reactive oxygen species (ROS) metabolism in the stele, indicating that this tissue may be an important center for ameliorating the toxic effects of high salinity (Dinneny et al., 2008). Irondeficiency, on the other hand, results in the activation of scavenging pathways such as metal–ion transport and nicotianamine (iron–chelator) biosynthesis in the epidermis. Nitrogen availability is known to affect root architecture and therefore is likely to have a cell-type-specific regulatory effect on tissues such as the pericycle. Consistent with this hypothesis, it was found that the pericycle was one of the most responsive tissues in this data set (Gifford et al., 2008). A clue as to how this regulation might occur came from the discovery that ARF8 was upregulated in the pericycle. From previous studies, it was known that ARF8 plays a role in shoot development (Wu et al., 2006) and as discussed before, auxin signaling is important for regulating LRI. When arf8 mutants were examined it was found that differential regulation of lateral root outgrowth by nitrogen is abolished. Because ARF8 is known to be repressed by a microRNA (miR167) (Wu et al., 2006), this suggested a potential mechanism for nitrogen regulation of ARF8 through miR167. To test this, Gifford et al. examined miR167 pre-microRNA expression and found it to be repressed by nitrogen treatment. Furthermore, plants that overexpressed miR167 are insensitive to nitrogen treatment similar to arf8 mutants. Gifford et al. followed up these experiments by transcriptionally profiling arf8 mutants after nitrogen treatment to identify likely targets of this TF. Thus, cell-type-specific expression profiling was useful for identifying an important transcriptional module regulating nitrogen-mediated developmental plasticity. In the evolutionary analysis of genomes, identification of DNA sequence that is conserved between diverse organisms typically indicates that the

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sequence is important for controlling some shared function. Likewise, the preservation of cell-type-specific expression between diverse environmental conditions may suggest that the biological function encoded by the gene is broadly important in that cell type. Based on this hypothesis, Dinneny et al. utilized the salt-stress and iron-deprivation data sets to screen through the many genes that show cell-type-specific expression under standard conditions to identify those genes that consistently define each cell layer (Dinneny et al., 2008). It was found that only 15% of the biological functions that define each cell layer under standard conditions are conserved across both stresses. Thus, environmental cues are crucial for determining the transcriptional outcome of cell fate decisions. This analysis also led to the identification of core biological functions that consistently define each cell layer (Fig. 12.4) and the finding that regulators of cell-fate decisions tend to exhibit environmentindependent expression patterns. These data suggest that, through the examination of the response of cell types to diverse environmental stimuli, it may be possible to identify additional genes that are important for determining the essential features of a cell type. Together, the work presented by Gifford et al. and Dinneny et al. supports the hypothesis that studying classical physiological stimuli from a developmental perspective not only provides rich information for better understanding how roots respond to a changing environment, but also benefit our understanding of the process of cell-type specification. 12.4.7

Potential uses of cell-type-specific expression data sets

The compendium of expression profiles for the various cell types of the root allows one to not only explore how gene expression is regulated in an organ, it also provides a resource that can be used to characterize genomic data sets obtained by other methods. For example, after identifying genes that are differentially expressed in a mutant of interest, the normal expression pattern of these genes can then be inferred from the compendium of sorted expression profiles. A short list of the best candidates to focus on for later experiments can then be obtained by identifying genes whose expression pattern overlaps with the expected domain. Finally, if the function of the gene of interest is unclear, the expression pattern of the candidate targets can help identify tissues that are most severely affected in the mutant, thus allowing a more targeted phenotypic analysis.

12.5

Building transcriptional networks II: exploring the role of transcriptional and posttranscriptional mechanisms in regulating TF activity

One caveat for all transcriptional profiling experiments is that mRNA expression does not perfectly correlate with the activity of the encoded protein.

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Root Development Posttranscriptional modifications of proteins, such as phosphorylation, ubiquitination and myristoylation, can all affect a protein’s activity. Furthermore, in plants, some proteins are capable of translocating to neighboring cells through the plasmodesmata, small channels that connect the cytoplasm of cells (Wu et al., 2002; Gallagher and Benfey, 2005). More recently, sequences within an mRNA have been discovered that can signal it for degradation via the microRNA degradation pathway (Chen, 2005; Vaucheret, 2006). To address the frequency of some of these processes in regulating gene activity, Lee et al. (2006) produced two types of gene fusions with GFP to monitor the transcriptional activity of a promoter separately from mRNA stability and protein movement (Lee et al., 2006). In the first set, 3000 base pairs of the upstream noncoding region was fused to endoplasmic reticulum localized GFP (erGFP). This set of 61 transcriptional fusions allowed the activity of the cloned promoter fragments to be monitored and the patterns of GFP expression to be compared with the mRNA expression profiles obtained through cell sorting. In the second set, the same promoter fragments were used as those used for the transcriptional fusions, but the coding sequence (cDNA) of the gene was also fused in-frame to a nontargeted GFP sequence. These translational fusions expressed the cDNA:GFP chimeric transcript in the same domain as the corresponding transcriptional fusions, but mRNA/protein stability and movement now could affect the pattern of GFP fluorescence detected. Thus, by comparing the GFP patterns between the transcriptional and translational fusions, posttranscriptional regulation of gene activity could be inferred from any differences observed. Furthermore, depending on the type of changes, different posttranscriptional regulatory mechanisms could be deduced. For example, if the translational fusion exhibited GFP fluorescence in a more expanded region of the root than that of the transcriptional fusion, the TF is probably capable of translocating between cells. On the other hand, if the domain of GFP fluorescence is reduced in the translational fusion, mRNA or protein degradation might regulate gene activity. To compare the microarray data obtained by cell sorting and the image data obtained from the GFP reporter lines using a confocal microscope, two methods were developed. The first method relied on visual inspection of the image data to determine the presence or absence of GFP expression in each tissue. The presence of GFP expression in the various tissues was then compared to the ranking of expression values obtained in the cell-sorting microarray experiments. Using this method, 80% of the 44 transcriptional marker lines were found to exhibit GFP expression patterns that matched the array data. The second method utilized image analysis software to extract intensity values for four different tissue regions of the root elongation zone (Mace et al., 2006). By these methods, image data from 24 transcriptional fusions were analyzed of which the majority correlated well with the sorted cell expression profiles. Thus, in most cases, 3 kb of upstream promoter sequence is sufficient to recapitulate the expression pattern of genes in Arabidopsis.

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An analysis of the translational fusions found that 39 out of 59 translational fusions generated displayed a consistent and observable GFP expression pattern. Of these, 10 lines displayed an expression pattern that differed from the transcriptional fusion, suggesting possible posttranscriptional mechanisms controlling expression. In two of these lines, the GFP expression for the translational fusion had a reduced domain compared to that of the transcriptional fusion. Both genes belong to a group of microRNA-regulated class III HD-ZIP TFs (McConnell et al., 2001; Emery et al., 2003; Mallory et al., 2004); thus, the reduction in the expression domain is probably caused by microRNA-regulated degradation in the tissue layers in which the transcriptional and translational fusions do not match. Eight translational fusions had an expanded GFP expression domain compared to that of the transcriptional fusion. For four of these genes, the transcriptional fusion produced an expression pattern that matched the mRNA expression based on cell sorting, thus the promoter fragment used was sufficient to drive expression in the normal domain. However, the expression domain for the translational fusions had expanded into the surrounding tissue layers. These cases might represent instances in which sequences in the coding region allowed for protein translocation between cells (Lee et al., 2006). Consistent with this interpretation, one of the four TFs, CAPRICE, has been previously documented as having the ability to move between nonhair cells and hair cells in the epidermis (Kurata et al., 2005).

12.6

Building transcriptional networks III: identifying direct targets of TFS – the SHORTROOT pathway

Genomic information sources have recently been combined to characterize the transcriptional network regulated by the SHORTROOT (SHR) TF (Levesque et al., 2006). SHR is a member of the GRAS family of putative TFs and regulates the specification of the QC (Helariutta et al., 2000). SHR also plays an important role in a specific set of stem cells, the CEI, where an asymmetric cell division is triggered to establish the two ground tissue layers. In the endodermal lineage, SHR additionally promotes the specialized features of this cell layer, such as the formation of the Casparian strip. SHR controls QC and CEI development in part by inducing the expression of another TF of the GRAS family, SCARECROW (SCR), which is expressed in the QC, CEI and endodermal cell layer (Di Laurenzio et al., 1996). SHR itself is expressed in tissue layers internal to the SCR domain, in a subsection of the stele. The SHR protein, however, has been discovered to have an expanded presence, moving from the stele into the QC, CEI and endodermal layer (Nakajima et al., 2001; Gallagher et al., 2004; Sena et al., 2004). Thus, SHR acts non-cell autonomously to promote SCR expression. To identify genes that are directly regulated by SHR, three microarray data sets have been developed that examine the effects of modulating SHR

