Plant Tropisms

Plant Tropisms

Plant Tropisms Edited by SIMON GILROY PATRICK H. MASSON

Simon Gilroy, Ph.D., is Professor of Biology at Pennsylvania State University. Patrick H. Masson, Ph.D., is Professor of Genetics at the University of Wisconsin. © 2008 Blackwell Publishing All rights reserved Blackwell Publishing Professional 2121 State Avenue, Ames, Iowa 50014, USA Orders: Office: Fax: Web site:

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Blackwell Publishing Ltd 9600 Garsington Road, Oxford OX4 2DQ, UK Tel.: +44 (0)1865 776868 Blackwell Publishing Asia 550 Swanston Street, Carlton, Victoria 3053, Australia Tel.: +61 (0)3 8359 1011 Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Blackwell Publishing, provided that the base fee is paid directly to the Copyright Clearance Center, 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license by CCC, a separate system of payments has been arranged. The fee codes for users of the Transactional Reporting Service are ISBN-13: 978-0-8138-2323-2/2007. First edition, 2008 Library of Congress Cataloging-in-Publication Data Plant Tropisms/edited by Simon Gilroy, Patrick H. Masson.-1st ed. p. cm. Includes bibliographical references. ISBN 13:978-0-8138-2323-2 (alk. paper) ISBN 10:0-8138-2323-4 1. Tropisms. 2. Growth (Plants) I. Gilroy, Simon. II. Masson, Patrick H. QK745.P59 2007 571.8⬘2—dc22 2007013116 The last digit is the print number: 9 8 7 6 5 4 3 2 1

Contents

List of Contributors Preface

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Chapter 1: Mechanisms of Gravity Perception in Higher Plants

3

ALINE H. VALSTER AND ELISON B. BLANCAFLOR 1.1 1.2

1.3

1.4 1.5 1.6 1.7 1.8

Introduction 3 Identification and characterization of gravity perception sites in plant organs 4 1.2.1 Roots 4 1.2.2 Hypocotyls and inflorescence stems (dicotyledons) 6 1.2.3 Cereal pulvini (monocotyledons) 8 The starch-statolith hypothesis 9 1.3.1 A variety of plant organs utilize sedimenting amyloplasts to sense gravity 9 1.3.2 Amyloplast sedimentation is influenced by the environment and developmental stage of the plant 11 The gravitational pressure model for gravity sensing 11 The cytoskeleton in gravity perception 12 Concluding remarks and future prospects 14 Acknowledgment 15 Literature cited 15

Chapter 2: Signal Transduction in Gravitropism

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BENJAMIN R. HARRISON, MIYO T. MORITA, PATRICK H. MASSON, AND MASAO TASAKA 2.1 2.2

2.3 2.4 2.5

Introduction 21 Gravity signal transduction in roots and aboveground organs 22 2.2.1 Do mechano-sensitive ion channels function as gravity receptors? 24 2.2.2 Inositol 1,4,5-trisphosphate seems to function in gravity signal transduction 2.2.3 Do pH changes contribute to gravity signal transduction? 27 2.2.4 Proteins implicated in gravity signal transduction 28 2.2.5 Global ‘-omic’ approaches to the study of root gravitropism 32 2.2.6 Relocalization of auxin transport facilitators or activity regulation? 37 2.2.7 Could cytokinin also contribute to the gravitropic signal? 38 Gravity signal transduction in organs that do not grow vertically 39 Acknowledgments 40 Literature cited 40

Chapter 3: Auxin Transport and the Integration of Gravitropic Growth

26

47

GLORIA K. MUDAY AND ABIDUR RAHMAN 3.1 3.2 3.3 3.4 3.5

Introduction to auxins 47 Auxin transport and its role in plant gravity response 47 Approaches to identify proteins that mediate IAA efflux 51 Proteins that mediate IAA efflux 51 IAA influx carriers and their role in gravitropism 53

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3.6

Regulation of IAA efflux protein location and activity during gravity response 55 3.6.1 Mechanisms that may control localization of IAA efflux carriers 56 3.6.2 Regulation of IAA efflux by synthesis and degradation of efflux carriers 58 3.6.3 Regulation of auxin transport by reversible protein phosphorylation 59 3.6.4 Regulation of auxin transport by flavonoids 61 3.6.5 Regulation of auxin transport by other signaling pathways 61 3.6.6 Regulation of gravity response by ethylene 64 3.7 Overview of the mechanisms of auxin-induced growth 65 3.8 Conclusions 67 3.9 Acknowledgements 68 3.10 Literature cited 68

Chapter 4: Phototropism and Its Relationship to Gravitropism JACK L. MULLEN AND JOHN Z. KISS 4.1 Phototropism: general description and distribution 4.2 Light perception 80 4.3 Signal transduction and growth response 82 4.4 Interactions with gravitropism 83 4.5 Importance to plant form and function 84 4.6 Conclusions and outlook 85 4.7 Literature cited 86

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Chapter 5: Touch Sensing and Thigmotropism

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GABRIELE B. MONSHAUSEN, SARAH J. SWANSON, AND SIMON GILROY 5.1 Introduction 91 5.2 Plant mechanoresponses 91 5.2.1 Specialized touch responses 92 5.2.2 Thigmomorphogenesis and thigmotropism 94 5.3 General principles of touch perception 95 5.3.1 Gating through membrane tension: the mechanoreceptor for hypo-osmotic stress in bacteria, MscL 98 5.3.2 Gating through tethers: the mechanoreceptor for gentle touch in Caenorhabditis elegans 99 5.3.3 Evidence for mechanically gated ion channels in plants 101 5.4 Signal transduction in touch and gravity perception 103 5.4.1 Ionic signaling 103 5.4.2 Ca2+ signaling in the touch and gravity response 103 5.5 Insights from transcriptional profiling 107 5.6 Interaction of touch and gravity signaling/response 110 5.7 Conclusion and Perspectives 113 5.8 Acknowledgements 114 5.9 Literature cited 14

Chapter 6: Other Tropisms and their Relationship to Gravitropism GLADYS I. CASSAB 6.1 6.2

Introduction 123 Hydrotropism 123 6.2.1 Early studies of hydrotoprism 124 6.2.2 Genetic analysis of hydrotropism 125 6.2.3 Perception of moisture gradients and gravity stimuli by the root cap and the curvature response 126

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6.3 6.4 6.5 6.6 6.7 6.8 6.9

6.2.4 ABA and the hydrotropic response 6.2.5 Future experiments 129 Electrotropism 129 Chemotropism 131 Thermotropism and oxytropism 132 Traumatropism 134 Overview 135 Acknowledgments 135 Literature cited 135

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Chapter 7: Single-Cell Gravitropism and Gravitaxis

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MARKUS BRAUN AND RUTH HEMMERSBACH 7.1 7.2 7.3 7.4 7.5 7.6 7.7 7.8 7.9 7.10 7.11 7.12 7.13 7.14 7.15 7.16 7.17

Introduction 141 Definitions of responses to environmental stimuli that optimize the ecological fitness of single-cell organisms 141 Occurrence and significance of gravitaxis in single-cell systems 142 Significance of gravitropism in single-cell systems 143 What makes a cell a biological gravity sensor? 144 Gravity susception—the initial physical step of gravity sensing 145 Susception in the statolith-based systems of Chara 145 Susception in the statolith-based system Loxodes 149 Susception in the protoplast-based systems of Euglena and Paramecium 150 Graviperception in the statolith-based systems of Chara 150 Graviperception in the statolith-based system Loxodes 151 Graviperception in the protoplast-based systems Paramecium and Euglena 151 Signal transduction pathways and graviresponse mechanisms in the statolith-based systems of Chara 153 Signal transduction pathways and graviresponse mechanisms in Euglena and Paramecium 154 Conclusions 155 Acknowledgements 156 Literature cited 156

Color Section Chapter 8: Space-Based Research on Plant Tropisms MELANIE J. CORRELL AND JOHN Z. KISS 8.1 8.2 8.3 8.4

Introduction—the variety of plant movements 161 The microgravity environment 162 Ground-based studies: mitigating the effects of gravity 165 Gravitropism 166 8.4.1 Gravitropism: gravity perception 166 8.4.2 Gravitropism: signal transduction 168 8.4.3 Gravitropism: the curving response 169 8.5 Phototropism 171 8.6 Hydrotropism, autotropism, and oxytropism 172 8.7 Studies of other plant movements in microgravity 174 8.8 Space flight hardware used to study tropisms 175 8.9 Future outlook and prospects 177 8.10 Literature cited 177

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Chapter 9: Plan(t)s for Space Exploration

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CHRISTOPHER S. BROWN, HEIKE WINTER SEDEROFF, ERIC DAVIES, ROBERT J. FERL, AND BRATISLAV STANKOVIC 9.1 9.2 9.3 9.4 9.5 9.6 9.7 9.8 9.9

Index

Introduction 183 Human missions to space 184 Life support 184 Genomics and space exploration 185 Nanotechnology 187 Sensors, biosensors, and intelligent machines Plan(t)s for space exploration 188 Imagine . . . 192 Literature cited 192

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List of Contributors

Elison B. Blancaflor Plant Biology Division The Samuel Roberts Noble Foundation 2510 Sam Noble Parkway Ardmore, OK 73401 USA Tel: (580) 224-6687 Fax: (580) 224-6692 E-mail: [email protected] Markus Braun Gravitationsbiologie Institut für Molekulare Physiologie und Biotechnologie der Pflanzen Universität Bonn 53115 Bonn Germany Tel: (49) 228-73-2686 Fax: (49) 228-732677 E-mail: [email protected] Christopher S. Brown Kenan Institute for Engineering, Technology & Science North Carolina State University Raleigh, NC 27695 USA Tel: (919) 513-2457 Fax: (919) 515-5831 E-mail: [email protected]

Gladys I. Cassab Department of Plant Molecular Biology Institute of Biotechnology National Autonomous University of Mexico P.O. Box 510-3 Cuernavaca, Mor. 62250 Mexico Tel: (52) 5556-22-7660 Fax (52) 7773-13-9988 E-mail: [email protected] Melanie J. Correll Department of Agricultural and Biological Engineering University of Florida 209 Frazier Rogers Hall P.O. Box 110570 Gainesville, FL 32611-0570 USA Tel: (352) 392-1864 Fax: (352) 392-4092 E-mail: [email protected] Eric Davies Department of Plant Biology North Carolina State University 1231 Gardner Hall Box 7612 Raleigh, NC 27695 USA Tel: (919) 513-1901 Fax: (919) 515-3436 E-mail: [email protected]

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LIST OF CONTRIBUTORS

Robert J. Ferl Department of Horticulture University of Florida Gainesville, FL 32611 USA Tel: (352) 392-1928 Fax: (352) 392-4072 E-mail: [email protected] Simon Gilroy Biology Department The Pennsylvania State University 208 Mueller Laboratory University Park, PA 16802 USA Tel: (814) 863-9626 Fax: (814) 865-9131 E-mail: [email protected] Benjamin R. Harrison Laboratory of Genetics (Room 3262) University of Wisconsin–Madison 425G Henry Mall Madison, WI 53706 USA Tel: (608) 265-8632 Fax: (608) 262-2976 E-mail: [email protected] Ruth Hemmersbach Institute of Aerospace Medicine DLR (German Aerospace Research Establishment) 51140 Köln Under Höhe Germany Email: [email protected] John Z. Kiss Department of Botany Pearson Hall Miami University Oxford, OH 45056 USA Phone: (513) 529-5428 Fax: (513) 529-4243 E-mail: [email protected]

Patrick H. Masson Laboratory of Genetics (Room 3262) University of Wisconsin–Madison 425G Henry Mall Madison, WI 53706 USA Tel: (608) 265-2312 Fax: (608) 262-2976 E-mail: [email protected] Gabriele B. Monshausen Biology Department The Pennsylvania State University 208 Mueller Laboratory University Park, PA 16802 USA Tel: (814) 863-9625 Fax: (814) 865-9131 E-mail: [email protected] Miyo T. Morita Graduate School of Biological Sciences Nara Institute of Science and Technology 8916-5 Takayama Ikoma, Nara 630-0101 Japan Phone: (81) 743-72-5487 Fax: (81) 743-72-5487 E-mail: [email protected] Gloria K. Muday Department of Biology Wake Forest University Winston-Salem, NC 27109-7325 USA Tel: (336) 758-5316 Fax: (336) 758-6008 E-mail: [email protected] Jack L. Mullen Department of Bioagricultural Sciences and Pest Management Plant Science Building, Room C 129 Colorado State University Fort Collins, CO 80523-1177 USA Tel: (970) 491-5261 E-mail: [email protected]

LIST OF CONTRIBUTORS

Abidur Rahman Biology Department University of Massachusetts 611 North Pleasant St. 106 Morrill 3 Amherst, MA 01003 USA Phone: (413) 545-2776 Fax: (413) 545-3243 E-mail: [email protected] Heike Winter Sederoff Department of Plant Biology North Carolina State University Raleigh, NC 27695 USA Phone: (919) 513-0076 Fax: (919) 515-3436 E-mail: [email protected] Bratislav Stankovic Brinks Hofer Gilson & Lione 455 N. Cityfront Plaza Drive, Suite 3600 Chicago, IL 60611-5599 USA Sarah J. Swanson Biology Department The Pennsylvania State University 208 Mueller Laboratory University Park, PA 16802 USA Tel: (814) 863-9625 Fax: (814) 865-9131 E-mail: [email protected]

Masao Tasaka Graduate School of Biological Sciences Nara Institute of Science and Technology 8916-5 Takayama Ikoma, Nara 630-0101 Japan Phone: (81) 793-72-5480 Fax: (81) 793-72-5489 E-mail [email protected] Aline H. Valster Plant Biology Division The Samuel Roberts Noble Foundation 2510 Sam Noble Parkway Ardmore, OK 73401 USA Tel (580) 224-6756 Fax (580) 224-6692 E-mail: [email protected]

xi

Preface

As sessile organisms, plants spend their entire lives at the site of seed germination. Consequently, they require a suite of strategies to survive very diverse environmental stresses. Part of this plasticity relies on the ability of most plant organs to grow in directions that are dictated by specific cues from the environment, seeking out better conditions to fulfill their primary functions. Typical guidance for the growth of plant organs is provided by gravity, light, touch, gradients in humidity, ions, oxygen, and temperature. Such directional growth, defined by vectorial stimuli, is called a tropism and is believed to significantly contribute to plant survival. The concept of tropism was introduced 200 years ago, when Knight (1806) postulated that a plant’s perception of gravity might modulate its ability to direct shoots to grow upward and guide roots downward. Eighty years later, Darwin (1880) made seminal contributions to the field by documenting a wide array of tropic responses and identifying regions of the root and shoot specialized for the perception of light and gravity. He also predicted the existence of auxin by proposing the presence of a plant growth regulator (hormone) whose gravity-induced redistribution across the tip of an organ might signal differential growth. Since these discoveries, our analysis of tropic growth has expanded to include measurements of responses to light, touch, and gradients in humidity, ions, chemicals, and oxygen. However, only recently have the data converged to provide a picture of the physiological, molecular, and cell biological processes that underlie plant tropisms. Thus, the last few years have witnessed a true renaissance in the analysis of tropic response, mainly driven by the marrying of modern tools and strategies in the fields of forward and reverse genetics, biochemistry, cell biology, expression profiling, and proteomics, to a very careful analysis of the growth process itself. When such analyses have been coupled with the utilization of model systems such as Arabidopsis thaliana and rice, where their entire genome has been sequenced, these strategies have provided an unprecedented power of resolution in our analysis of growth behaviors. Consequently, our conception of tropisms has evolved from their being considered as simple laboratory curiosities to becoming important tools/phenotypes with which to decipher basic cell biological processes that are essential to plant growth and development. Thus, current insight into tropisms is intimately involved in our understanding of auxin transport and response; cytoskeleton organization and its involvement in the control of anisotropic cell expansion; the perception and transduction of stimuli such as light, touch, humidity, ions, or oxygen; the biogenesis and function of organelles such as plastids and vacuoles; and even the control of vesicular trafficking, to name but a few (Blancaflor and Masson 2003). Of the tropic stimuli, our understanding of the mechanisms behind gravitropic and phototropic response has shown extremely rapid advances in the last few years and, in xiii

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Chapter 1, Valster and Blancaflor describe our current models of gravitropic sensing in plants, a theme further developed in Chapter 2, where Harrison and colleagues discuss the molecular mechanisms behind transduction of the gravity signal. In Chapter 7, Braun and Hemmersbach further explore sensing and signaling in plants by comparison to the wealth of data on how single-celled organisms detect and respond to gravity. Similarly, in Chapter 4, Mullen and Kiss describe the remarkably detailed knowledge we now have of the mechanisms whereby plants perceive light and translate that cue into a phototropic growth. Despite Darwin’s prediction of the action of auxin in tropic response as early as 1880, only recently have the mechanisms behind auxin transport and action been defined to the molecular level. For example, we now understand that the relocalization of auxin transporters is a central component regulating tropic response pathways and critical components of the auxin transport pathway have been defined with molecular precision. In Chapter 3, Muday and Rahman provide an overview of this extremely rapidly evolving field. Although individual tropic stimuli are often studied in a controlled laboratory setting, nature provides a harsh environment where multiple vectorial stimuli often signal conflicting information for a plant organ. An important step in our conceptualization of plant responses to such a complex environment has been the realization that organs not only perceive and respond to each one of these parameters, but they also have to integrate and interpret the corresponding environmental information into global “decisions” that manifest themselves into complex growth behaviors. The integration of other tropic stimuli with the gravitropic response has recently received intense analysis and, in Chapters 5 and 6, Monshausen and colleagues and Gladys Cassab describe the wealth of tropic responses in plants and specifically how responses to touch and moisture alter gravitropic response. Such integrated responses to combined environmental cues appear to involve complex intra- and intercellular communications. Recent analyses have uncovered some of these fascinating signaling events (Fasano et al. 2002), opening the possibility of, one day, being able to engineer plants that are capable of using a defined set of directional cues for growth guidance while being oblivious to other cues. Such engineering accomplishments could find applications in agriculture and in more futuristic endeavors such as space exploration. Indeed, spaceflight has offered researchers a unique opportunity to dissect tropic response in the absence of the effects of gravity. However, in space, in addition to exposure to microgravity, organisms also suffer from a lack of convection, growth-space limitations, lower light exposures, and increased radiation levels. Hence, the spaceflight environment appears quite unfavorable to plant success, and tropic responses are likely to be altered accordingly. Because plants have been identified as an ideal choice for utilization in bioregenerative life-support systems during long-term space exploration missions, there is a definite need for a better understanding of their growth behavior and sustainability during long-term exploration travels in order to prevent or overcome potential catastrophic system breakdowns in the midst of a mission. Recognition of this need recently fueled efforts at developing orbit-based experiments on plant growth behavior and gravitropic sensitivity, eventually leading to the design and building of the International Space Station where such studies can be carried out. Space

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experiments have added new information on plant growth responses to directional cues such as gravity, light, or oxygen gradients and, in Chapter 8, Correll and Kiss describe the opportunities that spaceflight has provided to understand how a range of such tropic responses operate. However, spaceflight experimentation has also been plagued by a variety of constraints that have diminished their potential scientific value. Hence, a combined approach, including both ground- and orbit-based research, is necessary to gain a better understanding of the behavior of plants and their organs under micro- or hypergravity environments in the hope of being able to, one day, engineer cultivars that are better-adapted to the conditions likely to be encountered during space exploration missions. Thus, the field of plant tropisms has received considerable attention in the last few years for its impact on both basic understanding of plant growth and development and applied aspects, such as crop response or application to spaceflight. We hope this book will provide a comprehensive yet integrated coverage of our current state of knowledge on the molecular and cell biological processes that govern plant tropisms, with major emphasis on gravitropism (one of the most extensively studied plant tropisms). Our understanding of tropic responses is rapidly increasing and, with each new insight, the potential to engineer new traits into plants moves closer. Therefore, for the last chapter of the book we asked Chris Brown and colleagues to present a vision for how our increasingly detailed understanding of these plant growth responses might translate into designing plants to sustain human endeavors in perhaps the most inhospitable environment for life imaginable— space. Simon Gilroy Patrick H. Masson Literature Cited Blancaflor EB and Masson P. 2003. Update on Plant gravitropism. Unraveling the ups and downs of a complex process. Plant Physiology 133: 1677–1690. Darwin C. 1880. The Power of Movement in Plants. London: John Murray. Fasano JM, Massa GD and Gilroy S. 2002. Ionic signaling in plant responses to gravity and touch. Journal of Plant Growth Regulation 21: 71–88. Knight T. 1806. On the direction of the radicle and germen during the vegetation of seeds. Philosophical Transactions of the Royal Society 99: 108–120.

Plant Tropisms

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Mechanisms of Gravity Perception in Higher Plants Aline H. Valster and Elison B. Blancaflor*

1.1 Introduction Plant growth and development is influenced by a multitude of exogenous and endogenous signals. Among the signals a plant encounters during its lifetime, gravity is one that remains constant throughout development. Since the plant needs to orient its organs to position itself within available environmental resources such as light and soil nutrients, the gravity stimulus is significant for its survival. From the moment the seed germinates, the seedling orients its emerging root such that it grows downward, toward the gravity vector, whereas it directs its shoot to grow upward, opposite the gravity vector. This phenomenon, referred to as gravitropism (geotropism in the older literature) requires the coordinated response and interaction of different cell types. Furthermore, an array of cellular structures and endogenous molecules, which in turn are modulated by a variety of environmental stimuli including light, moisture, oxygen, and touch, eventually determine the final manifestation of the gravity response (Blancaflor and Masson 2003; Morita and Tasaka 2004; Perrin et al. 2005; Esmon et al. 2005). Gravitropism has traditionally been divided into a series of events: gravity perception, signal transduction, and the growth response (Sack 1991; Kiss 2000). In higher plants, these events appear to take place in spatially distinct regions of the organ, in contrast to tip-growing cells such as rhizoids of the green algae Chara and protonemata of moss and Chara where, as discussed in Chapter 7, all phases of gravitropism occur within the same cell (Sievers et al. 1996; Schwuchow et al. 2002). Since gravity must ultimately work on a mass to exert its effect on a given biological system, it has been widely accepted that plants sense gravity through falling organelles (statoliths) within specialized cells (statocytes). Through the years, this model of plant gravity perception has been refined and alternative hypotheses have been proposed, including the possibility that the settling of the whole cell protoplast rather than sedimenting organelles is responsible for gravity sensing (Staves 1997). A number of excellent articles which provide a historical perspective on gravity perception in plants include Sack (1991, 1997) and Kiss (2000). The reader is referred to these articles for an in-depth discussion and critical analysis of the experimental data that have led to current models on how plants sense gravity. In this chapter, we revisit the topic of gravity perception mechanisms, focusing primarily on roots and shoots of higher plants. Although we occasionally refer to some of the older literature, this chapter will highlight recent findings that are leading to new, testable models explaining how plants sense gravity.

*Corresponding author

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1.2 Identification and Characterization of Gravity Perception Sites in Plant Organs Gravity has been shown to regulate the orientation of different plant organs such as roots, shoots (Fukaki et al. 1998; Morita and Tasaka 2004; Perrin et al. 2005), leaves (Mano et al. 2006), inflorescence stems (Weise et al. 2000), cereal pulvini (Perera et al. 2001), and peanut gynophores (Moctezuma and Feldman 1999). The response to gravity in the majority of these plant organs is manifested as differential cell growth between opposite flanks of the organ, leading to upward or downward bending. Since not all cells within the organ undergo differential growth (Sack et al. 1990), an important question in gravitropism research is how the different cell and tissue types within the organ contribute to the gravity response. A more specific question is whether the machinery for sensing gravity occurs in the same sites as the responding tissues. To address these questions, research spanning two centuries has focused on elucidating the spatial regulation of gravitropism. For example, work that began with Charles Darwin in the late 1800s and followed-up by several other investigators in the 1900s identified the cap as the major gravity perception site in roots (reviewed by Konings 1995; Boonsirichai et al. 2002). These early experiments showed that surgical removal of the root cap tissue inhibited the gravitropic curvature without affecting overall root growth. In this section, we briefly review experimental evidence that has further reinforced the existence of specific gravity-sensing sites, distinct from the responding tissues, in the best-studied multicellular plant organs, namely roots, dicot stems, and grass pulvini. 1.2.1 Roots As noted earlier, gravity must work on a mass to elicit a specific biological response. Therefore, cells that would be prime candidates for perceiving gravity are those which exhibit a distinct structural polarity with respect to the gravity vector. Indeed, detailed ultrastructural studies of the cap of vertically growing roots in a variety of plant species revealed that the central region of the cap (called the columella) contains cells with organelles that are consistently positioned at the bottom of the cell (reviewed in Sack 1991, 1997). These organelles, later identified as starch-containing plastids called amyloplasts (Figure 1.1A and Color Section), would rapidly change position (i.e., sediment) when the root was reoriented. The sedimentation of amyloplasts is the most widely accepted explanation for how plant organs sense gravity, a model currently known as the starch-statolith hypothesis (refer to The Starch-Statolith Hypothesis section later in this chapter). The identification of the cap, particularly the columella, as a major gravity-sensing site in roots led many researchers to utilize roots as a model system for studying mechanisms underlying plant gravitropic responses. This is because the apparent physical separation of the gravity-sensing cells in the cap from the responding cells in the elongation zone in angiosperm primary roots could, in theory, facilitate the analysis of individual phases of gravitropism. For instance, more than a century after Darwin first reported on the necessity of the cap for root gravitropism, laser ablation of Arabidopsis root cap cells allowed a more detailed spatial analysis of specific cells within the columella region that contributed most to the gravity response (Blancaflor et al. 1998). In this study, ablation of the most centrally located root cap cells, namely the second story (S2) columella cells

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Figure 1.1 (also see Color Section). A. Longitudinal section of the root cap of Medicago truncatula showing the centrally located columella cells (c) containing starch-filled plastids (a, amyloplasts). Note that the columella cells are highly polarized with the nucleus (n) located at the upper side of the cell and amyloplasts (a) sedimented on the bottom side. B. Hypocotyl of a Medicago truncatula seedling bends upward when positioned horizontally. Longitudinal section of the reoriented hypocotyl shows amyloplasts (a) sedimented to the new bottom side of the cell. White arrow indicates the direction of gravity.

Figure 1.2. Brightfield and corresponding fluorescence micrograph of an Arabidopsis root cap expressing the actin binding domain (ABD-2) of Fimbrin. Actin filaments in the centrally located columella cells, stories 2 and 3 (S2, S3) appear to have a finer structure than the peripheral cap (PC) and tip cells (TC). Bar = 20 µm.

(Figure 1.2), had the strongest inhibitory effect on the gravity response—identifying those specific cells of the cap as the most important for gravity sensing. Destroying the lower part of the cap in horizontally positioned roots with heavy-ion microbeams also inhibited gravitropism, possibly by interfering with the cap tissue responsible for transmission of the gravity signal from the columella (Tanaka et al. 2002). Another set of studies implicating the root cap in the gravitropic response employed a genetic approach to remove root cap cells. A protein synthesis inhibitor (diphtheria toxin A) was expressed under a root cap specific promoter in Arabidopsis, killing the expressing cells (Tsugeki and Fedoroff 1999). In addition to having altered morphology, the re-

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sulting roots were agravitropic, providing further evidence that the cap is the primary site of gravity perception in roots. Despite overwhelming evidence supporting the cap as a major gravity-sensing site in roots, there are sparse reports demonstrating that the root cap might not be the only tissue that is able to perceive gravity. Early experiments employing centrifugation methods suggested that the elongation zone might also be involved (reviewed in Boonsirichai et al. 2002). However, these experiments are difficult to interpret because the centrifugation technique itself possibly introduces mechanical effects that could contribute to the bending response of the root. More recently, Wolverton et al. (2002a) devised a method (named ROTATO) that allowed different regions of the root outside the cap to be maintained at a defined angle to the vertical (continuously gravistimulated). If a section of the elongation zone of the root was kept at a defined angle, curvature of the root persisted even after the root cap had reached its normal vertical position. From these experiments it was concluded that the elongation zone can contribute to gravitropic sensing, although to a lesser extent than the root cap. It was estimated that 20% of the total rate of curvature originates from the distal elongation zone or the apical portion of the central elongation zone. The finding that the elongation zone contributes to root gravitropic sensing might explain why roots sometimes curve past the vertical and why starchless mutants of Arabidopsis still have a residual gravitropic response (Wolverton et al. 2002a, b). In support of the notion that other tissues outside the cap can sense gravity was the recent observation that gravitropic curvature in decapped roots of maize can be restored by myosin and actin inhibitors. This indicates the existence of a mechanism for gravity sensing outside the cap that relies on a dynamic cytoskeleton (Mancuso et al. 2006; see The Cytoskeleton in Gravity Perception section). Although these new findings continue to support the conclusion made more than a century ago that the root cap is a major site for gravity perception, it appears that it may not be the sole site. The availability of techniques such as ROTATO should allow more detailed investigations into alternative gravity-sensing sites in roots. 1.2.2 Hypocotyls and Inflorescence Stems (Dicotyledons) In contrast to roots, shoots exhibit negative gravitropism, meaning that they grow upward. In shoots of dicots, sedimented amyloplasts were often observed in endodermal cells adjacent to the vasculature, leading to the proposal that the endodermis might be the primary gravity-sensing tissue in shoots (reviewed in Kiss 2000; Morita and Tasaka 2004; see Figure 1.1B). However, it was not until the late 1990s that a better appreciation of the importance of the endodermis for shoot gravitropism was realized. This was due to the fact that, unlike the cap in roots, which is easy to microsurgically remove, the endodermal cells in shoots are physically difficult to manipulate because of their internal location within the organ. The isolation of a series of Arabidopsis mutants with defects in shoot gravitropism (sgr) facilitated the genetic analysis of gravity-sensing and signaling mechanisms in shoots (Morita and Tasaka 2004). Of particular importance was the identification of two Arabidopsis mutants (sgr1 and sgr7) that were allelic to the radial pattern mutants scarecrow (scr) and short-root (shr), respectively. sgr1 and sgr7 were shown

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to encode transcription factors necessary for the normal development of the endodermis in roots and shoots. Both mutants failed to develop an endodermal cell layer and, as a result, lacked sedimentable amyloplasts in their shoots. However, both mutants had normal columella cells containing sedimentable amyloplasts, indicating that amyloplast formation or sedimentation itself was not impaired. Importantly, although shoot gravitropism was defective in these mutants, root gravitropism was not. Thus, it was concluded from these studies that the presence of a normal endodermal layer is important in shoot gravitropism (Fukaki et al. 1998). The importance of the endodermis for gravitropism was also recently demonstrated in other plant species. For example, an agravitropic mutant in the Japanese morning glory (Pharbitis nil) called weeping was shown to be defective in the formation of the endodermal layer, similar to the sgr1 mutant in Arabidopsis (Hatakeda et al. 2003). Interestingly, the disrupted gene in weeping was recently shown to be an ortholog of Arabidopsis scr. When the wild-type scr from morning glory (PnSCR) was introduced into the Arabidopsis scr mutants for complementation, the agravitropic phenotype of Arabidopsis was rescued (Kitazawa et al. 2005). Results from the above studies are also beginning to shed light on the relationship between gravitropism and circumnutation (i.e., oscillatory plant movements). It has long been debated whether gravitropism and circumnutation are causally related to each other (Kiss 2006). Interestingly, it appears that circumnutation and winding movement defects in weeping could be attributed to loss of endodermal function since the circumnutation phenotype in Arabidopsis scr1 could be rescued by scr from wild-type morning glory. Although this provided compelling evidence that other types of plant movements (such as circumnutation) require gravity-sensing cells, recent studies in rice coleoptiles indicate that the relationship between gravitropism and circumnutation may be more complex. For example, Yoshihara and Iino (2005) demonstrated that red light abolished circumnutation in rice coleoptiles without affecting the gravitropic response. Furthermore, coleoptiles of the rice mutant lazy do not circumnutate but do contain sedimentable amyloplasts, which suggests that mechanisms independent of gravitropism might operate in plant circumnutations (Yoshihara and Iino 2006). In addition to proving directly that the endodermis is a major gravity-sensing cell layer, the screen for shoot gravitropic mutants has helped uncover additional features of the endodermis that might be important for gravity perception. Although much attention has been given to sedimenting amyloplasts, other cellular compartments in the gravisensitive cells are likely important. For example, in the above-mentioned screen performed by the group of Tasaka, several other shoot gravitropic mutants were isolated. Of these, the sgr2, sgr3, and sgr4/zigzag mutants are notable because they are implicated in vacuolar membrane trafficking (Kato et al. 2002; Yano et al. 2003). The SGR2 gene encodes a phospholipase-like protein and the ZIG gene encodes a SNARE protein. Both are thought to be involved in vacuolar membrane dynamics because abnormal vesicular and vacuolar structures were observed in the endodermis and several other tissues of the mutant plants. In addition, GFP-SGR2 localizes to the vacuole and some small organelles. Interestingly, in the mutant plants, many amyloplasts do not sediment properly to the bottom of the cell and appear to maintain their localization in the peripheral cytoplasm of the endodermal cells (Morita et al. 2002).

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Further evidence for the role of vacuolar membrane dynamics in shoot gravitropism comes from the analysis of another mutant in Arabidopsis, namely the sgr3 mutant (Yano et al. 2003). The SGR3 gene encodes a syntaxin (AtVAM3) that also has been implicated in vacuolar transport. The SGR3 protein is localized to the prevacuolar compartment and the vacuole itself. The sgr3 mutant shows abnormal appearance of vacuoles in endodermal cells with irregular curves and aberrant membranous structures. This mutant also displays abnormal sedimentation of amyloplasts in the shoot endodermis, coupled with altered gravitropism in inflorescence stems (Yano et al. 2003). This study provides additional evidence for a role of vacuolar membrane dynamics and vacuolar biogenesis in proper amyloplast sedimentation and, by extension, graviperception. Further evidence for a role of membrane dynamics comes from yet another mutant. The gravitropism defective 2 mutant (grv2) in Arabidopsis exhibits a defect in shoot and hypocotyl gravitropism. The responsible gene, GRV2, appears to code for a protein that is similar to the RME-8 protein in Caenorhabditis elegans, which is important for endocytosis (Silady et al. 2004). Since the endodermal cell layers in shoots of grv2 mutant display defects in amyloplast sedimentation, endocytosis might be involved in the initial gravity perception steps as well as the membrane dynamics necessary for targeting of auxin efflux carriers such as PIN3 to their correct position within the columella cell (Friml et al. 2002; Chapter 3 in this book). 1.2.3 Cereal Pulvini (Monocotyledons) In contrast to roots and shoots in dicotyledons, the shoots of grasses have a specialized tissue that mediates gravitropism, namely the pulvinus. The pulvinus is a cushion-like swelling at the base of each internode and the vascular bundles within it are surrounded by bundle sheath cells that contain sedimenting amyloplasts. Upon reorientation of the monocotyledon plant within the gravity field, the amyloplasts sediment to the bottom cell wall in the same way as described for columella and endodermal cells (Allen et al. 2003). Also, in pulvini it is this sedimentation that is thought to set off the signal transduction cascade that ultimately regulates the reorientation of the stem of the plant. However, in contrast to the primary root, the processes of gravity perception, signal transduction, and growth response (i.e., the establishment of a gradient of cell elongation) all occur in the same tissue (Collings et al. 1998). In order for the cells in the pulvini to be gravicompetent, it appears that they must delay maturation. Maturation of the surrounding cells in the tissue of the stems involves lignification of the cell wall and rearrangement of microtubules from transverse to oblique after elongation is completed. Once these cellular processes have taken place, the pulvinus is no longer capable of responding to gravistimulation. During the onset of the gravitropic curvature, maturation occurs only on the upper side of the pulvinus, whereas the elongating cells in the lower side mature only after maximum bending capacity of the stem has been reached (about 30 degrees upward bending) (Collings et al. 1998). The pulvini are amenable to biochemical studies, as relatively large amounts of cells can be harvested. In maize and oat, upper and lower pulvinus tissue can even be collected and analyzed separately. Using these techniques, Perera et al. (1999, 2001) have been able to show that inositol 1,4,5-trisphosphate (InsP3) levels increase transiently

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within seconds to minutes after gravistimulation. As discussed in Chapter 2, the role for InsP3 as a likely universal signaling molecule in the gravity response was recently demonstrated by constitutively overexpressing a human type I polyphosphate 5-phosphatase in Arabidopsis. This enzyme specifically hydrolyzes InsP3. This experiment resulted in a 90% reduction in InsP3 levels and a 30% decline in the gravity response of roots, hypocotyls, and inflorescence stems (Perera et al. 2006). Genetic and biochemical studies such as these are yielding important information regarding the signal transduction pathways activated upon gravistimulation. However, demonstrating how the cells that sense gravity via sedimenting plastids convert this directional information into biochemical signals that include InsP3 transients remains a major challenge in gravitropism research.

1.3 The Starch-Statolith Hypothesis It is clear from the previous section that a common denominator of gravisensitive plant organs is the presence of statoliths/amyloplasts which sediment to the bottom of the graviperceptive cells and that, upon reorientation of these cells within the gravity field, the amyloplasts rapidly resediment to the new bottom of the cell. The starch-statolith hypothesis poses that the sedimentation of the statoliths within the gravisensitive cells is the event that translates the gravity-driven mechanical stimulus into a chemical signal that further regulates differential growth of the flanks of the reorienting organ. How the “falling” of the statoliths exactly converts into a chemical signal is not clear yet, but it is thought that possible interactions of amyloplasts with other cell components such as the endoplasmic reticulum, cytoskeleton, or vacuole are important. The starch-statolith hypothesis is now widely accepted as the major gravity-sensing mechanism despite the lack of knowledge about some details. 1.3.1 A Variety of Plant Organs Utilize Sedimenting Amyloplasts to Sense Gravity Evidence in support of a role for the starch-filled amyloplasts in the gravitropic response comes from several Arabidopsis mutants that are impaired in amyloplast formation. For example, the phosphoglucomutase mutant (pgm) is impaired in starch synthesis and as a result shows reduced gravitropic responses in both roots and shoots (Kiss et al. 1989; Kiss et al. 1997). Some controversy about the starch-statolith hypothesis came from a paper published in 1989 by Caspar and Pickard (1989) showing that an Arabidopsis mutant lacking plastid phosphoglucomutase activity still was able to respond to gravistimulation, albeit in a reduced manner. From this study it was concluded that starch is unnecessary for gravitropism. However, upon careful ultrastructural examination of this mutant, it appeared that not all starch formation was inhibited. Apparently, some starch grains were formed in a subset of the amyloplasts, explaining the reduced but not completely abolished gravitropic response (Saether and Iversen 1991). More recently, Fujihara et al. (2000) isolated another mutant with impaired amyloplasts. The endodermal-amyloplastless 1 (eal1) mutant has no amyloplasts in the hypocotyl endodermal cells while retaining normal plastids in the columella cells. Strong

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support for the starch-statolith hypothesis comes from the observation that gravitropism is impaired in shoots of the eal1 mutant but not in roots. Further compelling support comes from the laser ablation experiments mentioned earlier, where it was shown that a strong correlation exists between the maximum amyloplast sedimentation rates in the different cell layers of the root cap (S1, S2, and S3: Figure 1.2) and their involvement in the gravitropic response (Blancaflor et al. 1998). Moreover, a series of elegant experiments employing a high-gradient magnetic field to displace amyloplasts statoliths mimicked a gravitropic response (i.e., induced curvature) in roots and shoots, providing further strong support for the starch-statolith hypothesis (Kuznetsov and Hasenstein 1996; Weise et al. 2000). Recently, other less-studied plant organs have been shown to exploit sedimenting amyloplast for the purpose of gravity sensing. For example, Arabidopsis plants shift their leaves upward against the gravity vector when kept in the dark. This movement was shown to be a combination of nastic and gravitropic movement (Mano et al. 2006). Cells with sedimenting amyloplasts were observed in several cell layers around the vasculature of the proximal region of the petiole. Two mutants in amyloplast formation (pgm and sgr2-1) showed that abnormal distribution or absence of amyloplasts in the petioles resulted in reduced upward bending of Arabidopsis leaves, indicating that sedimenting amyloplasts are in part responsible for this process (Mano et al. 2006). An unusual example of gravitropism can be found in peanut gynophores (Moctezuma and Feldman 1999). The gynophore is a specialized organ which ensures that developing fruits are buried in the soil. This is a step that is essential to the life cycle of the peanut plant. Moctezuma and Feldman (1999) explored gravitropism in the peanut gynophore and found that sedimenting amyloplasts are present in the starch sheath cells that surround the vasculature in the gynophore. De-starching the plastids by incubation with gibberellic acid and kinetin in the dark did not affect overall growth but abolished 82% of the gravitropic response. Gravitropism in the peanut gynophore is interesting because the gravity-sensing mechanism seems similar to gravity sensing in shoots, yet the induced growth response is opposite (i.e., the gynophore grows down rather than upward). Thus, studies on the peanut gynophore might prove extremely helpful in addressing the question on how the positive versus negative gravitropic response is determined in roots and shoots. Another example where sedimenting plastids appear to mediate a particular gravitropic response is in peg formation in cucumber hypocotyls. Cucumber seedlings form a specialized protuberance called a peg on the concave side of the bending site between the hypocotyl and the root, which assists in shedding of the seed coat upon germination. The formation of the peg is gravity-dependent and the amyloplast-containing sheath cells of the vascular bundles in this area are responsible for gravity sensing (Saito et al. 2004). In this case, gravity is responsible for the inhibition of peg formation on the convex side of the bending zone as, under microgravity conditions, two pegs (one on either side of the bending site) will form (Takahashi et al. 2000). From the above, it appears that not only roots and shoots rely on sedimenting amyloplasts to perceive gravity, but that other plant organs do as well. These observations are consistent with, and testify to, the generality of the starch-statolith hypothesis proposed more than a century ago.

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1.3.2 Amyloplast Sedimentation Is Influenced by the Environment and Developmental Stage of the Plant To optimize exposure to the available environmental resources, it would be beneficial for the plant to regulate the gravitropic response of its organs to a certain degree. One can imagine that under certain circumstances it is in the plant’s best interest to suppress the gravitropic response. Indeed, evidence for such regulatory mechanisms does exist. For example, a downward-growing root (i.e., positively responding to the gravity stimulus) encountering a rock in the soil may benefit from temporarily suppressing its gravitropic response so that it can grow around the encountered obstacle. As discussed in Chapter 5, Massa and Gilroy (2003) have shown that the gravitropic response in Arabidopsis roots is suppressed after tactile stimulation of the peripheral cap cells. Interestingly, it seems that this suppression is correlated with the inhibition of normal amyloplast sedimentation in the statocytes. This finding indicates a complex and direct interaction between mechanisms of thigmo- and gravitropism allowing the roots to navigate the soil and grow around rocks and other obstacles. Another example of down-regulation of the gravitropic response comes from experiments in which it was shown that roots of Arabidopsis and radish show greater hydrotropism and reduced gravitropism after columella amyloplasts are degraded (Iino 2006, and references therein). The interaction between these two tropisms allows the root to inhibit the gravitropic response (by disintegrating amyloplasts) in favor of a search for water (reviewed in Chapter 6). Regulated gravitropism can also be found in lateral roots. Lateral roots need to grow out horizontally before becoming plagio-gravitropic (i.e., assume an oblique orientation relative to the gravity vector). It appears that lateral root gravitropism is delayed until amyloplasts mature and accumulate numerous starch grains in the lateral root columella cells (Kiss et al. 2002). In line with these observations is a recent study by Ma and Hasenstein (2006) that addresses the question of when the embryonic root is capable of sensing gravity. To this end, flax seeds were gravistimulated and allowed to germinate during clinorotation. The onset of gravisensing was determined as the time after which 50% of the emerging roots bent in the direction of the gravity vector during gravistimulation. It was found that the onset of graviperception was established 11 hours before root emergence at the time of germination (and 8 hours after imbibition) and, interestingly, coincided with the development of mature amyloplasts in embryonic columella cells (Ma and Hasenstein 2006). Taken together, the role of starch-statoliths in gravity perception seems undeniable. Almost all graviperceptive tissues and organs in higher plants (stems, leaves, roots, gynophores, and pegs) display sedimenting amyloplasts in their gravisensitive cells. As mentioned above, in some reported cases the development of mature amyloplasts coincides with the ability of cells to perceive gravity and the plant seems to be able to downregulate the gravity-sensing ability by disintegration of amyloplasts. 1.4 The Gravitational Pressure Model for Gravity Sensing Despite general support for plastid-based gravity perception in higher plants, one cannot discount the possibility that plants possess other means to sense gravity. An alternative

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proposal, called the gravitational pressure model, originated with studies in single internodal cells of Chara. Chara internodes display a gravity-dependent polarity of cytoplasmic streaming such that the downward velocity of streaming is 10% faster than the upward stream. However, when positioned horizontally, left and right streaming velocity of the internodes occur at the same rate. A series of studies in the 1990s suggest that this change in the polarity of streaming is a physiological response to gravity and thus reflective of gravity sensing (Wayne and Staves 1996). Because of the lack of sedimentable organelles in the internodes, the gravitational pressure model proposed that the cell perceives gravity by sensing the weight of the entire protoplast. Differential tension between the upper and lower part of the cell then activates receptors in the plasma membrane, initiating events relevant to the gravity response (reviewed in Staves 1997). Although the gravitational pressure model seems to adequately explain the gravityinduced changes in cytoplasmic streaming in giant internodal cells of Chara, the debate continues as to its applicability to gravity sensing in higher plants. One interpretation made by proponents of the gravitational pressure model is that amyloplasts contribute to gravity sensing by simply adding to the weight of the protoplast. They argue that if gravity sensing was dependent on the mass of the protoplast, then media of higher density should inhibit gravitropism. Indeed, when rice roots were grown in aqueous media of higher density due to the addition of various solutes, gravitropism was inhibited without affecting amyloplast sedimentation (Staves et al. 1997). However, these data have yet to be confirmed in other plant species. Instead, one report on protonemata of the moss Ceratodon purpureus contradicts the earlier studies with rice. In plastid-containing protonemata of Ceratodon, robust upward bending was shown to proceed despite growth in high-density media (Schwuchow et al. 2002). It has also been suggested that both plastid and protoplast pressure-based sensing may operate in regulating gravitropism. Indeed, although sedimenting plastids appear to accelerate a gravity response in Arabidopsis inflorescence stems, they are not required to elicit this response (Weise and Kiss 1999). There are earlier examples in the literature that support the gravitational pressure model for gravity sensing (Kiss 2000), but it seems that most of the cell biological and genetic studies in the last five years continue to point to sedimenting plastids as a primary mechanism to explain gravity perception in higher plants (Blancaflor and Masson 2003; Morita and Tasaka 2004).

1.5 The Cytoskeleton in Gravity Perception Although much of the experimental evidence to date continues to favor the starchstatolith hypothesis, it remains unclear how the physical falling of amyloplasts in the gravity-sensing cells is translated into a biochemical or physiological signal that eventually leads to differential organ growth. One cellular structure that has been implicated in modulating gravitropism is the cytoskeleton. Although recent review articles have touched on this topic quite extensively (e.g., Baluˇska and Hasenstein 1997; Volkmann et al. 1999; Blancaflor 2002; Perbal and Driss-Ecole 2003), we cover this area again briefly in light of new evidence pointing to a more complex role for the cytoskeleton in gravity sensing than was previously proposed.

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The plant cytoskeleton is a network of dynamic filamentous proteins that consist of microtubules, actin filaments (F-actin), and a number of accessory proteins (Blancaflor et al. 2006). In the mid- to late 1980s, interest in the cytoskeleton as a mediator of plant gravity sensing was ignited when Wolfgang Hensel published a series of papers describing the organization of the cytoskeleton in the root columella in an attempt to explain the highly polarized nature of this particular cell type (Hensel 1985, 1988, 1989). This was followed by a series of studies on the structure of the cytoskeleton in gravity-sensing cells of roots and shoots, which continued for several years (e.g., White and Sack 1990; Baluˇska et al. 1997; Collings et al. 1998; Collings et al. 2001; Driss-Ecole et al. 2000); (Figure 1.2). The widespread interest in describing the nature of the cytoskeleton in statocytes was likely triggered by the proposal of Andreas Sievers and colleagues in the late 1980s that the cytoskeleton may not only be involved in maintaining statocyte polarity but could also be a direct cellular target of sedimenting amyloplasts, raising the possibility that it might function as a gravity receptor. Sievers et al. (1991) suggested that sedimenting amyloplasts could pull on the cytoskeleton, particularly on F-actin networks, triggering a cascade of signaling events leading to differential organ growth. Indeed, this proposal has been supported by several studies demonstrating that amyloplast movement is dependent on the actin cytoskeleton since drugs that disrupt F-actin alter the dynamics and sedimentation of amyloplasts in space (Volkmann et al. 1999) and on the ground (Hou et al. 2004; Saito et al. 2005; Palmieri and Kiss 2005). In shoot endodermal cells, for example, amyloplast position is restricted by the large, centrally located vacuole, and amyloplasts have to traverse the thin cytoplasmic strands within the cell to sediment (Kato et al. 2002; Saito et al. 2005). As noted earlier, mutants impaired in shoot gravitropism have defects in vacuolar membrane dynamics, and it is possible that the cytoskeleton may function in shoot gravity sensing by regulating the trafficking of vesicles to the vacuole and indirectly influencing amyloplast sedimentation (Morita et al. 2002; Yano et al. 2003). Indeed, shoots treated with latrunculin B, a drug that disrupts F-actin, showed reduced amyloplast settling upon gravistimulation (Friedman et al. 2003; Palmieri and Kiss 2005; Saito et al. 2005). One would predict that if amyloplast sedimentation was impeded by F-actin disruption, the ability of shoots to bend after a gravity stimulus would be inhibited due to altered gravity sensing. Although one study in snapdragon inflorescence stems has demonstrated that this is indeed the case (Friedman et al. 2003), a number of other reports surprisingly show that altering the actin cytoskeleton with latrunculin B can actually promote gravitropism in shoots (Yamamoto and Kiss 2002; Palmieri and Kiss 2005; Saito et al. 2005). It is difficult to explain how impeding the mass translocation of amyloplasts in shoots by disrupting F-actin can promote gravitropism, considering the predictions of the starchstatolith hypothesis. However, one way to interpret these recent results is to view the actin-dependent saltatory movement of amyloplasts in the statocytes as background noise. Reduction of this noise by latrunculin B would essentially increase system sensitivity leading to enhanced gravitropism. Although a majority of amyloplasts in the endodermis are rendered immobile after latrunculin B treatment (Palmieri and Kiss 2005), the few “rogue” amyloplasts that still manage to sediment could still be sufficient to trigger a gravity signal (Saito et al. 2005). With the diminished background noise, the signal gen-

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erated by these “rogue” amyloplasts could be amplified several-fold, leading to a stronger gravitropic response. Alternatively, extended contact between amyloplasts and the vacuolar membrane surface in endodermal cells with a disrupted actin network could be responsible for the enhanced gravity response in shoots (Palmieri and Kiss 2005). Similar to shoots, latrunculin B has a promotive effect on root gravitropism. The enhanced gravity response in roots is clearly manifested as persistent curvature on a slowly rotating clinostat after a short gravistimulus is provided (Hou et al. 2003). In contrast to endodermal cells of shoots, actin disruption has consistently induced the enhancement of amyloplast sedimentation in the root columella upon gravistimulation (Baluˇska et al. 1997; Yoder et al. 2001; Hou et al. 2004). This could be explained by smaller vacuoles in the columella being less of an impediment to amyloplast movement. Nonetheless, like in shoots, the increased sensitivity of roots to gravity may result from diminished system noise and the amplified signals generated by rapidly falling amyloplasts (Hou et al. 2004). Although the above proposal is purely speculative at this point, the ability to manipulate cytoskeletal dynamics specifically in the gravity-sensing cells should provide a significant step toward further testing the relationship between the cytoskeleton and plastid-based gravity sensing in plants. There have been other explanations as to how the cytoskeleton and amyloplasts in the columella interact to generate a gravity signal, including a proposal that localized disruption of the actin network can produce a directional signal by altering the balance of forces acting on plasma membrane receptors (Yoder et al. 2001). Moreover, a recent report showing that actin disruption of decapped maize roots can partially restore gravitropism, which points to the intriguing possibility that actin-dependent gravity sensing may occur outside the root cap (Mancuso et al. 2006). Although it will require additional work to tease apart the different possibilities, results from recent cellular and pharmacological approaches are leading to new, testable hypotheses on how the cytoskeleton mediates in gravity perception in higher plants.

1.6 Concluding Remarks and Future Prospects Gravitropism is important for plant survival since it directs the growth of organs to locations that optimize the utilization of direct environmental resources such as light, water, and soil nutrients. It seems clear that the process of gravity sensing is mainly accomplished through sedimenting amyloplasts in gravisensitive cells of the columella in roots, the endodermis in dicotyledon shoots, and the pulvinus in monocotyledons. However, one cannot totally discount the experimental evidence pointing to alternative gravityperceiving sites, or that other cellular structures (including the whole cell itself) could function as gravitropic susceptors, in addition to falling plastids. Given the importance of gravitropism in plant development, it might not be surprising to find some redundancy in mechanisms of gravity sensing. Evolutionarily speaking, a single mutation that would wipe out amyloplast formation could severely compromise survival if the plant relies entirely on amyloplasts for gravity sensing. With this in mind, it is reasonable to imagine that the starch-statolith hypothesis and the gravitational pressure model are not necessarily conflicting, even though evidence for the starch-statolith hypothesis seems over-

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whelming. Both processes working in tandem might be part of a redundant mechanism to ensure the plant’s best chances for survival in its environment. In conclusion, although our view about gravity perception in higher plants continues to revolve around the idea of sedimenting plastids, one of the more pressing issues is how the gravity-induced mechanical stimulus of falling statoliths or, alternatively, the pressure exerted by the entire protoplast, is converted to a chemical signal that subsequently regulates a physiological response. As discussed in Chapter 2, studies involving the interactions between amyloplasts and other cell organelles such as the vacuole, the cytoskeleton, and the endoplasmic reticulum are fruitful avenues for research that will help address this issue. In addition, studies of gravitropism in other plant organs might provide useful additional information about the gravitropic response. For example, the peanut gynophore could be developed into a useful model to discern how the gravity signal is translated into a positive (i.e., downward) or negative (i.e., upward) response. Moreover, transcript and protein profiling are helping to identify additional molecular players in the gravitropic response (Moseyko et al. 2002; Kimbrough et al. 2004). An exciting avenue for future research will be to observe changes in transcript, protein, and metabolite levels specifically in the cells that are presumed to sense gravity. 1.7 Acknowledgment Funding from the Samuel Roberts Noble Foundation Inc. and the National Science Foundation (DBI-0400580) is gratefully acknowledged. 1.8 Literature Cited Allen NS, Chattaraj P, Collings D, and Johannes E. 2003. Gravisensing: Ionic responses, cytoskeleton and amyloplast behavior. Advances in Space Research 32:1631–7. Baluˇska F, and Hasenstein KH. 1997. Root cytoskeleton: Its role in perception of and response to gravity. Planta 203:S69–S78. Baluˇska F, Kreibaum A, Vitha S, Parker JS, Barlow PW, and Sievers A. 1997. Central root cap cells are depleted of endoplasmic microtubules and actin microfilament bundles: Implications for their role as gravity-sensing statocytes. Protoplasma 196:212–23. Blancaflor EB. 2002. The cytoskeleton and gravitropism in higher plants. Journal of Plant Growth Regulation 21:120–36. Blancaflor EB, Fasano J, and Gilroy S. 1998. Mapping the functional roles of cap cells in the response of Arabidopsis primary roots to gravity. Plant Physiology 116:213–22. Blancaflor EB, and Masson PH. 2003. Update on plant gravitropism. Unraveling the ups and downs of a complex process. Plant Physiology 113:1677–90. Blancaflor EB, Wang Y-S, and Motes CM. 2006. Organization and function of the cytoskeleton in developing root cells. International Review of Cytology 252:219–264. Boonsirichai K, Guan C, Chen R, and Masson PH. 2002. Root gravitropism: An experimental tool to investigate basic cellular and molecular processes underlying mechanosensing and signal transmission in plants. Annual Review of Plant Biology 53:421–47. Caspar T, and Pickard BG. 1989. Gravitropism in a starchless mutant of Arabidopsis: Implications for the starch-statolith theory of gravity sensing. Planta 177:185–97.

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Collings DA, Winter H, Wyatt SE, and Allen NS. 1998. Growth dynamics and the cytoskeleton organization during stem maturation and gravity-induced stem bending in Zea mays L. Planta 207:246–58. Collings DA, Zsuppan G, Allen NS, and Blancaflor EB. 2001. Demonstration of prominent actin filaments in the root columella. Planta 212:392–403. Driss-Ecole D, Vassy J, Rembur J, Giuvarch A, Proteau M, Dewitte W, and Perbal G. 2000. Immunolocalization of actin in root statocytes of Lens culinaris L. Journal of Experimental Botany 51:512–28. Esmon CA, Pedmale UV, and Liscum E. 2005. Plant tropisms: Providing the power of movement to a sessile organism. International Journal of Developmental Biology 49:665–74. Friedman H, Vos JW, Hepler PK, Meir S, Halevy AH, and Philosoph-Hadas S. 2003. The role of actin filaments in the gravitropic response of snapdragon flowering shoots. Planta 216: 1034–42. Friml J, Winiewska J, Benková E, Mendgen K, and Palme K. 2002. Lateral relocation of auxin efflux regulator PIN3 mediates tropism in Arabidopsis. Nature 415:806–9. Fujihara K, Kurata T, Watahiki MK, Karahara I, and Yamamoto KT. 2000. An agravitropic mutant of Arabidopsis, endodermal-amyloplast less, that lacks amyloplasts in hypocotyls endodermal cell layer. Plant Cell Physiology 41:1193–9. Fukaki H, Wysocka-Diller J, Kato T, Fujisawa H, Benfey PN, and Tasaka M. 1998. Genetic evidence that the endodermis is essential for shoot gravitropism in Arabidopsis thaliana. Plant Journal 14:425–30. Hatakeda Y, Kamada M, Goto N, Fukaki H, Tasaka M, Suge H, and Takahashi H. 2003. Gravitropic response plays an important role in the nutational movements of the shoots of Pharbitis nil and Arabidopsis thaliana. Physiologia Plantarum 118:464–73. Hensel W. 1985. Cytochalasin B affects the structural polarity of statocytes from cress roots Lepidium sativum L. Protoplasma 129:178–87. Hensel W. 1988. Demonstration by heavy meromyosin of actin microfilaments in extracted cress (Lepidium sativum L.) root statocytes. Planta 173:142–3. Hensel W. 1989. Tissue slices from living root caps as a model system in which to study cytodifferentiation of polar cells. Planta 177:296–303. Hou G, Mohamalawari DR, and Blancaflor EB. 2003. Enhanced gravitropism of roots with a disrupted cap actin cytoskeleton. Plant Physiology 131:1360–73. Hou G, Kramer VL, Wang Y-S, Chen R, Perbal G, Gilroy S, and Blancaflor EB. 2004. The promotion of gravitropism in Arabidopsis roots upon actin disruption is coupled with the extended alkalinization of the columella cytoplasm and a persistent lateral auxin gradient. Plant Journal 31:113–25. Iino M. 2006. Toward understanding the ecological functions of tropisms: Interactions among and effects of light on tropisms. Current Opinion in Plant Biology 9:89–93. Kato T, Morita MT, Fukaki H, Yamauchi Y, Uehara M, Niihama M, and Tasaka M. 2002. SGR2, a phospholipase-like protein, and ZIG/SGR4, a SNARE, are involved in the shoot gravitropism of Arabidopsis. Plant Cell 14:33–46. Kimbrough JM, Salinas-Mondragon R, Boss WF, Brown CS, and Sederoff HW. 2004. The fast and transient transcriptional network of gravity and mechanical stimulation in the Arabidopsis root apex. Plant Physiology 136:2790–805. Kiss JZ. 2000. Mechanisms of the early phases of plant gravitropism. Crit Rev Plant Sci 19:551–73. Kiss JZ. 2006. Up, down, and all around: How plants sense and respond to environmental stimuli. Proceedings of the National Academy of Sciences USA 103:829–30. Kiss JZ, Guisinger MM, Miller, AJ, and Stackhouse KS. 1997. Reduced gravitropism in hypocotyls of starch-deficient mutants of Arabidopsis. Plant Cell Physiology 38:518–25.

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Kiss JZ, Hertel R, and Sack FD. 1989. Amyloplasts are necessary for full gravitropic sensitivity in roots of Arabidopsis thaliana. Planta 177:198–206. Kiss JZ, Miller KM, Ogden LA, and Roth KK. 2002. Phototropism and gravitropism in lateral roots of Arabidopsis. Plant Cell Physiology 43:35–43. Kitazawa D, Hatakeda Y, Kamada M, Fujii N, Miyazawa Y, Hoshino A, Iida S, Fukaki H, Morita MT, Tasaka M, Suge H, and Takahashi H. 2005. Shoot circumnutation and winding movements require gravisensing cells. Proceedings of the National Academy of Sciences USA 102:18742–7. Konings H. 1995. Gravitropism of roots: An evaluation of progress during the last three decades. Acta Botanica Neerlandica 44:195–223. Kuznetsov OA, and Hasenstein KH. 1996. Intracellular magnetophoresis of amyloplasts and induction of root curvature. Planta 198:87–94. Ma Z, and Hasenstein KH. 2006. The onset of gravisensitivity in the embryonic root of flax. Plant Physiology 140:159–66. Mancuso S, Barlow PW, Volkmann D, and Baluˇska F. 2006. Actin turnover-mediated gravity response in maize root apices. Gravitropism of decapped roots implicates gravisensing outside of the root cap. Plant Signaling and Behavior 1:52–8. Mano E, Horiguchi G, and Tsukaya H. 2006. Gravitropism in leaves of Arabidopsis thaliana (L.) Heynh. Plant Cell Physiology 47:217–23. Massa GD, and Gilroy S. 2003. Touch modulates gravity sensing to regulate the growth of primary roots of Arabidopsis thaliana. Plant Journal 33:435–45. Moctezuma E, and Feldman LJ. 1999. The role of amyloplasts during gravity perception in gynophores of the peanut plant (Arachis hypogaea). Annals of Botany 84:709–14. Morita MT, Kato T, Nagafusa K, Saito C, Ueda T, Nakano A, and Tasaka M. 2002. Involvement of the vacuoles of the endodermis in the early process of shoot gravitropism in Arabidopsis. Plant Cell 14:47–56. Morita MT, and Tasaka M. 2004. Gravity sensing and signaling. Current Opinion in Plant Biology 7:712–18. Moseyko N, Zhu T, Chang H-S, Wang X, and Feldman, LJ. 2002. Transcription profiling of the early gravitropic response in Arabidopsis using high-density oligonucleotide probe microarrays. Plant Physiology 130:720–8. Palmieri M, and Kiss JZ. 2005. Disruption of the F-actin cytoskeleton limits statolith movement in Arabidopsis hypocotyls. Journal of Experimental Botany 419:2539–50. Perbal G, and Driss-Ecole D. 2003. Mechanotransduction in gravisensing cells. Trends in Plant Science 8:498–504. Perera IY, Heilmann I, and Boss WF. 1999. Transient and sustained increases in inositol 1,4,5triphosphate precede the differential growth response in gravistimulated maize pulvini. Proceedings of the National Academy of Sciences, USA 96:5838–43. Perera IY, Heilmann I, Chang SC, Boss WF, and Kaufman PB. 2001. A role for inositol 1,4,5triphosphate in gravitropic signaling and the retention of cold-perceived gravistimulation of oat shoot pulvini. Plant Physiology 125:1499–507. Perera IY, Hung CY, Brady S, Muday GK, and Boss WF. 2006. A universal role for inositol 1,4,5trisphosphate-mediated signaling in plant gravitropism. Plant Physiology 140:746–60. Perrin RB, Young L-S, Murthy N, Harrison BR, Wang Y, Will JL, and Masson PH. 2005. Gravity signal transduction in primary roots. Annals of Botany 96:737–43. Sack FD, Hasenstein KH, and Blair A. 1990. Gravitropic curvature of maize roots is not preceded by root cap asymmetry. Annals of Botany 66:203–9. Sack FD. 1991. Plant gravity sensing. International Review of Cytology 127:193–252. Sack FD. 1997. Plastids and gravitropic sensing. Planta 203:S63–S68.

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Saether N, and Iversen TH. 1991. Gravitropism and starch statoliths in an Arabidopsis mutant. Planta 184:491–7. Saito Y, Yamasaki S, Fuji N, Hagen G, Guilfoyle T, and Takahashi H. 2004. Isolation of cucumber CsARF cDNAs and expression of the corresponding mRNAs during gravity-regulated morphogenesis of cucumber seedlings. Journal of Experimental Botany 55:1315–23. Saito C, Morita MT, Kato T, and Tasaka M. 2005. Amyloplasts and vacuolar membrane dynamics in the living graviperceptive cell of the Arabidopsis inflorescence stem. Plant Cell 17:548–558. Schwuchow JM, Kern VD, and Sack FD. 2002. Tip-growing cells of the moss Ceratodon purpureus are gravitropic in high-density media. Plant Physiology 130:2095–2100. Sievers A, Buchen B, Volkmann D, and Hejnowicz Z. 1991. Role of cytoskeleton in gravity perception. In The cytoskeletal basis of plant growth and form, edited by Clive W. Lloyd, 169–82. London: Academic Press. Sievers A, Buchen B, and Hodick D. 1996. Gravity sensing in tip-growing cells. Trends in Plant Science 1:273–9. Silady RA, Kato T, Kukowitz W, Sieber P, Tasaka M, and Somerville CR. 2004. The gravitropism defective 2 mutants of Arabidopsis are deficient in a protein implicated in endocytosis in Caenorhabditis elegans. Plant Physiology 136:3095–103. Staves MP. 1997. Cytoplasmic streaming and gravity sensing in Chara internodal cells. Planta 203:S79–S84. Staves MP, Wayne R, and Leopold AC. 1997. The effect of external medium on the gravitropic curvature of rice (Oryza sativa, Poaceae) roots. American Journal of Botany 84:1522–9. Takahashi H, Kamada M, Yamazaki Y, Fujii N, Higashitani A, Aizawa S, Yoshizaki I, Kamigaichi S, Mukai C, Shimazu T, and Kukuki K. 2000. Morphogenesis in cucumber seedlings is negatively controlled by gravity. Planta 210:515–8. Tanaka A, Kobayashi Y, Hase Y, and Watanabe H. 2002. Positional effect of cell inactivation on root gravitropism using heavy-ion microbeams. Journal of Experimental Botany 53:683–7. Tsugeki R, and Fedoroff NV. 1999. Genetic ablation of root cap cells in Arabidopsis. 96:12941–6. Volkmann D, Baluˇska F, Lichtscheidl IK, Driss-Ecole D, and Perbal G. 1999. Statoliths motions in gravity-perceiving plant cells: Does actomyosin counteract gravity? FASEB Journal 13:S143–S147. Wang Y-S, Motes CM, Mohamalawari DR, and Blancaflor EB. (2004). Green fluorescent protein fusions to Arabidopsis fimbrin 1 for spatio-temporal imaging of F-actin dynamics in roots. Cell Motility and the Cytoskeleton 59:79–93. Wayne R, and Staves MP. 1996. A down-to-earth model of gravisensing or Newton’s Law of Gravitation from the apple’s perspective. Gravitational and Space Biology Bulletin 98:917–21. Weise SE, and Kiss J. 1999. Gravitropism of inflorescence stems in starch-deficient mutants of Arabidopsis. Int J Plant Sci 160:521–7. Weise SE, Kuznetsov OA, Hasenstein KH, and Kiss JZ. 2000. Curvature in Arabidopsis inflorescence stems is limited to the regions of amyloplasts displacement. Plant and Cell Physiology 41:702–9. White RG, and Sack FD. 1990. Actin microfilaments in presumptive statocytes of roots caps and coleoptiles. American Journal of Botany 77:17–26. Wolverton C, Mullen JL, Ishikawa H, and Evans ML. 2002a. Root gravitropism in response to a signal originating outside of the cap. Planta 215:153–7. Wolverton C, Ishikawa H, and Evans ML. 2002b. The kinetics of root gravitropism: Dual motors and sensors. Journal of Plant Growth and Regulation 21:102–12. Yamamoto K, and Kiss JZ. 2002. Disruption of the actin cytoskeleton results in the promotion of gravitropism in inflorescence stems and hypocotyls of Arabidopsis. Plant Physiology 128: 669–81.

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Yano D, Sato M, Saito C, Sato MH, Morita MT, and Tasaka M. 2003. A SNARE complex containing SGR3/AtVAM3 and ZIG/VTI11 in gravity-sensing cells is important for Arabidopsis shoot gravitropism. Proceedings of the National Academy of Sciences, USA 100:8589–94. Yoder TL, Zheng H-Q, Todd P, and Staehelin LA. 2001. Amyloplast sedimentation dynamics in maize columella cells support a new model for the gravity-sensing apparatus of roots. Plant Physiology 125:1045–60. Yoshihara T, and Iino M. 2005. Circumnutation of rice coleoptiles: Its occurrence, regulation by phytochrome, and relationship with gravitropism. Plant Cell and Environment 28:134–46. Yoshihara T, and Iino M. 2006. Circumnutation of rice coleoptiles: Its relationships with gravitropism and absence in lazy mutants. Plant Cell and Environment 29:778–92.

2

Signal Transduction in Gravitropism Benjamin R. Harrison, Miyo T. Morita, Patrick H. Masson*, and Masao Tasaka

2.1 Introduction As discussed in Chapter 1, most plant organs use gravity as a growth guide. However, different organs will interpret that information in different ways. Shoots grow upward toward light, aboveground environments in order to optimize photosynthesis, exchange gases, and perform their reproductive functions. Most roots, on the other hand, grow downward, into the soil, where they anchor the plant and take up water and nutrients necessary for plant growth, development, and reproduction. To understand how these organs interpret differently the information provided by gravity, we first need to understand the molecular mechanisms that govern gravity signal transduction in gravity-sensing cells, termed statocytes. Before we describe the current state of our knowledge on the mechanisms that govern gravity signal transduction in plants, it is important to understand that the gravityresponding organs of higher plants are diversified in their tissue structure and developmental origin. In cereal grasses, the graviresponsive coleoptile of seedlings is a hollow cylindrical sheath, whereas the pulvini of adult plants are swellings at the base of each internode. In dicots, hypocotyls, and epicotyls of young seedlings and leaf petioles and stems of adult plants are all graviresponsive. Similarly, both primary and lateral roots are graviresponsive in monocots and dicots, even though the site of root gravicurvature does not contain obviously differentiated statocytes. Hence, the morphology and cytology of different plant organs may affect the machinery that modulates their gravitropic responses. In spite of such diversity, all graviresponsive organs share two common features: they contain graviperceptive cells with sedimentable amyloplasts (Sack 1997), and they develop asymmetry in auxin concentration between their upper (lower concentration) and lower (higher concentration) flanks upon gravistimulation (Philippar et al. 1999; Muday and DeLong 2001; Friml et al. 2002; Long et al. 2002). Thus, within these organs, a gravitational signal perceived through the relocalization of amyloplasts within differentiated statocytes is converted into biochemical signal(s) that is (are) transmitted to adjacent cells, leading to the formation of a lateral gradient of auxin at the elongation zone, responsible for the gravitropic curvature (see also Chapter 3). Although seemingly similar in global terms, the physiological and biochemical events that accompany gravitropism in aboveground organs and roots differ substantially in the details. For instance, the site of gravity perception and signal transduction (endodermal cells) overlaps with the site of curvature response in shoots, which has been proposed to occur simultaneously and uniformly along the organs (Firn and Digby 1980). Roots, on the other hand, show a physical separation between the primary site of gravity perception and signal transduction (the root cap columella) and the site of curvature response (the *Corresponding author

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elongation zones) (Masson et al. 2002). Hence, in roots, the gravity-induced lateral auxin gradient generated across the cap has to be transmitted basipetally, from the site of perception and signal transduction to the site of response in order for a gravitropic curvature to develop (Figure 2.1A and Color Section). In aboveground organs, the signal has only to be transmitted laterally, across the organ. In the Arabidopsis shoot elongation zone, the epidermis, cortex, gravity-perceiving endodermis, and stele are arranged radially, in successive layers, and Arabidopsis stem segments dissected from any part of the elongation zone are gravitropic (Fukaki et al. 1996). Thus, the gravity signal perceived in the endodermal cell layer may be transmitted in an inner-to-outer fashion (laterally), leading to a unilateral, asymmetric auxin distribution between the lower and upper flanks that results in differential cell elongation and consequent gravitropic curvature (Fukaki et al. 1998; Tasaka et al. 1999). This centrifugal aspect of gravitropic signaling between cell types seems general in aboveground organs. However, we do not know whether all cells in the endodermal layer play equal roles in gravity perception and signaling (Figure 2.1B and Color Section). Recent physiological and molecular genetic studies have provided insights into the mechanisms that govern gravity signal transduction in roots, hypocotyls, shoots, and cereal pulvini. This information, summarized in this chapter, will provide the foundation for future research aimed at better understanding how this machinery might be modulated by endogenous and environmental cues.

2.2 Gravity Signal Transduction in Roots and Aboveground Organs Roots constitute an excellent system for analysis of gravity signal transduction in plants. The physical separation existing between the primary site of gravity perception (the root cap columella) and the site of differential cellular elongation that drives the corresponding curvature response (the elongation zones) allows for careful dissection of the molecular mechanisms involved in each phase of the process. Hence, it is not surprising that several teams have focused their research on this system to decipher some of the molecular mechanisms that govern gravity signal transduction. The gravity receptor remains unknown. However, several potential gravity signal transducers have recently been uncovered. They include cytoplasmic cations, inositol 1,4,5-trisphosphate (InsP3), and several proteins. Downstream effects of activation of the gravity signal transduction pathway in the root cap were recently uncovered. They include rapid changes in gene expression within the root tip (Moseyko et al. 2002; Kimbrough et al. 2004); a lateral polarization of the statocytes as reflected by accumulation of the PIN3 auxin efflux facilitator at the lower membrane of gravistimulated statocytes (Friml et al. 2002; Harrison and Masson 2006); and a preferential downward lateral transport of auxin across the cap, with accumulation at the lower flank (Young et al. 1990; Rashotte et al. 2001; Boonsirichai et al. 2003). Because the subsequent lateral auxin gradient at the cap is transported basipetally along the root tip, it will eventually direct differential cellular elongation between upper and lower root flanks at the elongation zones, which drives most of the gravitropic curvature (Chapter 3; reviewed in Ishikawa and Evans 1997).

Figure 2.1 (also see Color Section). Histological differences between roots and inflorescence stems translate into gravitropic signaling specificities in Arabidopsis thaliana. A. Root from a 4-day-old light-grown seedling (left) showing the cap, distal and central elongation zones (DEZ and CEZ), and the mature zone (MZ). The inset (right) represents a confocal image of a propidium iodide-stained root tip showing the root cap (with its L1, L2, and L3 layers of columella cells, and lateral cap cells, LRC), the promeristem (with its quiescent center cells, QC, surrounded by initials). The root proper is composed of several cell layers, including the epidermis (Ep); the cortex (C); the endodermis (En); and the stele (St). The primary site of gravity perception in roots is the root cap, with the statocytes located in the first two layers of columella cells, whereas the curvature response initiates in the DEZ. Therefore, a gravitropic signal has to be transmitted basipetally upon gravistimulation. B. Top of an inflorescence stem showing the stem ended by a shoot apical meristem (SAM), flowers, and siliques. The entire stem region located below the SAM in this picture is part of the elongation zone (EZ). The middle drawing corresponds to a region of shoot stem. Microscopical image of a longitudinal section of this region (indicated by the rectangle in the drawing) is represented on the right showing the epidermis (Ep), three layers of cortical cells (C), one layer of endodermis (En), and the stele (St). Sedimenting amyloplasts, located in the endodermal statocytes, are indicated by black arrowheads. Hence, the sites of gravity sensing and curvature response overlap in stems, and the gravitropic signal has to be transmitted outward, rather than basipetally, in order for a curvature response to develop. The American Society of Plant Biologists is the copyright holder for this figure, which first appeared in Masson et al. (2002).