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Root Development function in different ways (Levesque et al., 2006). Combining information from multiple independent data sets, a method termed meta-analysis increased the statistical confidence in the genes identified as SHR targets. The first data set was generated via a chemically inducible system in which the SHR-coding region was fused in-frame to the glucocorticoid receptor (GR). The GR fusion protein is normally sequestered in the cytoplasm, preventing the associated TF from entering the nucleus and activating transcription. The root can then be treated with a synthetic hormone, dexamethazone (DEX), which causes the fusion protein to enter the nucleus and regulate expression of target genes. The SHR:GR fusion was expressed in the normal expression domain of SHR (pSHR::SHR:GR, indicated as SHR:GR hereafter) and fully complemented the mutant phenotype of a shr null allele. SHR:GR-shr seedlings were first grown without DEX and then transferred to media with or without DEX. Cyclohexamide (CYC) was added to both media to prevent translation, ensuring that any transcriptional change occurring is due to the activity of SHR:GR, rather than to that of downstream TFs indirectly influenced by SHR:GR. Genes found to be regulated in this data set were designated as ‘direct targets’. The second data set was obtained by transcriptional profiling of shr mutant roots 5 days after germination, when phenotypic defects become apparent. Finally, the transcriptional effects of ectopic SHR activity were assayed with a transgenic line in which SHR:GFP is expressed from the WEREWOLF (WER) promoter (Lee and Schiefelbein, 1999), driving expression in cells of the epidermis, lateral root cap, and epidermal/LRC initial cells. GFP expressing cells were isolated using the cell-sorting method (Birnbaum et al., 2005) and compared to cells sorted from WER::GFP roots. Previous analysis of WER::SHR transgenic roots had found that ectopic SHR activity is sufficient to promote aspects of endodermal identity in the epidermal cell layer, which is consistent with the normal function of SHR in radial patterning (Sena et al., 2004). A common method for comparing microarray data sets is to identify genes that are differentially expressed in each individual experiment using a significance threshold. Subsets of genes are then identified with significant expression changes in multiple experiments. This method is conservative in that information between experiments is not used. For example, if a gene is differentially expressed in every experiment, but does not pass the significance cut-off in any one of them, it will not be identified. If, however, the consistency with which expression is regulated across all experiments is considered, the gene may pass the threshold. An inverse chi-squared method (Fischer, 1950) was used to generate a significance estimate for differential expression based on all three experiments (Levesque et al., 2006) and eight candidate targets were identified that are upregulated by SHR. Direct regulation of these candidates by SHR was tested using ChIP followed by RT-qPCR (reverse transcription followed by quantitative polymerase chain-reaction) to determine whether SHR is bound to their promoters under normal conditions. Of the eight candidate targets, SHR was found to directly bind to the

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promoters of four of them, including SCR, a gene encoding a receptor-like kinase (RLK), and two closely related genes encoding C2 H2 zinc-finger TFs, NUTCRACKER (NUC) and MAGPIE (MGP). The expression pattern of the SHR direct targets was analyzed using the sorted expression profiles for all of the genes and by in situ hybridization for NUC and MGP. For six of the eight genes identified as putative direct targets and for all the genes shown to be directly regulated by SHR using ChIP, significant expression enrichment in cell types that overlap with the SHR domain and the expression domain of the target genes was found, further strengthening the evidence that SHR regulates the expression of these genes. From this analysis many SHR targets were found to be expressed predominantly in the stele, which houses the vasculature (Levesque et al., 2006). These results suggest that while previous characterization of the shr mutant focused mainly on the role of SHR in root length and radial patterning of the cortex and endodermal tissue layers, SHR may also play an important role in patterning internal tissues. To examine this more closely, several GFP reporter lines that marked subpopulations of cells in the stele were examined. These observations, combined with measurements of the size of the shr stele demonstrated developmental defects in the stele. Thus, knowledge of the expression domain of transcriptional targets helped to identify new roles for SHR in root development. Finally, indirect targets of SHR were analyzed by GO category enrichment to obtain a picture of what kinds of developmental pathways SHR might ultimately regulate. This analysis showed that genes encoding proteins with TF activity or involved in phosphorylation processes are overrepresented in the list of SHR indirect targets. Interestingly, when the expression pattern of the indirect targets was characterized with the compendium of sorted expression profiles (Birnbaum et al., 2003), no clear enrichment in expression could be seen in cell types in which the SHR protein is present. Thus, while the immediate function of SHR is largely limited to the cell layers in which it is expressed, indirect regulation of secondary targets ultimately leads to SHR activity affecting all tissues of the root.

12.7

Exploring gene function using genomic variation: quantitative trait locus analysis of root growth

Knowledge of the complete sequence of a genome not only facilitates the cloning of typical laboratory-induced mutations affecting root development, but it also facilitates mapping and cloning genetic loci identified through the study of natural variation (quantitative trait loci, QTLs) (Alonso-Blanco et al., 2005). Many strains have been collected for the various model plant systems. In Arabidopsis, for example, ecotypes (strains) have been collected from around the world, each displaying variation in important developmental

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Root Development traits, such as flowering time and light sensitivity. To identify the molecular determinants of such variation, two ecotypes, typically with the largest discrepancy for the trait of interest, are crossed together to produce an F1 hybrid. The F1 plant is allowed to self-fertilize and several hundred F2 progeny are grown. Near isogenic lines (NILs) are generated through successive rounds of selfing and single seed descent. The end result is a collection of NILs that are homozygous for most loci in the genome. The NILs are genotyped with markers spaced throughout all the chromosomes. Once genotyped, the NILs can be phenotyped for any quantitative trait of interest. Since the NILs are essentially true breeding, they can act as a resource for QTL analyses studying many different types of traits. In Arabidopsis, studies on natural variation affecting root growth have largely focused on environmental stresses and only a few have focused on differences in root architecture under standard conditions (Mouchel et al., 2004; Loudet et al., 2005; Fitz Gerald et al., 2006). By studying the length of the primary root, Mouchel et al. (2004) found two- to threefold variation in root length among 40 Arabidopsis ecotypes. In particular, one ecotype, Umkirch-1 (Uk-1) developed the shortest roots that were measured. A segregation analysis of the F2 progeny of a cross between the Uk-1 and Slavice (Sav) ecotypes revealed that the Uk-1 short-root phenotype segregated in a 1:3 ratio, indicating that a single locus, designated BREVIS RADIX (BRX), was responsible for most of the variation in root length observed. The locus was mapped using genomic sequences to design molecular markers that were polymorphic between the two ecotypes used in the QTL analysis and BRX was found to encode a member of a novel plant-specific family of putative TFs (Mouchel et al., 2004). The Uk-1 allele of BRX, brx, was shown to confer shorter roots due to smaller cells and fewer cell divisions. A transcriptional analysis of brx roots was performed in order to understand the mechanism responsible for the reduction in root growth (Mouchel et al., 2006). Amazingly, the brx allele significantly affected the expression of ∼15% of the genome at least twofold. Of the most affected genes, the CONSTITUTIVE PHOTOMORPHOGENESIS AND DWARF (CPD) gene stood out as a possible determinant of the short phenotype of brx roots. CPD encodes an enzyme in a rate-limiting step of the synthesis of the plant hormone, brassinolide (BL) (Szekeres et al., 1996; Tanaka et al., 2005). Consistent with BL playing an important role promoting root growth, cpd mutants have shorter roots as do plants with a mutation in the BL receptor, BRI1 (Szekeres et al., ¨ 1996; Mussig et al., 2002). To test whether reduced BL levels might cause the brx phenotype, brx roots were grown in the presence of additional BL which resulted in a significant rescue of the root length (Mouchel et al., 2006). Most impressively, when brx roots were grown with added BL and transcriptionally profiled, nearly all of the transcriptional changes elicited by the brx allele were suppressed. BL is thought to partly modify the response to auxin, another plant hormone known to play an important role in root development (Nakamura et al., 2003, 2006; Wang et al., 2005). While previous studies

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showed that auxin had little effect on the growth of brx roots, analysis of the expression of an auxin reporter (DR5::GUS) determined that auxin response was strongly reduced in brx root apices. Importantly, addition of both BL and auxin to brx roots restored DR5::GUS expression to normal levels indicating that BRX-dependent production of BL is necessary for a strong auxin response. These data demonstrate a functional relationship between BL and auxin, with BL regulating the responsiveness of the root to auxin and BRX playing an important role regulating the crosstalk between these two hormonal pathways.

12.8

Future directions: from single gene biology to systems biology

What one gets out of using genome-scale data sets depends on the questions being asked. Having the full genomic sequence of an organism and the technologies to explore the function of any gene allows a reductionist biologist to ask many questions in a rapid manner. For example, a typical reductionist approach to understanding root patterning would entail genetic screens to identify mutants in which the normal development of cell types is altered. Having a sequenced genome allows one to map the mutation to a small region of the genome, sequence a list of candidate genes and identify the exact molecular lesion causing the mutant phenotype. It would be possible to use DNA microarrays to characterize the expression of genes misexpressed in the mutant or in a gain-of-function background. This type of experiment would typically lead to a list of candidate targets that could be studied by identifying insertion mutants in these genes. These mutant lines would then be characterized to understand subsequent steps in the pathway. Systems biology is in many ways a frontier field without defined methods or limits. Kirschner (2005) stated, ‘Systems Biology is the study of the behavior of complex biological organization and processes in terms of molecular constituents’. This definition might seem a bit vague and all encompassing. In some ways, a systems approach can best be understood in practical terms. In the above-mentioned example of a reductionist approach, the goal of the genome-scale experiments is to immediately reduce the scope of analysis to a few components of the system. However, there is richness in genome-scale information that can serve other goals. Thus, a systems approach would try to unravel the effect of a specific mutation by understanding its effect on the entire system. Another goal of systems biology is to integrate large data sets together to obtain a better understanding of the biological system as a whole. In addition to genome-based information, systems biology attempts to integrate metabolomics, proteomics and even ecological data sets. One of the best examples of a systems approach being used to study plant development comes from the characterization of the global effect of various

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Root Development hormones on gene expression in Arabidopsis (Nemhauser et al., 2006). Comparison of the gene sets that are regulated by the different hormones revealed that, while many hormones affect similar biological processes, there is little overlap in the specific genes they target. Thus, hormone signal-transduction pathways probably do not converge during the initial stages of hormone response. This model of the structure of hormone pathways in plants required both large-scale data sets and a nonreductionist analysis to be formulated. Systems biology, however, does not replace reductionism as data generated from this approach represent the foundation of biology. It is, nevertheless, a useful way of thinking about complex biological processes and an innovative way of doing large-scale biology.