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The same second messengers have been implicated in the modulation of gravity signal transduction in aboveground organs as in roots, including cytoplasmic pH (Johannes et al. 2001) and InsP3 (Perera et al. 1998, 1999, 2001). Furthermore, several proteins have been identified as potential gravity signal transducers in shoots (Yamauchi et al. 1997; Wyatt et al. 2002; Morita et al. 2006). Yet, there has been no indication of relocalization of auxin transporters within the statocytes in aboveground organs, even though evidence for asymmetric downward transport of auxin across gravistimulated organs also exists in these systems (Li et al. 1991; Kaufman et al. 1995; Philippar et al. 1999; Friml et al. 2002; Long et al. 2002; Abas et al. 2006). Below, we briefly discuss how these pieces of the gravity signal transduction puzzle might fit together to promote pathways that ultimately lead to the curvature of roots and aboveground organs. 2.2.1 Do Mechano-Sensitive Ion Channels Function as Gravity Receptors? As discussed in Chapter 1, several researchers have proposed that membrane-associated mechano-sensitive ion channels might function as gravity receptors in plant statocytes. The sedimentation of, and/or pressure/tension exerted by amyloplasts would trigger the opening of such channels at sensitive membranes. This would allow for a flux of Ca2+ ions within the statocytes, serving as second messengers to trigger a cascade of events that would ultimately lead to the lateral polarization of statocytes, as discussed above (Sievers et al. 1984; Sievers et al. 1989; Pickard and Ding 1993; Volkmann and Baluˇska 1999; Yoder et al. 2001). As discussed in Chapter 5, a number of pharmacological studies support a role for Ca2+ in gravity signal transduction. Agents that inactivate mechano-sensitive ion channels (i.e., Gd3+ or La3+) alter the function of Ca2+ regulatory proteins (calmodulin and Ca2+-ATPases), and Ca2+ chelators all inhibit gravitropism (reviewed in Sinclair and Trewavas 1997; Fasano et al. 2002). If Ca2+-selective, mechano-sensitive ion channels contribute to gravity signal transduction in the statocytes, one should be able to detect changes in cytosolic Ca2+ levels upon gravistimulation. In an attempt to detect such Ca2+ responses, Plieth and Trewavas (2002) generated transgenic Arabidopsis thaliana seedlings that constitutively express apoaequorin. This protein interacts with coelenterazine (an exogenously added compound that freely diffuses through membranes) to generate a cytoplasmic aequorin complex whose luminescence intensity is proportional to the levels of Ca2+ within the cytoplasm (Knight et al. 1991). They then analyzed the intensity of light emitted by groups of seedlings in response to gravistimulation. Under these conditions, gravistimulation promoted biphasic and transient peaks of aequorin luminescence, reflective of transient peaks in cytosolic Ca2+ levels. The first peak lasted a few seconds and was reminiscent of responses to mechano-stimulation. The second peak lasted several minutes, a duration that was related to the intensity of the stimulus (angle of plant reorientation within the gravity field). It was inhibited by treatments that alter polar auxin transport. Clinorotation (low-speed rotation within the gravity field to avoid constant stimulation), on the other hand, promoted a sustained rise in luminescence. It was concluded that a change in cytosolic Ca2+ levels follows gravistimulation and may contribute to gravity signal transduction in Arabidopsis seedlings (Plieth and Trewavas 2002).

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Unfortunately, the aequorin-derived signal could only be detected if hundreds of Arabidopsis seedlings were examined in bulk, and the luminescence was too weak for identification of its source. In the future, it will be necessary to use a similar detection strategy to analyze gravity-induced Ca2+ transients in seedlings that express aequorin specifically within the root and hypocotyl statocytes to determine whether the signal derives from these cells. It will also be important to determine whether such a signal disappears in mutants that are defective in early phases of gravity signal transduction, such as the starch-deficient and signal-transduction mutants described below. In separate experiments using either the system described above or Ca2+-sensitive fluorophores, other researchers were unable to detect changes in cytosolic Ca2+ levels in the statocytes upon gravistimulation (Sedbrook et al. 1996; Legue et al. 1997; Fasano et al. 2002; Massa and Gilroy 2003). Although these negative results cast doubt on the possible involvement of Ca2+ in gravity signal transduction within the statocytes, they may reflect a lack of sensitivity of the Ca2+ detection approaches used, or a highly localized, yet functionally significant, Ca2+ pulse undetectable by these sensors. It is interesting to note that even small and/or highly localized changes in cytosolic Ca2+ levels within the root statocytes might be functionally relevant because these cells express high levels of calmodulin (Sinclair et al. 1996; see also Chapter 5). Additional complexity arises in studies that investigate the role of cytosolic Ca2+ in gravity signal transduction within the root statocytes. As emphasized in Chapter 1, touch stimulation of the root tip promotes a fast increase in cytosolic Ca2+ levels within the stimulated peripheral cap cells, which propagates to surrounding cells to eventually reach the columella region. There, the corresponding Ca2+ wave appears to inhibit gravitropism by interfering with amyloplast sedimentation (Massa and Gilroy 2003). Hence, if a Ca2+ wave signals an inhibition of gravitropic sensitivity in the statocytes in response to a touch stimulus at the cap, a role for gravity-induced Ca2+ flux in gravity signal transduction within the statocytes would require a corresponding Ca2+ signal that displays a distinctive signature (Massa and Gilroy 2003). Models postulating the involvement of mechano-sensitive ion channels as gravity receptors in plant statocytes currently suffer from a major roadblock: such channels have not been identified. This may be because most of the ion channels identified as mechanosensitive in animals and yeast do not have obvious orthologs in plants (Barritt and Rychkov 2005; Haswell and Meyerowitz 2006; for a more detailed discussion, see Chapter 5). The recent discovery of a family of 10 Arabidopsis genes encoding membrane-spanning proteins related to the bacterial MscS channels, which serve to protect the bacteria against cellular lysis during osmotic downshock (Kung and Blount 2004), may constitute an important breakthrough (Haswell and Meyerowitz 2006). Indeed, initial studies on one of these Arabidopsis MscS-like proteins (MSL3) demonstrated its ability to rescue the osmotic sensitivity of an E. coli mutant lacking mechanosensitive ion channel activity. This suggests the channel is mechano-sensitive, at least in this bacterial system. Interestingly, MSL3 and the highly related MSL2 protein were targeted to the plastid membranes in plants, where they seemed to localize at focal sites associated with proteins implicated in plastid division. Furthermore, double mutants showed abnormal plastid morphology, suggestive of defects in plastid division. These exciting results were used to suggest an involvement of these related proteins in regulating

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the plastids’ osmotic pressure during division through mechanisms related to those of mechano-sensitive ion channels (Haswell and Meyerowitz 2006). As of now, there has been no direct evidence for an involvement of these two potential mechano-sensitive ion channels in gravity signal transduction (Haswell and Meyerowitz 2006). However, some of the eight remaining Arabidopsis MSL genes are predicted to encode proteins targeted to other cellular compartments. Even more strikingly, some of these are highly expressed in the root statocytes and are transcriptionally responsive to gravistimulation (Neal and Masson, unpublished data; Nawy et al. 2005). Careful functional analysis of this outstanding set of genes, along with that of other potential plant ion channels with as-yet poorly defined properties and functions (Schulz et al. 2006), may soon yield important new insights into the gravitropic response. It remains quite possible that gravity reception involves mechanisms that do not rely upon mechano-sensitive ion channels. It is quite exciting to note recent developments in the study of model systems that involve single-cell gravitropic responses. As discussed by Braun and Hemmersbach in Chapter 7 of this book, Chara rhizoids sense gravity through the sedimentation of BaSO4-containing statoliths. Elegant experiments with this system have demonstrated that the statoliths must sediment onto sensitive membranes at a subapical region of the cell for the signal to be perceived and transduced into a curvature response (Braun 2002). Interestingly, experiments involving short-term exposure to hypo- and hypergravity have indicated that simple contact of statoliths with the sensitive membrane is sufficient for gravity perception to occur; differential pressure or tension is not needed (Braun 2002; Limbach et al. 2005). This result led the authors to postulate that gravity signal transduction might be triggered by molecular interaction between ligands carried by the sedimenting statoliths and receptors located at the sensitive membranes (Limbach et al. 2005). A similar ligand-receptor model of gravity reception should not be excluded in higher plants. In fact, experiments involving starch-deficient mutants of Arabidopsis (which have lighter amyloplasts that do not appear to sediment), or seedlings exposed to drugs that destabilize the actin filaments in shoot statocytes—thereby disabling amyloplast sedimentation (see Chapter 1)—have suggested that a few “rogue” sedimenting amyloplasts might be responsible for the remaining gravitropic capability associated these systems (Kiss et al. 1997; Palmieri and Kiss 2005; Saito et al. 2005). This interesting model could be tested by subjecting higher plants to short-term hyper- and hypogravity treatments similar to those performed on Chara (Limbach et al. 2005). 2.2.2 Inositol 1,4,5-Trisphosphate Seems to Function in Gravity Signal Transduction A potential role for inositol 1,4,5-trisphosphate (InsP3) in gravity signal transduction was recently suggested from elegant physiological, biochemical, and genetic studies utilizing both aboveground and root model systems. These experiments demonstrated both the existence of gravity-induced changes in InsP3 levels in stimulated organs and a need for wild-type levels of InsP3 for full graviresponsiveness. Biphasic changes in InsP3 levels upon gravistimulation were first reported for pulvini of maize and oat (Perera et al. 1999; Perera et al. 2001). In this system, transient fluctuations in InsP3 levels were observed in both upper and lower pulvinus halves within 10 to 15 sec of gravistimulation. This was followed by a long-term, sustained increase that

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occurred only in the lower pulvinus halves and correlated with the bending response. These changes in InsP3 levels were functionally relevant because treatments with an inhibitor of phospholipase C, an enzyme that contributes to InsP3 synthesis, blocked the increase in InsP3 levels and inhibited the bending response in oat (Perera et al. 2001). Recently, the same investigating team detected similar biphasic changes of InsP3 levels upon gravistimulation in Arabidopsis inflorescence stems (Perera et al. 2006). In both cereal pulvini and dicot inflorescence stems, the second increase in InsP3 levels preceded visible bending responses, suggesting its involvement in early phases of the pathway. To further investigate the potential role of InsP3 in gravity signal transduction, transgenic Arabidopsis thaliana plants expressing human type I inositol polyphosphate 5-phosphatase, an enzyme that specifically hydrolyzes InsP3, were generated. These plants grew like wild type despite containing very low levels of InsP3 (10% of wild-type levels). However, they exhibited reduction in the kinetics of inflorescence-stem, hypocotyl, and root gravitropism, enhanced root gravitropic sensitivity to extracellular Ca2+, decrease in basipetal auxin transport along the root, and delay in the development of lateral auxin gradients upon gravistimulation, as deduced from expression analyses of auxin-sensitive reporter constructs (Perera et al. 2006). Because InsP3 is a soluble second messenger that propagates localized Ca2+ fluxes through the cell and to neighboring cells, and the levels of both molecules display parallel, biphasic increases upon gravistimulation, it appears likely that both Ca2+ and InsP3 contribute to gravity signal transduction in most or all organs of the plant, possibly by modulating auxin transport (Plieth and Trewavas 2002; Perera et al. 2006). Determination of the tissue(s) within responding organs where both InsP3 and Ca2+ changes occur upon gravistimulation should provide important insights into their mode of action. 2.2.3 Do pH Changes Contribute to Gravity Signal Transduction? Although an involvement of cytosolic Ca2+ as a second messenger in gravity signal transduction remains speculative, better evidence exists for a contribution of cytoplasmic pH in this phase of the response. If cytosolic pH contributes to gravity signal transduction, its level in the statocytes should change upon gravistimulation, and interference with such changes should affect the response. Indeed, both assumptions were recently validated. Rapid cytoplasmic pH changes upon gravistimulation were observed in the statocytes of both maize pulvini and Arabidopsis roots (Scott and Allen 1999; Fasano et al. 2001; Johannes et al. 2001; Boonsirichai et al. 2003; Hou et al. 2004; Young et al. 2006). By using longitudinal maize stem sections loaded with a pH indicator, Johannes and collaborators were able to monitor the cytoplasmic pH of both gravity-sensing bundle-sheath and parenchyma cells upon gravistimulation. They found that gravistimulation promotes a fast alkalinization of the bundle-sheath statocytes without altering the pH of parenchyma cells. They also found that the gravity-induced cytoplasmic alkalinization in the pulvinus statocytes occurs only in a restricted region of the cytoplasm where the sedimenting amyloplasts accumulated. Similar experiments in which the cytoplasmic pH of the root statocytes was measured during gravistimulation of live Arabidopsis seedlings indicated that a transient alkalinization occurs early after gravistimulation onset, peaks within 1 to 2 min, and returns to

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baseline levels within 8 to 10 min (Scott and Allen 1999; Fasano et al. 2001; Boonsirichai et al. 2003; Young et al. 2006). Although Scott and Allen suggested distinct kinetics in the cytoplasmic alkalinization of layer-2 cells (layers of columella cells are represented in Figure 2.1A) between upper and lower statocytes of gravistimulated roots, and an acidification of layer-3 cells, Fasano and collaborators described similar alkalinization in all statocytes within both columella tiers (Fasano et al. 2001). Gravity-induced alkalinization of the root statocytes was dramatically attenuated in pgm, a mutant that contains starchless plastids and displays altered gravisensitivity (Fasano et al. 2001). It was also accompanied by an acidification of the apoplast in wild-type roots, suggesting it might result from the activation of plasma membrane or vacuolar H+ transporters (Fasano et al. 2001; Li et al. 2005). Interestingly, altering the cytoplasmic pH of root statocytes by releasing preloaded caged protons resulted in a delay in the gravitropic response (Fasano et al. 2001). Furthermore, treating root caps with agents that acidify the apoplast and the cytoplasm at concentrations that do not alter the overall rate of root growth resulted in an enhancement of the gravitropic response, whereas treatments with alkalinizing agents delayed it (Scott and Allen 1999). Likewise, treatments with agents that disrupt actin filaments resulted in both sustained cytoplasmic alkalinization upon short periods of gravistimulation and enhanced gravicurvature, as discussed in Chapter 1 (Hou et al. 2004). Even though the initial studies differed in the details of their observations, current data converge to suggest an important role for cytosolic pH in gravity signal transduction in both coleoptiles and roots. It should be cautioned that the data obtained so far do not demonstrate an essential role for pH in this pathway, as none of the cytosolic pH manipulations performed so far have led to a complete elimination of gravitropism. In conclusion, cytoplasmic pH changes may have a universal role in the early signaling phases of gravitropism. What might they be doing in this process? We currently have no definite answer to this important question, partly because we have only a rudimentary understanding of the molecular mechanisms that govern it. We also have a limited knowledge of the locale of these gravity-induced pH changes within individual statocytes. It has been proposed that gravity-induced pH changes in the statocytes might facilitate auxin transport (Fasano et al. 2002). Indeed, such pH changes may be related to the asymmetric pH responses that were observed at the surface of gravistimulated roots by protonselective microelectrodes. These asymmetric surface-pH changes originated at the root cap and progressed along the root tip at a rate comparable with polar auxin transport (Monshausen and Sievers 2002). It is possible that surface-pH changes and polar auxin transport are related, and that the gravity-induced pH changes in columella cells regulate the activity or cellular distribution of auxin transporters in the statocytes (Fasano et al. 2002). In agreement with this model, mutant and transgenic Arabidopsis plants with altered expression of a H+-pyrophosphatase (AVP1) display altered auxin transport along with altered expression and mislocalization of the PIN1 auxin efflux facilitator (Li et al. 2005). 2.2.4 Proteins Implicated in Gravity Signal Transduction Recent genetic analyses of gravitropism have contributed to the identification of several gravity signal transducers. Although most gravitropism mutations turned out to affect as-

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pects of polar auxin transport or auxin response, a few have been obtained that affect earlier phases of gravity signal transduction that occur in the statocytes. For instance, mutations that affect starch biosynthesis, such as pgm, have been shown to affect gravitropism. As discussed in Chapter 1, this is not surprising since starch is a dense material that increases the weight of amyloplasts, enabling their sedimentation in the favorable environment presented by the statocytes’ cytoplasm (Kiss et al. 1989). Other mutations that affect gravitropism without altering phototropism, starch synthesis, amyloplast sedimentation, or growth responses to auxin, other phytohormones, or polar auxin transport inhibitors have also been identified. This class of mutants likely affects genes involved specifically in gravity signal transduction. The first such mutants isolated in Arabidopsis thaliana affected the ARG1 locus. The rhg/arg1 mutants displayed altered root and hypocotyl gravitropism while maintaining wild-type root-growth responses to phytohormones and polar auxin transport inhibitors, as well as normal phototropism (Fukaki et al. 1997; Sedbrook et al. 1999). The latter observation was particularly revealing because it indicated that the mutant organs retained their ability to curve in response to other directional cues. Hence, the defect was likely to lie in the early phases of gravity signal transduction. The ARG1 gene encodes a J-domain protein that is conserved between plants and the worm Caenorhabditis elegans, but is absent in yeast or other animals (Sedbrook et al. 1999). This protein was found to contain a J-domain at its N-terminus, a central hydrophobic region and a C-terminal domain predicted to form a coiled coil structure (typically involved in protein–protein interactions). The C-terminal region shares sequence similarity with proteins that interact with the cytoskeleton, although strong evidence for cytoskeleton interaction is currently lacking (Sedbrook et al. 1999; Boonsirichai et al. 2003). In other, better-characterized, J-domain proteins, the highly conserved J-domain directly interacts with the HSP70 chaperone, modulating its ATPase activity. The residues needed for this interaction are conserved in ARG1, suggesting that this protein might also function in association with HSP70 in the folding, trafficking, localization, and/or regulation of gravity signal transducers in the statocyte (Sedbrook et al. 1999). A combination of biochemical fractionation and functional GFP-fusion localization studies demonstrated that ARG1 is a peripheral membrane protein that associates with multiple components of the vesicle trafficking pathway in all plant cells. Targeting its expression to the root or hypocotyl statocytes of an arg1-2 null-mutant rescued the gravitropic phenotype of the corresponding organ (root or hypocotyls, respectively), demonstrating ARG1’s role in early phases of gravity signal transduction. A more thorough phenotypic analysis of arg1-2 demonstrated an inability for mutant root statocytes to respond to gravistimulation by cytoplasmic alkalinization and by relocalization of the PIN3 auxin efflux facilitator to their lower membrane, confirming a role for ARG1 in gravity signal transduction (Boonsirichai et al. 2003; Harrison and Masson 2006). Supporting this conclusion, the auxin-responsive DR5-GUS construct demonstrated an inability for mutant roots to develop a lateral auxin gradient across their cap upon gravistimulation (Boonsirichai et al. 2003). Taken together, these data suggest that ARG1 modulates the trafficking and/or activity of auxin efflux facilitators and/or other membrane-associated proteins needed for lateral auxin transport in the statocytes.

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Although the Arabidopsis thaliana genome contains more than 90 J-domain genes (Miernyk 2001), only two encode proteins that are similar to ARG1 throughout their lengths: ARG1-Like 1 (ARL1) and ARL2. A reverse genetic approach was used to demonstrate that ARL2 also contributes to early phases of gravity signal transduction, whereas ARL1 does not. In fact, physiological and molecular studies of arl2 mutant seedlings also showed defects in lateral auxin transport and PIN3 protein relocalization within the root statocytes upon gravistimulation (Guan et al. 2003; Harrison and Masson 2006). Hence, ARG1 and ARL2 appear to function in the same pathway. In agreement with this conclusion, arg1-2 arl2-1 double mutants show intermediate gravitropic defects that are similar to those of the corresponding single mutants (Guan et al. 2003). Analysis of double mutants between arg1-2, arl2-1, and pgm-1 also led to surprising results. Remember that pgm-1 affects phosphoglucomutase, an enzyme that contributes to starch biosynthesis. As discussed above, both pgm-1 and arg1-2 or arl2-1 display altered kinetics of gravitropism. However, their gravitropic defects are not complete and their organs still develop reasonably strong gravitropic responses, though with slower kinetics. If PGM and ARG1/ARL2 contribute to a linear gravity signal transduction pathway in the statocytes, double mutants should display a phenotype similar to that of single mutants. When arg1-2 pgm-1 and arl2-1 pgm-1 double mutants were analyzed, a surprisingly strong enhancement of the gravitropic defect was observed relative to that of single mutants (Boonsirichai et al., unpublished data; Guan et al. 2003). This result can be explained in several ways. First, it is possible that ARG1 and ARL2 function in a pathway that is distinct from the PGM pathway. In fact, as discussed in Chapter 1, several experiments have suggested the existence of more than one mechanism of gravity sensing in roots. Second, it is possible that these mutations affect partially their respective steps in a linear pathway. Indeed, ARG1 has been postulated to function as part of a chaperone complex that might modulate, but not be required for, the targeting or activity of membrane-associated proteins in the statocytes, whereas pgm-1 mutants are not completely defective in amyloplast sedimentation (Saether and Iversen 1991). In addition to revealing the possible topology of the gravisensing network, these double-mutant studies suggest alternative genetic strategies to search for novel gravity signal transducers. For instance, a screen for genetic enhancers of arg1-2/arl2-3 may lead to the identification of new gravity signal transducers in the “PGM genetic pathway,” whereas searching for enhancers of pgm will likely lead to the discovery of genes that function in the “ARG1/ARL2 genetic pathway.” The recent isolation and initial characterization of mutations falling in the first group (genetic enhancers of arg1-2) allowed the identification of an outer plastid membrane-associated protein as a possible gravity signal transducer, boding well for the success of this approach (Stanga et al. 2006). Genetic studies have also been quite effective at uncovering gravity signal transducers in aboveground organs through careful investigations of Arabidopsis mutants with defects in shoot gravitropism. As discussed in Chapter 1, a number of shoot gravitropism (sgr) mutants have been identified in Arabidopsis which exhibit reduced gravitropic responses in inflorescence stems (Yamauchi et al. 1997). Several of these mutants have been very useful for our understanding of gravity perception in shoots as they confirmed endodermal cells as shoot statocytes, re-emphasized the contribution of amyloplast sedi-

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mentation in gravity susception, and uncovered an important role for vacuolar biogenesis and function in gravity perception or signal transduction in shoots (Saito et al. 2005). In addition, other sgr mutants also identified new gravity signal transducers in Arabidopsis shoots. sgr5 and sgr6 are among the shoot gravitropism mutants that are most likely to be defective in early phases of gravity signal transduction within the shoot statocytes. Indeed, amyloplasts in mutant shoot endodermal cells sediment almost like wild type, indicating that gravity susception is not affected (Morita et al. 2006; Yano et al., unpublished results). SGR5 is a zinc-finger protein that is localized in the nucleus and is mainly expressed in the endodermis. In addition, endodermis-specific expression of wild-type SGR5 in the sgr5-1 mutant restored shoot gravitropism to wild-type levels. Hence, SGR5 is probably a transcription factor that contributes to early events of gravity perception and/or signaling in the statocytes (Morita et al. 2006). Further analyses, such as exploration of downstream target genes of SGR5, should clarify its function in shoot gravitropism. A similar analysis of sgr6 mutants suggested that the SGR6 protein is also involved in an early step of gravity signal transduction in stem statocytes, subsequent to amyloplast sedimentation. Unfortunately, the molecular function of SGR6 remains unknown, considering that its amino acid sequence is conserved with predicted orthologous proteins of unknown function in higher eukaryotes (Yano et al., unpublished results). An alternative screening approach was also developed to isolate additional shoot gravitropism mutants with defects in gravity perception or early phases of signal transduction (Wyatt et al. 2002). Arabidopsis inflorescence stems show no response to gravistimulation at 4°C. However, stems that are gravistimulated by horizontal placement at 4°C can execute a bending response to the cold gravistimulus if returned to the vertical position at room temperature within the next hour (Fukaki et al. 1996). It has been demonstrated that basipetal auxin transport is abolished in the inflorescence stems of wild-type plants at 4°C (Nadella et al. 2006). Taking advantage of this unusual behavior, Wyatt and collaborators (2002) isolated several gravity persistence signal (gps) mutants that exhibit normal shoot gravitropism at room temperature but display abnormal bending responses to stimuli provided in the cold. gps1 does not bend, gps2 bends in the wrong direction, and gps3 over-responds when returned to room temperature after cold gravistimulation (Wyatt et al. 2002). Amyloplasts sediment in the direction of gravity in all gps mutants during cold gravistimulation, indicating that gravity susception is not affected. To investigate a possible effect of the gps mutations on the ability of inflorescence stems to “remember” a cold stimulus by developing a lateral auxin gradient upon return to vertical position at room temperature, Wyatt and her collaborators studied expression of the auxin-dependent pIAA2::GUS gene in cold gravistimulated wild-type and gps inflorescence stems. As expected, wild-type stems showed asymmetrical activation of GUS expression on the lower side of a section of its stem elongation zone. On the other hand, gps mutant plants displayed patterns of GUS expression that were consistent with the characteristics of their respective bending defects: gsp1 stems displayed no evidence of asymmetrical activation of pIAA2::GUS expression after cold pretreatment, whereas gps2 showed enhanced GUS expression on the upper side of cold-gravistimulated stems. gps3

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displayed increased GUS expression on the lower side in an extended region along the elongation zone of its stems (Nadella et al. 2006). These results suggest that the gps mutants fail to properly establish a lateral auxin gradient across their inflorescence stems after cold gravistimulation, supporting a role for the corresponding genes in early phases of gravity signal transduction. Initial results in the molecular genetic analysis of three GPS genes appear to support their involvement in gravity signal transduction (Sarah Wyatt, personal communication). GPS1 encodes a cytochrome P450 of unknown function. Although GPS1 is not functional in the roots, a root-specific family member has been identified that is up-regulated in response to gravistimulation. Initial experiments indicate that these P450s may be involved in synthesis of flavonoids and, thus, the regulation of auxin transport through that pathway (Buer and Muday 2004; Withers and Wyatt, unpublished data). GPS2 encodes a hypothetical synaptobrevin/vesicle-associated membrane protein, v-SNARE (McCallister and Wyatt, unpublished data). GPS2 protein may be involved in transport of the PIN efflux carriers or other regulatory molecules in the inflorescence stem (see Chapter 5). Finally, GPS3 encodes a transcription factor with a B3 DNA-binding domain similar to auxin response factors (ARFs). However, GPS3 protein lacks the C-terminal dimerization domain common among ARF proteins (see Chapter 5). Initial subcellular localizations of GPS3 using a GFP fusion support nuclear localization for the protein, but its role in gravitropic signal transduction is as yet unknown (Nadella and Wyatt, unpublished data). Hence, these GPS genes hold great promise to further our understanding of the molecular mechanisms that govern gravity signal transduction in shoots. 2.2.5 Global ‘-omic’ Approaches to the Study of Root Gravitropism Although the genetic approach has been successful at identifying new gravity signal transducers, it also has limitations due to functional redundancy associated with frequent gene duplications in plants and from pleiotropy, both of which mask function in gravitropism. Trying to bypass such difficulties, several groups have recently used techniques derived from genomics and proteomics to identify genes or proteins whose expression varies early in response to gravistimulation. Their hope is that some of these candidates will contribute to gravitropism. Knowledge of the Arabidopsis thaliana genome sequence (Arabidopsis-GenomeInitiative 2000) allowed the design of partial- and whole-genome microarray chips that can be used for global analyses of gene expression in response to environmental stimuli. A first attempt at identifying genes whose expression varies early in response to gravistimulation was reported in 2002, when scientists at the University of California–Berkeley used expression profiling with an Arabidopsis thaliana 8,300-gene microarray to demonstrate that 1.7% of the genes analyzed are differentially expressed in entire seedlings within 30 min of a gravistimulus. Thirty-nine percent of these differentially expressed genes were also regulated by gentle mechanical perturbation, unveiling the extreme sensitivity of plants to mechano-stimulation (see Chapter 5). Most of the gravity-regulated genes fell into only a few functional categories, including Oxidative stress/Plant defense (22.7%); Metabolism (14.9%); Transcription (8.5%); Cell wall/Plasma membrane (7.1%), and Signal transduction (6.4%). Many of the gravity up-regulated genes con-

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tained short, conserved sequences in their promoters, suggesting a potential role for these DNA motifs as cis-elements in the regulation of gene expression by gravistimulation (Moseyko et al. 2002). Although this initial study provided exciting new information on transcriptional responses to gravistimulation in Arabidopsis seedlings, it suffered from the fact that entire seedlings were analyzed, including organs that respond in opposite ways to gravistimulation (hypocotyls and roots), and from probing only a fraction of the Arabidopsis genome. A second study used similar strategies to monitor transient changes in gene expression in primary root tips of Arabidopsis thaliana seedlings in a time course during the first hour of gravity- and/or mechano-stimulation. The whole-genome Affymetrix ATH1 microarray was used in this analysis (Kimbrough et al. 2004). This study helped uncover clusters of genes that show similar kinetics of expression change in the root tip upon gravistimulation. A vast majority of the differentially expressed genes (1,665 genes, or 96% of the regulated genes) were regulated by both gravity- and mechano-stimulation. Only 65 differentially expressed genes showed specific up-regulation in response to gravistimulation. Five were up-regulated by a factor of three or more within 2 min of gravistimulation, and remained high during the first 30 min of the response (Kimbrough et al. 2004). These fast graviresponding genes did not change their expression in response to gravistimulation in arg1-2 and arl2-3 mutant backgrounds, confirming the key role played by ARG1 and ARL2 in early phases of gravity signal transduction in Arabidopsis root tips, upstream of the transcriptional responses (Yester et al. 2006). Most of the genes found to be regulated by gravity- or mechano-stimulation again fell into only a few functional classes: Transcription (258 genes); Metabolism (144 genes); Protein fate (114 genes); and Signal transduction (97 genes). Only a minority of the differentially expressed genes fell in the Defense (31) and Stress (14) functional categories, which were the most highly represented classes in Moseyko et al. (2002). Furthermore, only three genes were found to be regulated by gravity and/or touch in both studies (Moseyko et al. 2002; Kimbrough et al. 2004). The difference in results between these two studies probably reflects differences in experimental procedures, as discussed in Kimbrough et al. (2004). Now that multiple genes have been identified whose expression varies in response to gravistimulation, they can be tested for a contribution to gravity signal transduction by reverse genetics (Kimbrough et al. 2004). Although global expression profiling is useful for identifying clusters of genes with similar expression profiles under defined conditions, it cannot uncover post-transcriptional regulatory processes. Hence, attempts have been made at identifying proteins whose abundance, localization, and/or post-translational modifications are altered by gravistimulation. That gravistimulation promotes changes in protein abundance or post-translational modification was first demonstrated in pioneering studies comparing protein profiles between vertical and gravistimulated Arabidopsis seedlings grown in the presence of 35Slabeled methionine. Gravistimulation was provided by rotating the dishes in which seedlings were growing 90 degrees and allowing the organs to reorient over a period of 24 hours. Subsequently, proteins were extracted from vertical control and gravistimulated

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samples, and separated by two-dimensional gel electrophoresis (2D-GE). This form of electrophoresis separates proteins based on their isoelectric point in the first dimension, and on their molecular weight in the second dimension. After electrophoresis the gels were autoradiographed, leading to protein-spot profiles for the extracts under investigation. When protein profiles from control and gravistimulated seedlings were compared, 2 out of approximately 600 detectable spots increased in intensity in the gravistimulated samples relative to control. A similar experiment testing the effect of continuous stimulation by rocking the dishes over a period of 24 hours led to the identification of 10 gravity up-regulated and 4 gravistimulation-specific protein spots (Sakamoto et al. 1993). To detect potential differences in protein phosphorylation upon stimulation, control and continuously rocked samples were labeled with 32P-orthophosphate during the stimulus. Proteins that were phosphorylated during the period of treatment should appear as radioactive spots on the 2D-GE gels. By comparing radioactive protein spots between control and continuously rocked samples, Sakamoto et al. (1993) were able to demonstrate that continuous rocking enhances the phosphorylation of two protein spots. Hence, an important conclusion of these studies is that gravity- and/or mechano-stimulation promote changes in 2D-GE protein spot intensity, reflective of changes in protein abundance and/or post-translational modification, and differential phosphorylation of specific proteins in Arabidopsis thaliana seedlings. Another attempt at establishing a role for protein phosphorylation in gravity signal transduction sought phosphoproteins with differential levels of expression between upper and lower flanks of gravistimulated oat pulvini (Chang and Kaufman 2000; Chang et al. 2003). These investigations uncovered two soluble and two membrane-associated proteins that are differentially phosphorylated in lower versus upper pulvinus halves in response to gravistimulation. Subsequent work defined more thoroughly the gravityinduced phosphorylation of one of the soluble oat proteins, demonstrating that it occurs as early as 5 min after initiation of gravistimulation and requires a newly synthesized protein. This time of initial phosphorylation correlates well with the minimal gravistimulation time needed to activate a productive transduction pathway leading to curvature response (presentation time), which is 5.2 min in oat pulvini. The differentially phosphorylated 50kD oat protein is itself a kinase, as demonstrated in autophosphorylation experiments. Altogether, these data indicate that the differential phosphorylation of this 50kD protein in graviresponding oat pulvini might contribute to gravity signal transduction in this system (Chang et al. 2003). In these early proteomic experiments, no attempts were made to identify the proteins present in the differentially represented 2D-GE protein spots (Sakamoto et al. 1993; Chang et al. 2003). However, the last decade witnessed amazing developments in mass spectrometry that truly revolutionized our ability to identify proteins based on their mass, on the mass of their proteolytic products, and on their amino acid content (Li and Assmann 2000). Taking advantage of this technological revolution, researchers are now able to identify differentially represented proteins as long as they are working with an organism whose genome has been completely sequenced. It is not surprising that recent proteomic studies have identified a number of Arabidopsis proteins whose abundance or modification varies in response to gravistimulation. First, Kamada et al. (2005) identified proteins associated with the cytoskeleton and the

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Ca2+ signal transduction pathway as being differentially represented in the root tip of Arabidopsis thaliana seedlings upon 0.5 and 3 hours of gravistimulation. Three other proteins were found to change transiently their molecular weight, but not their pI, upon gravistimulation, suggesting post-translational modification (Kamada et al. 2005). The stimulation times used in this study allowed significant gravitropic curvature, implying that the identified proteins have the potential of functioning in any phase of gravitropism, from gravity perception or signal transduction to the curvature response. In an attempt to focus on early phases of gravity perception and signal transduction, another study analyzed the protein profiles of 12-min gravistimulated Arabidopsis roottip samples (Murthy, Young, Sabat, and Masson, in preparation). This time point was chosen because it is sufficient to promote productive gravity signal transduction (as determined by the ability of 12-min gravistimulated root tips to develop tip curvatures after subsequent clinorotation; see Chapter 1), but insufficient for curvature initiation. Control and 12-min gravistimulated root tips were dissected, and proteins were extracted using a three-step fractionation procedure. Protein fractions were subjected to 2DGE, followed by silver-staining of the corresponding gels. The protein profiles of control and gravistimulated samples were compared. Fifty-seven protein spots were uncovered whose staining intensity was altered after 12 min of gravistimulation relative to unstimulated controls, and the corresponding proteins were identified by mass spectrometry. An additional control was included in this experiment, in which Arabidopsis seedlings were gently rotated to the horizontal, then immediately returned to the vertical for an additional 12 min, as a way to control for the mechano-stimulus that accompanies gravistimulation. Only 4 of the 57 graviresponding proteins also showed differential regulation in response to the mechano-stimulus control. Hence, a vast majority of these proteins responded specifically to gravistimulation. Most of the proteins identified in the latter study fell into the following functional categories: Unknown function (24%); Metabolism (17%); Stress and detoxification (13%); Defense (10%); and Energy (10%) (Murthy et al., in preparation). It is striking that only one of these differentially represented proteins is encoded by a gene also found to be transcriptionally regulated by gravistimulation (Kimbrough et al. 2004). This difference between root-tip transcriptional- and proteomic-response profiles may again reflect differences in the experimental procedures. Indeed, transcriptional profiling was carried out on dark-grown seedlings, whereas analysis by Murthy et al. involved light-grown material (Kimbrough et al. 2004; Murthy et al. 2007). On the other hand, 6 of the 16 proteins identified by Sakamoto et al. (2005), or their paralogs, were also identified as differentially represented by Murthy and collaborators, indicating some consistency between independent proteomic studies. Among the 53 gravity-responding root-tip proteins identified by Murthy and collaborators, 3 function in the S-adenosylmethionine (AdoMet) methyl-donor pathway. This pathway generates precursors for ethylene and polyamine synthesis, and provides methyl groups for transmethylation reactions that target a number of plant regulatory molecules such as auxin, cytokinin, jasmonate, salicylate, etc. This result suggested an involvement of the AdoMet cycle in gravity signal transduction (Young et al. 2006). A reverse genetic approach was used to investigate this possibility. One of two Arabidopsis genes encoding adenosine kinase (ADK1) was shown to contribute to root

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gravitropism. A mutation in this gene resulted in plants displaying altered kinetics of root graviresponse and longer presentation time relative to wild type. Root cap morphology was also altered, probably as a consequence of altered auxin accumulation in the root cap. Upon gravistimulation, adk1-1 mutant seedlings failed to relocalize the auxin efflux facilitator PIN3 to the lower membrane of their root statocytes, confirming a role for this gene in early phases of gravity signal transduction (Young et al. 2006). Surprisingly, adk1-1 roots displayed wild-type cytoplasmic alkalinization of their statocytes in response to gravistimulation. Furthermore, when expression of three of the five fast gravity-responsive genes described above was analyzed, one (At4g23670) still increased in expression in adk1-1 root tips upon gravistimulation as it did in wild type, whereas the two other genes (At5g38020 and At5g48010) did not respond to gravistimulation in adk1-1 (Yester et al. 2006). Hence, adk1-1 affected differently the expression response to gravistimulation of three fast gravity-responsive genes, even though arg1-2 and arl2-1 obliterated completely the response of all three, as discussed above (Yester et al. 2006). It is interesting to note here that At5g38020, whose expression response to gravistimulation is obliterated by adk1-1, encodes an enzyme whose predicted biochemical function (AdoMet-dependent methyltransferase activity) is directly associated with the AdoMet cycle (Kimbrough et al. 2004; Schoor and Moffatt 2004; Yester et al. 2006). Together, these fascinating results can be interpreted in several ways. For instance, gravity signal transduction could involve a single linear pathway in which ADK1 functions downstream of ARG1 to mediate gravity-induced PIN3 relocalization and differential expression of At5g38020 and At5g48010 in the statocytes, with gravity-induced cytoplasmic alkalinization and differential expression of At4g23670 requiring only the presence of functional ARG1 (Figure 2.2A). Alternatively, it is possible that the gravity signal transduction pathway is bifurcated, with the ADK1-dependent branch of the pathway leading to PIN3 relocalization and up-regulation of At5g38020 and At5g48010 whereas the other branch would lead to cytoplasmic alkalinization and activation of At4g23670. In this case, ARG1 would function upstream of ADK1, before the point of pathway bifurcation (Figure 2.2B). Further genetic analysis of double and multiple mutants should help resolve this ambiguity. The data discussed above support a role for ADK1 in early phases of gravity signal transduction in roots. However, the molecular mechanisms underlying its contribution remain uncharacterized. Does adenosine, the main substrate of ADK that feedback inhibits the AdoMet cycle (Schoor and Moffatt 2004), function in cellular signaling like it does in animal systems (Nishizaki 2004)? Or do other AdoMet cycle-derived regulatory compounds modulate gravity signal transduction? A systematic study on the contribution of distinct biochemical branches derived from the AdoMet pathway in different phases of the root gravitropic response will undoubtedly yield new insights into these long-ranging questions. Preliminary results suggest a role for spermine, an AdoMet-derived polyamine, in the signal-transmission or curvature-response phases of root gravitropism (Young et al. 2006). In conclusion, the genes and proteins identified through the genomic and proteomic screens described above constitute good candidates as gravity signal transducers. They belong to only a few predicted functional categories, thereby identifying specific biochemical pathways (such as the AdoMet cycle) as potential contributors to the process.

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Figure 2.2. Linear (A) and bifurcated (B) models of gravity signal transduction in the root statocytes.

They also constitute an outstanding list of markers that can be used to better define the pathways involved in gravity signal transduction, as illustrated by the expression analysis of fast gravity-responsive genes in adk1-1, arg1-2, and arl2-1 mutant root tips. It is likely that a combination of reverse genetics and marker-gene-expression analysis will yield important insights into the molecular mechanisms that govern gravity signal transduction in plants. 2.2.6 Relocalization of Auxin Transport Facilitators or Activity Regulation? As reviewed in Chapter 3, activation of the gravity signal transduction pathway in roots results in a relocalization of the PIN3 auxin efflux facilitator to the bottom membrane of the statocytes (Friml et al. 2002). PIN3, a member of the PIN family of transmembrane auxin efflux facilitators, is expressed in the statocytes of both roots (columella cells) and shoots (endodermal cells). It localizes to the plasma membrane and to vesicles that cycle between plasma membrane and endosome. Mutations in the PIN3 gene result in altered root and hypocotyl gravitropism, supporting a role for this protein in gravity signal transduction (Friml et al. 2002). In statocytes of vertical roots, PIN3 is positioned symmetrically at the plasma membrane. It rapidly relocalizes laterally, to the bottom membrane, upon gravity stimulation (Friml et al. 2002). PIN3 relocalization initiates within 2 min of gravistimulation, therefore preceding establishment of a lateral auxin gradient across the root tip. As discussed above, PIN3 relocalization requires the presence of fully functional ARG1, ARL2, and ADK1 (Harrison and Masson, unpublished data; Young et al. 2006), suggesting that it functions downstream of these proteins in the gravity signal transduction pathway in roots. Because at least ARG1 (and probably ARL2) has been associated with the vesicular trafficking pathway (see above), it appears reasonable to postulate that gravity signal transduction modulates the vesicular trafficking of PIN3 and probably of other membrane-

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associated molecules within the root statocytes (Boonsirichai et al. 2003). It is interesting to note that several of the molecules suggested to function as second messengers in gravity signal transduction, including Ca2+, proton flux, and phosphoinositides, have been shown to function in the regulation of vesicular trafficking in other cell types, thus arguing for a direct connection between gravity signal transduction and vesicular trafficking (reviewed in Blancaflor and Masson 2003). However, we have not yet eliminated the possibility that gravity-induced PIN3 relocalization in the root statocytes is a consequence of transporter activity regulation right at the plasma membrane. Indeed, changes in auxin level have been shown to result in changes in the cycling and polar localization of PIN proteins in other cell types (Blilou et al. 2005; Paciorek et al. 2005). More work is needed to resolve this ambiguity. The situation is even more complex if one considers potential effects of gravity signal transduction on the polarity of vesicular trafficking and on the localization of auxin transporters in shoot statocytes. Indeed, PIN3 is also an ideal candidate for molecular linkage between gravity perception and asymmetric auxin distribution in shoots. Here, PIN3 is localized to the plasma membrane of the inner longitudinal and bottom sides of endodermal cells in vertically oriented organs (Friml et al. 2002), and no data currently exist to support or contradict a possible intracellular relocalization of PIN3 upon gravistimulation. Careful immunolocalization or GFP-PIN fusion expression studies are needed to determine whether such a relocalization also occurs in shoot statocytes, and to establish whether it involves other auxin transporters. It is essential to establish whether gravistimulation regulates the activity of auxin transporters on the outer side of lower flank statocytes, or directly regulates the vesicular trafficking of auxin transporters. As reviewed in Chapter 3, the transduction pathway could also lead to differential phosphorylation of auxin transport facilitators, or it could regulate the levels of small-molecule effectors of auxin transporters, thereby contributing to the regulation of lateral auxin transport and lateral gradient formation in the absence of transporter relocalization. These possibilities will have to be investigated carefully in order to gain a better understanding of the molecular mechanisms that govern gravity signal transduction in both roots and shoots. 2.2.7 Could Cytokinin Also Contribute to the Gravitropic Signal? Although lateral gradients of auxin have long been considered the biochemical signals that inform the tissues involved in the differential cell elongation of gravitropic signals perceived by the statocytes, cytokinin was recently proposed as an alternative signal mediating early phases of gravitropic curvature in roots (Aloni et al. 2004). Careful kinematical studies of early phases of root gravitropic curvature have long puzzled researchers (Ishikawa and Evans 1993; Baluˇska et al. 1996; Wolverton et al. 2002). These investigations have revealed a complexity of the early root-growth response to gravistimulation that could not be explained by the simple model of auxin redistribution described above. Instead, it revealed a first step where overall growth was inhibited at both the upper and lower sides of gravistimulated roots, followed by a second, auxin-gradient-independent phase where the gravitropic curvature is initiated by an enhancement in the rate of cellular elongation on the upper flank of the distal elongation zone (reviewed in Wolverton et al. 2002).

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Recent studies provide potential auxin-related explanations for the surprising characteristics of the initial phases of root gravicurvature, including early gravity-induced increases in the levels of flavonoids at the root tip, which inhibit general auxin transport (Buer and Muday 2004); and an indirect effect of decreased auxin levels on the vesicular trafficking and degradation of the AGR1/EIR1/PIN2/WAV6 auxin efflux facilitator at the top flank (Abas et al. 2006). However, independent observations suggest that cytokinin might also contribute to at least some aspects of root gravicurvature. Using a cytokinin-sensitive ARR5-GUS reporter construct to indirectly follow changes in cytokinin levels within the root tip, investigators were able to demonstrate that gravistimulation promotes a fast asymmetrical increase in reporter expression within the bottom lateral cap flank, potentially reflecting differential increases in cytokinin levels at that flank (Aloni et al. 2004). The same authors also showed that application of exogenous cytokinin to one side of the elongation zone of vertically oriented roots results in curvature development in the direction of cytokinin application. Hence, they suggested that gravistimulation might promote a lateral transport of cytokinin across the cap, with accumulation on the lower flank and consequent differential cellular elongation resulting in initial curvature response (Aloni et al. 2004). Although this model is attractive at first glance, it should be cautioned that genetic support for it is currently lacking. The cytokinin-deficient mutants analyzed so far have not shown dramatic changes in root gravitropism. However, it is true that none of these mutants showed complete obliteration of cytokinin sensitivity (Aloni et al. 2004). Furthermore, expression of one of the genes that contribute mainly to cytokinin biosynthesis in the Arabidopsis root cap, isopentenyl transferase 5 (IPT5), is auxin-sensitive (Miyawaki et al. 2004), suggesting the possibility that the asymmetrical activation of ARR5-GUS expression on the lower flank of gravistimulated root caps might simply be a consequence of lateral auxin transport, rather than reflecting an effect of gravity signal transduction on the lateral transport of cytokinins across the cap. The kinetics of differential ARR5-GUS expression across gravistimulated root tips appear to precede the asymmetrical activation of the auxin-sensitive DR5-GUS reporter (Aloni et al. 2004). However, these two genes are indirect reporters of cytokinin and auxin levels, respectively, functioning at the end of their respective pathways. Hence, data derived from comparative analyses of their expression kinetics upon gravitimulation should be interpreted with great caution. Future work will be needed to test asymmetrical activation of ARR5-GUS expression across the root tip in different mutants, such as arg1-2, arl2-1, adk1-1, or pgm, in order to test whether this response lies in one of the known gravity signal transduction pathways. Investigation of this response in a variety of mutants affected in diverse aspects of cytokinin synthesis and response is also crucial. For instance, an analysis of the simple, double, and multiple mutants carrying defects in the three cytokinin receptor genes found in Arabidopsis thaliana (Higuchi et al. 2004) should be carried out to evaluate their relative contributions to the initial phases of the root gravitropic curvature.

2.3 Gravity Signal Transduction in Organs that Do Not Grow Vertically The organs discussed in this chapter tend to grow vertically, either upward or downward. However, other plant organs may display distinct growth behavior. For instance, lateral

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organs will tend to grow at a defined angle from the vertical (called gravity set point angle or GSA) after emerging from the primary organ. This distinct growth behavior, termed plagiogravitropism, is pushed to its extreme in organs that grow horizontally (diagravitropism). For instance, stolons and rhizomes grow horizontally, exploring a plant’s neighborhood and colonizing it through vegetative reproduction at their nodes. Interestingly, GSA can also vary depending on the environmental status of a growing organ. As discussed in Chapters 5 and 6, touch or lateral humidity gradients can temporarily inhibit gravitropism, thereby allowing a root to change its growth pattern to reach better environments despite the influence of gravity (Massa and Gilroy 2003; Takahashi et al. 2003). Similarly, light can modulate gravitropism in addition to promoting distinct tropic responses called phototropism (Kiss et al. 2002; Kiss et al. 2003). The developmental stage of a plant organ will also influence its GSA. For instance, the peanut gynophore will switch from negative (upward) to positive (downward) gravitropism upon fertilization, driving the developing fruit into the sand where it has to be located in order complete its developmental and maturation program (Moctezuma and Feldman 1998). Hence, not only is it necessary for a plant organ to perceive gravity, it is also important for that organ to transduce the corresponding vectorial information into a defined growth pattern that will ultimately allow it to reach environments that are better suited for plant growth and development. How is this accomplished? Although new information hints at some of the molecular mechanisms that allow distinct directional stimuli (such as touch and humidity gradients) to affect gravitropism in roots (discussed in Chapters 5 and 6), very little is known about how a specific GSA is actually set for a defined organ based on its developmental program or surrounding environment. Yet, the tools used in the study of organ growth behavior in plants are becoming increasingly sophisticated, such that we can anticipate a future that will shed light on the regulatory mechanisms that tune gravity signal transduction to the characteristics of an organ’s endogenous and external environment, thereby modulating overall growth and morphogenesis.

2.4 Acknowledgments We thank John Stanga, Laura Vaughn, Jessica Will, Elison Blancaflor, and Simon Gilroy for critical comments on this manuscript. This work was supported by grants from NSF, NASA, UW College of Agriculture and Life Sciences Hatch funds, and UW Graduate School Grant-in-Aid to PHM. 2.5 Literature Cited Abas L, Benjamins R, Malenica N, Paciorek T, Wirniewska J, Moulinier-Anzola J, Sieberer T, Friml J, and Luschnig C. 2006. Intracellular trafficking and proteolysis of the Arabidopsis auxin-efflux facilitator PIN2 are involved in root gravitropism. Nat Cell Biol Advance Online Publication: 1–8. Aloni R, Langhans M, Aloni E, and Ullrich C. 2004. Role of cytokinin in the regulation of root gravitropism. Planta 220:177–82.

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Arabidopsis-Genome-Initiative. 2000. Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408:796–814. Baluˇska F, Hauskrecht M, Barlow P, and Sievers A. 1996. Gravitropism of the primary root of maize: A complex pattern of differential cellular growth in the cortex independently of the microtubular cytoskeleton. Planta 198:310–18. Barritt G, and Rychkov G. 2005. TRP as mechanosensitive channels. Nature Cell Biol 7:105–7. Blancaflor, EB, and Masson PH. 2003. Plant gravitropism. Unraveling the ups and downs of a complex process. Plant Physiol 133: 1677–90. Blilou I, Xu J, Wildwater M, Willemsen V, Paponov I, Friml J, Heidstra R, Aida M, Palme K, and Scheres B. 2005. The PIN auxin efflux facilitator network controls growth and patterning in Arabidopsis roots. Nature 433:39–44. Boonsirichai K, Sedbrook J, Chen R, Gilroy S, and Masson PH. 2003. ARG1 is a peripheral membrane protein that modulates gravity-induced cytoplasmic alkalinization and lateral auxin transport in plant statocytes. Plant Cell 15:2612–25. Braun M. 2002. Gravity perception requires statoliths settled on specific plasma membrane areas in characean rhizoids and protonemata. Protoplasma 219:150–9. Buer C, and Muday G. 2004. The transparent testa4 mutation prevents flavonoid synthesis and alters auxin transport and the response of Arabidopsis roots to gravity and light. Plant Cell 16:1191–1205. Chang S, Cho M, Kim S-K, Lee J, Kirakosyan A, and Kaufman P. 2003. Changes in phosphorylation of 50 and 53 kDa soluble proteins in graviresponding oat (Avena sativa) shoots. J Exp Bot 54:1013–22. Chang S, and Kaufman PB. 2000. Effects of staurosporine, okadaic acid and sodium fluoride on protein phosphorylation in graviresponding oat shoot pulvini. Plant Physiol Biochem 38:315–23. Fasano J, Massa G, and Gilroy S. 2002. Ionic signaling in plant responses to gravity and touch. J Plant Growth Reg 21:71–88. Fasano J, Swanson S, Blancaflor E, Dowd P, Kao T, and Gilroy S. 2001. Changes in root cap pH are required for the gravity response of the Arabidopsis root. Plant Cell 13:907–21. Firn R, and Digby J. 1980. The establishment of tropic curvature in plants. Ann Rev Plant Physiol 31:131–48. Friml J, Wisniewska J, Benkova E, Mendgen K, and Palme K. 2002. Lateral relocation of auxin efflux regulator PIN3 mediates tropism in Arabidopsis. Nature 415:806–9. Fukaki H, Fujisawa H, and Tasaka M. 1996. Gravitropic response of inflorescence stems in Arabidopsis thaliana. Plant Physiol 110:933–43. Fukaki H, Fujisawa H, and Tasaka M. 1997. The RHG gene is involved in root and hypocotyl gravitropism in Arabidopsis thaliana. Plant Cell Physiol 38:804–10. Fukaki H, Wysocka-Diller J, Kato T, Fujisawa H, Benfey P, and Tasaka M. 1998. Genetic evidence that the endodermis is essential for shoot gravitropism in Arabidopsis thaliana. Plant J 14:425–30. Guan C, Rosen E, Boonsirichai K, Poff K, and Masson P. 2003. The ARG1-LIKE2 (ARL2) gene of Arabidopsis thaliana functions in a gravity signal transduction pathway that is genetically distinct from the PGM pathway. Plant Physiol 133:100–12. Harrison B, and Masson P. 2006. Molecular functions of ARL2 and ARG1 in Arabidopsis gravitropism. In Proceedings of the 17th International Conference on Arabidopsis Research, Madison, WI, Abstract, 387. Haswell E, and Meyerowitz E. 2006. MscS-like proteins control plastid size and shape in Arabidopsis thaliana. Curr Biol 16:1–11. Higuchi M, Pischke M, Mahonen A, Miyawaki K, Hashimoto Y, Seki M, Kobayashi M, Shinozaki K, Kato T, Tabata S, Helariutta Y, Sussman M, and Kakimoto T. 2004. In planta functions of the Arabidopsis cytokinin receptor family. Proc Natl Acad Sci USA 101:8821–6.

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3

Auxin Transport and the Integration of Gravitropic Growth Gloria K. Muday* and Abidur Rahman

3.1 Introduction to Auxins Auxins have been implicated in controlling elongation, branching, and development of plant organs (as reviewed in Woodward and Bartel 2005), as well as the asymmetric growth known as tropisms, which is the focus of this chapter. Of the plant hormones, auxin best resembles the canonical concept of a messenger, being synthesized in one place and acting in another. Indole-3-acetic acid (IAA) is the most studied and abundant natural auxin, yet in some plants indole-3-butyric acid (IBA) is at almost equivalent levels (as reviewed in Ludwig-Muller et al. 1993). Auxins also include synthetic compounds such as 1-naphthaleneacetic acid (1-NAA) and 2,4-dichlorophenoxyacetic acid (2,4-D), which have been used in many studies because they are not susceptible to photolysis from blue and ultraviolet lights, like the native auxin IAA (Stasinopoulos and Hangarter 1990). The synthetic auxins have differences in activity (as reviewed in Woodward and Bartel 2005) and in transport properties (Delbarre et al. 1996), which make them useful for designing experiments to test specific aspects of auxin function. 3.2 Auxin Transport and Its Role in Plant Gravity Response Auxin moves through plants by a unique cell-to-cell polar transport mechanism, from the shoot meristem and young leaves (Ljung et al. 2002) toward the base of stems (as reviewed in Blakeslee et al. 2005). Figure 3.1 contains a diagram summarizing the movements of IAA in a seedling. Polar auxin transport results in an auxin gradient down the length of the stem or hypocotyl, with the highest auxin concentrations found in the regions of greatest elongation (Ortuno et al. 1990). Auxin is also synthesized in the root tip (Ljung et al. 2005), where auxin transport is more complex, with two distinct polarities. Shoot-derived IAA moves acropetally (toward the root apex) through the central cylinder, and basipetally (from the apex toward the base) through the outer layers of root cells (Tsurumi and Ohwaki 1978). Arabidopsis roots also have a tip-focused IAA gradient (Casimiro et al. 2001) and basipetal transport of radiolabeled auxin applied to the root tip moves only within the apical centimeter of the root tip (Rashotte et al. 2000; Geisler et al. 2005). It is this basipetal IAA transport movement that is specifically linked to root gravitropism (Rashotte et al. 2000). In addition to polar transport down the length of plant tissues, the Cholodny-Went hypothesis suggests that the lateral transport of auxin across gravity-stimulated plant tissues drives differential gravitropic growth, as indicated in Figure 3.1B (as reviewed in Evans 1991; Trewavas 1992; Muday 2001; see also Chapters 1 and 2). Such a lateral redistrib*Corresponding author 47

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Figure 3.1. Auxin transport is polar in Arabidopsis and other plants. A. In an upright hypocotyl, inflorescence, and other stem tissues, auxin moves in single direction, from the shoot apex toward the base (basipetal). In roots, movement from the shoot into the root is from the base toward the root apex (acropetal) through cells in the central cylinder. In roots, auxin also moves from the root tip toward the base in a basipetal direction through cells of the cortex and/or epidermis. B. In a plant reoriented 90 degrees relative to gravity, although auxin transport continues in a polar fashion, lateral auxin transport also occurs. In shoot tissues, this transport may occur across the hypocotyl, whereas in roots, redirection of auxin transport is believed to be controlled from the root cap. The regions in which gravitropic bending will occur are also indicated. This figure is reprinted from Muday (2001) with permission of Springer Publishing.

ution of auxin also appears to be responsible for the curvature response to lateral light stimulation (phototropism; see Chapter 4). Gravity perception in stems occurs in the starch sheath parenchyma tissues that run the length of the hypocotyl (Fukaki et al. 1998). Lateral auxin transport then is believed to occur in multiple cells along the hypocotyl, and the elevated levels of auxin on the lower flank of the hypocotyl stimulate cellular elongation and allow upward growth (Blancaflor and Masson 2003). In contrast, roots sense gravity very locally in the columella cells in the root cap (Blancaflor et al. 1998), and auxin is redistributed from the root tip to the lower side of the root after gravity stimulation, rather than being laterally transported across the root (as reviewed by Blancaflor and Masson 2003; see Chapters 1 and 2). As elevations in auxin concentration generally inhibit root growth, this redistribution would result in slower growth on the lower side relative to the upper side, resulting in downward root growth. Asymmetric redistribution of radiolabeled IAA has been measured in both shoots (Parker and Briggs 1990) and roots (Young et al. 1990), preceding differential gravitropic growth (Parker and Briggs 1990). Additionally, gradients in endogenous free IAA have been observed across gravity-stimulated oat and maize pulvini and maize coleoptiles (Kaufman et al. 1995; Philippar et al. 1999; Long et al. 2002). Growth of seedlings on IAA efflux inhibitors (as reviewed in Rubery 1990) leads to a rapid inhibition of the gravity response in a number of plant species under conditions where growth still occurs

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(Katekar and Geissler 1980; Muday and Haworth 1994; Rashotte et al. 2000). Recently, synthetic and naturally occurring inhibitors of auxin influx have been identified and these compounds also inhibit gravitropic bending (Rahman et al. 2001a; Parry et al. 2001a). Although the validity of the Cholodny-Went hypothesis has been debated (Trewavas 1992), molecular and genetic evidence has provided significant support to this hypothesis (as reviewed in Blancaflor and Masson 2003). As discussed in Chapters 1 and 2, one powerful test of this hypothesis has been the examination of auxin-induced gene expression across gravity-stimulated plants. Transgenic plants carrying several different auxinresponsive promoters driving the expression of the gene encoding ß-glucuronidase (GUS) or green fluorescent protein (GFP) have now been used to show asymmetric auxin-induced gene expression across gravity-stimulated shoots (McClure and Guilfoyle 1989; Li et al. 1991; Li et al. 1999) or roots (Larkin et al. 1996; Luschnig et al. 1998; Rashotte et al. 2001; Ottenschläger et al. 2003). Although GUS and GFP reporters indirectly measure changes in auxin accumulation, the ability to easily observe the expression with high spatial resolution makes this a powerful approach to explore the role of auxin in tropisms. The asymmetric expression of the DR5::GUS reporter in Arabidopsis roots that are vertical and roots reoriented 90 degrees relative to gravity is shown in Figure 3.2 (also see Color Section. The inhibition of gravitropic bending and differential auxin-regulated gene expression by IAA efflux inhibitors, as shown in Figure 3.2C (Li et al. 1991; Rashotte et al. 2001; Paciorek et al. 2005), indicates that lateral auxin transport is required for differential gene expression. It is likely that this asymmetric gene expression also requires a change in auxin sensitivity (Salisbury et al. 1988), perhaps through activation of transcription factors necessary for auxin-induced gene expression (as reviewed in Leyser 2006 and discussed below). A critical question about these expression studies is whether they support the idea that gradients in auxin precede differential growth in response to gravity. In roots, particularly of Arabidopsis, the spatial and temporal character of the response to gravity stimulation is well characterized. Root gravitropic bending is narrowly constrained to the distal elongation zone (DEZ) (as reviewed in Evans 1991; Wolverton et al. 2002), which in a gravity-stimulated Arabidopsis root is localized on the lower side to a region between 100 and 300 microns from the tip (Mullen et al. 1998). The DR5::GUS reporter detects asymmetric auxin-induced gene expression in the same region as the differential growth in the DEZ (Rashotte et al. 2001), as shown in Figure 3.2B. This epidermal localization is consistent with previous evidence suggesting that the gravity signal in roots moves through cells of the epidermis and/or cortex (Yang et al. 1990; Björkman and Cleland 1991). Yet, gradients in DR5::GUS and GH3::GUS expression are detected after roots have begun to bend (Larkin et al. 1996; Rashotte et al. 2001). In DR5rev::GFP transgenic similar asymmetric gene expression is visible by 2 to 3 hours after gravity stimulation, and even earlier asymmetries in auxin-induced gene expression in the root cap are detectable at ~1.5 hours after gravity stimulation (Ottenschläger et al. 2003; Paciorek et al. 2005), which temporally precedes the asymmetry in the epidermal/cortical cells that is shown in Figure 3.2. This timing of this root cap asymmetry more closely parallels the initiation of gravitropic curvature.

Figure 3.2 (also see Color Section). Expression of auxin-responsive reporters is elevated on the lower side of gravity-stimulated wild-type roots consistent with redistribution of auxin at the root tip. A–C. GUS expression was visualized in 7-day-old seedlings homozygous for the DR5-GUS reporter construct (D) or for a DR5-GFP reporter. Root tips are shown for (A), a vertically grown seedling on control media, and (B and C) seedlings 6 hours after gravity stimulation on (B) control media, (C) 1 µM NPA, or (D) 3 hours after gravity stimulation on control media. The scale bar is equal to 100 microns. A model showing the mechanism of IAA redistribution and gradient formation at the root tip is shown in E and F. To simplify the model, both PIN proteins, which are part of the IAA efflux protein complex, and AUX1, an auxin influx protein, have been shown to be asymmetrically localized in the diagram, although the localization of AUX1 was reported to be axial. For gradients in IAA to form across a horizontal root, there are likely to be mechanisms to increase IAA transport to lower side of the root and decrease IAA transport to the upper side that act to change the activity and abundance of IAA efflux proteins. Panels A through C reprinted from Rashotte et al. (2001), with kind permission of the American Society of Plant Biologists.

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3.3 Approaches to Identify Proteins that Mediate IAA Efflux Biochemical and genetic approaches have identified a number of proteins associated with auxin transport. Auxin transport inhibitors have been used in both of these approaches. Auxin efflux inhibitors reduce polar IAA transport, efflux of auxin from membranes and cells, and inhibit physiological processes dependent upon auxin transport, such as gravitropic curvature (as reviewed in Rubery 1990). In particular, the inhibitor, naphthylphthalamic acid (NPA), has been used in biochemical experiments, since a tritiated form of this molecule can be used to follow the activity of one class of auxin transport proteins, termed NPA binding proteins (Muday et al. 1993). These proteins have been biochemically characterized using their NPA binding activity (Dixon et al. 1996; Butler et al. 1998; Hu et al. 2000, as reviewed in Muday 2000). Recently, NPA affinity chromatography has been used to identify a number of proteins that bind NPA (Murphy et al. 2002) with some proteins identified by both this method and genetic approaches (Noh et al. 2001), as described below. More recently, auxin influx inhibitors have also been identified (Imhoff et al. 2000; Rahman et al. 2001a; Parry et al. 2001a), but as yet these inhibitors have been used to characterize previously identified proteins, not to identify new proteins. Genetic approaches have been the most productive in identifying candidates for auxin transporters and associated regulatory proteins. The screens that have identified these proteins have included altered responses to auxin (Maher and Martindale 1980) or to auxin transport inhibitors (Ruegger et al. 1997), altered growth and developmental processes that are dependent upon auxin transport, including gravity response (Chen et al. 1998), and a number of developmental processes (Okada et al. 1991; Noh et al. 2001). Many of the mutant genes have been identified and the functions of the encoded proteins have been linked to IAA influx or efflux, as described below.

3.4 Proteins that Mediate IAA Efflux Experimental evidence suggests that IAA efflux carriers are made of multiple protein subunits, which are encoded by gene families with unique expression patterns that mediate the distinct polarities of IAA movement in different tissues (as reviewed by Blakeslee et al. 2005). The first gene family linked to IAA efflux encodes PIN proteins (as reviewed by Friml 2003). Arabidopsis mutants with defects in PIN1 were isolated due to their pinformed inflorescences and were shown to have reduced IAA transport in this tissue (Okada et al. 1991). PIN1 encodes a protein with 10 membrane-spanning domains, which is asymmetrically localized to the basal end of the membrane of inflorescence cells, consistent with a role in mediating basipetal IAA transport (Gälweiler et al. 1998). Several other screens isolated pin2/agr1/eir1/wav6 mutants based on altered root growth in response to gravity, waving, or ethylene and found that these mutant genes encoded a protein with extensive sequence similarity to PIN1 (Chen et al. 1998; Müller et al. 1998; Luschnig et al. 1998; Utsuno et al. 1998). These mutants have reduced basipetal IAA transport in roots (Chen et al. 1998; Rashotte et al. 2000) and the PIN2 protein is ex-

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pressed in roots and is localized asymmetrically in the plasma membrane, consistent with a role in mediating basipetal IAA transport (Müller et al. 1998). A recent report suggests that PIN function is redundant, using an eir1/pin2 mutant line transformed with a PIN1::GFP variant that has localization consistent with PIN2 but opposite to PIN1 (Wisniewska et al. 2006). This construct restored gravitropic bending and lateral IAA transport to eir1/pin2, whereas wild-type PIN1::GFP did not, indicating that the polar localization of PIN proteins is sufficient to direct IAA movement with appropriate polarity (Wisniewska et al. 2006). Finally, the overexpression of PIN7 in tobacco cell cultures significantly enhanced IAA, 1-NAA, and 2,4-D efflux, but not the tryptophan movement (Petrasek et al. 2006), conclusively demonstrating that PIN proteins can mediate IAA efflux. A second family of proteins has also been suggested to participate in IAA efflux by both genetic and biochemical approaches. The multidrug resistance/P-glycoprotein (MDR/PGP) gene family encodes proteins with sequence similarity to genes in the ATP Binding Cassette (ABC) transporter superfamily, which transport a variety of molecules in addition to the cytotoxic compounds for which they were initially isolated (Geisler and Murphy 2006). AtMDR1/PGP19, AtPGP1, and AtPGP2 proteins have been shown to bind to an NPA affinity column and expression of AtMDR1 in yeast increases NPA binding activity (Noh et al. 2001), consistent with the possibility that these proteins are the target for IAA efflux inhibitors. The mdr1/pgp19 mutant has reduced basipetal IAA transport in inflorescence (Noh et al. 2001) and roots (Geisler et al. 2005), and has phenotypes consistent with altered auxin transport (Noh et al. 2001; Lin and Wang 2005; Geisler et al. 2005), including enhanced gravitropic responses in both inflorescences (Noh et al. 2003) and roots (Lin and Wang 2005). The pgp1 mutant has a weak phenotype that enhances the mdr1/pgp19 mutant phenotype (Noh et al. 2001; Noh et al. 2003; Lin and Wang 2005), suggesting partially redundant functions for these two proteins. The pgp4/mdr4 mutant also has reduced root basipetal IAA transport and gravitropic response (Terasaka et al. 2005). The ability of PGP1 and PGP4 to mediate IAA and NAA efflux in heterologous systems (Geisler et al. 2005; Terasaka et al. 2005) further links these proteins to the process of IAA transport (Blakeslee et al. 2005). Surprisingly, current evidence supports a role of PGP4 in control of IAA influx, not efflux (Terasaka et al. 2005). Additionally, PGP19/ MDR1 has also been shown to mediate IAA efflux from plant tissue culture (Petrasek et al. 2006), although it has been difficult to demonstrate its function in heterologous systems (Geisler and Murphy 2006). PGP1, PGP4, and MDR1/PGP19 all have membrane localizations that are asymmetrically distributed across auxin transporting cells, suggesting that their localization could convey directional control of auxin transport (Geisler et al. 2005; Terasaka et al. 2005; Geisler and Murphy 2006). Additionally, AtPGP1 and 19 have been shown to participate in protein complexes with TWISTED DWARF1, a unique plasma membrane-anchored, immunophilin-like protein, which is required for maximal auxin transport and appropriate plant development (Geisler et al. 2003). An important question is whether PIN and MDR proteins interact in vivo to modulate IAA movements or whether they represent two distinct IAA transport pathways. The expression patterns of PGP1 and PGP19/MDR1 overlap with AtPIN1, whereas the PGP1 and PGP4 patterns overlap with AtPIN2 in other tissues, consistent with possible com-

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plex formation (as reviewed in Geisler and Murphy 2006). The altered localization of PIN1 in the mdr1/pgp19 mutant is consistent with a protein complex (Noh et al. 2003) that is altered in the absence of MDR/PGP proteins. The possibility of cooperativity between PIN1 and PGP1 and MDR1/PGP19 was examined in pgp1/pgp19 double mutants that overexpressed PIN1 under the control of an estradiol response promoter (Petrasek et al. 2006). PIN1 overexpression in both wild-type and pgp1/pgp19 mutants was sufficient to induce agravitropic root growth in both cases, suggesting that PIN1 action did not require PGP1 and PGP19 protein (Petrasek et al. 2006). The recent demonstration that the potassium carrier, TRH1, can also mediate IAA efflux in vivo and in vitro further suggests the complexity of proteins that mediate IAA efflux (Vicente-Agullo et al. 2004). A complete understanding of the complex assembly of auxin efflux carriers awaits further experimentation. The specificity of IAA transport machinery for naturally occurring auxins other than IAA has also been examined. IBA transport has been examined in several species (Ludwig-Muller 2000; Rashotte et al. 2003). In Arabidopsis, plants that have either the aux1 or pin2/eir1/agr1 mutation exhibit wild-type levels of IBA transport (Rashotte et al. 2003). Similarly, IBA transport is not affected by auxin efflux inhibitors such as NPA (Rashotte et al. 2003). The Arabidopsis rib1 mutant has altered IBA, but not IAA, transport, suggesting that the RIB1 protein may participate in IBA specific transport (Poupart et al. 2005). Similarly, in rice roots, the arm2 mutant has altered IBA uptake but wildtype levels of IAA uptake (Chhun et al. 2005). These results suggest that there may be additional as-yet unknown mechanisms that mediate transport of other auxins.

3.5 IAA Influx Carriers and Their Role in Gravitropism For many years the existence of an IAA influx carrier was questioned based on the chemiosmotic model of IAA transport (as reviewed in Goldsmith 1977). This model posits that because of the low pH of the extracellular space and a pKa for IAA of 4.8, some extracellular IAA should be protonated and the hydrophobicity of uncharged IAA should allow it to passively enter plant cells (as reviewed in Goldsmith 1977). Yet, the majority of IAA will be at pH above the pKa, so carrier-mediated uptake of the IAA anion would increase IAA accumulation. The demonstrations that IAA uptake was saturable in suspension cells (Rubery and Sheldrake 1974) and that there is substrate-specific uptake of auxins, with IAA and 2,4-D but not 1-NAA, moving into plant cells by carriermediated uptake (Delbarre et al. 1996) further supported the concept of protein-mediated IAA uptake. Our understanding of auxin influx has been extensively improved through the molecular and functional characterization of AUX1’s activity as an IAA influx protein (as reviewed by Parry et al. 2001b; Blakeslee et al. 2005) and identification of compounds which function as auxin influx inhibitors (Imhoff et al. 2000; Rahman et al. 2001a). The best-characterized protein with a role in auxin influx is AUX1. The aux1 mutant of Arabidopsis was identified in a screen for seedlings with altered elongation in the presence of 2,4-D (Maher and Martindale 1980). The aux1 mutant has an agravitropic root phenotype, as well as alterations in other auxin transport-dependent processes such as

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root branching and root hair formation (Maher and Martindale 1980; Marchant et al. 2002; Rahman et al. 2002). Consistent with a role in mediating IAA influx, roots of aux1 show a selective resistance to the auxins whose uptake appears to be carrier-mediated, IAA and 2,4-D, but not to the membrane-diffusible auxin, 1-NAA (Delbarre et al. 1996; Marchant et al. 1999; Yamamoto and Yamamoto 1998). The uptake and transport of IAA are also reduced in aux1 mutants (Rahman et al. 2001a; Rashotte et al. 2003). Molecular cloning of AUX1 supported the idea that this protein mediates IAA transport as the gene is similar in sequence to the ATF (amino acid transporter family) of proteins (Bennett et al. 1996; Young et al. 1999; Ortiz-Lopez et al. 2000). In the Arabidopsis genome, three other genes showed a high degree of sequence similarity to AUX1 and are termed the LAX (Like AUX1) gene family (Parry et al. 2001b). However, the function of the other family members has not yet been reported. AUX1 encodes a membrane protein of 48 KD composed of 11 transmembrane (TM)-spanning domains, with a cytoplasmic facing N-terminal domain (Swarup et al. 2004). The functional characterization of aux1 alleles revealed that the central region of AUX1 appears particularly important for protein function as nine of the missense mutations cluster between TM VI and VII. On the other hand, studies of the sole conditional allele aux1-7 mutant suggest that the Cterminal region of AUX1 may perform a regulatory function (Rahman et al. 2001a; Swarup et al. 2004). Finally, a recent report provided direct evidence that AUX1 indeed functions as an auxin influx career (Yang et al. 2006). Expression of wild-type AUX1 protein increased auxin uptake in Xenopus oocytes. In contrast, expression of three independent point mutants of AUX1, which abrogate the AUX1 function in planta, did not mediate auxin influx in oocytes. The substrate specificity of AUX1 in heterologous expression system is similar to the auxin specificity in plants, showing movement of IAA and 2,4-D, but not NAA and IBA (Yang et al. 2006). The tissue-specific expression pattern of AUX1 provides insight into its developmental role in planta. In the lateral root cap cells, AUX1 is localized without polarity, whereas in epidermal cells it is mainly axial, localized at both upper and lower sides (Marchant et al. 1999; Swarup et al. 2005). This expression pattern has been proposed to facilitate the basipetal transport of auxin between the gravity-sensing columella cells and the gravityresponsive cells of the distal elongation zone, but suggests that AUX1 does not specify the basipetal polarity (Swarup et al. 2005). Consistent with this model, aux1 mutations disrupt basipetal auxin transport and lead to agravitropic growth (Rashotte et al. 2003; Swarup et al. 2005). This idea is further confirmed by the restoration of gravitropic response of aux1 by tissue-specific expression of AUX1 (Swarup et al. 2005) in plants transformed with constructs expressed in cells of the lateral root cap and the epidermal cells of the elongation zone (Swarup et al. 2005). Collectively, these results suggest that localization of AUX1 is important for its role in plant development, so understanding the mechanisms that specify AUX1 localization is necessary. AXR4 functions as a specific regulator of AUX1 localization in epidermal and protophloem cells (Dharmasiri et al. 2006), but not for other membrane proteins such as PIN1, PIN2, or PM H+-ATPase, whose localization remain unaltered in the axr4 mutant (Dharmasiri et al. 2006). These findings are consistent with the similar aux1 and axr4 mutant phenotypes, which include agravitropic and auxin-insensitive growth (Dharmasiri et al. 2006). The AXR4 gene encodes a transmembrane protein with sequence similarity

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to the ␣/ß hydrolyase super-family and the protein localizes to the ER membrane, consistent with the accumulation of AUX1 protein in the ER of the axr4 mutant (Dharmasiri et al. 2006). One additional study suggested that brefeldin A (BFA), a molecule that blocks vesicle transport (described in detail below), may prevent the localization of AUX1 to the appropriate membrane in protophloem cells (Grebe et al. 2002). Identification of IAA influx inhibitors has also enhanced our understanding of the process of IAA influx. Imhoff et al. (2000) screened a large number of aryl and aryloxyalkylcarboxylic acids for their ability to block IAA influx in suspension-cultured tobacco cells. Two compounds, 1-napthoxyacetic acid (1-NOA) and 3-chloro-4hydroxyphenylacetic acid were found to inhibit auxin influx at micromolar concentrations (Imhoff et al. 2000) and to inhibit polar IAA transport in plants (Parry et al. 2001a). These influx inhibitors phenocopy the differential auxin resistance as well as agravitropic phenotypes of aux1, with 1-NOA having a specific effect on IAA influx (Parry et al. 2001a; Rahman et al. 2002). The ability of 1-NOA to specifically inhibit AUX1-mediated IAA influx was confirmed when AUX1 was expressed in Xenopus oocytes, as described above (Yang et al. 2006). A naturally occurring plant secondary metabolite, chromosaponin I (CSI), reduces IAA influx and phenocopies the agravitropic root growth of the aux1 mutant (Rahman et al. 2001a). In most alleles of aux1, where the mutation lies in the central domain or in the N-terminal domain (to date, 13 alleles tested), CSI either completely inhibited the gravitropic response in weak alleles or did not have any effect on the already agravitropic root growth in strong alleles (Swarup et al. 2004). Interestingly, CSI had a very different effect in aux1-7, which carries a mutation in the C-terminal domain. CSI rescued the gravitropic response and auxin uptake defects in aux1-7, indicating that CSI may directly interact with AUX1 protein (Rahman et al. 2001a) via the C-terminal domain (Swarup et al. 2004). The restoration of gravitropic response by CSI in an engineered transgenic line expressing HA-aux1-7 in a null allele (aux1-22) background confirmed this direct interaction (Swarup et al. 2004). The finding that chromosaponin acts as a naturally occurring influx inhibitor parallels the identification of flavonoids as regulators of auxin efflux, described below. The identification of these two molecules suggests that regulation of auxin movements by endogenous small molecules may be an important and general mechanism to control auxin transport and dependent physiological processes.