Acknowledgments We thank Miriam L. Gifford and Kenneth D. Birnbaum for sharing unpublished results. We also thank S. M. Brady, J. Y. Lee, T. Long and A. Pillai for careful review of this manuscript. Funding for work on genomic approaches to understanding root development in the authors’ laboratory comes from grants from the National Science Foundation and the National Institutes of Health to PNB and the National Research Foundation of Signapore to JRD.

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INDEX

β-expansin, 187 β-glucans (β-glu− mutants), 245 β-glucuronidase (GUS), 22 staining, 86, 99–100, 106 γ -amino butyric acid, 138 γ -glutamylcysteine synthase, 25 1-aminocyclopropane-1-carboxylate (ACC), 141, 268, 300 1-N-naphthylphtalamic acid (NPA), 3, 87, 91, 93–4, 98, 100, 102, 204, 222, 227, 265, 332 1-naphthalene acetic acid (NAA), 96, 100, 104, 163, 332 1-naphtoxyacetic acid (1-NOA), 227 2-(4-carboxyphenyl)-4,4,5,5tetramethylimidazole-1-oxyl-3-oxide (CPTIO), 99 2,3-norbornadiene, 141 2,4-dichlorophenoxyacetic acid (2,4-D), 3, 144 resistant mutant 109, 185 3’-PHOSPHOINOSITIDE-DEPENDENT KINASE-1, PDK1, 74 3-hydroxy-3-methylglutaryl CoA reductase 1 (HMGR1), 252 3,4,5-triiodobenzoic acid (TIBA), 102, 140, 222, 227, 265 9-cis-epoxycarotenoid dioxygenase 1, LeNCED1, 145 14-3-3-like protein, 137 aba2-1, 102 aba3-1, 102 abi2, 103 ABI3, abi3, 103 ABI4, abi4, 308 ABI5, abi5, 103, 308 abi8, 103 abiotic factors, 131 stresses, 338 abscisic acid (ABA), 98, 102–4, 106, 141, 269, 308 acid phosphatases, 89 acidification of cell walls, 72 acropetal transport, 92 actin-depolymerizing drugs, 73 actinomycete, 223 actinorhizae, 209 actinorhizal symbioses, 228 adaxialization/abaxialization concept, 53 adenosine diphosphate (ADP)-ribosylation factor-GTPase, 47 adhesin-like polypeptides, 220 ADP-ribosylation factor 1, 74 adventitious rooting related enzyme-1, ARRO-1, 144

352

adventitious roots, 128, 176 ADVENTITIOUS ROOTLESS 1, ARL1, arl1, 146, 182–3 aerenchyma, 226 aerial roots, 289 affected lateral root formation 3, alf3, 85, 89, 101, 106–7, 110 affected lateral root formation 4, alf4, ALF4, 85, 89, 97 Affymetrix, 57, 328, 329, 332 AFLP markers, 143 AGAMOUS LIKE 16, AGL16, 309 AGAMOUS LIKE 21, AGL21, 309 agave, 290 AGL42 MADS-box, 335 agravitropism, 258 root phenotype, 165 Agrobacterium, 222 rhizogenes, 136 AHP6, AHP6, 51 alfalfa. See Medicago sativa Alfin1 Allium, 263 cepa, 107, 129 porrum, 215–17 sativum, 129, 215 Allocasuarina verticillata, 223, 227 almond (Prunus dulcis), 137 Alnus, 224–5, 229 glutinosa, 227–8 altered carbon allocation, 105 ALTERED MERISTEM PROGRAM 1, AMP1, amp1, 7 ALTERED PHLOEM DEVELOPMENT, 53 ALTERED RESPONSE TO GRAVITY, ARG1, ARG1, 160 AMF colonization, 215 aminoethoxyvinylglycine, 141, 262 amiRNA, 327 ammonium, 243, 302–6, 308–9 amyloplast, 241, 243, 266 anaphase-promoting complex/cyclosome (APC/C), 256 Annexin1, Ann1, 258 ANR1, 309 ANTHRANILATE SYNTHASE b1, ASB1, 27 anticlinal divisions, 87, 92, 332 AP2 class, 296 AP2 domain transcription factor, 11, 106 AP2/EREBP (Apetala2/ethylene-responsive element-binding protein), 101

Index
353

AP2/ERF (Apetala 2/ethylene response factor), 254 apical meristem, 226 activation, 89 maintenance, 89 APL, 53, 57 apple (Malus domestica), 131, 137, 139, 140, 143, 144 appressoria, 214 APS1, 315 aquaporin, 144, 255 AR, 128 Arabidopsis (Arabidopsis thaliana), 2, 40–42, 68, 75, 85–8, 90–92, 96, 98–9, 101, 103–4, 107–111, 114–15, 130, 133, 135, 139–41, 143, 145–6, 157–9, 163–4, 167–8, 177, 179–80, 193, 196, 199, 202, 204–5, 213, 217, 241, 257, 259, 261, 267, 294–8, 300–302, 304, 306–9, 311–15, 326–9, 340, 343–4, 346 AtEPX18, 74 embryos, 337 EXPANSIN 7, AtEXP7, 74 genome sequencing, 325 HISTIDINE KINASE2, AHK2, 51, 55, 56 HISTIDINE KINASE3, AHK3, 51, 55 HISTIDINE KINASE 4, AHK4, ahk4, 49, 51, 142, 267 K TRANSPORTER 1, AKT1, 311 SULFATE TRANSPORTER 68, AST68, 315 XYLOGEN-PRODUCING, 49 arbuscular mycorrhiza (AM), 210, 215, 218, 223, 290 Arum-type, 214 Paris-type, 214 arbuscular mycorrhizal fungi (AMF), 210–12, 216–17, 229 arbuscules, 214 architecture, 109, 111–12, 114–15, 222, 228, 291, 293, 302 ARF, auxin response factor ARF1, 74 ARF7, 95, 165, 297 arf7, 165, 333 ARF7/ARF19, 95, 297 arf7/arf19 double mutants, 95, 165, 297 arf7/myb77 double mutants, 95 ARF8, arf8, 338 ARF10, 21 ARF16, 21 ARF17, 146 ARF19, 95, 165, 297, 333 ARF-GAP, 47 ARF-GEF, 47, 160 ARG1-like 2, ARL2, 160 ARGONAUTE1, ago1, 134, 144 arm2, 92, 96 ARR4, 267 ARR5, Arabidopsis response regulator 5, 267 artificial miRNAs, 327 Asplenium, 202 astray, 260–61 asymbiotic stage, 211

asymmetric division, 15, 87, 179, 341 ASYMMETRIC LEAVES2/LATERAL ORGAN BOUNDARIES, AS2/LOB, 146 ATAF1/2, 179 AtGP19, 167 ATHB14, 53 ATHB8, 53, 55 AtHTH18, 166 AtHTH19, 166 ATML1, 66 ATP sylfurylase, APS, 314 AtSIZ1, 301–2 AUX1, aux1, 2, 90–93, 100, 162–4 AUX1-like, LAX1, 265 AUX1-mediated loading of auxin, 94 auxin, 54, 75, 90, 92, 99, 101, 107, 110–11, 132, 139–40, 160, 162, 166, 184, 204, 217, 219, 222, 264–5, 296–7, 311, 314, 332–3, 335–6 biosynthesis, 335–6 carrier, 46, 159, 268 distribution, 71 efflux, 72, 300 and ethylene signalling, 75 gradients, 46 homeostasis, 145, 333 influx, 227 influx and efflux carrier, 163 influx carrier, 227 maximum, 10, 101 receptor, 184 reflux loop, 23 response element (AuxRE), 146, 165, 183 response factor, ARF, 46, 94–6, 146, 165, 183, 296–7, 333 response gradient, 159 signaling, 71, 94 signal transduction, 183 transport, 68, 93, 94, 102, 311, 336 transport inhibitor (ATI), 87, 258, 265 treatment, 89 routes, 5 auxin-binding protein 1 (ABP1), 184 auxin-conjugating enzymes, 336 auxin/indole-3-acetic acid (AUX/IAA), 3, 46, 94–6, 165, 168, 204, 333 auxin-induced gene expression, 86 genes, 95 auxin insensitive, axr2, 313 auxin resistant 4, axr4, 92 axr6, 7 auxin resistant axr3-1, 164 axialization, 180 AXR2/IAA7, 98 AXR3/IAA17, AXR3/IAA17, 96, 98 axr4, 309 AXR5/IAA1, 96 axr6, 7 Azolla, 199, 201, 202 BABY BOOM, BBM, 11 bacteria, 303 bacteroid, 242–3, 245, 256, 263