3.6 Regulation of IAA Efflux Protein Location and Activity during Gravity Response For a better understanding of changes in auxin transport, two important mechanisms need to be clarified. First, the initial establishment of polarity of auxin transport, and second, how this polarity is changed in response to gravity stimulation. The presence of multiple PIN and MDR/PGP and LAX genes, with distinct expression patterns and subcellular localizations, suggests that changes in synthesis and localization of IAA transporters are adequate to mediate these changes (as reviewed in Benjamins et al. 2005; Blakeslee et al. 2005). An interesting theme in this regulation is the ability of IAA to positively regulate the efflux of auxin through regulation of synthesis, targeting, and proteolytic degradation

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of IAA efflux proteins. Efflux carrier activity and/or synthesis may be regulated by phosphorylation and by regulatory molecules, such as flavonoids (as reviewed by Muday and DeLong 2001; Benjamins et al. 2005). Finally, increasing evidence suggests that an interplay between hormonal signaling pathways may regulate gravity response, with interactions between auxin and ethylene signaling being the best developed. 3.6.1 Mechanisms that May Control Localization of IAA Efflux Carriers The polar localization of PIN proteins has been suggested to be mediated by dynamic cycling of these proteins between internal compartments and the plasma membrane (Geldner et al. 2001; Geldner et al. 2003; as reviewed in Murphy et al. 2005). A recent report has used live imaging of PIN1::GFP in the Arabidopsis inflorescence meristem to identify rapid changes in protein localization that modulate auxin transport directionality and that are linked to floral meristem initiation (Heisler et al. 2005). Pharmacological approaches also support the idea that PIN protein localization is dynamic. The drugs monensin and brefeldin A (BFA), which are inhibitors of vesicle movements, were shown to reduce auxin transport though alterations in auxin efflux in the absence of protein synthesis (Wilkinson and Morris 1994; Morris and Robinson 1998; Delbarre et al. 1998). The asymmetric plasma membrane localization of PIN1 was altered by treatment with BFA, which led to accumulation of PIN1 in internal compartments termed BFA bodies (Geldner et al. 2001; Geldner et al. 2003). These BFA bodies contain endosomal markers (Geldner et al. 2003) and PIN1 protein accumulation in these structures is fully reversible upon removal of BFA (Geldner et al. 2001), suggesting that dynamic cycling of PIN1 between endosomes and the plasma membrane could control the localization of this and other auxin transport proteins, as shown in Figure 3.3. BFA treatment has now been shown to cause accumulation of PIN2, PIN3, and PIN4 in BFA bodies (Paciorek et al. 2005), suggesting that multiple IAA efflux proteins use similar mechanisms to reach their appropriate localization on the plasma membrane. Animal cells possess BFA-sensitive ARF-GEFs (ADP ribosylation factor-guanine nucleotide exchange factors) that direct vesicle movements through several pathways (Donaldson and Jackson 2000). The Arabidopsis gnom mutant, which has a defect in a gene encoding an ARF-GEF, exhibits altered PIN1 localization in developing embryos (Steinmann et al. 1999). GNOM was later shown to be the target for BFA in PIN cycling, as the GNOM transgenic plants with a mutated BFA binding site (GNOMM–L-myc) showed resistance to BFA-regulated, auxin-mediated developmental processes such as root gravitropic bending and lateral root formation and to accumulation of PIN1-GFP in BFA bodies (Geldner et al. 2003). Together, these results indicate that GNOM is a target in BFA inhibition of PIN1-dependent auxin transport, and suggest a mechanism for differential localization of IAA efflux carriers. Accordingly, mutations in SCARFACE (SFC), a gene recently shown to encode an ARF-GAP (GTPase activating protein) that enhances cleavage of ARF bound GTP to GDP, thereby negatively regulating ARF activity (Randazzo and Hirsch 2004), result in altered BFA-dependent PIN1::GFP cycling and defects in auxin transport and dependent physiological processes (Sieburth et al. 2006). An important question is whether cycling of a PIN protein may allow changes in auxin transport polarity that is needed to mediate gravitropism. Experimental evidence suggests

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Figure 3.3. A model of interactions of BFA and auxin efflux inhibitors with PIN1 cycling and auxin efflux. A magnification of vesicular cycling in the basal portion of a polarized plant cell is shown. A. PIN1 cycles between the endosome and plasma membrane in untreated, polarized, IAA-transporting cells. B. Interruption of PIN1 cycling in BFA-treated cells results in PIN1 accumulation in an endomembrane compartment, which reduces IAA transport. BFA binds to GNOM, which co-localizes with endomembrane aggregations of PIN1. BFA washout restores PIN1 cycling and IAA efflux. This figure has been modified from Muday et al. (2003) and reprinted with permission of Elsevier Publishing.

that the PIN3 protein may function in this way (Friml et al. 2002; see also Chapter 2). PIN3 is expressed in root columella cells and PIN3 localization changes in response to reorientation of roots relative to gravity, consistent with redistribution of IAA from the root tip (Friml et al. 2002). This protein also cycles in a BFA-dependent manner (Friml et al. 2002). Therefore, relocalization of PIN3 in the root cap may redirect auxin flow at the tip of roots reoriented relative to gravity. Consistent with this role, pin3 mutant roots respond to gravitropic reorientation with a slower response than wild-type (Friml et al. 2002). Recent work has examined the physiological significance of the cycling of PIN proteins in controlling auxin transport polarity, with a specific focus on the role of this process in changing the auxin transport polarity in gravity-responding roots. Paciorek et al. (2005) tested the possibility that IAA controls the cycling of PIN proteins. In their experimental system, the active auxins IAA, NAA, and 2,4-D prevent the BFA-induced accumulation of PIN1 into BFA bodies, as did the yucca mutant, which has elevated endogenous IAA levels (Paciorek et al. 2005). In addition, it was shown that the auxins inhibit uptake of FM 4-64, a fluorescent dye taken up by cells during endocytosis (Paciorek et al. 2005). To determine whether the cycling of PIN2 is affected by the local IAA concentrations, Paciorek et al. (2005) used the endogenous gradients in IAA across a gravity-stimulated root and found that on the lower side of the root, where IAA levels are higher, there is less accumulation of PIN2 in BFA bodies and less endocytosis of FM 4-64. Treatment with NPA, which prevents formation of the IAA gradient, resulted in a similar PIN2 accumulation in BFA bodies and levels of endocytosis on the two sides of

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the root (Paciorek et al. 2005). Taken together, these results suggest a mechanism by which IAA may reinforce its own changing polarity during root gravitropism. The pharmacological approach using BFA, described above, provides strong evidence for the cycling of PIN proteins as a mechanism to change auxin transport polarity during gravity response, and has uncovered some intriguing potential regulatory mechanisms. Yet several aspects of these studies need to be carefully considered. First, the dose of BFA needed to cause PIN1 accumulation in BFA bodies (Geldner et al. 2001) is much higher than that needed to reduce root growth, gravity response, and lateral root formation (Geldner et al. 2001; Geldner et al. 2004) and to alter vesicular movements (Niu et al. 2005; Parton et al. 2003). Second, these BFA bodies are not observed in untreated cells and, although they contain endosomal markers (Geldner et al. 2003), the biological significance of these structures is not yet clear. Aside from changes in endocytosis of FM 464, which correlate with changes in BFA sensitivity to BFA body formation (Paciorek et al. 2005), there has been no evidence of regulated endosomal cycling of PIN proteins in the absence of BFA. Finally, although BFA has been shown to alter IAA transport in inflorescence tissues (Geldner et al. 2003) and cultured plant cells (Morris and Robinson 1998), the effect of BFA on IAA transport in the tissues in which BFA affects PIN cycling and growth and development have not been reported. These points highlight the importance of additional studies to understand the intriguing idea of endosomal cycling as a regulatory mechanism to control IAA transport polarity. 3.6.2 Regulation of IAA Efflux by Synthesis and Degradation of Efflux Carriers Changes in abundance of IAA efflux proteins may also enhance the effects of PIN protein cycling to amplify gradients in IAA. Although PIN protein cycling has been shown to occur in the absence of protein synthesis (Geldner et al. 2001), other experiments have also shown that there are transcriptional controls of efflux carriers that accompany change in auxin transport (Peer et al. 2004; Vieten et al. 2005). In particular, the expression of the PIN1-6 genes has been examined and shown to be controlled by changing auxin levels as a result of application of auxins and auxin efflux inhibitors (Peer et al. 2004; Vieten et al. 2005). These results are consistent with transcriptional controls of efflux carriers that may alter the capacity of plant tissues to transport auxin, and with an additional level of feedback of auxin on its own transport. Induced synthesis of IAA transport proteins in cells on the lower side of roots in response to gravistimulation could then serve to enhance transport on the lower side of horizontal roots, whereas decreased expression and enhanced turnover of auxin transporters in cells of the upper side could reduce auxin transport on this upper side. A schematic diagram of auxin transport changes at the root tip is shown in Figure 3.2E, F. Just as enhanced synthesis of efflux carriers can increase IAA transport to the lower side of a gravity-stimulated root, proteolytic degradation of existing carriers that mediate IAA transport to the upper side of horizontal roots would also facilitate formation of asymmetries in IAA concentrations (Sieberer et al. 2000; Abas et al. 2006). Differential proteolysis of PIN2 protein across gravity-stimulated roots has been shown for endogenous PIN2 and EIR1/PIN2 fusions to GUS and GFP (Sieberer et al. 2000; Abas et al. 2006). This proteolytic cleavage is prevented by proteosome inhibitors (Abas et al. 2006).

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Addition of auxin prevents the loss of EIR1/PIN2 protein and PIN2-GFP fusions (Abas et al. 2006). Similarly, in gravity-stimulated roots, the loss of PIN2-GFP on the upper side of the root can be detected with kinetics that parallel the gravitropic response (Abas et al. 2006). Together, these results suggest that efflux carriers are regulated at the level of synthesis, breakdown, and localization. 3.6.3 Regulation of Auxin Transport by Reversible Protein Phosphorylation The activity of many highly regulated proteins is controlled by reversible phosphorylation. Therefore, it is not surprising that changes in localization and/or activity of auxin transport proteins due to protein phosphorylation may also regulate auxin transport (as reviewed in DeLong et al. 2002; Muday et al. 2003 ). Inhibitor studies have implicated protein kinases in regulating auxin transport and its sensitivity to auxin transport inhibitors (Bernasconi 1996; Delbarre et al. 1998). Genetic evidence for phosphorylation control of auxin transport comes from studies of the pinoid (pid) and roots curl in NPA1 (rcn1) mutants, which have defects in genes encoding a protein kinase and a protein phosphatase regulatory subunit, respectively (as reviewed in DeLong et al. 2002). The PID gene encodes a member of the AGC-family of serine/threonine kinases (Christensen et al. 2000), and pid mutants exhibit altered auxin transport in inflorescences and a floral development defect resembling that of the pin1 mutant (Bennett et al. 1995; Christensen et al. 2000; Benjamins et al. 2001). Overexpression of PID reduced elongation of roots and hypocotyls, DR5:GUS expression in the root tip, gravitropism, and lateral root initiation (Christensen et al. 2000; Benjamins et al. 2001). Additionally, the main root meristem was also found to collapse after a few days of germination, followed by the emergence of lateral roots (Benjamins et al. 2001). Treatments with the IAA efflux inhibitors, NPA and TIBA, increased root elongation and prevented the collapse of the primary root meristem, suggesting that auxin transport is increased in 35S::PID seedlings and auxin transport inhibitors help to reduce auxin transport to normal levels in these seedlings. Finally, tissue-specific overexpression of PID in the shoot led to increased lateral root initiation, which was blocked by the application of NPA at the root shoot junction, consistent with PID regulating auxin flow from the shoot into the root (Benjamins et al. 2001). Consistent with this finding, overexpression of PID in root hair and tobacco cells enhanced auxin efflux (Lee and Cho 2006). In inflorescences, pinoid loss-of-function and PINOID overexpression have been suggested to have opposite effects on the polar targeting of the PIN1 auxin efflux facilitator protein (Friml et al. 2004), consistent with the hypothesis that reversible protein phosphorylation by PID may act at the level of protein targeting (Muday and Murphy 2002). Analysis of the rcn1 mutant has shown that protein phosphatase 2A (PP2A) activity regulates root auxin transport and gravitropic curvature. The RCN1 gene encodes a regulatory A subunit of PP2A, and the rcn1 mutant has reduced PP2A activity in vivo and in vitro (Garbers et al. 1996; Deruère et al. 1999; Muday et al. 2006). Roots of rcn1 seedlings have elevated basipetal auxin transport and exhibit a significant delay in gravitropism (Rashotte et al. 2001). Reduced PP2A activity causes the phenotypes observed in rcn1 roots and hypocotyls because these effects can be mimicked by treating wild-type seedlings with low doses of protein phosphatase inhibitors (Deruère et al. 1999; Rashotte et al. 2001; Larsen and Cancel 2003; Shin et al. 2005).

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Hypocotyls of rcn1 also exhibit altered gravity response and auxin transport (Muday et al. 2006). As in the root tip, RCN1-controlled PP2A activity appears to act as a negative regulator of basipetal auxin transport (Rashotte et al. 2001). Paradoxically, loss of RCN1 function impedes gravitropic response in roots but enhances curvature in hypocotyls. Consideration of the differences in gravity response mechanisms in these two tissues suggests a hypothesis to explain the apparent contradiction. As discussed in Chapters 1 and 2, roots sense gravity very locally in the columella cells in the root cap (Blancaflor et al. 1998), and auxin is redistributed from the root tip to one side of the root after gravity stimulation rather than being laterally transported across the root tip (as reviewed in Blancaflor and Masson 2003). In roots, uniformly increased basipetal transport may impede the redistribution of auxin at the root tip, which is required to form a lateral auxin gradient and to achieve maximal gravitropic bending (Rashotte et al. 2001). Consistent with this hypothesis, treatment of rcn1 roots with low doses of NPA reduces auxin transport and enhances gravity response to wild-type levels (Rashotte et al. 2001). In contrast, gravity perception in stems occurs in the starch sheath parenchyma tissues that run the length of the hypocotyl (Fukaki et al. 1998; see also Chapters 1 and 2). Lateral auxin transport then is believed to occur in multiple tissues along the length of the hypocotyl (Blancaflor and Masson 2003). Increased basipetal auxin transport would provide more auxin to the lateral transport stream and would thereby increase gravitropic bending. In contrast, the mdr1 mutant has reduced hypocotyl IAA transport (Noh et al. 2001), but has enhanced gravi- and phototropic responses. These differences may be due to specific effects of the mdr1 mutation on transporter localization or function (Noh et al. 2003), rather than the rcn1 mutation which affects bulk polar auxin flow. The possibility that the rcn1 gravitropic phenotype was due to altered ethylene response was examined (Muday et al. 2006), but this possibility is not consistent with several results. The rcn1-2 allele was identified in a screen for increased ethylene response in etiolated seedlings and was originally designated eer1 (enhanced ethylene response) (Larsen and Chang 2001; Larsen and Cancel 2003). Enhanced ethylene response in rcn1 is a hypocotyl-specific phenotype and is accompanied by ethylene overproduction (Larsen and Chang 2001). The rcn1 hypocotyl gravitropic phenotype was found to be ethylene-independent as the rcn1-2 etr1-1 and rcn1-2 ein2-1 mutants showed gravity responses that are identical to the rcn1 single mutant (Muday et al. 2006). Additionally, although silver treatment of wild-type seedlings reduces the gravity response, silver treatment of rcn1 seedlings further enhanced the gravity response, consistent with the enhanced gravitropic phenotype of rcn1 being independent of ethylene signaling (Muday et al. 2006). These results indicate that an intact ethylene signaling pathway is not required for the enhancement of gravity response in rcn1 hypocotyls. The etiolated growth phenotype of rcn1 is likely due to the elevated ethylene synthesis that is only found in dark-grown seedlings (Muday et al. 2006). Further experiments will be required to determine the mechanism by which PP2A affects auxin transport in roots, and to identify the targets of kinase and phosphatase regulation in auxin transport. Localization of the PIN2/AGR1/EIR1 protein appears to be normal in rcn1 root tips (Shin et al. 2005), and neither PIN2/AGR1/EIR1 nor AUX1 is required for the rcn1 transport phenotype (Rashotte et al. 2001). The recent identification of other auxin carriers that function in control of auxin transport in the root, and for

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which mutant phenotypes include altered gravity response, suggest other targets of phosphorylation beyond the PIN proteins. 3.6.4 Regulation of Auxin Transport by Flavonoids Prime candidates for endogenous auxin transport inhibitors are flavonoids. These phenolic compounds displace the binding of synthetic IAA efflux inhibitors, such as NPA, during in vitro assays (Jacobs and Rubery 1988). The flavonols quercetin and kaempferol had the greatest activity, suggesting that specific members of this chemical family function as auxin transport inhibitors (Jacobs and Rubery 1988). The role of flavonoids as regulators of auxin transport have been examined by in vivo studies in tt4 mutants, which have a defect in the CHS gene encoding chalcone synthase, the first enzyme in flavonoid synthesis. These tt4 mutants have elevated auxin transport in young seedlings, roots, or inflorescences, consistent with the absence of an endogenous negative auxin transport regulator (Murphy et al. 2000; Brown et al. 2001; Buer and Muday 2004; Peer et al. 2004; Buer et al. 2006). Roots of multiple alleles of tt4 mutants exhibit a lag in gravitropic curvature compared to wild-type roots (Buer and Muday 2004; Buer et al. 2006). Chemical complementation of tt4(2YY6) by naringenin reinstated flavonoid production and restored a wild-type gravity response, consistent with a role for flavonoids in controlling the flow of auxin needed for root gravitropism (Buer and Muday 2004). Mutations that alter flavonoid synthesis affect the abundance of the mRNA encoding members of the PIN gene family (Peer et al. 2004; Lazar and Goodman 2006), suggesting that flavonoids may regulate synthesis of auxin transport proteins, not just the activity of existing proteins. Consistent with flavonoid abundance affecting transcription, recent evidence indicated that flavonoid biosynthetic enzymes and flavonoid products accumulate in the nucleus (Saslowsky et al. 2005). Changing environmental conditions modulate flavonoid synthesis (Winkel-Shirley 2002) and these changes in flavonoid accumulation may regulate plant growth and development, including gravity responses (as reviewed in Taylor and Grotewold 2005). Reorientation of plants relative to gravity leads to enhanced flavonoid accumulation in the epidermal tissues of Arabidopsis root tips (Buer and Muday 2004), which are the site of basipetal IAA transport. Images of roots and the time course of this induction are shown in Figure 3.4. This induction is on both the upper and lower sides of gravistimulated roots, suggesting that its function may be to uniformly reduce the activity or abundance of a set of efflux carriers, and thereby accentuate the formation of a gradient of IAA across the root resulting from enhanced transport on the lower side. Therefore, induction of flavonoid synthesis in response to environmental stimuli may alter auxin transport to facilitate plant gravity response. 3.6.5 Regulation of Auxin Transport by Other Signaling Pathways Most of the mechanisms for altering IAA transport polarity during root gravitropism utilize IAA as a signal to enhance the gradient, consistent with multiple levels of positive feedback. It is critical to consider the gravity-signaling events that precede any IAA asymmetries and act as the initial signals to initiate auxin transport changes. As discussed

Figure 3.4. Flavonoid accumulation spikes during gravitropic stimulation in wild-type Arabidopsis. A. The relative flavonoid fluorescence determined by dividing average maximum flavonoid accumulation across the 40 µm from the root tip of vertically-grown controls versus gravity-stimulated roots measured over time is reported. Arrows indicate times of gravity-stimulated root bending in Col and tt4(2YY6). B–C. Col root tips of 2.25 h vertical controls and (D–E) gravity-stimulated roots. The optical slices in C and E are approximately 15 µm below the root-tip surface that is shown in B and D. The scale bar = 40 µm. This figure is reprinted from Buer and Muday (2004), with kind permission of the American Society of Plant Biologists.

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in Chapter 2, biochemical and cell biological approaches have identified several signaling molecules that are produced in response to a changing gravity vector and may modulate auxin transport. It has long been suspected that calcium is a signaling molecule that changes in concentration in response to changes in the vector of gravity (as reviewed in Sinclair and Trewavas 1997). Attempts to demonstrate changes in cytoplasmic calcium concentration using calcium ratio imaging in Arabidopsis roots did not detect calcium concentration changes in response to gravitropic stimulation (Legue et al. 1997). Evidence in support of calcium as a signal in gravitropic response comes from young seedlings of Arabidopsis expressing aequorin in the cytoplasm (Plieth and Trewavas 2002). When a population of seedlings is reoriented relative to gravity, there is an enhanced aequorin signal consistent with elevated cytoplasmic calcium concentration (Plieth and Trewavas 2002). Several experiments have suggested that gravity stimulation may be amplified by cascades involving Ca2+/calmodulin (Sinclair et al. 1996; Lu and Feldman 1997), and a number of older studies suggested a relationship between auxin transport and calcium concentration (dela Fuente 1984; Allan and Rubery 1991). Therefore, the possibility that calcium signals are integral to gravitropic response remains intriguing but requires additional experimental test. The role of several other signaling molecules has been more clearly demonstrated, as discussed in Chapter 2. Changes in pH have been observed after gravitropic stimulation in both Arabidopsis roots (Scott and Allen 1999; Fasano et al. 2001; Boonsirichai et al. 2003) and the maize pulvinus (Johannes et al. 2001). Proton movements have been tied to auxin transport through examination of mutants and transgenics with altered expression of the H+-pyrophosphatase, AVP1 (Li et al. 2005). Altered AVP1 expression changes vacuolar pH and alters IAA transport and PIN1 localization (Li et al. 2005) Additionally, inositol lipids have been implicated in the gravity signal transduction pathway in maize and Arabidopsis. A transient increase in the InsP3 lipid signal have been observed in gravity-stimulated maize and oat pulvini (Perera et al. 1999; Perera et al. 2001). Gravity-stimulated pulvini undergo rapid initial changes in InsP3 levels on both sides, followed by a greater and more persistent elevation on the lower side (Perera et al. 1999). This later InsP3 elevation on the lower side is necessary for gravitropic bending of the pulvinus, as treatment with phospholipase C inhibitors prevent formation of the InsP3 gradient and reduced gravitropic bending (Perera et al. 1999). In maize, free IAA has been measured and shown to develop an asymmetry across the gravity-stimulated pulvinus that follows the changes in InsP3 levels (Long et al. 2002). More recently, additional support for the role of InsP3 comes from transgenic studies in Arabidopsis with plants constitutively expressing the human type I inositol polyphosphate 5-phosphatase (InsP 5-ptase), an enzyme that specifically hydrolyzes InsP3 (Perera et al. 2006). In the transgenic plants, basal InsP3 levels are reduced by greater than 90% compared to wild-type plants. With gravistimulation, InsP3 levels in inflorescence stems of transgenic plants show no detectable change, whereas in wild-type plant inflorescences, InsP3 levels increase approximately threefold within the first 5 to 15 min of gravistimulation, preceding visible bending (Perera et al. 2006). Gravitropic bending of the roots, hypocotyls, and inflorescence stems of InsP 5-ptase transgenic plants is reduced (Perera et al. 2006). These transgenics also exhibit reductions in root basipetal IAA transport and delays in formation of lateral asymmetries in auxin-induced gene expression as

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compared to the controls (Perera et al. 2006). Together, these results suggest that InsP3 synthesis may be a signal that modulates auxin transport to allow differential growth in response to changes in the gravity vector. 3.6.6 Regulation of Gravity Response by Ethylene Increasing evidence suggests that ethylene may regulate gravitropism either directly or through modulation of auxin’s role in this process. Exogenous application of ethylene gas or the ethylene precursor 1-aminocyclo-propanecarboxylic acid (ACC) has been used to test the role of ethylene in gravity response. In some experiments ethylene or ACC treatment clearly reduced the early phase of gravitropic response of roots and shoots (Wheeler and Salisbury 1981; Wheeler et al. 1986; Lee et al. 1990; Kiss et al. 1999; Madlung et al. 1999; Buer et al. 2006), whereas others showed no effect (Kaufman et al. 1985; Harrison and Pickard 1986; Woltering 1991). A third set of experiments suggested that ethylene positively regulated shoot gravitropism (Chang et al. 2004). The experiments that identified a role for ethylene in gravity response examined the initial rate of gravitropic curvature (Wheeler and Salisbury 1981; Wheeler et al. 1986; Lee et al. 1990), suggesting that the absence of kinetic data at early times after gravitropic stimulation in several of the experiments may explain their negative results (Kaufman et al. 1985; Harrison and Pickard 1986; Woltering 1991). It has been shown that the application of ethylene inhibitors such as AVG/AgNO3 also reduced the initial gravitropic curvature (Wheeler et al. 1986; Lee et al. 1990; Muday et al. 2006) both in roots and shoots. The similar effect observed by the ethylene precursor and inhibitors suggests that ethylene may both positively and negatively regulate gravitropism. Mutants altered in ethylene signaling and/or synthesis have been used to examine the role of ethylene in gravitropic curvature. The gravitropic responses of the etr1 roots (Buer et al. 2006) and hypocotyls (Muday et al. 2006) and ein2-1 roots are wild type (Roman et al. 1995; Rahman et al. 2001b; Buer et al. 2006). The gravitropic response of shoots of the tomato mutants, Never-Ripe (Nr) and epi, which have reduced ethylene response and enhanced synthesis, respectively, were examined. Both mutants exhibit delays in shoot gravitropic response but with only a small reduction in Nr (Madlung et al. 1999), consistent with a role for ethylene in the early events of gravitropic response. The study of Madlung et al. (1999) revealed a concentration-dependent modulation of shoot gravitropism by ethylene, with Nr being insensitive to the effect of exogenous ethylene on hypocotyl gravitropism. Similarly, etr1 and ein2 roots and hypocotyls are insensitive to the inhibition of gravitropism by ACC treatment (Buer et al. 2006; Muday et al. 2006). These results suggest that ethylene negatively regulates gravity response, and that for plants grown on agar the endogenous levels of ethylene are low enough that there are no detectable differences between wild-type and ethylene-insensitive mutants. The one exception to this conclusion are the agravitropic hypocotyls of ein2, although this phenotype may be linked to EIN2 activities that are ethylene-independent (Muday et al. 2006). One mechanism by which auxin and ethylene may interact is at the level of hormone synthesis with auxin-inducing ethylene synthesis and/or ethylene-inducing auxin synthesis. Auxin is a positive regulator of ethylene biosynthesis in many plants, including Arabidopsis (Yang and Hoffman 1984; Woeste et al. 1999; Harper et al. 2000). The rate-

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limiting step in ethylene synthesis is catalyzed by ACC synthase, which is encoded by the ACS gene family with some family members being auxin-inducible in dark-grown seedlings and plants (Abel et al. 1995; Yamagami et al. 2003). Recently, one ACC synthase gene was shown to be asymmetrically induced across a gravity-stimulated snapdragon flower spike, suggesting that asymmetric synthesis of ethylene may be part of gravitropic response in some tissues (Woltering et al. 2005). The regulation of auxin synthesis by ethylene has been uncovered by the mutants designated weak ethylene insensitive (wei2 and wei7). The WE12 and WE17 genes encode ethylene-regulated enzymes of Trp synthesis, whose activity positively regulates IAA synthesis (Stepanova et al. 2005). Another explanation for the interaction between auxin and ethylene is that ethylene may inhibit IAA transport. In some plant species, ethylene has been shown to inhibit the polar IAA transport in shoot tissues (Morgan and Gausman 1966; Suttle 1988), but a direct effect of ACC treatment on IAA transport was not detected in Arabidopsis hypocotyls (Muday et al. 2006) or roots (Buer et al. 2006). Lateral IAA transport has also been found to be inhibited in both shoots (Burg and Burg 1966) and gravity-stimulated corn roots (Lee et al. 1990). These results suggest that ethylene-mediated inhibition of auxin transport may play an important role in regulating the gravity response. A recent article from Buer et al. (2006) asked whether ethylene might inhibit gravity response through induction of flavonoid synthesis resulting in reduced IAA transport. Several alleles of the flavonoid-deficient mutant tt4 exhibit a delayed gravity response in roots and are insensitive to the inhibition of gravitropism at early time points. More interestingly, ACC has been shown to induce flavonoid accumulation in Arabidopsis roots through a mechanism that requires EIN2 and ETR proteins (Buer et al. 2006). Taken together, these results suggest that the ethylene regulation of root gravity response may occur through altering flavonoid synthesis, assuming that enhanced flavonoid accumulation will reduce IAA transport. However, Buer et al. (2006) did not find any effect of ACC on root basipetal auxin transport, indicating that this interaction may be more complex in nature or too difficult to detect in the tips of the small roots of Arabidopsis. The most direct evidence for interaction between ethylene signaling and auxin transport comes from the studies of Arabidopsis mutants having mutations in auxin transport proteins. Both the auxin influx mutant, aux1, and the auxin efflux mutant, agr1/ eir1/pin 2/wav6, have ethylene-insensitive root elongation (Roman et al. 1995; Pickett et al. 1990). The restoration of ethylene sensitivity in both aux1 and eir1 by exogenous application of NAA and IAA indicates that cytoplasmic auxin is needed at sufficient levels for ethylene response (Rahman et al. 2001b).

3.7 Overview of the Mechanisms of Auxin-Induced Growth This chapter has focused on the mechanisms by which asymmetries in auxin are established in response to changing orientation of plants relative to gravity. Yet, to understand how these auxin gradients control growth, a brief discussion of auxin signal transduction is required, although this topic has been reviewed in greater detail elsewhere (Leyser 2002, 2006). Plants respond to changing levels of auxin by dramatic changes in transcription, with

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microarray experiments identifying hundreds of genes whose expression is rapidly modified in response to auxin treatment (Pufky et al. 2003). The most rapidly expressed genes are induced even in the absence of protein synthesis. These primary auxin-responsive genes include SAURs (small auxin up-regulated), GH3s, and AUX/IAAs (Auxin/IAA inducible genes) gene families (as reviewed in Hagen and Guilfoyle 2002). Consensus promoter elements were found in these auxin-responsive genes and used to identify proteins, named Auxin Response Factors (ARFs), that bind to them (as reviewed in Hagen and Guilfoyle 2002). The in vivo function of these proteins has been demonstrated by isolation of Arabidopsis mutants altered in expression or function of ARF and AUX/IAA genes. These mutants have a diversity of auxin-dependent phenotypes, including agravitropic roots and/or hypocotyls (as reviewed in Liscum and Reed 2002). The nph4 (nonphototropic hypocotyl) mutant, which has a defect in the ARF7 gene, exhibits reduced hypocotyl phototropic and gravitropic responses (Stowe-Evans et al. 1998; Harper et al. 2000). Several mutants with defects in IAA genes also have agravitropic phenotypes, including iaa14/slr1-1, iaa17/axr3-1, and iaa19/msg2 (as reviewed in Liscum and Reed 2002). ARF proteins contain DNA binding domains and dimerization domains, and have been shown to act as transcriptional regulators (Tiwari et al. 2003). AUX/IAA proteins have similar dimerization domains and form homo- and hetero-dimers with ARF proteins, thereby modulating the ability of ARFs to bind to the promoter of auxin-responsive genes (Kim et al. 1997). Until relatively recently, it has been unclear how the auxin signal influences the formation of transcription factor complexes that are needed to modulate the expression of auxin-responsive genes. Recent experiments have demonstrated that auxin-dependent proteolytic destruction of AUX/IAA proteins is a critical factor (as reviewed in Leyser 2006). Specifically, mutations in the TIR1, AXR1, and AXR6 genes lead to auxin-resistant plants. The proteins encoded by these genes have now been shown to be part of protein complexes that ubiquitinylate substrate proteins, thereby targeting them for destruction (as reviewed in Leyser 2006). The current model for this process is that TIR1 binding to auxin facilitates formation of complexes with AUX/IAA proteins, resulting in their ubiquitination. Ubiquitinylated proteins are then targeted for destruction by the proteosome. Consequently, ARF proteins are released from inhibitory complexes with AUX/IAA and bind to the promoter elements of auxin-responsive genes, regulating their expression (Dharmasiri et al. 2005; Kepinski and Leyser 2005). Although this mechanism was rather unexpected, the data clearly show that destruction of regulatory transcription factor subunits is an efficient system for rapid gene expression changes in response to changing auxin levels. An additional question that remains to be answered is: What kinds of genes are induced in response to auxin? Microarray experiments have shown that expression of a large number of genes changes in response to auxin application (Pufky et al. 2003) or gravitropic reorientation (Moseyko et al. 2002; Kimbrough et al. 2004; Esmon et al. 2006). Auxininduced gene products include enzymes that may loosen the cell wall to facilitate growth, such as expansins (Esmon et al. 2006) or invertase (Long et al. 2002), or transporters that allow ion flow across the membrane to alter membrane polarization, turgor, and characteristics of the cell wall (as reviewed in Becker and Hedrich 2002), such as H+-ATPase (as reviewed in Hager 2003) and potassium channels (Philippar et al. 1999). The most strik-

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ing study, which ties auxin-induced gene expression to differential growth during phototropism and gravitropism, utilized Brassica oleracea (Esmon et al. 2006). These seedlings were of sufficient size to isolate opposite flanks of hypocotyls after exposure to lateral light stimulation or after being placed horizontally. mRNA was isolated from each flank and used to hybridize an Arabidopsis microarray. A number of genes were shown to be differentially expressed on the two sides of the stimulated hypocotyls. Additionally, the expression of these same genes was enhanced in the hypocotyls of auxin-treated etiolated Arabidopsis seedlings in a NPH4/ARF7-dependent fashion (Esmon et al. 2006). Several of these genes were shown to encode proteins that are directly tied to growth, including two expansin genes, EXPA1 and EXPA8 (Esmon et al. 2006). Finally, the functional significance of differences in auxin-regulated gene expression across gravity-stimulated tissues needs to be evaluated in the context of differences in gravity response between roots and shoots. In light-grown shoot tissues, auxin is generally limiting for growth (Yang et al. 1993; Gray et al. 1998), so the elevated auxin concentrations on the lower side of shoots reoriented relative to gravity could operate to induce transcription of genes encoding proteins that enhance growth, consistent with the report described above (Esmon et al. 2006). In contrast, although roots also redirect auxin to the lower side, the opposite response is initiated, resulting in growth with the gravity vector. Although root growth is negatively regulated by auxin under most conditions (Pickett et al. 1990), it is not completely clear whether this growth response is due to the elevated auxin on the lower side or the reduced auxin levels on the upper side. Detailed kinetic analysis of roots after gravitropic reorientation in many species indicates that most exhibit enhanced growth on the upper side, rather than the predicted growth inhibition on the lower side (as reviewed in Wolverton et al. 2002). These kinetic studies also revealed that shortly after gravitropic reorientation, there is enhanced growth on both sides of the root, which is sustained on the upper side of the root, followed by a reduced growth rate on the lower side of the root once curvature initiates (Buer and Muday 2004). The complexity of this response has led to the suggestion that parts of the response may be auxin-independent (Wolverton et al. 2002), but it is also possible that the signaling mechanisms (including transcription changes described above) could control this process. Since auxin positively and negatively regulates the expression of responsive genes, depending on the complexes of ARF and AUX/IAA proteins involved (Hagen and Guilfoyle 2002), scenarios can be envisioned to support this more complex response of roots. The lower auxin levels on the upper side of a reoriented root could relieve auxin’s repression of genes that encode proteins that positively regulate growth, just as the enhanced auxin levels on the lower side could repress synthesis of growth-inducing proteins. Additional experiments will be needed to determine the complex interplay of signals that change in response to gravity stimulation and their mechanisms for controlling the complex process of root gravitropism.