354
Index bald root barley, brb, 186–7 barley. See Hordeum vulgare basal auxin maximum, 9, 20 basal cell lineage, 5, 7 basal meristem, 110 basal root growth, 164 basic helix-loop-helix, bHLH, 57, 312 basipetal auxin transport, 23, 94, 102 Betulaceae, 223 BIG/TIR3, 106 bioinformatic framework, 335 bioinformatics, 166 biotic and abiotic components, 77 biotic and abiotic soil environment, 178 biotic factors, 136 blackberry (Rubus), 128 bodenlos, BDL, bdl, 6–10, 28 brace roots, 176 Bradyrhizobium japonicus, 264 branching hairs, 73 branching infection threads 1/poodle, bit1/pdl, 254 Brassicaceae, 228 Brassica juncea, 45 Brassica napus, 142 Brassica oleracea, 314 brassinolide (BL), 344 application, 98 brassinosteroid (BR), 26, 54, 98, 162, 270 receptor (BRI1), 56, 335 BRASSINOSTEROID INSENSITIVE1, 56 brefeldin A, BFA, 5, 100 BREVIS RADIX, BRX, brx, 26, 344–5 BRI1-LIKE1, 56 brushy1, bru1, 21 Bryophyllum, 128 bryophytes, 289 bulb, 130 burr medic. See Medicago polymorpha bZIP transcription factor, 261 C-S lyase, 145 C2 H2 zinc finger putative transcription factor, ZPT2-1, Zpt2-1, 15, 256, 263 cactus, 290 CADMIUM SENSITIVE2, 25 CAF-1, 22 calcineurin B-like calcium-sensing proteins, 74 calcium (Ca, Ca2+ ), 72, 131–2, 212, 253 calmodulin, 213 calmodulin dependent protein kinase (Ca/CaM-DPK), 252 dependent protein kinases, 74 spiking, 251–3, 270, 268 calmodulin, 184, 253 calyptrogen, 177 CaMV 35S, 49 canalization of auxin, 94 model/theory, 45, 47 CAN OF WORMS1, COW1, 74 CAPRICE, CPC, 69, 75, 107, 299, 341 carbon (C), 132

energy partioning, 106 nitrogen metabolism, 256 carrot, 107 Casparian strip, 341 Castanea sativa, 137, 146 CASTOR, 213, 251 Casuarina, 226 glauca, 225, 227, 229, 312 richardii, 195–200, 202, 204–5 thalictroides, 200 Casuarinaceae, 223 CCAAT-box binding factor (CBF/NF-Y/HAP), 256 Ccs52A, 256 Ceanothus, 226 CEGENDUO, CEG, 103 cell cycle, 98, 186, 216, 297 regulation, 332 regulatory proteins, 97 switch 52 kDa protein, 256 cell differentiation, 68, 332 cell division, 100, 185, 334 cell laser ablation, 205 cell lineage, 68–9 cell morphogenesis, 77 cell position, 68 cell proliferation, 333 cell shape, 337 cell sorting, 326, 334–5, 337, 342 cell-to-cell movement, 70, 77 cell type, 185 specification, 178, 339 cellulase, 299 cell wall, 73 central cylinder, 185 Ceratopteris, 201 richardii, 194 Cercocarpus, 226 cereal embryo, 177 genetics, 176 mutant, 187 root system, 177 cereals, 177, 182, 184 Cg12, 225 CgAUX1, 227 cGMP, 166 chalcone synthase, CHS, 139, 266 chelate Fe (III), 312 cherry. See Prunus avium chestnut. See Castanea sativa chitin, 246 chromatin assembly factor 1, 21–2 chromatin immunoprecipitation (ChIP), 328–9, 342–3 chromatin-remodelling factors, 21 chrysanthemum (Chrysanthenum), 135 citrate cycle, 185 citrus, 306 class III HD-ZIP, 52, 260, 341 CLAVATA1, CLV1, 261 CLAVATA3, CLV3, 18

Index
355

clonal deletion, 16 clonal lineage experiments, 87 clonal propagation, 129 CLV-like pathway, 19 CLV3-like peptide, CLE, 19 CO2 levels, 105 cochleata, coch, 258 coffee, 113 coiled hyphae, 214 colchicine, 87 cold storage, 135 coleoptilar embryo, 180 coleoptilar node, 183–4 coleorhiza, 177, 180 collateral vascular bundles, 182 columella, 19, 67, 158, 160, 166, 178 columella-like region, 200 common bean. See Phaseolus vulgaris common gardenia. See Gardenia jasminoides Comptonia, 225–7 confocal microscopy, 213, 326 CONSTITUTIVE PHOTOMORPHOGENESIS AND DWARF, CPD, cdp, 26, 344 Coomassie blue-stained two-dimensional gels, 183, 185 COP1, 105 corms, 130 corn, 197 cortex, 88, 177, 182, 332 cortex/endodermis, 15 cortex/endodermis initial (CEI), 89, 179, 203, 334, 341 cortical and endodermal tissue, 164 cortical cell, 68 cell type, 185 elongation, 186 cotton (Gossypium), 215 cottonwood. See Populus trichocarpa × deltoides CRE-family receptors, 55 crinkle, 260 cRNA probes, 334 crop, 115 growth, 76 improvement, 115 cross-table clustering, 333 CROWN ROOTLESS CROWN ROOTLESS 1, CRL1, crl1, 146, 182 CRL1/ARL1, 146, 183 crownrootless2, crl2, 183 crown roots, 176, 184, 305 and brace roots, 182 and lateral root development, 186 Cryptomeria, 129 CUC2, 179 cucumber (Cucumis sativus), 141 CULLIN, 8 currant (Ribes), 130 cuttings, 130 cyclin-dependent kinase (CDK), 20 CDKA;1, CdkA, 86, 97 CDKB2;1, 98 cyclins, 97

CYCA2;1, 86 CYCA2;4, 98 CYCB1;1, 22, 86, 97 CYCD, 20 CYCD3, 99 CYCD3;1, 97 cyclohexamide, 342 cyclophilin, 96 cyclosporin A, 96 Cyp83B1, 145 cytochrome P450, 145, 255 cytokinin (CK), 28, 49, 54, 99, 139, 141, 217, 219, 225, 264, 266–7, 305, 314 receptor, 49, 142 response 1, CRE1, 49, 51, 55, 260 CYTOKININ OXIDASE, 51 cytoplasm, 255, 298, 332 cytoskeleton, 73, 160 ER, 213 cytosol, 251, 310 alkalinization, 160 Datisca glomerata, 223 daughter merophytes, 198 de-differentiation, 195, 204 defense, 185 deficiency, 300–301 deglycosylated xylogen, 49 derivative merophytes, 199 desert plants, 290 determinate nodules, 242, 269 developmental plasticity, 84, 338 dexamethazone (DEX), 342 diageotropic, dgt, DGT, 96–7 diagravitropic, 157 diarch, 107 dicotyledonous, 128 differential display, 254 differentiation zone, 184 digital in situ, 334 distal elongation zone, 164 root stem cell maintenance, 18 stem cell fate, 21 DNA metabolism, 333 methylation, 330 microarray, 328–9 synthesis, 333 Does not Make Infection, DMI, dmi, 212–14, 251 DMI1, dmi1, 217, 251, 264 DMI1/DMI2, 213 DMI2, dmi2, 218, 251–2, 264 DMI2/SymRK, 229 DMI3, DMI3, dmi3, 213, 218, 251–3, 264 DR5 reporter line (synthetic auxin-responsive promoter), 3, 46, 86, 92, 99–100, 103, 110, 160, 227, 345 drought avoidance, 113 resistance, 114 stress, 112–14