3.8 Conclusions The ability of plants to respond to gravity is tied to the redistribution of auxin. In the last decade, the proteins that are involved in auxin redistribution have been elucidated by a

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combination of genetic and biochemical approaches. These findings offer tremendous insight into the novel mechanisms that control the synthesis, targeting, activity, as well as the proteolytic degradation of proteins that act in concert to drive auxin redistribution. Finally, the understanding of how the auxin signal is transduced to control synthesis of proteins needed for asymmetric growth rounds out our understanding of how the asymmetries in auxin induce differential growth. 3.9 Acknowledgments Several grants facilitated this work, with GKM supported by the U.S. Department of Agriculture (No. 2006-35304-17311) and AR supported by the National Science Foundation (IBN 0316876). 3.10 Literature Cited Abas, L., R. Benjamins, N. Malenica, T. Paciorek, J. Wisniewska, J. C. Moulinier-Anzola, T. Sieberer, J. Friml and C. Luschnig. 2006. Intracellular Trafficking and Proteolysis of the Arabidopsis Auxin-Efflux Facilitator PIN2 Are Involved in Root Gravitropism. Nat Cell Biol 8: 249–56. Abel, S., M. Nguyen, W. Chow and A. Theologis. 1995. Asc4, a Primary Indoleacetic AcidResponsive Gene Encoding 1-Aminocyclopropane-1-Carboxylate Synthase in Arabidopsis thaliana. J Biol Chem 270: 19093–99. Allan, A. C. and P. H. Rubery. 1991. Calcium Deficiency and Auxin Transport in Cucurbita pepo L. Seedlings. Planta 183: 604–12. Becker, D. and R. Hedrich. 2002. Channelling Auxin Action: Modulation of Ion Transport by Indole-3-Acetic Acid. Plant Mol Biol 49: 349–56. Benjamins, R., N. Malenica and C. Luschnig. 2005. Regulating the Regulator: The Control of Auxin Transport. Bioessays 27: 1246–55. Benjamins, R., A. Quint, D. Weijers, P. Hooykaas and R. Offringa. 2001. The PINOID Protein Kinase Regulates Organ Development in Arabidopsis by Enhancing Polar Auxin Transport. Devel 128: 4057–67. Bennett, M. J., A. Marchant, H. G. Green, S. T. May, S. P. Ward, P. A. Millner, A. R. Walker, B. Schulz and K. A. Feldmann. 1996. Arabidopsis Aux1 Gene: A Permease-Like Regulator of Root Gravitropism. Science 273: 948–50. Bennett, S. R. M., J. Alvarez, G. Bossinger and D. R. Smyth. 1995. Morphogenesis in Pinoid Mutants of Arabidopsis thaliana. Plant J. 8: 505–20. Bernasconi, P. 1996. Effect of Synthetic and Natural Protein Tyrosine Kinase Inhibitors on Auxin Efflux in Zucchini (Cucurbita pepo) Hypocotyls. Physiol Plant 96: 205–10. Björkman, T. and R. Cleland. 1991. The Role of Extracellular Free Ca2+ Gradients in Gravitropic Signalling in Maize Roots. Planta 185: 379–84. Blakeslee, J. J., W. A. Peer and A. S. Murphy. 2005. Auxin Transport. Curr Opin Plant Biol 8: 494–500. Blancaflor, E. B., J. M. Fasano and S. Gilroy. 1998. Mapping the Functional Roles of Cap Cells in the Response of Arabidopsis Primary Roots to Gravity. Plant Physiol 116: 213–22. Blancaflor, E. B. and P. H. Masson. 2003. Plant Gravitropism. Unraveling the Ups and Downs of a Complex Process. Plant Physiol 133: 1677–90.

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Sieberer, T., G. J. Seifert, M. T. Hauser, P. Grisafi, G. R. Fink and C. Luschnig. 2000. PostTranscriptional Control of the Arabidopsis Auxin Efflux Carrier EIR1 Requires AXR1. Curr Biol 10: 1595–8. Sieburth, L. E., G. K. Muday, E. J. King, G. Benton, S. Kim, K. E. Metcalf, L. Meyers, E. Seamen and J. M. Van Norman. 2006. SCARFACE Encodes an ARF-Gap That Is Required for Normal Auxin Efflux and Vein Patterning in Arabidopsis. Plant Cell 18: 1396–411. Sinclair, W., I. Oliver, P. Maher and A. Trewavas. 1996. The Role of Calmodulin in the Gravitropic Response of the Arabidopsis thaliana agr3 Mutant. Planta 199: 343–51. Sinclair, W. and A. Trewavas. 1997. Calcium in Gravitropism. A Re-Examination. Planta 203: S85–S90. Stasinopoulos, T.C. and R.P. Hangarter. 1990. Preventing Photochemistry in Culture Media by Long-Pass Light Filters Alters Growth of Cultured Tissues. Plant Physiol 93: 1365–9. Steinmann, T., N. Geldner, M. Grebe, S. Mangold, C. L. Jackson, S. Paris, L. Galweiler, K. Palme and G. Jurgens. 1999. Coordinated Polar Localization of Auxin Efflux Carrier PIN1 by GNOM ARF GEF. Science 286: 316–8. Stepanova, A. N., J. M. Hoyt, A. A. Hamilton and J. M. Alonso. 2005. A Link between Ethylene and Auxin Uncovered by the Characterization of Two Root-Specific Ethylene-Insensitive Mutants in Arabidopsis. Plant Cell 17: 2230–42. Stowe-Evans, E. L., R. M. Harper, A. V. Motchoulski and E. Liscum. 1998. NPH4, a Conditional Modulator of Auxin-Dependent Differential Growth Responses in Arabidopsis. Plant Physiol 118: 1265–75. Suttle, J. C. 1988. Effect of Ethylene Treatment on Polar IAA Transport, Net IAA Uptake and Specific Binding of N-1-Naphthylphthalamic Acid in Tissues and Microsomes Isolated from Etiolated Pea Epicotyls. Plant Physiol. 88: 795–9. Swarup, R., J. Kargul, A. Marchant, D. Zadik, A. Rahman, R. Mills, A. Yemm, S. May, L. Williams, P. Millner, S. Tsurumi, I. Moore, R. Napier, I. D. Kerr and M. J. Bennett. 2004. Structure-Function Analysis of the Presumptive Arabidopsis Auxin Permease AUX1. Plant Cell 16: 3069–83. Swarup, R., E. M. Kramer, P. Perry, K. Knox, H. M. Leyser, J. Haseloff, G. T. Beemster, R. Bhalerao and M. J. Bennett. 2005. Root Gravitropism Requires Lateral Root Cap and Epidermal Cells for Transport and Response to a Mobile Auxin Signal. Nat Cell Biol 7: 1057–65. Taylor, L. P. and E. Grotewold. 2005. Flavonoids as Developmental Regulators. Curr Opin Plant Biol 8: 317–23. Terasaka, K., J. J. Blakeslee, B. Titapiwatanakun, W. A. Peer, A. Bandyopadhyay, S. N. Makam, O. R. Lee, E. L. Richards, A. S. Murphy, F. Sato and K. Yazaki. 2005. PGP4, an ATP Binding Cassette P-Glycoprotein, Catalyzes Auxin Transport in Arabidopsis thaliana Roots. Plant Cell 17: 2922–39. Tiwari, S. B., G. Hagen and T. Guilfoyle. 2003. The Roles of Auxin Response Factor Domains in Auxin-Responsive Transcription. Plant Cell 15: 533–43. Trewavas, A. J. 1992. Forum: What Remains of the Cholodny-Went Theory? Plant Cell and Environ 15: 759–94. Tsurumi, S. and Y. Ohwaki. 1978. Transport of 14C-Labeled Indoleacetic Acid in Vicia Root Segments. Plant Cell Physiol 19: 1195–1206. Utsuno, K., T. Shikanai, Y. Yamada and T. Hashimoto. 1998. AGR, an Agravitropic Locus of Arabidopsis thaliana, Encodes a Novel Membrane-Protein Family Member. Plant Cell Physiol 39: 1111–8. Vicente-Agullo, F., S. Rigas, G. Desbrosses, L. Dolan, P. Hatzopoulos and A. Grabov. 2004. Potassium Carrier TRH1 Is Required for Auxin Transport in Arabidopsis Roots. Plant J 40: 523–35.

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4

Phototropism and Its Relationship to Gravitropism Jack L. Mullen and John Z. Kiss*

4.1 Phototropism: General Description and Distribution That sunlight can affect the development of plants has been known for thousands of years, and one of the more striking effects is the growth of plants toward areas of more intense light. This response, phototropism, is caused by a difference in growth rate across part of the plant with the direction of the growth differential determined by the gradient in light intensity. Phototropism is widespread among plants and is found in algae, mosses, and ferns, as well as seed plants. In flowering plants, stems are generally positively phototropic, curving toward the direction of highest light intensity. However, there are some species, including climbing plants with tendrils, which have negatively phototropic shoots, allowing them to grow toward neighboring plants (Darwin 1875; Strong and Ray 1975). Some recent reviews of phototropism include Liscum and Stowe-Evans (2000), Kimura and Kagawa (2006), and Whippo and Hangarter (2006). Leaves are also frequently phototropic, though the effect of light on leaf growth is more complex than for stems. Leaves of many species engage in a diurnal phototropic response, following the movement of the sun. Much of the curvature occurs in the leaf petiole, which is capable of sensing the light directly or, if shaded, responding to light sensed by the leaf blade (Haberlandt 1914). Some species have specialized turgor-driven motor tissues, called pulvini, which allow for greater reversibility in the response. The leaf blade itself may also reorient to be normal to the incident light and can track the sun across the sky in this position (Lang and Begg 1979; Koller and Levitan 1989). Since the direction of the light is nearly perpendicular to the leaf surface throughout the day, this response requires impressive sensitivity to changes in photostimulation. Although its importance is not clear, phototropism has also been observed in root systems, with roughly half of species examined showing some response (Hubert and Funke 1937). Root phototropism could be useful in positioning lateral roots near the soil surface, where light may penetrate, though the response is generally smaller than the gravitropic response of roots (Kiss et al. 2002). The majority of roots showing a response are negatively phototropic (Hubert and Funke 1937; Okada and Shimura 1992). However, roots of Arabidopsis can respond either positively or negatively to unidirectional light, depending on the light quality (Kiss et al. 2003). Thus, it appears that multiple lightsignaling pathways interact with other growth responses such as gravitropism, even belowground in the root systems.

*Corresponding author

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4.2 Light Perception The light environment in which plants naturally live and grow can be a very heterogeneous one, with both diffuse and directional light varying throughout the landscape. Spatial differences in the light intensity impinging upon a plant manifest as a gradient in light intensity across the particular plant organ, due to scattering and absorption by the plant tissue (Vogelmann et al. 1996). This leads to a gradient in photoreceptor activation in the plant organ, providing directional information regarding the light stimulus. The presence of screening pigments, such as carotenoids, in the plant can increase the light gradient and have been found to increase the magnitude of phototropic responses (Piening and Poff 1988). In leaves of solar-tracking plants, however, gradients in light intensity across the organs do not play an important role in the phototropic response. In these plants, the leaf surface is perpendicular to the direction of the light beam, and phototropic leaf reorientation occurs when the direction of the light beam becomes oblique (Schwartz and Koller 1978). This vectorial phototropic response is sensed by cells above the veins of the leaves, and the ability to sense light direction is related to the angle between the light beam and the directions of the major axes of the veins. Sensing of the direction of light, in this case, has been postulated to be due to localization of photoreceptors to the end walls of the cells along the leaf veins (Koller et al. 1990). Plants are generally bathed in a full spectrum of light, but nevertheless they have different photoreceptor pigments to sense specific qualities of light (Figure 4.1). The action spectrum for phototropism in flowering plants clearly shows that blue light is the most effective in this process. Although this observation was known for more than a century (reviewed in Whippo and Hangarter 2006), it was only in the 1990s that the specific bluelight-absorbing photoreceptor was identified by the use of nph (non-phototropic hypocotyl) mutants of Arabidopsis. This family of photoreceptors, renamed phototropins, has two members found in Arabidopsis, PHOT1 and PHOT2 (Christie et al. 1999; Sakai et al. 2001). The phot1 and phot2 holoproteins have different sensitivities to light intensity, with phot2 being functionally active only at the relatively higher light intensities of greater than approximately 10 µmol m–2 s–1 (Sakai et al. 2001). The phototropins are autophosphorylating serine/threonine kinases with two LOV (light-, oxygen-, or voltagesensitive) domains, a motif that is present in a wide diversity of organisms including plants, fungi, and bacteria. The LOV domains bind flavin mononucleotide as chromophores for the photoreceptor (Christie et al. 1999). However, although both LOV domains are evolutionarily conserved, only the second domain (LOV2) appears to be functionally important for photoperception (Christie et al. 2002; Cho et al. 2007). The phototropins are localized to the plasma membrane, though following stimulation with blue light a fraction dissociate from the membrane (Sakamoto and Briggs 2002; Kong et al. 2006). In parenchyma cells near the vascular tissue, phot1 is asymmetrically distributed, with most of the photoreceptor found along the end walls normal to the major axis of the veins (Sakamoto and Briggs 2002). This localization is consistent with the proposed mechanism for vectorial light sensing (Koller et al. 1990). Despite their name, phototropins are not specific to the phototropic response. They can also function in other responses such as chloroplast movement in leaves and stomatal opening (Briggs and Christie 2002; Kimura and Kagawa 2006).

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Figure 4.1 (also see Color Section). Light quality and phototropic responses. Several photoreceptors are important in phototropic responses, sensing light in the blue, red, and far-red ranges of the visible spectrum. The signaling pathways from these photoreceptors interact to regulate the blue-light-based phototropic responses. The phytochromes separately also regulate some other phototropism-related responses, including red-light root phototropism and alteration of stem gravitropism, which can enhance phototropic responses. Some ferns and algae have a chimeric photoreceptor, neochrome, which can mediate phototropism. BL, blue light; Dk, darkness; RL, nondirectional red light; WL, white light.

Although the phototropins are the primary photoreceptors responsible for initiating phototropic responses, another family of blue-light-absorbing photoreceptors in plants, the cryptochromes, is also involved in regulating the response (Figure 4.1). The principal roles of cryptochromes are in photomorphogenesis and the entrainment of circadian rhythms (Cashmore 2003), but they have also been found, in mediating blue-lightinduced stomatal opening, to act synergistically with phototropins (Mao et al. 2005). Cryptochromes are not necessary for the induction of phototropic responses (Lasceve et al. 1999; Sakai et al. 2000). However, at low fluence rates, cryptochrome-deficient mutants have reduced phototropic responses (Ahmad et al. 1998; Lasceve et al. 1999; Whippo and Hangarter 2003). Cryptochromes are also important in regulating light-induced growth inhibition in hypocotyls, together with the phototropins (Parks et al. 2001). Because phototropism is caused by complex changes in growth rate, alterations in growth inhibition by cryptochromes is likely to intersect phototropic signaling. Indeed, Whippo and Hangarter (2003) have shown that at high light intensities, the cryptochromes, along with the phototropins, inhibit phototropism. Thus, the cryptochromes may interact with phototropins to modulate the phototropic pathway (Figure 4.1 and Color Section) and also control the inhibition of growth of hypocotyls in a fluence-rate-dependent manner, which can limit the potential phototropic response.

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In most cases, red light does not induce phototropism in flowering plants. Red and farred light are sensed by the phytochromes (Figure 4.1)—photoreceptors which are involved in numerous aspects of the plant life cycle, including seed germination, flowering, and circadian rhythms (Schepens et al. 2004). The phytochrome gene family has five members (PHYA–E) in Arabidopsis. Although unidirectional red light does not induce phototropism in Arabidopsis hypocotyls (Liscum and Briggs 1996), red-light pretreatments are known to greatly enhance blue-light phototropism, in a process mediated by phytochromes (Janoudi et al. 1992; Liu and Iino 1996; Hangarter 1997). Phytochromes can also absorb blue light and they modulate blue-light phototropic responses, even in the absence of a red-light pretreatment (Correll et al. 2003). At low fluence rates of unidirectional blue light, phytochromes (particularly PHYA and PHYB) reduce the latent period and enhance the magnitude of the phototropic response (Janoudi et al. 1997; Hangarter 1997). However, at higher fluence rates of blue light, phytochrome (primarily PHYA) causes an attenuation in the response (Whippo and Hangarter 2004). Phytochromes can interact with the cryptochromes (Ahmad et al. 1998; Mas et al. 2000); and like the cryptochromes, phytochromes also regulate light-induced growth inhibition in hypocotyls (Folta and Spalding 2001). Therefore, in addition to a possible role in directly modulating the phototropism signaling pathway, phytochromes also appear to act in coordination with the cryptochromes and phototropins (Figure 4.1) to regulate shoot growth rate more generally in response to light stimuli, a process that also modulates the overall phototropic response observed. Some algae, mosses, and ferns engage in red-light phototropism, mediated by phytochrome (Wada and Sei 1994; Esch et al. 1999), in addition to blue-light phototropism. The green alga Mougeotia and a group of ferns have independently evolved a chimeric photoreceptor that is a hybrid between phytochrome and phototropin, termed neochrome (Figure 4.1), which controls the red-light phototropism in these plants (Kawai et al. 2003; Suetsugu et al. 2005). However, neochromes have not been found in mosses or flowering plants. This chimeric photoreceptor broadens the response by allowing strong absorption of both red light and blue light. The signaling of neochrome for the two wavelengths is synergistic, so that the photoreceptor has increased sensitivity to weak white light; this allows plants with these pigments (e.g., polypodiaceous ferns) to sense and respond to low-light signals in their naturally shaded light environment (Kanegae et al. 2006). There have been a few reports of phototropism mediated by normal phytochromes in flowering plants. In roots of Arabidopsis, there is a positive red-light phototropic response controlled by phytochromes A and B, in addition to the negative blue-light response (Kiss et al. 2003). In shoots, there are reports of phytochrome-regulated phototropism in mesocotyls of maize (Iino et al. 1984) and negative far-red phototropism in cucumber (Ballare et al. 1992, 1995), as well as positive far-red phototropism in the parasitic plant Cuscuta planiflora (Orr et al. 1996).

4.3 Signal Transduction and Growth Response Although there has been considerable progress in understanding the cell and molecular biology of the primary photoreceptors including phototropin, cryptochrome, and phytochrome, relatively little is known about the downstream signaling events (i.e., signal

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transduction) following light perception. Nevertheless, in recent years, numerous mutants in light-signaling intermediates, particularly in the phytochrome pathways, have been isolated and used to investigate light signaling in plants (Møller et al. 2002). In terms of signaling intermediates that are downstream from the phototropins, RPT2 (root phototropism) and NPH3 (non-phototropic hypocotyl) have been shown to bind to PHOT1 and to function very early in some blue-light-based signaling pathways (Sakai et al. 2000; Inada et al. 2004). For instance, RPT2 is involved in phototropism and stomatal opening but not in chloroplast movements, and although NPH3 is involved in phototropism, it is not needed for stomatal opening or chloroplast relocation. RPT2 and NPH3 are considered to be in the same biochemical family, but the exact structure and precise physiological functions of these proteins are not known. Although NPH3-deficient mutants of Arabidopsis showed no phototropism in shoots or roots (Sakai et al. 2000), the cpt1 mutant of rice, orthologous to NPH3, retained some root phototropism, suggesting that other members of the gene family may also play a role in this plant (Haga et al. 2005). The growth response of phototropism involves differential elongation on opposite sides of a plant organ, which eventually leads to phototropic curvature. It is wellestablished that the plant hormone auxin plays an integral role in the differential growth that results in curvature. In fact, the discovery of auxin and the classical Cholodny-Went model for auxin transport have been tied to research in tropisms, especially phototropism. The NPH4 locus, which is important for both phototropism and gravitropism, encodes an auxin response factor involved in auxin-sensitive transcriptional regulation (Harper et al. 2000). Much recent focus in auxin research, especially in conjunction with understanding the role of auxins in tropisms, has centered on the auxin transport proteins in the PIN (termed such because of the pin-shaped inflorescence stem in mutants) family (Blakeslee et al. 2005; Paponov et al. 2005; see also Chapter 3 of this book). Understanding the link among auxin transport proteins, NPH3/RPT2, and the actin cytoskeleton will be important in determining the precise molecular role of these molecules in linking light perception in phototropism to the differential growth effects mediated by auxin (Maisch and Nick 2007).

4.4 Interactions with Gravitropism Once a phototropic stimulus causes curvature in a plant, the orientation of the particular plant organ will begin to change in relation to the gravity vector. This generally will lead to a countering gravitropic response following the initial phototropic curvature. The two signaling pathways thus need to be integrated into an overall growth response. Experiments examining simultaneous gravitropic and phototropic stimulation have found that the equilibrium growth response can be more complex than simple additivity (Nick and Schäfer 1988; Galland 2002). Interpretation of these experiments is made somewhat difficult by differences in the kinetics of gravitropic and phototropic responses. For example, the responses can have different latent periods, and there may be adaptation to the stimulus during the responses, depending on stimulus strength, such that the magnitude of curvature decreases with time (Iino 1988; Mullen et al. 2002; Kiss et al. 2003). In the case of phototropism, part of the adaptation is likely due to changes in photoreceptor lev-

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els, as expression of PHOT1 (Sakamoto and Briggs 2002; Kong et al. 2006) and several phytochromes (Sharrock and Clack 2002) is down-regulated by light. In the case of gravitropic adaptation, however, it is unclear where in the signaling pathway the modulation occurs. Part of the reduction in curvature may also be due to autotropic straightening, a reversal of curvature which can occur following tropistic responses (Orbovic and Poff 1991; Stankovic et al. 1998). There can also be differences in the spatial distribution of the responses, adding complexity to the overall growth pattern. In roots, phototropic curvature develops farther from the root tip than gravitropic curvature (Mullen et al. 2002; Kiss et al. 2003), whereas in shoots, there can be differences in the extent of the region of curvature between gravitropic and phototropic responses (Tarui and Iino 1999). Nondirectional light, more generally, may modulate gravitropic responses apart from the integration of gravitropic and phototropic growth responses. There have been observations of red light sensitizing hypocotyls to gravitropic stimulation (Britz and Galston 1982; Woitzik and Mohr 1988). However, it appears more common for red light to inhibit gravitropism. In hypocotyls of Arabidopsis, red light acting via phytochrome appears to inhibit gravitropism, causing a randomization of shoot orientation (Figure 4.1; Liscum and Hangarter 1993; Poppe et al. 1996; Robson and Smith 1996). This inhibition of a countering gravitropic response may explain how red light enhances blue-light phototropism (Hangarter 1997; Parks et al. 1996). Red-light attenuation of shoot gravitropism has also been observed in pea and tobacco (McArthur and Briggs 1979; Hangarter 1997). And in mosses, at least, red light does not act to inhibit gravitropism at the level of perception, as red-light treatments did not repress amyloplast sedimentation (Kern and Sack 1999). This signaling pathway may also be acting during blue-light phototropism, as phytochrome absorption of blue light can also cause randomization of shoot orientation (Lariguet and Fankhauser 2004). Light inhibition of gravitropism has also been observed in leaves (Mano et al. 2006), although whether it is also mediated by phytochrome is unclear.

4.5 Importance to Plant Form and Function At a whole-plant level, the positioning of branches and leaves through phototropic and gravitropic responses will play an important role in the overall functioning of the organism. Yet tests of the roles of specific phototropic responses at specific stages of the life cycle of a plant remain limited, although it appears that a key function of phototropism is to help the plant maximize photosynthesis. Experiments with Arabidopsis phot mutants suggest that PHOT1 and PHOT2 may be important at different developmental stages, consistent with their different sensitivities to light intensity (Galen et al. 2004; Galen et al. 2007). In young seedlings, one of the important roles of phototropism may be orienting the root system away from the surface to aid in dealing with dry conditions (Galen et al. 2007). As plants develop, positioning of leaves becomes increasingly important, allowing them to exploit light gaps or orient at the correct angle for light capture from the sun. For solar-tracking plants, phototropism allows for increased photosynthesis in the morning and evening. This appears to be particularly important for plants found in habitats with short growing seasons (Ehleringer and Forseth 1980). However, because solar tracking

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increases light interception by leaves, it also increases the heat load for these plants, which could be disadvantageous, particularly if water is limited. In fact, some solartracking species can change their leaf orientation from being perpendicular to the direction of light to parallel to the light, depending on water stress (Shackel and Hall 1979; Forseth and Ehleringer 1980). Also, in many arid environments the leaves of plants that do not solar track become more vertical with increasing light intensity, apparently as a protective mechanism against photodamage from excess light (King 1997; Valladares and Pugnaire 1999; Falster and Westoby 2003). In young seedlings, the primary shoot generally grows upward in a seemingly straightforward integration of positive phototropism and negative gravitropism, and the primary root grows downward in a similarly straightforward combination of positive gravitropism and negative phototropism (Okada and Shimura 1992). However, the bulk of a mature plant is made up of lateral organs—branches and leaves in the shoot and lateral and adventitious roots belowground—which grow at nonvertical orientations even in the absence of a phototropic stimulus. It is the growth of these lateral branches that gives plants their characteristic form, and they have large effects on plant productivity. Because these nonvertical orientations appear to be actively maintained via gravitropic responses, Digby and Firn (1995) have termed the growth orientation of these organs to be their gravitropic set-point angle, or GSA (see also Chapter 2 of this book). The GSA of both shoots and roots can be altered by red light (Gaiser and Lomax 1993; Digby and Firn 2002; Mullen and Hangarter 2003). Thus, the fine-tuning of organ positioning in mature plants by directional light cues requires a more complex integration of growth responses, involving not only phototropism, gravitropism, and their interactions, but also light-dependent changes in GSA. Because the GSA is a developmentally regulated variable (Digby and Firn 1995), light regulation of GSA allows for differences in response depending on the age of the specific organ. This may allow different parts of a mature plant to tailor their growth in an appropriate manner for the specific environmental conditions they encounter.

4.6 Conclusions and Outlook Tremendous progress in understanding the mechanisms of phototropism has been made in recent years. Although phototropism has been intensively studied since the time of Darwin’s classic experiments, it is only within the past decade that we have identified the phototropins as the primary pigments involved in light perception in phototropism. It has become increasingly obvious that the other two major groups of photoreceptors, the cryptochromes and phytochromes, interact with phototropins and play both direct and indirect roles in phototropic responses. Much of the current research focuses on a better understanding of the cellular and molecular events downstream from the primary photoreceptors. There is increased recognition that the interaction between and among the primary photoreceptors is important in phototropism and other important light-regulated developmental processes (Mas et al. 2000; Folta and Spalding 2001; Whippo and Hangarter 2003; Lariguet and Fankhauser 2004; Kumar and Kiss 2007). One recent approach we have used to better understand the relationship between pho-

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totropism and gravitropism is to use the microgravity conditions during spaceflight (Correll et al. 2005; Kiss et al. 2007; see also Chapter 8 of this book). In these experiments on the international space station, we plan to study the role of phytochromes in phototropism and to determine whether red light affects phototropism directly or indirectly by the attenuation of gravitropism. However, through the increased use of the new tools of molecular and systems biology in the next decade, we should gain an even better understanding of the basic mechanisms of phototropism, its relationship to gravitropism, and its importance to plants.

4.7 Literature Cited Ahmad, M., J.A. Jarillo, O. Smirnova, and A.R. Cashmore. 1998. Cryptochrome blue-light photoreceptors of Arabidopsis implicated in phototropism. Nature 392: 720–723. Ballare, C.L., A.L. Scopel, S.R. Radosevich, and R.E. Kendrick. 1992. Phytochrome-mediated phototropism in de-etiolated seedlings: occurrence and ecological significance. Plant Physiol. 100: 170–177. Ballare, C.L., A.L. Scopel, M.L. Roush, and S.R. Radosevich. 1995. How plants find light in patchy canopies. A comparison between wild-type and phytochrome-B-deficient mutant plants of cucumber. Funct. Ecol. 9: 859–868. Blakeslee, J.J., W.A. Peer, and A.S. Murphy. 2005. Auxin transport. Curr. Opin. Plant Biol. 8: 494–500. Briggs, W.R., and J.M. Christie. 2002. Phototropins 1 and 2: versatile plant blue-light receptors. Trends Plant Sci. 7: 204–210. Britz, S.J., and A.W. Galston. 1982. Light-enhanced perception of gravity in stems of intact pea seedlings. Planta 154: 189–192. Cashmore, A.R. 2003. Cryptochromes: enabling plants and animals to determine circadian time. Cell 114: 537–543. Cho, H.-Y., T.-S. Tseng, E. Kaiserli, S. Sullivan, J.M. Christie, and W.R. Briggs. 2007. Physiological roles of the Light, Oxygen, or Voltage domains of phototropin 1 and phototropin 2 in Arabidopsis. Plant Physiol. 143: 517–529. Christie, J.M., M. Salomon, K. Nozue, M. Wada, and W.R. Briggs. 1999. LOV (light, oxygen, or voltage) domains of the blue-light photoreceptor phototropin (nph1): binding sites for the chromophore flavin mononucleotide. Proc. Natl. Acad. Sci. (USA) 96: 8779–8783. Christie, J.M., T.E. Swartz, R. Bogomolni, and W.R. Briggs. 2002. Phototropin LOV domains exhibit distinct roles in regulating photoreceptor function. Plant J. 32: 205–219. Correll, M.J., K.M. Coveney, S.V. Raines, J.L. Mullen, R.P. Hangarter, and J.Z. Kiss. 2003. Phytochromes play a role in phototropism and gravitropism in Arabidopsis roots. Adv. Space Res. 31: 2203–2210. Correll, M.J., R.E. Edelmann, R.P. Hangarter, J.L. Mullen, J.Z. Kiss. 2005. Ground-based studies of tropisms in hardware developed for the European Modular Cultivation System (EMCS). Adv. Space Res. 36:1203–1210. Darwin, C.R. 1875. “The movements and habits of climbing plants.” John Murray, London. Digby, J., and R.D. Firn. 1995. The gravitropic set-point angle (GSA): the identification of an important developmentally controlled variable governing plant architecture. Plant Cell Environ. 18: 1434–1440. Digby, J., and R.D. Firn. 2002. Light modulation of the gravitropic set-point angle (GSA). J. Exp. Bot. 53: 377–381. Ehleringer, J., and I. Forseth. 1980. Solar tracking by plants. Science 210: 1094–1098.

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Esch, H., E. Hartmann, D. Cove, M. Wada, and T. Lamparter. 1999. Phytochrome-controlled phototropism of protonemata of the moss Ceratodon purpureus: physiology of the wild type and class 2 ptr-mutants. Planta 209: 290–298. Falster, D.S., and M. Westoby. 2003. Leaf size and angle vary widely across species: what consequences for light interception? New Phytol. 158: 509–529. Folta, K.M., and E.P. Spalding. 2001. Opposing roles of phytochrome A and phytochrome B in early cryptochrome-mediated growth inhibition. Plant J. 28: 333–340. Forseth, I., and J.R. Ehleringer. 1980. Solar tracking response to drought in a desert annual. Oecologia 44: 159–163. Gaiser, J.C., and T.L. Lomax. 1993. The altered gravitropic response of the lazy-2 mutant of tomato is phytochrome regulated. Plant Physiol. 102: 339–344. Galen, C., J. Huddle, and E. Liscum. 2004. An experimental test of the adaptive evolution of phototropins: blue-light photoreceptors controlling phototropism in Arabidopsis thaliana. Evolution 58: 515-523. Galen, C., J.J. Rabenold, and E. Liscum. 2007. Functional ecology of a blue light photoreceptor: effects of phototropin-1 on root growth enhance drought tolerance in Arabidopsis thaliana. New Phytol. 173: 91-99. Galland, P. 2002. Tropisms of Avena coleoptiles: sine law for gravitopism, exponential law for photogravitropic equilibrium. Planta 215: 779-784. Haberlandt, G. 1914. “Physiological plant anatomy.” Macmillan, London. Haga, K., M. Takano, R. Neumann, and M. Iino. 2005. The rice COLEOPTILE PHOTOTROPISM1 gene encoding an ortholog of Arabidopsis NPH3 is required for phototropism of coleoptiles and lateral translocation of auxin. Plant Cell 17: 103-115. Hangarter, R.P. 1997. Gravity, light and plant form. Plant Cell Environ. 20: 796-800. Harper, R.M., E.L. Stowe-Evans, D.R. Luesse, H. Muto, K. Tatematsu, M.K. Watahiki, K. Yamamoto, and E. Liscum. 2000. The NPH4 locus encodes the auxin response factor ARF7, a conditional regulator of differential growth in aerial Arabidopsis tissue. Plant Cell 12: 757-770. Hubert, B., and G.L. Funke. 1937. The phototropism of terrestrial roots. Biologisch Jahrboek 4: 286-315. Iino M. 1988. Desensitization by red and blue light of phototropism in maize coleoptiles. Planta 176: 183-188. Iino, M., W.R. Briggs, and E. Schäfer. 1984. Phytochrome-mediated phototropism in maize seedling shoots. Planta 160: 41-51. Inada, S., M. Ohgishi, T. Mayama, K. Okada, and T. Sakai. 2004. RPT2 is a signal transducer involved in phototropic response and stomatal opening by association with phototropin 1 in Arabidopsis thaliana. Plant Cell 16: 887-896. Janoudi, A.K., R. Konjevic, P. Apel, and K.L. Poff. 1992. Time threshold for second positive phototropism is decreased by a preirradiation with red light. Plant Physiol. 99: 1422-1425. Janoudi, A.K., R. Konjevic, G. Whitelam, W. Gordan, and K.L. Poff. 1997. Both phytochrome A and phytochrome B are required for the normal expression of phototropism in Arabidopsis thaliana seedlings. Physiol. Plant. 101: 278-282. Kanegae, T., E. Hayashida, C. Kuramoto, and M. Wada. 2006. A single chromoprotein with triple chromophores acts as both a phytochrome and a phototropin. Proc. Natl. Acad. Sci. (USA) 103: 17997-18001. Kawai, H., T. Kanegae, S. Christensen, T. Kiyosue, T. Sato, T. Imaizumi, A. Kadota, M. Wada. 2003. Responses of ferns to red light are mediated by an unconventional photoreceptor. Nature 421: 287-290. Kern, V.D., and F.D. Sack. 1999. Irradiance-dependent regulation of gravitropism by red light in protonemata of the moss Ceratodon purpureus. Planta 209: 299–307.

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Kimura, M., and T. Kagawa. 2006. Phototropin and light-signaling in phototropism. Curr. Opin. Plant Sci. 9: 503–508. King, D.A. 1997. The functional significance of leaf angle in Eucalyptus. Austral. J. Bot. 45: 619–639. Kiss, J.Z., P. Kumar, R.N. Bowman, M.K. Steele, M.T. Eodice, M.J. Correll, R.E. Edelmann. 2007. Biocompatibility studies in preparation for a spaceflight experiment on plant tropisms (TROPI). Adv. Space Res., in press. Kiss, J.Z., K.M. Miller, L.A. Ogden, and K.K. Roth 2002. Phototropism and gravitropism in lateral roots of Arabidopsis. Plant Cell Physiol. 43: 35–43. Kiss, J.Z, J.L. Mullen, M.J. Correll, and R.P. Hangarter. 2003. Phytochromes A and B mediate redlight-induced positive phototropism in roots. Plant Physiol. 131: 1411–1417. Koller, D., and I. Levitan. 1989. Diurnal phototropism in leaves of Lavatera cretica L. under conditions of simulated solar tracking. J. Exp. Bot. 40: 1059–1064. Koller, D., S. Ritter, W.R. Briggs, and E. Schäfer. 1990. Action dichroism in perception of vectorial photo-excitation in the solar-tracking leaf of Lavatera cretica L. Planta 181: 184–190. Kong, S.-G., T. Suzuki, K. Tamura, N. Mochizuki, I. Hara-Nishimura, and A. Nagatani. 2006. Blue light-induced association of phototropin 2 with the Golgi apparatus. Plant J. 45: 994–1005. Kumar P., and J.Z. Kiss. 2007. The SHL1 and SHL5 genes influence both red- and blue-light-based phototropism in Arabidopsis thaliana. Environ. Exp. Bot. 60: 284–289 Lang, A.P.G., and J.E. Begg. 1979. Movements of Helianthus annuus leaves and heads. J. Applied Ecol. 16: 299–305. Lariguet, P., and C. Fankhauser. 2004. Hypocotyl growth orientation in blue light is determined by phytochrome A inhibition of gravitropism and phototropin promotion of phototropism. Plant J. 40: 826–834. Lasceve, G., J. Leymarie, M.A. Olney, E. Liscum, J.M. Christie, A. Vavasseur, and W.R. Briggs. 1999. Arabidopsis contains at least four independent blue-light-activated signal transduction pathways. Plant Physiol. 120: 605–614. Liscum, E., and W.R. Briggs. 1996. Mutations of Arabidopsis in potential transduction and response components of the phototropic signaling pathway. Plant Physiol. 112: 291–296. Liscum, E., and R.P. Hangarter. 1993. Genetic evidence that the Pr form of phytochrome B plays a role in Arabidopsis thaliana gravitropism. Plant Physiol. 103: 15–19. Liscum, E., and E. Stowe-Evans. 2000. Phototropism: a “simple” physiological response mediated by multiple interacting photosensory-response pathways. Photochem. Photobiol. 72: 273–282. Liu, Y.T., and M. Iino. 1996. Phytochrome is required for the occurrence of time-dependent phototropism in maize coleoptiles. Plant Cell Environ. 19: 1379–1388. Maisch, J., and P. Nick. 2007. Actin is involved in auxin-dependent patterning. Plant Physiol. 143: 1695–1704. Mano, E., G. Horiguchi, and H. Tsukaya. 2006. Gravitropism in leaves of Arabidopsis thaliana (L.) Heynh. Plant Cell Physiol. 47: 217–223. Mao, J., Y.C. Zhang, Y. Sang, Q.H. Li, and H.Q. Yang. 2005. A role for Arabidopsis cryptochromes and COP1 in the regulation of stomatal opening. Proc. Natl. Acad. Sci. (USA) 102: 12270–12275. Mas, P., P.F. Devlin, S. Panda, and S.A. Kay. 2000. Functional interaction of phytochrome B and cryptochrome 2. Nature 408: 207–211. McArthur, J.A., and W.R. Briggs. 1979. Effect of red light on geotropism in pea epicotyls. Plant Physiol. 63: 218–220. Møller, S.G., P.J. Ingles, and G.C. Whitelam. 2002. The cell biology of phytochrome signalling. New Phytol. 154: 553–590. Mullen, J.L., and R.P. Hangarter. 2003. Genetic analysis of the gravitropic set-point angle in lateral roots of Arabidopsis. Adv. Space Res. 31: 2229–2236.