356
Index drought inhibition of lateral root growth 3, dig3, 103 early auxin-responsive genes, 184 early nodulin, Enod, 244, 254 Enod2, 245, 265 ENOD11, Enod11, 212, 253, 254 Enod12, 265 Enod40, 245, 255–6 early post-embryonic root development, 184 eastern larch. See Larix decidua EC fungi, 223 ecotypes, 343 ectendomycorrhiza, 218 ectomycorrhiza (ECM), 134, 136, 210, 218, 221–2, 290 efflux carriers, 100 eir1-1, 300 Elaeagnaceae, 223 Eleagnus, 226 electron microscopy, 259 elg3, ELG3, 299 elongation, 186, 334 zone, 22, 166, 326 EMBO30/GN, 47 embryo, 70, 176 embryogenesis, 3, 65, 70, 177–8 embryonic phase, 176 roots, 176, 184 embryophytes, 289 emergence meristem activation stage, 103, 307 stage, 85 endocytic marker, 74 endodermis, 88, 177, 185, 332 endogenous glucose, 105 endoplasmic reticulum, 340 endosomal trafficking, 5 endosperm, 178 ENHANCER OF GLABRA 3, EGL3, 299 ENHANCER OF TRY AND CPC1, 69 enhancer-trap, 335 environmental regulation, 332 response pathways, 109 epidermis, 67, 68, 88, 165, 178, 185, 332, 338, 341–2 specification, 14 Equisetum, 193–4 era1, 103 ericoid associations, 290 ERF required for nodulation ERN1, 254 ERN3, 254 ETC1, 69 ethylene, 75, 139–40, 245, 261, 266, 268–9, 313 inhibitor, 262 precursor, 300 signaling, 27

ethylene insensitive, EIN2, 268 ethylene response 1, etr1, 269 Eucalyptus, 129, 137, 140 globulus, 131–5, 143 grandis, 134, 143 saligna, 133–5 sideroxylon, 133 tereticornis, 143 urophylla, 143 evolutionary adaptation, 288 exocyst, 186 exocytotic vesicles 186 exo− mutants, 245 exopolysaccharides, 245 expansins, 187, 299 expansion, 100 extraradical hyphae, 216, 218 F-box protein, 94, 103, 255 fasciata1, fas1, FAS1, 21 FAS2, fas2, 21 Fe deficiency, 313 starvation, 312 Fe-DEFICIENCY INDUCED TRANSCRIPTION FACTOR 1, FIT1, 312 FER, FER, fer, 312–13 ferns, 193 fertilizer, 76 ferulic acid, 139 FEZ, 14 Filicophyta, 193 FIT1, fit1, 312–13 fixation zone, 226 flacca, 145 flavin monooxygenase, 145 flavonoid, 245 pathway, 256 flavonone, 224 flowering, 186 time, 344 fluorescence activated cell sorting (FACS), 59, 180, 333–4 fluorescent DR5 reporter expression, 5 fluorescent marker lines, 326 fluorophore, 333 FM4-64, 74 forest trees, 137 formative divisions, 200 forward genetics, 178 founder cells, 86, 94, 110, 307 fountain model, 100 four-layered structure, 88 Frankia, 210, 223–7, 229 Fraxinus angustifolia, 138–9 mandshurica, 296 FRO2, FRO2, 312 fructose, 105, 133 fruit trees, 137 Fucus, 67 fungi, 210–11, 290, 303

Index
357

G-protein, 184 Rab type, 256 G1 stage (2C), 86 G2 stage (4C), 86 G2-to-M transition, 22, 99 galactolipid synthesis, 294 garden nasturtium. See Tropaeolum majus Gardenia jasminoides, 139 Garlic. See Allium sativum GDIs, 72 GEF, 47 Gene Ontology (GO), 336 category, 343 genetic engineering, 314 genome-scale strategies, 327 genomic data, 339 GH3, 222, 227 GH3-like, GHR3-like, 144, 146 gibberellic acid (GA), 18, 27, 141, 269, 335 biosynthesis, 335 gibberellin, 139, 217 gl3-egl3 double mutant, 299 GLABRA, 69 GLABRA2 (GL2), GL2, gl2, 69, 75, 299–301 GLABRA3 (GL3), 69, 299 globular embryo4, gle4, 179–80 Glomus caledonium, 215 intracadices, 217 mosseae, 216–17 glucocorticoid receptor (GR), 342 glucose, 105, 133 glutamate, 304 glutamine, 306 glutathione, 25 redox couple, 26 glycerophospholipids, 300 gnom/emb30, gn, 5 GNOM, GNOM, 47, 160 Golgi apparatus, 74 GPI-anchored proteins, 313 graded auxin distribution, 24 grapevine. See Vitis vinifera GRAS family transcription factors, 12, 179, 253, 341 gravistimulation, 110–11 gravitropic response, 182, 311 root growth, 164 signal attenuation, 166 gravitropism, 337 gravity, 157 stimulus, 159 green fluorescent protein (GFP), 333–4, 340–41, 343 marker line, 180 translational construct, 46 Grevillea robusta, 294 ground tissue, 341 patterning, 15 GS52, Glycine soja 52 kDa protein, 256 GSH (reduced form, oxidized form), 25–6 guanine nucleotide exchange factor, 47

H+ -pyrophosphatase AVP1, 115 H2 O2 , 138 hair (H) cells, 65, 298–300, 341 hair curling, hlc, 246 HAK family of potassium transporters, 72 halted root, hlr, 19 HAP2-1, 256 har1-1, 218 Hartig net, 218, 221–2 heart stage, 10 Hebeloma cylindrosporum, 222 hexose, 144 high cambial activity, hca, 56 high-resolution spatial maps, 331 high-resolution transcriptional profiling, 337 high salinity, 338 high-throughput sequencing technologies, 329 histidine phosphor, 51 hobbit, hbt, 7 homeodomain-leucine zipper transcription factor, 55 homeostatic redox buffer, 26 Hordeum vulgare, 76, 186–7, 217, 292, 306 hormone signal-transduction, 346 horse gram (Macrotyloma uniflorum), 132 HRGPnt3, 227 HvEXPB1, 187 hy4, 133 HY5, long hypocotyl 5, hy5, 56, 105, 261 hybridization to microarrays, 328 hydrogen peroxide (H2 O2 ), 166 hydrolytic enzymes, 89 hydrophobins, 220 hydroponic conditions, 309 hypaphorine, 219 hyper-geometric distribution, 336 hyperinfected 1, hit1, 267 hypernodulation aberrant root 1, har1, 218, 241, 259–60, 262, 270 hyperpolarization-activated calcium channel, 73 hyphae, 211, 213–14, 218, 220–21, 226–7, 225 hyphophodium, 213 hypocotyl, 41, 69, 99, 138, 261, 332 hypophysis, 6, 8 specification, 8 IAA, 91, 93, 97, 100–101, 104–5, 138, 140, 144, 163, 222, 225 IAA/Aux, 146 IAA14/SLR1, 165, 204, 297 IAA28, IAA28, 96, 98 IAA3/SHY2, iaa3/shy2-2, 165 ign1, ineffective greenish nodules, 257 image analysis software, 340 INCOMPLETE ROOT HAIR ELONGATION, 74 indeterminate nodule, 242, 266, 268 Indo-1, 72 indole-3-acetic acid (IAA), 90, 132, 204, 217, 308 indole-3-butyric acid (IBA), 92, 96, 132, 134, 138–40, 144, 204, 217

358
Index INDUCED BY PHOSPHATE STARVATION 1, IPS1, 302 infection thread, 225, 241–2, 252–4, 260, 269–70 zone, 226 initial, 67 initiation site, 71 stage, 85 inorganic phosphate, 65 inositol triphosphate phosphatase, 313 insertion alleles, 327 interfascicular fiberless1, ifl1, 55 internal Pi, 293 intrinsic pathways, 84, 109 invasion region, 242 inverse chi-squared method, 342 invertase, 105 IP3, 166–7 IRE, 74 iron (Fe) 65, 75–6, 131, 311, 338–9 deficiency, 75, 312 and phosphate stress, 75 IRON TRANSPORTER 1, IRT1, 312 IRREGULAR XYLEM, IRX1, 57 Isoetes, 194 isoflavone, 266 isopentenyl adenosine, iPA, 225 JACKDAW, JKD, 13, 17 jasmine (Jasminum), 130 jasmonic acid (JA), 217, 270 biosynthesis, 335 Juglans regia, 137, 138 K (potassium), 292, 310 availability, 310–11 deficiency, 310 deprivation, 311 patches, 310 KANADI, KAN,, 53 kinase, 334 Kip-related protein (KRP), 20, 97 KRP2, KRP2, 86, 97–8 kiwi (Actinidia sinensis), 137 klavier, klv, 260–61 KNOLLE, KN, kn, 67 KOJAK, KJK, 73, 75 LACS2, 103, 105 Lactarius deterrimus, 220 larch, 219 Larix decidua, 137 laser laser capture microdissection (LCM), 185–6, 205, 337 laser-induced ablation of QC, 13 laser-induced cell ablation, 18 late globular embryos, 10 late heart stage, 41 late nodulin, Nod, 254 lateral auxin gradient, 162, 165

LATERAL ORGAN BOUNDARIES (LOB), 183 lateral root (LR), 128, 176, 183–5, 193, 289–90, 296, 332 apical mother cell (LRMC), 203–4 associated nodule (LRAN), 243 base (LRB) nodulation, 243 cap (LRC), 158, 338 development, 241 elongation, 307 formation, 85, 182, 203, 218, 260–61, 264–5, 269, 297, 308 founder cells, 184 initation (LRI), 182, 184–5, 227, 257, 266, 307, 311, 332–3 primordia (LRP), 83, 92, 227, 259, 267, 296, 307, 332, 336 emergence, 88, 101, 106, 108, 111 initiation, 85, 87, 93, 111, 306 LATERAL ROOT DEVELOPMENT 2, LRD2, lrd2, 103, 105 lateralrootless1, lrt1, 184, 217 lateral root organ-defective, latd, LATD, 103, 218, 259–60, 269 latrunculin B, 75, 167 LAX1 (AUX1-like), 258 LAX2, 258 leaf borne roots, 196–7 heteroblasty, 196 LEAFY PETIOLE, LEP, 56 lectin, 220, 255 leek. See Allium porrum leghemoglobin, 255–6 legume, 212, 241, 262–3, 265, 268–9, 306 nodulation, 209 Rhizobium symbiosis, 213 lemon (Citrus medica), 130 leucine-rich-repeat (LRR), 213 receptor kinases, 261 LEUCINE-RICH REPEAT EXTENSIN1, LRX1, 73, 75 LRX2, 73 lrx1/lrx2, 74 light, 105, 133, 265, 344 lignin biosynthesis, 185 lin1, 307 LIPID TRANSFER PROTEIN1, 66 lipids, 74 lipopolysaccharides (lps− mutants), 245 liquid chromatography electrospray ionisation tandem mass, LC-ESI MS/MS, 183, 185 LOB domain, LBD, 95 LBD16, 95 LBD19, 95 loblolly pine. See Pinus taeda lodgepole pine. See Pinus contorta lodging resistance, 182 LONESOME HIGHWAY, LHW, 57 long-day conditions, 133 longitudinal

Index
359

axis, 334 position, 108–9 zones, 334 lopped, lop1, 68 Lotus Histidine Kinase 1, LHK1, 267 Lotus (Lotus japonicus), 213–14, 218–23, 241, 245, 251–7, 260–62, 264–5, 267, 269–70, 328 lowland rice, 113 LPD-specific inhibitor 1-butanol, 164 LRB nodulation, 269 LRB16, 95 LRC cells, 162 LRI gene set, 333 LTP1, 66 lupin (Lupinus albus), 115, 296, 315 Lycopodium, 193–4 Lycopodophytes, 194 LYK complexes, 251 LYK2, 246 LYK4, 246 LYK8, 251 LYK-related, LYR, LYR1, 251 LysM RLK genes, 246 lysophosphatidylcholine, 214 macronutrient, 338 MADS box family, 309 MAGPIE, MGP, 15, 343 maize (Zea mays), 88, 90, 107, 129, 112–14, 142, 176–7, 179–80, 182, 184–5, 196–7, 214–15, 217, 289, 294, 304, 306, 328, 337 MALDI-ToF, 185 manganese (Mn), 131 mannitol, 104 MAPK, 74 MAPKK, 5 Marsilea quadrifolia, 87, 204 maturation zone, 326 mature embryo, 42 MDR1, 102 MDR4, mdr4, 92, 102 Medicago, 103, 252–6, 259–61, 263, 266–7, 269–70, 315 polymorpha, 267 sativa, 74, 241, 245, 258, 263, 266 truncatula, 213–15, 217–18, 223, 246, 264, 268 Melilotus alba, 265 membrane permeable analogue 8-Br-cGMP, 166 meristem, 294 activation, 25, 101 proximal and distal, 177 region, 242 tissue, 101 transition zone, 28 meristematic zone, 226, 326 MERISTEM LAYER1, 15, 66 merophyte, 193, 197 mesocotyl, 176 Mesorhizobium loti, 251, 264 meta-analysis, 342 metabolomics, 345 metal-ion transport, 338

metaphase, 216 metaphloem, 42 metaxylem, 42 mgoun3, mgo3, 21 microarray, 144 analysis, 180, 293, 312, 330 data, 340–42 experiment, 186, 330 microarray analyses, 185 microdissection, 335 microfilaments, 73 micronutrient, 338 microorganisms, 114, 209, 262, 303 micro-RNA, 302, 315, 338 degradation pathway, 340 miR164, 96 miR167, 338 miR395, 314–15 miR399, 302 microsatellites, 143 microtubule, 73, 202 microtubule organisation 1/gemini pollen 1, MOR1/GEM1, 146 middle cortex, 18 mineral nutrition, 131 mitogen-activated protein kinase kinase, 5 mitotic cell cycle, 333 mitotic index, 88–9 ML1, 15 molecular markers, 180 monocotyledonous plants, 128, 187 MONOPTEROS, MP, mp, 6, 47 Monterey pine. See Pinus radiata movement of SHR, 17 mRNA stability, 340 msbA2 (multicopy suppressor of htrB mutations A), 245 MSG2/IAA19, 96 multiple testing, 330 mung bean. See Vigna radiata mutant phenome, 327 MYB77, 95 Myc, 229 factor, 212–13, 214, 251 myconodules, 229 mycorrhical fungi, 262, 290 interactions, 252 mycorrhizae, 209–10, 291 mycorrhization, 215–18 MYR1, 57 Myrica, 225–6 Myricaceae, 223 N availability, 304, 309 cells, 299–300 starvation, 303 NAC gene family, 179 OsNAC7, 180 ZmNAC5, 179–80 NADPH oxidase homolog AtrbohC, 73

360
Index NahG, 270 NAM, 179 NAP1-RELATED PROTEIN1, NRP1, NRP2, 22 narrowleaf ash. See Fraxinus angustifolia natural variation, 343 near isogenic lines (NILs), 344 negative feedback loop, 333 never ripe, 141 Nicotiana benthamiana, 253 tabacum, 49, 131, 137, 141, 213, 217, 227, 304, 306, 327 nicotianamine, 338 Nimblegen, 329 nitrate, 100, 102, 106, 111, 245, 262, 265, 302, 304, 306, 308, 309 availability, 307 patches, 111 reductase, 306 sensor, 99 transporter, 99–100, 307–8 nitrate tolerant symbiosis, nts, 259, 262, 267, 382 nitric oxide, 99, 132, 166 nitrilase 3, 314 nitrogen (N), 99, 105, 111, 131–2, 210, 223, 226, 228, 262, 292, 302–3, 338 availability, 306 deficiency, 133 fixing nodule, 241, 244 fixing symbioses, 210 salts, 104 nod− , 246 nod3, 261 Nod factor, 212, 218, 225, 229, 245, 251, 265–8, 270 perception, 254 signalling, 246 Nod factor perception, NFP, nfp, 218, 246, 251, 264 Nod factor receptor, NFR1, nfr1, 246, 251 NFR5, nfr5, 246, 251 nod mutants, 225 nodal roots, 176 nodulation, 218, 246, 267 in the absence of Rhizobium (NAR), 243 nodulation receptor kinase, NORK, 213, 251, 252 nodulation signalling pathway 1, nsp1, NSP1, 253–4, 264 NSP2, nsp2, 253 NSP3, 254 nodule, 224, 226, 251, 253, 257, 263, 269 formation, 260, 265 lobe primordium, 226 primordium, 241–2, 251, 253, 257, 261, 264, 268–9 nodule autoregulation receptor kinase, nark, 260 nodule inception, NIN, nin, 253–4, 267, 270 nodulin, 223, 254–6, 258, 266 non-hair (N) cells, 68, 298, 341 non-nitrogen fixing mutant (fix− ), 257 non-polar localized, 163 NONPHOTOTROPIC HYPOCOTYL 4, 11 notabilis, 145

Notch, 253 Nothofagus antarctica, 139 NPA resistant, 97 sensitive basipetal transport, 93 treated roots, 166 nuclear organization, 334 speckles, 255 nucleoporin 252 NUP133, NUP133, nup133, 213, 252 NUTCRACKER, NUC, 15, 343 nutrient, 77, 157, 211, 215, 228, 289, 290, 304 deficiency, 76, 105 oak. See Quercus robur Olea europeaea, 216 oligopeptide transporters, 144 onion. See Allium cepa open reading frames (ORFs), 136 organic acids, 114 organogenesis, 77 origin recognition complex, orc, 10 ornithine decarboxylase (ODC), 222 Orobanche, 211 orthostichies, 198 Oryza sativa, 93, 96, 112–15, 129, 140–43, 176–7, 179–80, 182–3, 185–6, 213, 294, 306, 315, 327 PINFORMED1, OsPIN1, 140 osmotic stress, 102, 104–5 OXIDATIVE SIGNAL-INDUCIBLE1, OXI1, 74, Papaver somniferum, 327 Parasponia, 257 parenchyma, 177, 243, 253 pattern formation, 77 Paxillus tinctorius, 136 PDF PDF1, 66 PDF2, 15, 66 Pea. See Pisum sativum peanut (Arachis hypogaea), 257 pectate lyase (PL), 89, 104 pedunculate oak. See Quercus robur pelargonium (Pelargonium), 132–3 Penicillium nodositatum, 229 pentarch symmetry, 107 PEP carboxylase, 256 peptidoglycan-binding motif, 246 peptidyl-proline cis-trans isomerase activity, 96 peribacteroid membranes, 263 periclinal divisions, 332 pericycle, 41, 86, 185, 227, 240–41, 290, 296–7, 332–4, 307, 338 cell, 93, 108, 176, 184 and cortical cells, 241 specific marker line Rm1007, 86 permeability, 105 peroxidase, 138 pH, 72, 76, 160, 167, 211, 223, 262, 308 PHABULOSA, PHB, 52