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Mullen, J.L., C. Wolverton, H. Ishikawa, R.P. Hangarter, and M.L. Evans. 2002. Spatial separation of light perception and growth response in maize root phototropism. Plant Cell Environ. 25: 1191–1196. Nick, P., and E. Schäfer. 1988. Interactions of gravi- and phototropic stimulation in the response of maize coleoptiles. Planta 173: 213–220. Okada, K., and Y. Shimura. 1992. Mutational analysis of root gravitropism and phototropism of Arabidopsis thaliana seedlings. Aust. J. Plant Physiol. 19: 439–448. Orbovic, V., and K.L. Poff. 1991. Kinetics for phototropic curvature by etiolated seedlings of Arabidopsis thaliana. Plant Physiol. 97: 1470–1475. Orr, G.L., M. A. Haidar, and D.A. Orr. 1996. Smallseed dodder (Cuscuta planiflora) phototropism toward far-red when in white light. Weed Sci. 44: 233–240. Paponov, I., W. Teal, M. Trebar, K. Blilou, and K. Palme. 2005. The PIN auxin efflux facilitators: evolutionary and functional perspectives. Trends Plant Sci. 410: 170–177. Parks, B.M., P.H. Quail, and R.P. Hangarter. 1996. Phytochrome A regulates red-light induction of phototropic enhancement in Arabidopsis. Plant Physiol. 110: 155–162. Parks B.M, K.M. Folta, and E.P. Spalding. 2001. Photocontrol of stem growth. Curr. Opin. Plant Biol. 4: 436–440. Piening, C.J., and K.L. Poff. 1988. Mechanism of detecting light direction in first positive phototropism in Zea mays L. Plant Cell Environ. 11: 143–146. Poppe, C., R.P. Hangarter, R.A. Sharrock, F. Nagy, and E. Schäfer. 1996. The light-induced reduction of the gravitropic growth-orientation of seedlings of Arabidopsis thaliana (L.) Heynh. is a photomorphogenic response mediated by the far-red-absorbing forms of phytochrome A and B. Planta 199: 511–514. Robson, P.R.H., and H. Smith. 1996. Genetic and transgenic evidence that phytochromes A and B act to modulate the gravitropic orientation of Arabidopsis thaliana hypocotyls. Plant Physiol. 110: 211–216. Sakai, T., T. Kagawa, M. Kasahara, T.E. Swartz, J.M. Christie, W.R. Briggs, M. Wada, and K. Okada. 2001. Arabidopsis nph1 and npl1: blue light receptors that mediate both phototropism and chloroplast relocation. Proc. Nat. Acad. Sci. (USA) 98: 6969–6974. Sakai, T., T. Wada, S. Ishiguro, and K. Okada. 2000. RPT2: a signal transducer of the phototropic response in Arabidopsis. Plant Cell 12: 225–236. Sakamoto, K., and W.R. Briggs. 2002. Cellular and subcellular localization of phototropin 1. Plant Cell 14: 1723–1735. Schepens, I., P. Duek, and C. Fankhauser. 2004. Phytochrome-mediated light signalling in Arabidopsis. Curr. Opin. Plant Biol. 7: 564–569. Schwartz, A., and D. Koller. 1978. Phototropic response to vectorial light in leaves of Lavatera cretica L. Plant Physiol. 61: 924–918. Shackel, K.A., and A.E. Hall. 1979. Reversible leaflet movements in relation to drought adaptation of cowpeas, Vigna unguiculata (L.) Walp. Aust. J. Plant Physiol. 6: 265–276. Sharrock, R.A., and T. Clack. 2002. Patterns of expression and normalized levels of the five Arabidopsis phytochromes. Plant Physiol. 130: 442–456. Stankovic, B., D. Volkmann, and F.D. Sack. 1998. Autotropism, automorphogenesis, and gravity. Physiol. Plant. 102: 328–335. Strong, D.R., and T.S. Ray. 1975. Host tree location behavior of a tropical vine (Monstera gigantea) by skototropism. Science 190: 804–806. Suetsugu, N., F. Mittmann, G. Wagner, J. Hughes, and M. Wada. 2005. A chimeric photoreceptor gene, NEOCHROME, has arisen twice during plant evolution. Proc. Nat. Acad. Sci. (USA) 102: 13705–13709. Tarui, Y., and M. Iino. 1999. Gravitropism and phototropism of oat coleoptiles: post-tropic autostraightening and tissue shrinking during tropism. Adv. Space Res. 24: 743–753.

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5

Touch Sensing and Thigmotropism Gabriele B. Monshausen, Sarah J. Swanson, and Simon Gilroy*

5.1 Introduction The sessile nature of the plant lifestyle requires exquisite sensitivity to environmental signals. These stimuli provide the information that controls much of plant behavior, ranging from decisions about when to grow or reproduce to whether to mount a defense response against a pathogen. The directional cue of gravity is clearly a central component of this wealth of information that regulates normal plant development. However, other mechanical stimuli, ranging from the buffeting by wind and rain, impedance of the soil, and even the weight of an organ itself provide similarly important information that governs plant morphogenesis (Figure 5.1). Indeed, plants are highly sensitive to mechanical cues in their environment. This is perhaps most obvious when seeing the rapid mechanoresponse in organs specialized for touch sensing, such as the closing of the Venus fly trap (Dionaea muscipula) triggered by the mechanical stimulus of an insect alighting on the leaf, or the twining of a tendril in response to the touch signal from contacting a support. However, plants exhibit many and varied responses to touch. In general, plants grown with mechanical stimulation develop a shorter stature, more robust and stronger support tissues, and altered organ architecture and growth habit (Braam 2005 and references therein). Similarly, the directional cues offered by mechanical stimulation (be it touch or gravity) lead to highly controlled directional growth responses manifested as thigmo- and gravitropism. Considering the physical nature of both the gravity and touch stimulus, it seems likely that they share common mechanotransduction elements. It has even been proposed that gravity sensing is derived from an ancestral touch perception apparatus (Trewavas and Knight 1994). In this chapter we will therefore describe some of the broad classes of mechanoresponse seen in plants, discuss some of the models for how mechanosensing is likely operating at a cell and molecular level, and then ask how the plant integrates multiple stimuli, in this case touch and gravity, to generate the appropriate tropic response. 5.2 Plant Mechanoresponses Plant mechanoresponses can be divided into two broad categories: those associated with highly specialized mechanosensory organs such as tendrils or the traps of carnivorous plants, and those reflecting a more ubiquitous mechanosensory system that seems to affect most parts of the plant. The specialized touch sensory systems have clearly evolved to trigger a single specialized response, such as the twining of a tendril. The more “general” touch sensitivity, however, may well relate to the systems required to monitor and control the mechanical stresses inherent in normal turgor-driven cell expansion. *Corresponding author 91

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Figure 5.1. Plants experience a complex mechanical environment that differs significantly above- and belowground. The root system tends to be supported in the soil to offset the effects of the weight of the organs, whereas aerial parts of the plant must support their own weight. The effects of incidental contact with animals, herbivory, cultivation, and obstacles to the current direction of growth are experienced throughout the mature plant. The effects of soil impedance are limited to roots and the seedling shoot prior to emergence from the soil. The direct effects of wind and rain are limited to the aerial parts of the plant. The mechanical stimulus from all these factors has been shown to significantly affect plant form and to have important real-world implications, for example, placing limits on agricultural productivity (Mitchell 1996; Zheng et al. 2000).

5.2.1 Specialized Touch Responses Many plants have highly specialized mechanosensory structures capable of remarkably sensitive and selective mechanical signaling. Carnivorous plants show some of the most dramatic examples of this class of touch responses. In the Venus fly trap, leaves are modified into a bilobed trap that closes in less than a second to capture insects for subsequent digestion (Darwin 1893; Simons 1992). This is a thigmonastic response in that the direction of the response is inherent to the structure of the organ and is not entrained by the direction of the touch stimulus. It is thought that the trap closes by rapid growth at the

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hinge region between the lobes (Fagerberg and Allain 1991), rapid turgor loss in motor cells in this region (Hill and Findlay 1981), and an inherent tendency of the lobes to “snap” shut due to the elasticity and geometry of the trap itself (Forterre et al. 2005). The sensor that triggers this closure is a series of modified hairs on the inner trap surface. Activating the trap requires multiple stimulations on these hairs as the insect climbs across the leaf. The requirement for several signals is likely a safeguard to prevent closure of the trap by random mechanical stimuli, such as the impact of a raindrop, which tend to be solitary events. The multiple stimulations then trigger action potentials that are transmitted to the hinge region to effect closure (Jacobsen 1965; Simons 1981). The continued struggling of the insect, likely supplemented with some chemical sensory events, then closes the trap even tighter, forming a chamber in which the insect is digested (Fagerberg and Allain 1991). Similar mechanosensory triggers are seen in other carnivorous plants, such as the suction trap of the bladderworts (Utricularia) and the leafrolling sticky trap of the sundews (Drosera) (Darwin 1893; Lloyd 1942). In this latter case the mechanosensor is responsive to microgram stimuli, yet is able to ignore the presumably much larger mechanical signals from the impact of rain or wind (Darwin and Darwin 1880; Darwin 1893). Although these plants have received much attention due to their dramatic carnivorous responses, we are largely ignorant of how their mechanosensors operate. The rapidity of triggering the response seen in Dionaea and Utricularia (which responds in the millisecond range) strongly suggests the involvement of a mechanosensory ion channel. Indeed, the inherent speed and signal amplification of channels makes them the top candidates for most rapid mechanosensory responses (Gillespie and Walker 2001, and see below). However, at present the molecular identity of plant mechanosensors remains unknown. Identification of this initial signaling system will be an important step toward answering the many perplexing questions raised by the touch response in these carnivorous plants. For example, how does Dionaea suppress its response until multiple touch signals have been received, and how can Drosera tell the difference between an insect and a raindrop impacting on the leaf? A similarly highly specialized touch response is seen in the thigmonastic leaf movements of the sensitive plant (Mimosa pudica) and its relatives. In these plants touch stimulation leads to rapid folding of the stimulated leaflet, possibly as an anti-herbivory/ defense response (Simons 1981). This collapse of the leaflets propagates along the leaf and, depending on the strength of stimulation, may lead to folding of the petiole and even induce a response in adjacent leaves. Similar to the carnivorous plants, the mechanisms driving the movements have been described but the nature of the mechanosensor or early signaling events remain undetermined. Thus, electrical signals, either a propagating action potential or a slow wave potential (Simons 1981; Fromm and Eschrich 1988; FleuratLessard et al. 1997), perhaps coupled to changes in hydraulic pressure in the vasculature (Malone 1994) and a chemical messenger (Schildknecht and Meier-Augenstein 1990; Varin et al. 1997), likely trigger the local and systemic movements. These signals then elicit ion efflux and loss of turgor in motor organs (pulvini) located at the base of each leaflet and the petiole (Simons 1981; Fromm and Eschrich 1988). The possibility that plants measure xylem pressure as a systemic signal remains an intriguing idea potentially allowing rapid communication throughout the plant via mechanosensors located around

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the vasculature. Again, we are still ignorant of the identity of the initial mechanosensor and associated signaling events that should be coupled to such a hydraulic signaling system. One general theme emerging from all these studies on thigmonastic responses is that the most rapid changes monitored upon mechanical stimulation are generally the electrical changes thought to transmit the touch information to the motor response. These electrical changes reflect ion transport phenomena at the plasma membrane, highlighting the central role of ionic signaling in the early phase of a plant mechanoresponse. 5.2.2 Thigmomorphogenesis and Thigmotropism Even in plants with no obvious specialized mechanosensory structures, mechanical perturbation leads to morphogenetic changes that are highly adaptive. Thus, in shoots, mechanical stimulation causes inhibition of elongation and radial swelling (Biddington 1986; Telewski and Jaffe 1986). Such changes can be local; for example, trees will strengthen regions of the trunk or branches undergoing compression or tension with the production of reaction wood (compression and tension wood; Hellgren et al. 2004). These modified areas are rich in cells exhibiting strengthened, often highly lignified cell walls to reinforce mechanically stressed tissues. However, changes in growth habit upon localized mechanical perturbation can also be systemic (Biddington 1986; Coutand et al. 2000), suggesting that a mobile signal is integrating the overall thigmomorphogenetic response. Although mechanosensitive channels, Ca2+-dependent signaling cascades (see below), ethylene, and perhaps GA (Mitchell 1996) are strong candidates for components of the mechanosensory signal transduction system regulating and integrating these changes in growth, we still have remarkably little molecular data on how these thigmomorphogenetic responses occur. In addition to this general change in form upon mechanical stimulation, plants also show highly oriented changes in growth where the direction of response is determined by the direction of the stimulus (i.e., a thigmotropic response). Thigmotropism can be seen in many parts of the plant but is perhaps most familiar in the specialized touchresponsive organs known as tendrils. Many plants use leaves or shoots modified into tendrils to secure themselves to supports to allow increased height without the need for extensive deposition of metabolically expensive strengthening agents such as lignin. These tendrils are highly touch-sensitive, responding to stimuli of as little as 250 µg (Simons 1992). Upon sustained mechanical stimulation they rapidly (often within seconds) begin to exhibit differential growth across the organ, leading to coiling around the contacted object (Jaffe and Galston 1968). In Bryonia dioica, octadecanoids and auxin seem to regulate this cell expansion (Stelmach et al. 1999). The direction of coiling is often determined by the direction of the mechanical stimulus, leading to a thigmotropic response of the organ. However, thigmotropism is not limited to such highly specialized touchsensitive organs. For example, thigmotropism is also exhibited by roots growing into obstacles in the soil, a response we will discuss in more detail later in this chapter. Again, although there is an extensive literature describing the physiology and development of the growth response in tendrils and roots, we are largely ignorant of the initial mechanosensory events making these organs so responsive to touch stimulation. However, there are many clues from other organisms as to the basic features that we should

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expect from such a sensor. Therefore, in the next section we will discuss the principles of mechanoperception that can be derived from the touch sensors seen across the kingdoms and ask how well the molecular insight gained from these animals and bacteria might translate to identification of a plant mechano- or gravisensory system.

5.3 General Principles of Touch Perception Any protein embedded in a membrane experiences mechanical force exerted by the lipid bilayer. All bilayers have a characteristic lateral pressure profile, with outward (positive) directed pressure in the hydrophobic core of the membrane and large tension at the membrane–water interfaces, though the specific distribution and magnitude of pressures vary considerably with lipid composition (Poolman et al. 2004) (Figure 5.2). In the resting state, a transmembrane protein adopts a conformation that is at equilibrium with the surrounding mechanical forces of the lipid bilayer. However, when the membrane is stretched or deformed by a mechanical stimulus, the distribution of forces is altered and the equilibrium perturbed. In a mechanosensitive protein, this change in force is thought to trigger a conformational change, which results in activation (or deactivation) of the protein (Hamill and Martinac 2001; Janmey and Weitz 2004; Kung 2005) (Figure 5.2). Though in-plane membrane forces may conceivably activate any number of transmembrane proteins, so far only the mechanosensitive ion channels of bacteria and the TRPC1 channel of Xenopus oocytes have been shown to be directly gated by membrane tension (Maroto et al. 2005; Moe and Blount 2005) (see below). However, several other candidate mechanosensitive ion channels from animals and yeast have recently been identified (TRPY1, TREK1 and 2) (Bang et al. 2000; Chemin et al. 2005; Zhou et al. 2005). How relevant is this model of mechanoperception for plant cells where the flexion of the plasma membrane must go hand-in-hand with the deformation of the cell wall? The cell wall is under hydrostatic pressure from the protoplast, so that cell wall deformation would require the application of an external force large enough to overcome turgor pressure. Given that turgor is in the range of 1 to 40 bars (Tomos and Leigh 1999; Franks 2003), it may at first glance seem that only extreme forms of mechanical stimulation would provide sufficient force to be detected via changes in membrane tension. However, even subtle stresses such as a gentle breeze can make leaves or stems sway and such organ bending can only occur when at least a subset of cells change shape (i.e., when the cell wall and plasma membrane are deformed). This is possible because parts of the plant act as levers. In this way, even very moderate mechanical forces can be sufficiently amplified and focused onto the responding cells to exceed turgor pressure, flex the plasma membrane, and thus directly gate transmembrane proteins via changes in lipid force distribution. A mechanosensitive protein can also be displaced within the pressure profile of the membrane by tethering it to force-transmitting elements of the cytoskeleton or the extracellular matrix (ECM). When these components are deformed during a mechanical stimulation, they could pull on the attached transmembrane protein and move it relative to the lipid bilayer (elevator model) (Kung 2005; Orr et al. 2006). Although the physical repositioning of the protein is thus achieved via a different mechanism than that described

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Figure 5.2. Forces exerted through the membrane can gate ion channels. A. The force distribution throughout the depth of the membrane leads to a steep gradient in gating force on the interface between the channel protein and the lipid head groups. Membrane tension can favor channel opening if the cross-sectional area of the open state of a channel is larger than the closed (B), or if membrane tension leads to thinning of the bilayer and the conformational change leading to the open state of the channel reduces the profile of the membrane-spanning, hydrophobic residues on the channel surface (C). Redrawn from Kung (2005).

above, the end result—the altered force distribution at the protein–lipid interface and ensuing activation of the mechanosensitive protein—is the same in both models (Kung 2005). According to another model, tether-mediated activation of membrane proteins occurs when stress-deformed cytoskeletal/ECM components pull on the protein and thereby directly change the protein conformation [e.g., by shifting an autoinhibitory domain which releases block and triggers activation (trapdoor model) (Hamill and Martinac 2001; Kung 2005)] (Figure 5.3) or by partially unfolding the attached protein and thus revealing previously hidden catalytic/binding sites (Janmey and Weitz 2004; Vogel 2006). Several mechanosensitive ion channels of vertebrate and invertebrate organisms are proposed to

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Figure 5.3. Mechanical force may elicit biochemical changes through inducing conformational changes in target proteins. Gating may be directly, through force transmissive linkages (e.g., in the “trap door” model where mechanical force directly opens a gating domain in the channel) (a), or more indirectly though other signaling proteins or cytoskeletal interactions. Shear at the membrane surface may drag channels relative to their tethers, with the stretching of the tether providing the mechanical gating force (b). Similarly, a separate mechanoresponsive protein may trigger conformational change and gating in the channel (c). Mechanotransduction does not a priori require channels; for example, a mechanosensor may induce structural changes in proteins bringing enzyme and substrate into close proximity (d), or revealing binding sites on target transduction elements. Force may also act to unravel a protein, domain-by-domain, revealing new activation or binding sites (e).

be activated by mechanical force transmitted through their linkage to the cytoskeleton (e.g., TRPA1) (Lin and Corey 2005) and/or the ECM (e.g., MEC channel complex; see below), though the precise mechanism of gating is not yet fully understood. In plant cells, any force large enough to overcome turgor and flex load-bearing elements of the cell wall and alter cellular shape will also deform the cytoskeleton. However, some plant organs are known to be extraordinarily sensitive to mechanical stimuli, responding to as little as a 10-µN weight (~1 mg) with altered growth (Jaffe and Galston 1968). Such weak forces are unlikely to change cell shape but may conceivably deform non-load-bearing elements embedded in the wall matrix, such as cell wall proteins. If these elements are linked via transmembrane proteins to the cytoskeleton, force may be transmitted from the cell exterior not only to plasma membrane mechanosensors, but— because the cytoskeleton is ideally suited to long-distance transfer of stresses—to mechanosensitive proteins in endomembranes as well (Ingber 2003, 2006). Most research has focused on ion channels as the principal transducers/sensors of mechanical stress because of their capacity to rapidly amplify a signal. Such amplification occurs because a single channel can transport millions of ions in seconds, altering the electrical (membrane potential) and chemical (ion concentration) cellular environment. However, there is increasing evidence for a role of nonchannel proteins in mechanosens-

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ing. Thus, the cytoskeleton may not only act to convey forces to interior regions of the cell by tugging on an attached mechanosensor, but may itself translate stress information into biochemical reactions via altered binding to associated proteins (Figure 5.3d). For example, specific cytoplasmic proteins show differential binding to the cytoskeleton dependent on the state of cytoskeletal tension (Sawada and Sheetz 2002; Tamada et al. 2004). Similarly, stretching of the cytoskeleton promotes activation of a tyrosine kinase (Scr) via interaction with an actin-binding protein (AFAP) (Han et al. 2004). Straininduced alterations in binding affinity of cytoskeletal proteins are thought to be mediated via conformational changes, especially the unfolding of protein domains and resulting exposure of cryptic binding (or catalytic) sites. Indeed, mechanically induced partial unravelling of proteins is a potentially widespread signalling mechanism. By using force spectroscopy to investigate the mechanical properties of multimodular proteins, such as the giant muscle protein titin and the ECM adhesion protein fibronectin, it was shown that, when exposed to increasing levels of tension, the modules unfolded one-by-one in a sequence determined by the mechanical stability of each domain (Zhuang and Rief 2003; Vogel 2006) (see Figure 5.3). The stability should be dictated by features such as disulfide and hydrogen bonds that determine the secondary structure of the protein. These features will in turn be modulated by cellular environment such as pH and ionic strength and local redox potential (Vogel 2006). Thus, the magnitude of stress experienced by the protein may well be encoded in the degree of protein unfolding and signalled to the cell by way of revealing different recognition (binding or catalytic) sites in each unravelled module (Vogel 2006). This is also an intriguing possibility for plant cells. If deformation of cell wall components results in exposure of new binding sites, interactions with plasma membrane receptors may be newly formed or broken to trigger signalling to the cell interior. In addition to such direct effects on protein conformation, two other themes that emerge from this overview of mechanosensory channel function are that gating through tension in the lipid bilayer and through tethering to either the ECM and/or the cytoskeleton are the most prominent modes regulating the opening and closing of mechanosensory channels. We will therefore describe in more detail two of the most intensively studied channel types, the MscL channels of Escherichia Coli and the MEC channels of Caenorhabditis elegans, to explore the molecular mechanisms that underlie each of these modes of channel regulation. These channels may well provide clues to the structure of the elusive plant mechanosensory complex. 5.3.1 Gating through Membrane Tension: The Mechanoreceptor for Hypo-osmotic Stress in Bacteria, MscL Free-living bacteria are exposed to very sudden changes in the osmolarity of their environment. When a sudden drop in external osmolarity results in rapid influx of water and threatens to cause lytic rupture of the cell, mechanically gated channels act as emergency valves. Two of these channels, named mechanosensitive channels of small (MscS) and large (MscL) conductance, have been characterized to be activated by increasing membrane tension and enable the cell to rapidly jettison large amounts of osmolytes (Perozo and Rees 2003).

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The MscL of E. coli is perhaps the best understood of these mechanosensors. Mechanically gated MscL activity was first observed in patch clamp experiments conducted on giant E. coli spheroplasts, and subsequent purification of MscL and reconstitution in artificial lipid bilayers demonstrated that the channel retained its characteristic mechanosensitive conductance (~3 nS) and gating kinetics even in a cell-free environment (Sukharev et al. 1993; Sukharev et al. 1994). These findings convincingly established that changes in membrane tension alone can be sufficient to activate a mechanosensitive ion channel. Sequence analysis revealed that the E. coli (Eco-)MscL encodes a 136 amino acid membrane protein with cytoplasmic N- and C-terminal regions and two ␣-helical transmembrane domains (designated TM1 and TM2) connected by a periplasmic loop (Blount et al. 1996). Determination of the crystal structure of a homolog of Eco-MscL, the MscL of Myobacterium tuberculosis (Chang et al. 1998), showed that five MscL subunits form a homopentamer, with TM1 domains forming the funnel-shaped permeation pathway and TM2 helices interacting with the lipid bilayer and surrounding the central barrel of TM1 domains (Figure 5.4a). On the cytoplasmic side of the channel, the five C-terminal helices assemble into a bundle not required for gating but presumed to act as a sizeexclusion filter (Anishkin et al. 2003). Though unresolved in the crystal structure, the five amphiphilic N-terminal (S1) domains were predicted to organize as ␣-helices, interacting to form another bundle situated between the channel pore and the C-terminal “hanging basket” (Sukharev et al. 2001b; Sukharev et al. 2001a). How do these different channel domains contribute to the mechanical gating of MscL? According to a current model based on computer simulations and experiments using cysteine substitutions to stabilize specific domain interactions, the channel is activated by the sequential tension-induced opening of two gates (Sukharev et al. 2001b). Upon stretch of the membrane, the channel pore increases in diameter from ~0.2 nm to about 3.5 nm (Doyle 2004). This dramatic change in channel conformation is thought to occur as the transmembrane domains TM1 and TM2 undergo significant tilting, thereby swinging away from the central channel axis. This iris-like expansion is considered the initial gating event because it would open the hydrophobic constriction at the narrow end of the funnel-shaped TM1 pore acting as a barrier to ion permeation. However, removal of this constriction is not sufficient to elicit full conductance of the channel. In fact, the initial conformational change only seems to draw the channel into a low subconducting state (Anishkin et al. 2005) (Figure 5.4a and Color Section). Further extension is required to activate the channel completely. The transition to the fully open state is thought to depend on the disruption of the N-terminal bundle connected to the TM1 domains via linker regions. Only when the TM barrel fully expands at close to lytic tensions is force transmitted via the linkers to this bundle, pulling it apart and thereby opening the second gate to release huge amounts of ions and small solutes (Sukharev et al. 2001b). 5.3.2. Gating through Tethers: The Mechanoreceptor for Gentle Touch in Caenorhabditis elegans The nematode C. elegans responds to gentle touch (as little as 10-µN force) with an avoidance reaction, moving backward when the stimulus is applied to the head and mov-

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Figure 5.4 (also see Color Section). Structures of mechanosensitive channels gated by membrane tension (MscL) and tethering to the cytoskeleton and extracellular matrix (Mec). In the bacterial MscL channel (a), five subunits form the channel, each contributing two transmembrane domains (green). In the closed state, the S1 domain (red) sits close to the inner face of the pore. When the membrane experiences tension, the transmembrane helices tilt and the pore expands in an iris-like fashion. S1 is drawn closer to the membrane, partially occluding the pore. As the channel fully opens, the S1 domains are dispersed, leaving the pore unobstructed. In the Mec channel of C. elegans (b), the Mec-4 and Mec-6 subunits form the conducting pore, which is gated through a complex of proteins that interact with the extracellular matrix and the cytoskeleton. Structures in (a) were rendered using MMDB from NCBI (Chen et al. 2003), and models of MscL opening and Mec channel structure (b) were redrawn from www.life.umd.edu/biology/faculty/sukharev and O’Hagan et al. (2005) respectively.

ing forward when the tail is touched. Laser ablation experiments have revealed that six touch receptor neurons are responsible for sensing these mechanical stimuli (Chalfie et al. 1985). These six neurons have processes that lie embedded in the hypodermis near the cuticle of the worm and seem to be attached to the hypodermis via extensive connections to the ECM (Garcia-Anoveros and Corey 1997). By patch-clamping these neurons in vivo while gently touching the nematode cuticle,

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O’Hagan et al. (2005) were able to show that mechanical stimulation triggers a rapidly activating mechanoreceptor current that is carried mostly by Na+ and depolarizes the plasma membrane. The membrane depolarization is then thought to elicit an increase in cytosolic Ca2+ by activating voltage-sensitive L-type Ca2+ channels (Suzuki et al. 2003), highlighting that a mechanosensory channel need not be directly Ca2+-permeable to elicit the touchinduced Ca2+ increases proposed to transduce the mechanical signal in plants (see below). Saturating mutagenesis analysis has identified 18 genes required for this C. elegans mechanosensitivity (named mechanosensory abnormal, or MEC proteins; Ernstrom and Chalfie 2002), some of which encode transcription factors, cytoskeletal proteins, or proteins of the ECM (Tavernarakis and Driscoll 1997). At least four of the MEC proteins (MEC-2, -4, -6, and -10) form the channel complex producing the mechanoreceptor current (O’Hagan et al. 2005) (Figure 5.4b and Color Section). MEC-4 and MEC-10 are poreforming channel subunits and belong to the DEG/ENaC (degenerin/epithelial sodium channel) family of amiloride-sensitive Na+-conducting channels found exclusively in animals (Lai et al. 1996; Goodman et al. 2002; Kellenberger and Schild 2002). MEC-6 is a singlepass transmembrane protein with a cytoplasmic N-terminus and large extracellular Cterminal domain (Chelur et al. 2002), whereas MEC-2, a monotopic protein with a stomatinlike region, does not span the plasma membrane but is associated with the cytoplasmic side of the lipid bilayer (Goodman et al. 2002). Both MEC-2 and MEC-6 interact with MEC-4 and MEC-10 to regulate channel conductance (Chelur et al. 2002; Goodman et al. 2002). Interestingly, no mechanically induced activation of current was observed when the wild-type channel complex was heterologously expressed in Xenopus oocytes (Goodman et al. 2002). This observation suggests that mechanical gating is not accomplished by changes in membrane tension alone, but requires additional force-transmitting elements such as proteins of the ECM and the microtubule cytoskeleton physically tethered to the channel complex. It has been proposed that the touch-induced movement of the two putative tethering sites (ECM-channel extracellularly and microtubule-channel intracellularly) relative to each other provides gating tension and activates the channel (reviewed by Tavernarakis and Driscoll 1997). Indeed, both the pore-forming channel subunits MEC-4 and MEC-10 as well as MEC-6 have large extracellular regions thought to be important for interaction with the ECM proteins MEC-1, MEC-5, and MEC-9 (Chelur et al. 2002; Emtage et al. 2004). The precise role of the prominent microtubule cytoskeleton in mechanosensation is less clear. Mutations in the genes MEC-7 and MEC-12 cause loss of mechanoresponse. These genes encode ß- and ␣-tubulins, respectively, which are required to form microtubule protofilaments. These microtubules are proposed to be tethered to the channel complex via MEC-2 (Tavernarakis and Driscoll 1997) (Figure 5.4b). However, mutants in MEC-7 still show (attenuated) touch-triggered mechanoreceptor currents, suggesting that direct linkage of the channel complex to microtubules is not an absolute requirement for gating (O’Hagan et al. 2005). 5.3.3 Evidence for Mechanically Gated Ion Channels in Plants Although no plant mechanosensor has been unequivocally identified at the molecular level, there is strong evidence for mechanically gated ion channels in the plasma mem-

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brane from a variety of cell types derived from different tissues in different plant species. Applied pressures of 1 to 10 kPa have been shown to trigger activation of channel currents in Arabidopsis, Allium, Vicia, Valonia, Lilium, and others. Apart from this common mechanism of gating, however, the observed channel activities are quite diverse with regard to selectivity, conductance, or voltage dependence. Some channels are unselective (Spalding and Goldsmith 1993) and others have been demonstrated to show preference for anions (Falke et al. 1988; Cosgrove and Hedrich 1991; Heidecker et al. 1999; Qi et al. 2004) or cations (Garrill et al. 1994), whereas certain channels show high selectivity for K+ or Ca2+ (Cosgrove and Hedrich 1991; Ding and Pickard 1993; Liu and Luan 1998; Dutta and Robinson 2004). Measured conductances range from 3 to 15 pS for Ca2+selective channels (Cosgrove and Hedrich 1991; Ding and Pickard 1993; Dutta and Robinson 2004) to 13 to 97 pS for anion channels (Falke et al. 1988; Cosgrove and Hedrich 1991). Although most channels are seemingly independent of membrane potential (Falke et al. 1988; Garrill et al. 1994; Liu and Luan 1998; Heidecker et al. 1999; Dutta and Robinson 2004; Qi et al. 2004), others show significantly reduced open probabilities at either negative or positive voltages (Cosgrove and Hedrich 1991; Spalding and Goldsmith 1993), whereas some are active only at higher voltages, irrespective of the polarity (Ding and Pickard 1993). Unfortunately, most of these channels have not been analyzed beyond the initial characterization. Thus, even though the molecular identity remains elusive, information on how the regulation of these mechanosensors are modulated by other factors such as pH, lipid environment, or the cytoskeleton could greatly enhance our understanding of how plant cells tune their mechanosensitivity during differentiation or in response to changing environmental conditions. Despite all the differences shown by the plant mechanosensitive conductances described above, one interesting common feature of all these channels is that their activation is feasible in excised membrane patches devoid of cell wall or intact cytoskeleton. This observation suggests that plant mechanosensory channels are sensitive to changes in lipid bilayer tension and their gating does not absolutely depend on tethering to force transmitting elements, though both cell wall and cytoskeleton may have a role in modulating membrane tension in vivo. The pressures used to open mechnosensitive conductances in all these plant experiments are also comparable to those that gate the bacterial MscL and MscS channels, consistent with a role for membrane tension in their regulation. Indeed, there are homologs of the MscS channels in plant genomes with, for example, 10 MscS-like (MSL) proteins in Arabidopsis and 6 in rice (Haswell and Meyerowitz 2006). The MSLs represent very strong candidates for mechanosensitive channels in plants, but to date there is no direct electrophysiological evidence that these proteins are indeed channels and show mechanosensitivity. In Arabidopsis, MSL2 and 3 are localized to plastids where they play a redundant role in plastid division (Haswell and Meyerowitz 2006), perhaps reflecting conservation of these channels from the endosymbiotic origin of the plastid. The other MSLs of Arabidopsis localize to other regions of the cell, but we must await analysis of their roles at the genetic and electrophysiological levels to know whether they represent the elusive plant plasma membrane mechanosensory channel.

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5.4 Signal Transduction in Touch and Gravity Perception 5.4.1 Ionic Signaling From the above discussion, it is clear that a major theme emerging from our current understanding of mechanosensors in animals and microbes is that channels and ionic fluxes play a central role in the transduction of external force to intracellular biochemical signal. There is also an increasing body of evidence that ionic signals are intimately associated with the touch and gravity responses in plants. Principal among these mechanically related signals are changes in Ca2+ and pH.

5.4.2. Ca2+ Signaling in the Touch and Gravity Response Changes in the levels of cytosolic Ca2+ are recognized as a ubiquitous regulatory system and are also among the most widely reported initial changes in response to mechanostimulation (Gillespie and Walker 2001). For plants there is a wide body of literature supporting the idea that mechanical stimuli, ranging from wind and rain to localized mechanostimulation of a single cell, elicit complex patterns of Ca2+ change. Most of these measurements have been made noninvasively using transgenic plants expressing the luminescent Ca2+-sensitive protein aequorin. Such analyses confirm touch-related Ca2+ increases in plants ranging from Arabidopsis and tobacco (Knight et al. 1991; Knight et al. 1992; Knight et al. 1993; Plieth and Trewavas 2002) to the moss Physcomitrella patens (Haley et al. 1995; Russell et al. 1996) and even the characean algae (Blancaflor and Gilroy, unpublished), suggesting the presence of an ancient mechanosensory system. Ca2+ chelation attenuates the stunting of growth associated with thigmomorphogenesis (Jones and Mitchell 1989), tentatively suggesting a functional role for such Ca2+ changes. The site of these Ca2+ fluxes (trans-plasma membrane versus release from intracellular stores) remains to be unequivocally determined. Thus, Ca2+ chelators and channel blockers that should inhibit influx at the plasma membrane have been reported to either block or fail to affect touch-induced Ca2+ increases (Haley et al. 1995; Legue et al. 1997). Further, evidence for release from intracellular stores comes largely from the ability of ruthenium red to inhibit Ca2+ changes (Knight et al. 1992; Legue et al. 1997). Ruthenium red is thought to block channel-mediated Ca2+ release from mitochondria and the ER (Denton et al. 1980; Campbell 1983) but its action on Ca2+ channels in plants is not well characterized. Ruthenium red is, however, known to affect other plant processes; for example, it binds strongly to unesterified pectins (Moffatt et al. 2002). In addition, in some reports (Haley et al. 1995) no effect of ruthenium red was observed on the mechanically induced Ca2+ transients. Despite this uncertainty as to its precise source, the mechanically induced Ca2+ increase has been confirmed in plants using sensors other than aequorin. Thus, plants expressing the Ca2+-sensitive, green fluorescent, protein-based sensor cameleon (Allen et al. 1999) show mechanically induced Ca2+ transients (Figure 5.5 and Color Section), as do plants loaded with the Ca2+-sensitive dye Indo-1 (Legue et al. 1997). In this latter study, the apical cells of the root cap were found to be approximately twice as sensitive to touch stimulation as those more basal to the tip, suggesting that touch sensitivity may vary dependent on cell

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Figure 5.5. Touch- and cold-induced calcium signatures in root cap cells. a. An outline of the cells in an Arabidopsis root tip; box indicates area of the root cap observed in (b) and (c). b. Changes in cytoplasmic calcium in a root cap peripheral cell following a touch stimulus. c. Changes in cytoplasmic calcium in the root cap following cold shock. In both (b) and (c), cells were expressing YC 3.6, a GFP-based calcium indicator, and fluorescence was monitored using a Zeiss 510 confocal microscope. Scale bars = 10 microns.

type. The high sensitivity of the surface cells of the root cap fits well with their position, being the first cells to encounter obstacles as the root forces itself through the soil. It is important to note that many other stimuli have been shown to elicit increases in intracellular Ca2+, ranging from cold shock to symbiont elicitors (reviewed in Hetherington and Brownlee 2004). These Ca2+ changes often occur in exactly the same cell as touch-induced Ca2+ transients (Figure 5.5). One possible explanation for this response to multiple stimuli is that Ca2+ is simply acting as an “on switch” informing the cell that something has happened, with the specificity of the response being encoded by other signaling systems (Scrase-Field and Knight 2003; Plieth 2005). Alternatively, the spatial and/or temporal footprint of the Ca2+ change (the so-called Ca2+ signature) may encode information about the stimulus evoking the response. There has long been literature supporting the informational content of Ca2+ changes in animal cells (Dolmetsch et al. 1997, 1998) and there is some evidence supporting a similar phenomenon in plants. For example, in stomatal guard cells, Ca2+ spiking is seen in response to stimuli that induce closure of the stomatal aperture (McAinsh et al. 1995; Allen et al. 1999). The frequency of spiking appears to be critical, with Ca2+ transients that occur either too frequently or too slowly being less effective in triggering the response (Allen et al. 2001). These observations suggest that, at least in the guard cell, the temporal character of a Ca2+ increase is important for the response that is elicited. Similarly, in the touch response the magnitude of Ca2+ increase has been reported to correlate with the magnitude of mechanical stimulation (Haley et al. 1995), consistent with the idea that the Ca2+ change could be carrying information about the kind of mechanical stimulation that the cell is experiencing. However, we clearly need more detailed analysis to distinguish a role for Ca2+ in the “signature” versus “on switch” modes of action in plants in general and the mechanoresponse in particular. A role for Ca2+ in signaling touch response appears to extend to the specialized touchsensitive systems such as the tendril described at the beginning of this chapter. For example, a Gd3+-sensitive, voltage-dependent Ca2+ release channel (BCC1) has been electrophysiologically identified in ER isolated from the tendrils of Bryonia dioica (Klusener et