Index
361

pharmacological condition, 332 Phaseolus, 138 aureus, 134 vulgaris, 114, 132, 258, 262, 294 PHAVOLUTA, PHV, 52 phenolic compound, 139 phenotypic plasticity, 288 phenylpropanoid, 219, 228 pathway, 255 Philodendron, 197 phloem, 39, 91, 94, 259, 304, 334 pole pericycle, 336 phloroglucinol, 139 PHO2, PHO2, pho, 302 phosphate (P, Pi), 75, 106, 111, 216–18, 262, 292 availability, 292–4, 296, 298, 300 binding sequence, 301 deficiency, 111, 114 stress, 75 deprivation, 292, 299, 302 responses, 106 starvation, 295, 297, 300–2 transporter, 214, 293, 297, 301 AtPT1, 297 AtPT2, 297 LjPT3, 214 MtPT4, 214 uptake, 115 phosphate starvation response 1, PHR1, 301 phosphatidic acid, 300 phosphatidylcholine, 301 phosphatidylinositol 3-kinase, 166 transfer protein, 74 phosphocholine, 301 phosphoenolpyruvate carboxylase, 255 phosphoethanolamine, 301 phosphoglycoprotein (P-glycoprotein, PGP), 94, 162 AtPGP1, pgp1, 94, 164 AtPGP4, pgp4, 94, 162 pgp19, mdr1, 94, 167 PHOSPHOLIPASE D Z1, PLDZ1, phospholipase Dζ 1, 74, 300–301 PHOSPHOLIPASE D Z2, PLDZ2, phospholipase Dζ 2, 164, 301 phospholipid, 293 degradation, 301 hydrolysis, 294 phosphorus, 105, 131, 157, 228, 262, 292 deficiency, 75, 262 stress, 114 phosphorylation, 343 photomorphogenesis, 261 photosiderophores, 312 photosynthesis, 105 phytase, 114 phytochromes A, B, D, 105 phytohormone, 54, 139, 261–2, 266, 268, 270 Picea abies, 134 PICKLE, 95 Pi deficiency response 2, pdr2, 107

PIN family of auxin efflux carriers, 90, 100 PIN, PIN, pin, 2, 5, 23, 45, 94, 100, 106, 160, 265, 297 pin1/pin3 double mutants, 94 PIN1, PIN1, pin1, 5, 45–6, 55, 90, 93, 100, 268 localization, 6, 9 mediated polar auxin transport, 91 PIN2, PIN2, pin2, 91–3, 100–101, 162–4, 167, 265, 268 PIN3, pin3, 94, 100, 159–60, 167 PIN4, 46, 100, 160 PIN6, 100 PIN7, pin7, 5, 46, 94, 100, 160 PINFORMED, pin-formed, 2, 45 PINHEAD/ZWILLE-like, 144 Pinus, 137 contorta, 143–4 pinaster, 222 radiata, 137, 141, 143, 146 strobus, 130 sylvestris, 135–6, 140 taeda, 137, 142 virginiana, 136, 138 Pisolithus tinctorius, 135, 219 Pisum sativum, 107, 141, 241, 252–3, 258, 260–61, 263, 267, 269 plagiotropic, 157 plant hormones, 29, 219 plasma membrane, 252, 310 plasmodesmata (PD), 259, 340 plastids, 251 Platanus acerifolia, 215 pleiotropic phenotypes, 96 PLETHORA, PLT, plt, 3, 89, 106, 296 PLETHORA1, PLT1, 11, 296–7 PLT2, 11, 296–7 PLT3, 11 plt1/plt2 double mutants, 106 plt1-4/plt2-2 double mutant, 296 protein fusions, 24 PL genes, 104 PLA1, 89, 104 PLA2, 89, 104 ploidy, 180 PLS, 27 PODs, 138 polar exocytosis, 186 flow of IAA, 90 transport, 90, 264 polar auxin transport (PAT), 45, 265–6, 268 inhibitor, 332 POLARIS, 27 POLLUX, 213, 251, 254 polyamines (PAs), 138, 222, 301 polymerase chain-reaction, 342 poplar. See Populus poppy. See Papaver somniferum Populus, 129–30, 132, 140, 215–16, 219 tremula L. × P. tremuloides, 138 trichocarpa, 327 trichocarpa × deltoides, 142–4

362
Index positional information, 69 signal, 68 postembryonic phase, 176 root type, 176 posttranscriptional modifications, 340 potato, 214 PP2A, 9 prenodule, 226 prepenetration apparatus, 214 presymbiotic phase, 211 primary homorhizy, 195 root, 176, 180, 182–3, 185–6, 305 meristem, 107 tip, 88 primordia, 182 formation, 11 primordiamorph, 87 procambium, 41, 180 proembryo, 6, 8, 179 layer, 4 prohibitin, 186 proliferative cell divisions, 200, 202 prop root, 289 Proteaceae, 228 proteasome machinery, 20 protein movement, 340 phosphatase, 9 protein serine/threonine kinase PINOID, 9 proteome analyses, 185 proteomics, 167, 183, 345 protoderm, 65, 66 PROTODERM FACTOR2, 15, 66 protodermal development, 67 markers, 67 protophloem, 42 cells, 90–91 poles, 107 protoplasted cells, 334 protoplasting, 334 protoxylem, 42 poles, 261 Prunus avium, 138 cerasifera, 215–16 pseudonodules, 227 Psilophyta, 193 Psilotum, 193 Pteridophyta/Pteridophytes, 193–4 PUCHI, puchi, 101 putative chromatin remodeling factor, 95 putrescine, 138, 222 QTL analyses, 344 quantitative trait loci (QTLs), 104, 112–13, 115, 143, 343 Quercus robur, 137, 143

quiescent center (QC), 2, 43, 67, 177, 179, 294, 296, 334–6, 341 identity, 295 QUIESCENT-CENTER-SPECIFIC HOMEOBOX gene, QHB, 179 Rab Rab1, 256 Rab7, 256 RabA4b, 74 radial organization, 14 pattern, 67, 88, 178 position, 107, 109 radicle, 65 radicleless1, ral1, RAL1, 180 rapeseed. See Brassica napus Raphanus sativus, 204 ratiometric fluorescent calcium marker, 72 reaction-diffusion model/theory, 45, 47 reactive oxygen species (ROS), 73, 166, 311, 338 receptor-like kinase, RLK, 343 RECEPTOR-LIKE PROTEIN KINASE1, RPK1, 15 red clover (Trifolium pratense), 263 red raspberry (Rubus strigosus), 130 redox homeostasis, 25 reduced mycorrhizal colonization, rmc, 214 reduced root length reduced root length 1, rrl1, 186 reduced root length 2, rrl2, 186 regeneration, 14 of radial pattern, 179 regulatory hormones, 101 reporter lines, 333 repressor of GA signaling and root growth, 27 resistant to IBA 1, rib1, 96, 97 response regulator, 51 response to auxin, 3 RETINOBLASTOMA (RB), 20 RETINOBLASTOMA-RELATED, RBR, 20 REVOLUTA, REV, 52 RGA, 27 rhizobia, 210, 241, 260, 262–3, 267–8, 270 Rhizobium, 223, 225, 229, 245, 251, 254, 259 etli, 258 Rhizobium induced peroxidase, MtRIP1, 253 rhizome, 130 rhizosphere, 176 Rho-related GTPase, ROP, 72 Ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco), 303 rice. See Oryza sativa RIL, 115 RNA in situ hybridization, 334 interference (RNAi), 246, 254, 256, 265, 300 binding protein, RBP1, 255 root apex, 184 apical cell (RAC), 194, 197–9, 205 apical meristem (RAM), 177, 180, 185 apical mother cell (RAMC), 195–6, 198

Index
363

architecture, 262, 264, 271 338 cap, 67, 88, 177, 180 depth, 113 development, 176 elongation, 186 epidermis, 65 gravitropism, 158, 160, 167 inhibitors, 168 hair, 70 initiation, 336 length, 75 morphogenesis, 65 morphology, 2, 75 heteroblasty, 196, 202 inducer (RIND), 258 initial, 130 meristem, 65, 67, 129, 297 nodules, 210 stele, 107 system architecture, 175 root growth defective, rgd1, rgd2, rgd3, 145 root hair curling (RHC), 252–3, 263 nodulation, 269 ROOT HAIR DEFECTIVE RHD1, 75 RHD2, RHD2, rhd2, 73, 75–6, 311 RHD3, 75 RHD4, 75 RHD6, rhd6, 72, 75–6 roothairless roothairless1.a, rhl1.a, 186–7 roothairless2, rth2, 186 roothairless3, rth3, 186 root hairless 1, rhl1, 241, 264 root hair primordia, rhp1.a, 187 ROOT INITIATION DEFECTIVE 5, RID5, rid1-rid5, 145–6 rootless1, rt1, 182 rootless concerning crown and seminal roots, rtcs, RTCS, 180, 182–4 rootless with undetectable meristems 1, rum1, 98, 180, 182, 184–5, 337 root loci, rol, 136 rolA, 136–7 rolB, 136 rolC, 136–7 rolD, 136 root meristemless1, rml1, 25 root mutant 1, rm1, 186 root mutant 2, rm2, 186 root primordium defective1, rpd1, 145 ROOT REDIFFERENTIATION1, RRD1, 145 RRD2, 145 RRD4, 145 rootstock, 137 rooty (rty)/aberrant lateral root formation (alf1)/superroot1, 92 rose (Rosae), 141 RTCS-LIKE, RTCL, 183 RT-qPCR (reverse transcription followed by quantitative PCR), 342

Rumex palustris, 141 rye. See Secale cereale, 76 S-acyl transferase, 74 S-adenosyl-L-methionine:phosphoethanolamine N-methyltransferase (PEAMT), 301 salicylate hydoxylase, 270 salicylic acid (SA), 270 Salix, 129–30 Salk Institute Genomic Analysis Lab (SIGnAL), 327 salt, 262, 339 stress, 263, 338 SAUR, 184 SCARECROW-LIKE, SCR-LIKE, 146 SCARECROW, SCR, scr, 12, 15, 89, 106, 146, 178–9, 202–3, 341, 343 AtSCR, 146, 179 OsSCR, 179 ZmSCR, 179–80 SCR::GFP, 334 SCFTIR1/AFB ubiquitin-ligase complex, 165 schizoriza, SCZ, scz, 68 SCN1/RhoGDI, 72 Scots pine. See Pinus sylvestris SCRAMBLED, SCM, 69 scutellar node, 176–7 SEC3, 186 Secale cereale, 76 secondary homorhizy, 193, 197 thickening, 65 vascular tissues, 44 secretory vesicles, 186 sedimentation of statoliths, 159 seedlings, 102 segregation analysis, 344 Selaginella, 193–4 seminal crown and brace roots, 182 roots, 176, 305 senescence zone, 226 serial analysis of gene expression, (SAGE), 255 serine/threonine kinase, 74 Sesbania rostrata, 243, 264, 268–9 sheath, 218 shoot apical meristem, 93 biomass, 76 excision, 93 heteroblasty, 196 shoot-borne root, 176, 182, 193–5, 199 initiation, 182–3 primordia, 183 system, 185 shoot-to-root transport, 99 shortlateralroots1, slr1, 184 shortlateralroots2, slr2, 184 SHORTROOT/SCARECROW, 3

364
Index SHORTROOT, SHR, shortroot, shr, 12, 15, 89, 106, 203, 341–3 short root 5, srt5, 186 short root 6, srt6, 186 SHR:GR, 342 SHY2/IAA3, 96 sickle, skl, SKL, 259, 261, 268 signalling cascade, 73 signal transduction, 186 silver thiosulfate, 141 SIMK, 74 SINAT5 E3 ligase, 96 Sinorhizobium meliloti, 245, 255, 264 site of hair outgrowth, 72 Slavice (Sav), 344 Solanum esculentum, 88, 96, 107, 145, 214–15, 217, 252, 304, 309, 310, 327 SOLITARY ROOT, SLR, solitary root, slr/IAA14, 85–6, 89, 95–8, 204, 297, 333 SOMBRERO, 14 sorted cells, 334 SORTING NEXIN1, SNX1, 6 soybean, 113–15, 212, 227, 256, 260–64, 266, 269 specification of the hyphosis, 9 of stem cell-organizing QC, 11–12 spectrometry, 185 spermidine, 138 spermine, 138 Sphenophyta/Sphenophytes, 193–4 spontaneous nodules formed1, snf1, 253 snf2, 267 sporangia, 226 spotted cDNA/oligonucleotide microarray, 328, 332 spruce, 220 SREBEP transcription factors, 253 SSR loci, 143 starch, 105 deficient mutants, 159 statolith hypothesis, 158 statistical analyses, 330 statolith, 159 sedimentation, 160, 167 stelar parenchyma, 88 stele, 13, 44 cells, 88 endodermis, 13 stem-borne roots, 195–7, 200 stem cell niche specification, 12–13 organizing center, 335 STEROL METHYLTRANSFERASE1, STM1, stm1, 10, 71, 94, 102 stomatal movement, 310 storeys, 158 strawberry (Fragaria), 128 strawberry tree (Arbutus unedo), 139 Striga, 211 subtilisin-like protease, 225 subtractive hybridization, 187 sucrose, 105, 133, 308

N ratio, 307 synthase, 256 transporter, 53 sucrose transporter 2, SUC2, 53, 57, 259 sugar, 105 signaling, 106 sulfate, 144, 313–14 deprivation, 314 starvation, 314 transporter, SULTR1;1, 314 sulfolipid, 293 synthesis, 294 sulfur, 292, 313 SULTR1;2, 314 SUMO E3 ligase, 301 sumoylation, 301 SUNN/NARK/SYMRK receptor kinase, 266 super numerous nodules, sunn, 259, 261, 266 SUPERROOT SUPERROOT 1, SUR1, sur1, 92, 140, 144–5 SUPERROOT 2, SUR2, sur2, 92, 134, 140, 144–5 SUPERROOT1/SUPERROOT2, 335 surface epitopes, 44 suspensor, 65 symbiosis, 210, 229 symbiosis locus 2, SYM2, sym2, 246 sym3, 252 sym8, 251 SYM19, 252 sym29, 260 sym35, 253 sym77, 261 sym78, 214, 218 sym79, 260 SYMBIOSIS RECEPTOR-LIKE KINASE, SYMRK, symrk, 213, 229, 252 symbiosome, 242–3 symbiotic associations, 291 phase, 213 symmetric distribution, 159 synteny, 143 systems biology, 325, 329, 345–6 taxol, 73 Taxus baccata, 139 tebichi, teb, 22 temperature, 135, 145 terfestatin A (TrfA), 168 terrestrial environment, 77 TEs, 49 tetrahedral/pyramidal apical cell (LRAC), 203–4 tetrarch, 107 three-dimensional cellular geometries, 168 three-layered structure, 88 tiling array, 329 TILLING (Targeting-Induced Local Lesions in Genomes), 328 TINY ROOT HAIR 1, TRH1, trh1, 71–2, 75, 311 tip growth, 73 TIP GROWTH DEFECTIVE 1, 74

Index
365

TIP1, TIP1, 74–5 TOADSTOOL2, TOAD2, 15 Tobacco. See Nicotiana tabacum Tomato. See Solanum esculentum tonsoku, tsk, 21 tornado tornado1, TRN1, trn1, 68 TRN2, trn2, 68 tracheary element, 42, 57 transcription, 186 factor (TF), 95, 224, 296, 327, 330–39, 343 transcriptional activator, 96 analysis, 344 fusion, 340–41 network, 329–31, 339, 341 transcriptome, 185 profiling, 255 transcriptomics, 167, 245 transition stage, 180 translational fusions, 340–41 transmembrane receptor kinase, 70 transparent testa, tt4, 139 TRANSPARENT TESTA GLABRA, TTG, TTG, ttg, 75, 299–300 TTG1, 69 transport inhibitor response 3, tir3/big, 92 TRANSPORT INHIBITOR RESPONSE1, TIR1, tir1, 46, 92, 94, 168 triarch, 107 trichoblast, 326 Trifolium repens, 243, 258, 269 triphosphatase (GTPase)-activating protein, 47 Triticum aestivum, 76, 129, 213 Tropaeolum majus, 252 tropisms, 109 TRYPTICHON, TRY, 69, 299 tuber, 130 tulip (Tulipa), 129 two-dimensional gel electrophoresis, 144 ubiquitin, 301 conjugating E2 enzyme (UBC), 302 uidA, 86 Umkirch-1 (Uk-1), 344 upland rice, 112 vacuolar protein sorting 29, VPS29, 6 vascular bundles, 14, 39, 44, 240, 243, 253 cambium, 40, 334 patterning, 180 primordium, 41 system, 39 vascular network defective1, VAN1, van1, 47 van6, 47 VASCULAR TISSUE SIZE, VAS, 56 VASCULAR-RELATED NAC-DOMAIN PROTEIN1, VDN1, 54 VND6, 54 VND7, 54 vasculature, 90

vegetative propagation, 129 VEIN PATTERNING, VEP1, 56 vesicles, 214 vesicular-arbuscular structures, 290 vetch. See Vicia sativa Vicia sativa, 264–5 Vigna radiata, 134, 138, 141 viral-induced silencing (VIGS), 327 Virginia pine. See Pinus virginiana virtual root system, 24 Vitis vinifera, 215 Walnut. See Juglans regia water, 77, 289–90 stress tolerance, 113 wav6-52, 163 WEAK ETHYLENE INSENSITIVE2/ANTHRANILATE SYNTHASE a1, WEI2/ASA1, 27 WEREWOLF, WER, WER, wer, 69, 75, 299–300, 342 WER::GFP, 342 WER::SHR, 342 Wheat. See Triticum aestivum white clover. See Trifolium repens white pine. See Pinus strobus white sweetclover. See Melilotus alba whorls, 182 willow. See Salix WOODEN LEG, WOL, wooden leg, wol, 49, 51, 54–5 wooden leg-3, wol-3, 142 WOL/CRE1, 55 WOL::GFP, 334 WOX transcription factor, 4 WRKY9 transcription factor, 311 WUSCHEL RELATED HOMEOBOX 5, AtWOX5, 18, 179 WUSCHEL-related HOMEODOMAIN, 3 WUSCHEL-type homeobox gene family, 179 WUSCHEL, WUS, WUS, 11–18, 66 xipotl, xpl, 301 xylem, 39, 86, 108, 227, 332, 334, 336 arms, 107 parenchyma, 90 pole (XP), 85, 109 pole pericycle, 85, 87, 92–8, 108 specific promoters, 86 XYLEM NAC DOMAIN1, XND1, 57 yeast two-hybrid, 70 yew. See Taxus baccata yield, 76, 112–15 YUCCA1, yucca, 145 zeatin O-glycosyltransferase, ZOG1, 142 zinc (Zn), 131 Zinnia elegans, 49, 57 ZWILLE/PINHEAD, 15 zygote, 65