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al. 1995). Touch-induced tendril coiling can be inhibited in this system by Gd3+ and erythrosine B, a putative inhibitor of the Ca2+-ATPases that pump Ca2+ into the ER. Thus, there is some tentative evidence placing a Ca2+ release channel involved in mechanoresponse on an intracellular membrane in this system. Interestingly, BCC1 may also be regulated by cytosolic pH and levels of reactive oxygen species (ROS) (Klusener et al. 1997). Considering the proposed roles for these two agents in response to mechanical perturbation and gravistimulation (see below), BCC1 may be hinting at one mechanism whereby gravity and touch signaling may modulate each other. Although the link between touch and Ca2+ is supported by a wealth of experimental data, an equivalent connection between Ca2+ and gravity signaling is less clear. There is much circumstantial evidence linking Ca2+ to the gravity response. For example, application of Ca2+ chelators such as EGTA and BAPTA abolishes the growth component of the graviresponse (Lee et al. 1983a, 1983b; Björkman and Cleland 1991) and the auxin fluxes that accompany tropic curvature (Young and Evans 1994; see also Chapter 3). Similarly, a range of pharmacological agents thought to disrupt diverse aspects of Ca2+ signaling have been reported to alter gravitropism (Fasano et al. 2002; Massa et al. 2003). There is also extensive evidence implicating the Ca2+-dependent regulatory protein calmodulin (CaM) in the graviresponse. For example, CaM levels are enriched in the root tip (the site of gravity perception) (Allan and Trewavas 1985; Stinemetz et al. 1987), are enhanced upon gravistimulation (Sinclair et al. 1996), and the CaM levels in the root tip correlate with the responsiveness of the organ to gravity (Stinemetz et al. 1987). Calmodulin transcripts have also been shown to be recruited into polysomes on the lower side of the gravistimualted pulvinus (Heilmann et al. 2001), suggesting that gravistimulation should change the abundance of CaM across the stimulated organ. Calmodulin antagonists inhibit the asymmetric Ca2+ and proton fluxes associated with graviresponse (Lee et al. 1983b, 1984; Björkman and Leopold 1987) and impair gravisensing and tropic response at levels that do not inhibit growth (Stinemetz et al. 1992; Sinclair et al. 1996). Although it is always important to view such pharmacological data with caution due to unknown targets and side effects of the antagonists used, this body of data, taken with the other evidence for a role for CaM described above, does point toward an important role for this Ca2+-dependent protein in gravisignaling and response, and therefore, by implication, a role for Ca2+. The identification of a possible role for inositol-1,4,5-trisphosphate (InsP3) in gravisignaling/response is also consistent with Ca2+ playing an important role in this process. Classically, the activation of the phospholipids-cleaving enzyme phospholipase C is thought to produce the second messengers diacylglycerol and InsP3. The precise signaling role for diacylglycerol in plants is still unclear but InsP3 seems to play a similar role to its function in animal cells in triggering signaling-related Ca2+ release from intracellular stores (Wang 2004). In the graviresponsive pulvinus from maize and oat, InsP3 becomes elevated within minutes of gravistimulation, although a clear asymmetry in levels between upper and lower side of the organ takes several minutes to appear (Perera et al. 1997, 1999, 2001). Phosphatidyl-inositol-phosphate (PIP) kinase activity similarly increased, suggesting that levels of the substrate for phospholipase C (phosphatidylinositol4,5-bisphosphate) might be fuelling the elevated InsP3 levels. Perera et al. (2006) showed that InsP3 also exhibits a three-fold increase within the first 15 minutes of gravsitmula-

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tion in Arabidopsis inflorescences. They then constructed Arabidopsis plants expressing a human inositol polyphosphate 5-phosphatase that should attenuate these InsP3 levels. These plants showed 90% reduction in basal InsP3 content and a disruption of gravitropic response kinetics, further implicating InsP3, and so perhaps Ca2+ release, in gravitropic signaling/response. Despite this strong circumstantial evidence for a role of Ca2+ in gravisignaling, direct measurements for this change remain ambiguous. Although Gehring et al. (1990) reported gravistimulation-induced Ca2+ increases in maize coleoptiles, it has been difficult to unequivocally relate these changes to Ca2+ increases associated with the gravity response (Firn and Digby 1990). Legue et al. (1997), using plants loaded with the Ca2+sensing fluorescent dye Indo-1, and Sedbrook et al. (1996), using plants expressing aequorin, were unable to detect Ca2+ increases upon gravistimulation, yet Legue et al. (1997) saw clear touch-induced changes under identical conditions. In contrast, Plieth and Trewavas (2002) have reported Ca2+ increases induced by gravistimulation. These measurements were made in Arabidopsis seedlings transformed with the Ca2+ sensor aequorin and gravistimulated by rotation through 135 degrees. The kinetics of these changes were different from plants rapidly rotated through 360 degrees, which was used to provide a control for the mechanical stimulation inherent in the rotation. However, considering the exquisite sensitivity of plants to mechanical stimulation, it still remains possible that the Ca2+ increases associated with the gravistimulation were also reflecting the mechanical stimulation of the rotation to 135 degrees. These kinds of caveats about experimental results highlight the difficulties of separating touch from gravity signaling and response. This problem is not only limited to experimental design but also to the biological responses to these stimuli, a theme we will discuss in more detail in the section describing transcriptional responses to touch and gravity below. If the circumstantial evidence so strongly points to a role for Ca2+ signaling in the gravity response, why has it been so hard to clearly demonstrate the change? One possibility is suggested by the experiments of Plieth and Trewavas (2002). In order to detect Ca2+ changes upon reorientation of their aequorin-expressing plants, these researchers had to resort to making simultaneous measurements on 500 to 1,000 seedlings and reconstituting the aequorin with the most sensitive version of its cofactor known (Cpcoelentrazine). The need for such high sensitivity and numbers suggests the Ca2+ signal is either localized to a very few cells and/or localized within those sensory cells. The elevated levels of CaM and CaM-like proteins in the root cap gravisensory cells will likely sensitize them to very small changes in Ca2+ that may be at the limits of current Ca2+ detection systems. Similarly, in animal cells it is well characterized that highly localized Ca2+ fluxes, which are extremely difficult to detect, can elicit dramatic effects on cellular response. For example, in neurons Ca2+ flux specifically through L-type Ca2+ channels at the plasma membrane allows activation of MAP kinase and the CREB transcription factor cascade. Such activation is mediated by CaM tightly bound to the inner face of the pore of the channel (Dolmetsch et al. 2001). This spatially restricted signaling system means that very small Ca2+ fluxes are funneled to the appropriate signal transduction chain. In this case, large-scale changes in cytosolic Ca2+ cannot even trigger the response. Thus, it may well be that limitations on the resolution of current Ca2+ measurement technology to detect

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such localized fluxes remain the reason no Ca2+ signal has been localized to the gravisensing cells of the plant. Alternatively, the circumstantial evidence for Ca2+ signaling may be misleading and gravisignaling may reside in some other transduction pathway. Also as a note of caution, the coelentrazine cofactor for aequorin required to reconstitute the active Ca2+ sensor is itself exquisitely sensitive to ROS. In response to ROS it generates a signal similar to that expected of an increase in Ca2+ (Lucas and Solano 1992; Plieth 2005; Molecular Probes 2006). Therefore, experiments using this approach require careful controls for possibly confounding effects of ROS production. Such controls are especially important as there are reports that transient changes in ROS production are associated with the auxin fluxes generated during the graviresponse (Joo et al. 2001; Joo et al. 2005). Similarly, sustained mechanical stress induces ROS production in suspension-cultured soybean and parsley cells within 4 to 10 min (Yahraus et al. 1995; Gus-Mayer et al. 1998), suggesting that ROS may also complicate aequorin-based measurements involving mechanical stimulation. ROS are now thought to participate in many plant response systems (Mori and Schroeder 2004), and the finding that they in turn affect ROS-gated Ca2+ channels (Pei et al. 2000; Demidchik et al. 2003; Foreman et al. 2003) makes them strong candidates for regulators in mechano-/graviperception. To make the story even more complex, ROS may well be involved both at the level of intracellular signaling elements (Mori and Schroeder 2004) and also as factors modulating cell wall properties associated with growth (Campbell and Sederoff 1996; Brady and Fry 1997; Coelho et al. 2002; Kerr and Fry 2004).

5.5 Insights from Transcriptional Profiling The initial identification of touch-responsive (TCH) genes in Arabidopsis was achieved through differential cDNA screening of plants stimulated by touch or wind (Braam and Davis 1990). The identity of many of these genes as encoding Ca2+-binding proteins reinforced the theme of Ca2+-dependent signaling in the mechanical response of the plant. Thus, TCH1 encodes CaM2 (one of the Arabidopsis CaMs) and TCH2 and TCH3 encode CML24 and CML12, both CaM-like proteins (Braam and Davis 1990; Sistrunk et al. 1994; Khan et al. 1997; McCormack and Braam 2003). Similar analysis has now identified a range of genes showing mechanosensitive expression, including other CaMs (Ling et al. 1991; Perera and Zielinski 1992; Gawienowski et al. 1993; Botella and Arteca 1994; Ito et al. 1995; Botella et al. 1996; Oh et al. 1996). However, the expression levels of many non-Ca2+-related genes have also been shown to be touch-responsive. For example, TCH4 codes for a cell wall-modifying enzyme, a xyloglucan endotransglucosylase/hydrolase (Xu et al. 1995). Indeed, the genome-wide view afforded by microarray transcript profiling has revealed a remarkably rapid and widespread alteration in the spectrum of mRNA present after both touch and gravistimulation. For example, Kimbrough et al. (2004) reported approximately 1,700 transcripts changing in abundance after either mechanical or gravity stimulation of Arabidopsis roots, representing approximately 7% of the genome. Many of the transcriptional changes were found to be common to both touch and gravistimulation, perhaps reflecting the sim-

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ilarity in the growth responses these stimuli elicit and/or the multiple pathways within which each protein operates. Indeed, many of the transcripts identified as touch- and/or gravity-responsive are also known to be regulated by other stimuli such as cold, light, and pathogens. However, a small subset were found to be either touch- or gravity-specific. For example, these researchers found 65 genes that changed rapidly and selectively in response to gravistimulation, with changes in five being evident within 1 to 2 minutes (Kimbrough et al. 2004). Similarly, 26 genes showed a mechanostimulation-specific profile. These gravity- or touch-specific genes covered a wide range of functional categories, from transcription factors to transporters and wall-modifying proteins. Correspondingly similar studies profiling mechanoresponsive genes in seedlings (Moseyko et al. 2002) or aerial tissues (Lee et al. 2005) led to similar conclusions. Thus, Moseyko et al. (2002) found that of the 8,300 genes they probed, 183 were significantly altered after 30 minutes of gentle mechanical stimulation, with a significant overlap to those regulated by gravistimulation. When comparing transcriptional profiles in aerial tissues after 30 minutes of touch or darkness, Lee et al. (2005) found that although 2.5% of the total genes were touch-inducible (589 genes had touch-inducible expression; 171 had reduced expression), 53% were also altered upon transfer of seedlings to darkness. Indeed, all but 3 of the 68 genes most strongly up-regulated by darkness were also touch-inducible. Again, the range of gene functions in these groups was diverse, including putative signaling elements, wall modification, and defense responses. These widespread changes in transcription suggest an exquisitely sensitive mechanical response system that feeds into much of the physiology and developmental pathways that are shared by many other signal/response systems. It seems likely that changes in mRNA stability as well as transcriptional regulation are playing some role in governing message abundance, especially over the very short (1- to 2-min) time frames where Kimbrough et al. (2004) reported alterations in transcript level. Indeed, Gutierrez et al. (2002) found that although only approximately 1% of Arabidopsis genes have unstable transcripts, touch-induced genes were among the most highly represented group in their analysis. Unstable transcripts are thought to be the hallmark of genes requiring rapid changes in steady-state transcript abundance, consistent with the rapid and widespread transcriptional changes seen in response to touch. The alterations in the five most rapidly changing gravity-responsive transcripts were abolished in plants where InsP3 signaling is likely curtailed through ectopic expression of a human inositol 5-phosphatase (Salinas-Mondragon et al. 2005). However, the other widespread changes in gene expression induced by gravistimulation were unaffected in these plants. Thus, there may well be a functional link between rapid InsP3-dependent signaling (Perera et al. 1997, 1999, 2001) and very rapid mRNA abundance changes seen upon gravistimulation. There must also be an alternative pathway acting to modulate the expression of the large number of other responsive genes. Equivalent analysis with respect to touch-related gene expression has yet to be reported. These extensive microarray data now also offer the possibility to look for promoter elements related to responsiveness to touch and gravity. By analyzing the five most highly gravitropically up-regulated transcripts, Kimbrough et al. (2004) identified TCATTAA as a potential gravity regulation motif in the promoter sequences of these genes. However, functional testing of this possibility has yet to be reported. Similarly, a 102-base-pair region of the TCH4 promoter confers touch, dark, cold, heat, and brassinosteroid sensitiv-

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ity to expression (Iliev et al. 2002), as does a similar sequence for CBF2 (a transcriptional activator with a central role in cold response; Zarka et al. 2003). The microarray data described above now present the possibility of assessing how widespread these touch- and gravity-related transcriptional regulatory motifs might be. Such analysis might provide one avenue to start to dissect the transcriptional machinery responsible for these genomewide changes in expression in response to either gravity or mechanical perturbation. The general conclusion from all these transcriptional profiling analyses is that both mechanical- and gravistimulation induce very rapid and extensive changes in mRNA profiles. In addition, these stimuli share a huge overlap in the transcriptional changes they cause, likely reflecting the similarity in developmental responses they elicit. This is in contrast to recent work on plant hormonal regulation of growth and development where Nemhauser et al. (2006) concluded that these regulators controlled similar developmental outcomes through largely non-overlapping transcriptional responses. The other striking feature is the differences in the precise suites of genes reported as being under mechanical or gravitational regulation in each study. This may in part reflect differences in timing and tissues under analysis. However, for the case of mechanical stimulation it is important to note that touch is a complex stimulus to quantify and apply, and so the variation is almost certainly also reflecting the different ways mechanical stimulation was conducted in each study. It seems likely that the transcriptional response is highly tailored to the kind of mechanical stimulation, hinting at a signaling system capable of encoding information such as magnitude, duration, and location of the touch stimulation coupled to an extremely adaptable response circuit. An additional theme from such studies is that genes for putative Ca2+-binding proteins are disproportionately up-regulated upon touch stimulation (Lee et al. 2005), suggesting an alteration in the Ca2+ response system, perhaps as part of an adaptation mechanism to the initial Ca2+ signals associated with touch sensing. In general, as the levels of Ca2+ signaling components are elevated, two outcomes are likely: (1) the sensitivity of the system to future Ca2+ increases should be increased, and (2) the emphasis of Ca2+-dependent cellular responses will be shifted toward those involving the now-elevated Ca2+ response elements. Thus, the touch history of the plant may well shift both its sensitivity and precise response to future touch stimulation. However, as a note of caution, although CML24 mRNA has been shown to be highly induced by touch (nine-fold increase at 30 minutes after stimulation) and to be expressed in regions of the plant likely experiencing mechanical strain (such as branch points and organs undergoing rapid elongation), recent analysis indicates no detectable change in protein abundance upon touch stimulation (Delk et al. 2005). As an alternative approach to monitor protein changes related to touch and gravistimulation, Young et al. (2006) have used proteomic profiling of the gravistimulated root tip. This analysis confirms rapid changes in protein levels/modifications. For example, adenosine kinase changes 1.8-fold during the first 12 minutes of a graviresponse but not in response to mechanostimulation. Interestingly, one of the early gravitropic response genes identified by Kimbrough et al. (2004) is S-adenosyl-L-methionine:carboxyl methyltransferase. Both of these enzymes contribute to the AdoMet cycle, which is involved in the synthesis of a host of cell regulators and components ranging from ethylene and IAA to lignin (Schoor and Moffatt 2004). Thus, again this analysis suggests that

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Figure 5.6. Response of a root as it grows into a barrier (coverglass). Initial bending of the root within 20 min of contact with the barrier occurs in the central elongation zone (CEZ). A second bend forms in the distal elongation zone (DEZ) as the root traverses the barrier, with the tip tracking along the surface at a fixed angle (in wild-type Arabidopsis, this angle is 42 degrees). Scale bar = 1 mm.

the gravity signal is rapidly transduced to a broad range of response elements. Interestingly, a similar proteomic analysis of the mechanoresponse does not show an extensive overlap in the proteins regulated at the post-transcriptional level in the mechanical and gravity response systems (see Chapter 2). Thus, one possibility is that post-transcriptional modification represents one theme of how specificity and selectivity may be imposed on the touch versus gravity response and possibly how these response systems may interact.

5.6 Interaction of Touch and Gravity Signaling/Response Evidence for touch and gravity signaling cross-talk is readily apparent in the gross morphological response of a root upon hitting an obstacle to growth. When encountering a barrier, the root forms a step-like growth habit due to the curvature at two sites (Massa and Gilroy 2003a) (Figure 5.6). The first is convex and initially occurs in the central elongation zone (CEZ) of the root, whereas the second is concave and located at the distal elongation zone (DEZ), causing the root cap to track along at a fixed angle to the barrier. Thus, the root body extending behind the DEZ aligns parallel to the barrier as growth

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progresses. Only a few of the peripheral cells of the root cap are in contact with the barrier and these must be transmitting touch information to the systems that control root growth to suppress gravitropic response (Massa and Gilroy, 2003a; Figure 5.6). Upon reaching the end of the obstruction and the mechanical stimulation, gravitropism dominates and the root resumes normal downward growth. The root tip-to-barrier angle is intermediate between vertical and flat (44 degrees in wild-type Arabidopsis; Massa and Gilroy 2003b), suggesting a compromise between rapidly moving around the object by growing across its surface and the gravitropic response sending the tip of the root vertically downward. A complete down-regulation of the gravity-sensing system during contact with the barrier would result in growth flat against its surface. Therefore, it is likely that the intermediate angle forms as an adaptive compromise between the graviresponse and the touch stimulation to successfully allow navigation around an obstacle (Fasano et al. 2002). Indeed, a brief touch to the root cap desensitizes roots to gravity, as assayed by inhibition of subsequent gravitropic growth on a clinostat (Massa and Gilroy 2003a). Likewise, Mullen et al. (2000) showed that mechanical stress during gravistimulation can delay the development of gravitropic curvature. Further data in support of the idea that gravisignaling is modulated by mechanosignaling come from experiments observing root growth along a barrier where some or all of the cells in the root cap are killed via laser ablation, thereby removing the gravisensing cells. Ablating the whole cap or only the graviperceptive columella cells causes a loss of the DEZ curvature phase upon encountering the barrier, while the initial CEZ curvature is unaffected (Massa and Gilroy 2003a). Such observations suggest that the differential growth in the DEZ comes about because of a modified gravitropic response, as this is dependent on the presence of an intact columella. However, an intact root cap is not needed for the initial CEZ curvature, suggesting that the differential growth in this region occurs as a result of cells sensing strain caused by the compressive forces that occur after initial contact (Evans 2003; Massa and Gilroy 2003a). That is, the cells in the CEZ that respond by producing tropic curvature may also be the cells experiencing the mechanical stimulation, rather than secondarily responding to a signal produced in the cap, where the direct touch stimulation is occurring. Although the precise mechanism for the interaction between touch and gravity signaling during these responses remains to be determined, there are clues to potential components responsible for such signal processing/integration in the physical machinery of the gravity sensing system. The first step in the graviresponse is the perception of the gravity vector by the plant. There are two schools of thought about how this initial event is achieved: the statolith theory and the gravitational pressure theory (see Chapters 1 and 2 for a complete description). The statolith theory states that intracellular sedimenting particles are responsible for sensing gravity. In higher plants, statoliths are dense, starch-filled amyloplasts inside specialized cells (Sack 1997; Kiss 2000). In contrast, the gravitational pressure theory states that the entire protoplast acts as the gravity sensor and the tension and compression by the protoplast against the extracellular matrix initiates the graviresponse (Wayne and Staves 1996; Staves et al. 1997a, 1997b). At present the preponderance of experimental data support the statolith theory of gravisensing (see Chapter 1) and indeed, starch statolith motility likely plays a central role in the integration of touch and gravity responses. Thus, when the plant is mechani-

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cally stimulated, the motility of the sedimenting statoliths is reduced (Massa and Gilroy 2003a). This reduced motility would provide a system whereby touch stimulation could reduce gravitropic response by simply reducing the sedimentation rate to the gravity responsive component of the sensory cell. However, it appears that such a model of sensing based on large-scale amyloplast sedimentation may be too simplistic a view of these initial sensory events. Thus, perception and presentation times (measures of the time to generate and export the initial gravitropic signal) fall in the seconds to < 1 minute range, times much less than required for amyloplasts in the sensory cells to completely sediment to the new, lower face (Hejnowicz et al. 1998; Perbal et al. 2002; Perbal and Driss-Ecole 2003). Such rapid generation of signal implies that the amyloplasts are in contact with some network that rapidly converts the force of their sedimentation to a biochemical signal. The cytoskeleton remains a prime candidate for such a network in most gravisensing models (Blancaflor 2002; see also Chapters 1 and 2). Filamentous actin in particular has been proposed to play a critical role in gravisensitivity, based on its abundant presence in columella cells (Yoder et al. 2001; Blancaflor 2002). The “restrained gravisensing” model (Baluˇska and Hasenstein 1997; Hejnowicz et al. 1998; Driss-Ecole et al. 2000; Perbal et al. 2002) suggests that amyloplasts are physically connected to the actin network, which then in turn connects to downstream components of the signaling cascade. Alternatively, in unrestrained gravity sensing, any connections between the cytoskeleton and the statoliths are nonexistent, too weak, or too transient to provide direct mechanotransduction. In this model, the dense actin web in a columella cell is deformed locally by sedimenting amyloplasts, resulting in distant effects in the cell such as activation or inactivation of mechanoreceptors on the plasma membrane. The reduced amyloplast motility induced by mechanical stimulation would therefore deliver less force to the actin network and so reduce gravitropic signal generation. Recent data suggest that this view of the role of actin may also be too simple. For example, disrupting the actin network (Blancaflor and Hasenstein 1997; Staves et al. 1997a) does not block the gravitropic response but, rather, enhances organ bending (Blancaflor and Hasenstein 1997; Yamamoto and Kiss 2002; Blancaflor and Masson 2003; Hou et al. 2003; Hou et al. 2004), suggesting that actin may operate to down-regulate gravitropic signaling. In this case mechanostimulation may actually be enhancing the interactions of actin with statoliths to inhibit gravitropic response. The graviresponse is also accompanied by an immediate cytoplasmic pH increase after reorientation that is required for maximal gravitropic bending (Scott and Allen 1999; Fasano et al. 2001). This pH increase is extended by treatments which disrupt actin filaments (Hou et al. 2004). It is possible that actin regulates transport processes at the plasma membrane, leading to the down-regulation of the pH signal, facilitating a resetting of the gravitropic signaling system (Hou et al. 2004). The pH change itself should have far-reaching effects in the cell, as a change in pH will alter the activity of most proteins in the cytoplasm. Thus, pH changes may represent a way to effect a large-scale change of cell activities upon gravistimulation, and touch may be acting through actin to modulate this system. The gravity-related pH changes may also alter auxin distribution by changing the chemiosmotic driving force for auxin uptake/redistribution into the columella (Friml et

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al. 2002; Muday and Murphy 2002; Friml 2003). Auxin plays an important role in tropic response (see Chapter 3) and actin may affect polar auxin transport by changes in cytoplasmic pH. Actin may also operate through altering the distribution, targeting, and turnover of auxin efflux and uptake transporters. How might touch stimulation affect all these features of the gravity signaling system, such as actin dynamics or pH fluxes? The touch-induced Ca2+ increase described above has the potential to directly affect many cellular activities, including actin structure (Blancaflor 2002), proton pumping (Kinoshita et al. 1995), and auxin transport/signal transduction (Benjamins et al. 2003). Thus, the ionic signaling associated with both touch and gravity signaling may well form a nexus at which information from both systems is incorporated to control pH and auxin flux and so generate an integrated tropic response.

5.7 Conclusion and Perspectives It is clear that plants integrate a tremendous amount of environmental information to dictate the appropriate growth response. Under laboratory settings, the gravitropic response can represent an extremely powerful and often dominating influence on growth habit. However, the integrative nature of plant signaling means that many other factors will likely influence growth in the field. Thus, thigmotropic (this chapter) and hydrotropic (Chapter 6) signals are known to modulate gravitropic response through reduction in gravitropic sensitivity. For roots, where touch stimulation is likely almost constant, these other stimuli may dominate, with gravitropism perhaps providing a default directional cue to orient growth in the absence of other stimuli. Indeed, a putative gravitropic sensor reported in the elongation zone of the maize root (Wolverton et al. 2002; also see Chapter 1) could well reflect a mechanical strain sensor eliciting a thigmotropic response to the stress from the mass of the unsupported root in these experiments. Although much work has been directed to analysis of the interactions of gravitropism with these other stimuli in primary roots, it will be very informative to understand how signals such as touch interact with the mechanisms that define gravitational set-point angles of lateral organs where growth is not simply directed straight up or down (Chapter 2). One other major question remaining about thigmotropic response is the molecular identity of the plant mechanosensors(s). It seems highly likely that the touch receptor is a mechanosensitive channel, and there is a wealth of electrophysiological data suggesting that stretch-activated channels exist in the plant plasma membrane. However, apart from the MSL genes, there are no other clear homologs of mechanosensitive channels from other kingdoms represented in the sequenced plant genomes. The clear challenge for those working on touch and thigmotropic response is to robustly define the primary sensor with molecular precision. At present, the MSL proteins hold great promise in defining putative mechanosensors, but we must await their eletrophysiological characterization to know whether the elusive plant mechanosensors have truly been found. Once these sensors are defined at the molecular level, we can anticipate an equivalent rapid increase in understanding of plant mechanotransduction as that which accompanied the identification of the Msc channels in bacteria or the Mec channel complex in C. elegans.

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5.8 Acknowledgments The authors gratefully acknowledge support by grants from NSF (MCB 02-12099, IBN 03-36738, and DBI 03-01460) and NASA (NAG2-1594). We also thank Greg Richter for critical reading of the manuscript.

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6

Other Tropisms and Their Relationship to Gravitropism Gladys I. Cassab*

6.1 Introduction Plants have evolved an elaborate and sophisticated set of growth responses to the environment that allow them to survive adverse conditions. The degree to which plants depend on environmental cues to orchestrate growth and development is unmatched in the animal kingdom. Of all of the environmental signals, gravity—which is a constant factor on Earth—profoundly impacts the form, structure, and function of plants (Hangarter 1997). However, gravitropic response of plant organs can be affected by the abundance of incoming signals from the environment, and therefore plants modify their growth accordingly, taking into account all these variables. Responses of plants to environmental stimuli such as moisture (hydrotropism), temperature (thermotropism), oxygen gradients (oxytropism), electric fields (electrotropism), touch (thigmotropism; see Chapter 5), and wounding (traumatropism) are short-term tropic responses that are commonly accomplished within a few hours by differential growth. These tropisms have been mostly observed and analyzed in roots, although there are a few studies on pollen tubes, coleoptiles, and shoots. Most of these tropisms were documented more than 100 years ago (Hart 1990); nevertheless, research in this field has so far received little attention despite the biological significance of these growth behaviors in plant survival. 6.2 Hydrotropism Even though the lack of sufficient water is the single most crucial factor influencing world agriculture, interest in hydrotropism has fluctuated over the years. Studies on hydrotropism have been scarce since Knight and von Sachs (in 1811 and 1872, respectively) showed that roots move toward water (Takahashi 1997). In particular, von Sachs (1877) demonstrated that in seedlings grown in a freely hanging sieve basket, the emergent roots became diverted from the vertical and grew along the bottom of the basket (wet substrate) (von Sachs 1887) (Figure 6.1). Around that time, Darwin, Pfeffer, and Weisner (who introduced the term hydrotropism) were all convinced that moisture gradients affected root orientation (Hart 1990). Interestingly, the idea that plant roots penetrate the soil in search of water to maintain their growth was first presented as the explanation for the downward orientation of roots (Dodart, around 1700, reviewed in Hart 1990). However, in comparison to studies on the roles of other directional signals (such as gravity and light) on the general orientation of plant organs, studies on hydrotropism have been surprisingly sparse. In fact, genetic analysis of hydrotropism lagged 19 years behind the first reports of Arabidopsis agravit*Corresponding author 123

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Figure 6.1. Hydrotropism in roots. Von Sachs (1887) demonstrated for the first time with the “hanging basket” technique that roots developed a hydrotropic curvature in response to moisture gradients against the gravity vector.

ropic mutants (Eapen et al. 2003; Olsen et al. 1984). Here, we discuss the potential of a genetic approach for understanding the molecular mechanisms governing root hydrotropism and their interaction with gravitropism. 6.2.1 Early Studies of Hydrotoprism Hydrotropism studies have always been hard to interpret because both thigmotropism and gravitropism interact with hydrotropism. Mechanical stimuli can generally be avoided (see Chapter 5), but gravity is ubiquitous on Earth. Consequently, several tools, such as those involving agravitropic mutants, clinorotation, or microgravity in space have been utilized to differentiate the hydrotropic from the gravitropic response (Takahashi 1997; see also Chapter 9). Significantly, experiments with the pea mutant ageotropum, whose roots were agravitropic but responded positively to hydrotropism, indicated that there are independent sensing and response pathways for these two tropisms (Jaffe et al. 1985). Hence, ageotropum was a model system for the study of hydrotropism for many years. In particular, roots of ageotropum responded to a gradient in water potential as small as 0.5 MPa (Takano et al. 1995). Darwin (1881) studied the location of the sensory system for root hydrotropism. He covered the apical 1 or 2 mm of roots from different species with a hydrophobic mixture of olive oil and lampblack and found that they no longer responded to a moisture gradient. When root caps from ageotropum or corn roots were removed, they failed to curve

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hydrotropically but their growth was not affected (Jaffe et al. 1985; Takahashi and Scott 1993). Cytoplasmic Ca2+ has been postulated to be a transducer for the gravity signal (Plieth and Trewavas 2002). However, changes in this ion after amyloplast sedimentation have not been properly documented (Blancaflor and Masson 2003; see also Chapters 2 and 5). Even so, hydrotropic response was completely blocked in ageotropum roots with a Ca2+ chelator and lanthanum, a Ca2+ channel blocker (Takano et al. 1997). These observations suggest that Ca2+ may function in the hydrotropic response, independently of amyloplast sedimentation. It has been proposed that mechanotransductive Ca2+ channels (Chapter 5) might be triggered by temperature, gravity, touch, and water stress (Pickard and Ding 1993), which might regulate tropic modification of growth (Pickard and Ding 1992). Pollen tube guidance on the stigma has been also considered the most frequently occurring hydrotropic response in higher plants (Lush et al. 1998). Guidance toward the stigma by a water gradient may be the first step in a multistage process of guidance to the ovules. 6.2.2 Genetic Analysis of Hydrotropism Up to now, various screening procedures have been implemented to isolate mutants affected in response to gravity, light, and obstacle touching. However, hydrotropism has not been common in genetic studies because of the complexity of establishing a large-scale screening system that offers an appropriate stimulus–response interaction (Eapen et al. 2005). For this reason, the design of a screening method for the isolation of Arabidopsis mutants with abnormal responses to a water potential gradient is noteworthy (Eapen et al. 2003). The screening method consists of a vertically oriented square Petri dish with a normal nutrient medium (NM) in the upper part, in which Arabidopsis seeds are plated, and a water stress medium (WSM) in the lower part. A gradient in water potential develops over time, and wild-type Arabidopsis roots stopped their downward growth and developed a hydrotropic curvature when the water potential was 0.53 MPa. By developing this hydrotropic response, Arabidopsis roots avoided the substrate with lower water potential; that is, they never reached the area containing the WSM and consequently arrested their gravitropic growth (Fig. 6.2A). Mutants were selected on two conditions: by their continuous root gravitropic response into the medium with lower water potential (lack of hydrotropic response), and by their inability to sustain continuous growth into the severe water-deficit conditions of the WSM (Fig. 6.2A). With this selection, hydrotropic mutants were distinguished from mutants resistant to severe water deficit conditions. The initial screening resulted in the isolation of two negative hydrotropic mutants, which were named no hydrotropic response (nhr). Importantly, in a different system with an air humidity gradient, nhr1 roots responded negatively to this stimulus, developing a curvature in response to gravity instead, confirming that their directional growth toward water is impaired (Eapen et al. 2003). Plants in habitats such as dunes or deserts develop extensive root systems that penetrate large volumes of soil in search of moisture, either by means of widely spreading roots or by roots stimulated to grow toward the water table (Larcher 1995). Hence, many plants likely possess the capacity for such an enhanced responsiveness to gradients in

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Figure 6.2 (also see Color Section). Genetic approaches for examining root hydrotropism in Arabidopsis. The screening systems in (A) and (B) are composed of a vertically oriented Petri dish containing two different media. In (A), a NM is poured in the upper part and a WSM [containing 2.5% (v/v) glycerol, 0.5% (w/v) alginic acid] in the lower part. The water potential (wp) in the upper part of the dish gradually dropped off due to diffusion of the glycerol over time, becoming more negative in positions closer to the WSM. Roots of wild-type plants usually develop a curvature in the NM when the water potential declines from –0.4 to –0.5 MPa after 5 to 6 days. The asterisks denote putative no hydrotropic response mutants. In (B), WSM is poured in the upper part and a NM in the lower part, which also develops a gradient in water potential over time, but in opposite direction to that in (A). Roots of wild type plants stop growing after 4 days, but putative super hydrotropic mutants (suh) grow into the NM after 8 to 9 days (asterisks). Roots of suh mutants continue their downward growth, which is never arrested.

water availability. Therefore, we assumed it feasible to screen for super-hydrotropic response mutants in Arabidopsis. For this approach, we used the model of the screening system for nhr mutants, but the WSM is placed in the upper part and the NM in the lower section of the dish. Roots from one mutant line isolated in this system named super hydrotropic response (suh1) continuously grow under water deficit for 10 days in order to reach the moderate water potential conditions present in the lower section of the dish (Saucedo and Cassab, unpublished results; Fig. 6.2B and Color Section). Thus, both the screen for impaired and enhanced hydrotropic response appear to be fruitful avenues of research toward dissecting the complex signaling phenomena behind the hydrotropic response. 6.2.3 Perception of Moisture Gradients and Gravity Stimuli by the Root Cap and the Curvature Response The sensitivity of the root cap and the root curvature response to hydrostimulation implies that roots have moisture gradient receptors which transform this information into a physiological signal, and so trigger differential growth. Furthermore, the course followed by roots through the soil is directed by extraordinarily complex and diverse stimuli and, thus, the root cap needs to integrate multiple signals to ultimately generate the most advantageous growth direction. However, neither the sensing cells for moisture gradients nor the molecular mechanism that transduces the stimulus to a signal has been character-

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ized as yet. In addition, the mechanisms that the root cap utilizes to intermingle different stimuli to resolve in which direction root growth should occur are unknown. Nonetheless, the recent isolation of hydrotropic mutants, in combination with analysis of a number of gravitropic mutants, have provided some hints of the signal transduction mechanisms of these tropisms. For example, the gravitropic and waving response of the nhr1 mutant is increased, suggesting that the lack of a hydrotropic response results in an enhancement of these other root growth responses (Eapen et al. 2003). Moreover, roots from the starchless mutant pgm1-1, which has a reduced gravitropic response (Kiss et al. 1989), showed an enhanced responsiveness to moisture gradients (Takahashi et al. 2003). Gravistimulated nhr1 seedlings also attained a root curvature of 80 degrees 6 hours before wild-type plants (Eapen et al. 2003). In contrast, pgm1-1 roots reached a 20-degree hydrotropic curvature 1 hour before the wild-type (Takahashi et al. 2003). These observations suggest that once the sensing system of the root cap is affected for one stimulus, the integration and assessment mechanism for other signals is enhanced, and thus these other root growth responses can occur faster. However, the differences in the rate of gravitropic bending of nhr1 versus hydrotropic bending of pgm1-1 roots seem also to reflect variations in the timing of perception between both stimuli. It has been shown that the perception time for a gravity stimulus can be as short as 1 second (Hejnowicz et al. 1998). In contrast, the perception time for osmotic stimulation is up to 2 minutes (Stinemetz et al. 1996). The variability observed in the perception time of both tropisms might be the consequence of their distinctive mechanisms of perception. It is widely accepted that perception of gravity occurs in specialized cells of the root cap (statocytes or columella), which contain motile amyloplasts that can sediment in response to gravity and can therefore elicit gravisensing (Sack 1997; see also Chapter 1). The capacity of the root cap to perceive and respond to moisture gradients apparently produces a dominant signal that abates the gravity response. Recently, it has been found that this signal triggers the degradation of amyloplasts in columella cells of both Arabidopsis and radish, and hence roots exhibit hydrotropism with fewer impediments from gravitropism (Takahashi et al. 2003). Transient touch stimulation of Arabidopsis root tips likewise restrains gravitropic growth but, in this case, by limiting amyloplast sedimentation in columella cells (Massa and Gilroy 2003; see also Chapter 5). Therefore, columella cells can integrate the signaling triggered by moisture gradients, touch receptors, and possibly even other stimuli in order to generate the appropriate tropic response. Columella cells also produce the initial gravity-induced lateral auxin gradient in the root cap. These cells contain one putative component of the auxin efflux carrier complex (PIN3) which shows rapid relocalization upon gravistimulation and is thought to drive asymmetrical auxin transport responsible for tropic bending (Friml et al. 2002; see also Chapters 2 and 3). Conceivably, columella cells might utilize the same signaling components that drive differential growth (such as auxin efflux) for all sensory systems, and so work like a funnel taking in many stimuli and transducing these toward a single set of response elements in order to synchronize the various tropistic responses (Eapen et al. 2005). Clearly, more remains to be learned about the functional interactions between sensing mechanisms that take place during gravity, moisture, and touch perception. Nonetheless, hydrotropic stimulation exerts a more dramatic effect than touch upon the primary mechanism for gravireception, which might be associated with the importance of water in the life of most plants.

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Interestingly, and unlike wild-type plants, hydrostimulated nhr1 roots maintained their amyloplasts in the columella (Ponce and Cassab, unpublished observations). This might suggest that the negative hydrotropic response of nhr1 is related to the absence of the signaling cascade that triggers amyloplast degradation during hydrotropic stimulation. In the screening system for the isolation of super-hydrotropic response mutants, wild-type seedlings ceased growing after six days in the WSM and lacked amyloplasts in their columella cells, supporting the observation made by Takahashi et al. (2003) that water-stressed roots also degrade amyloplasts. In contrast, suh1 roots exhibited amyloplasts in the columella, indicating that the root cap seems to combine both a hydrotropic and a gravitropic response in order to reach the medium with higher water potential in the bottom part of the plate. Root hydrotropic responsiveness has also been studied by using an agar KCl system in some Arabidopsis agravitropic, auxin-insensitive, and ABA-related mutants (Takahashi et al. 2002). In this study, Arabidopsis wild-type roots began hydrotropic curvature against the gravity vector after 30 minutes of stimulation and reached 80 to 100 degrees within 24 hours. In contrast, roots of axr1-3 and axr2-1 mutants showed a greater hydrotropic response compared with those of wild type. Both mutants are insensitive to auxin and show altered root gravitropism, with axr2 roots being particularly agravitropic (Lincoln et al. 1990; Nagpal et al. 2000). axr2 roots developed a curvature even in the absence of a moisture gradient, which might suggest that this response is a consequence of their random root growth direction. Additionally, mutants affected in basipetal polar auxin transport from the root cap to the root elongation zone, such as wav6 and aux1 (Blancaflor and Masson 2003; Swarup et al. 2005), showed hydrotropic curvature. However, it was previously shown that an auxin transport inhibitor blocked the hydrotropic curvature of ageotropum roots (Takahashi 1994). Therefore, this analysis suggests that auxin may regulate gravitropism and hydrotropism differently, although both tropisms might depend on the formation of an asymmetric auxin gradient for differential growth. Furthermore, the relatively random root growth direction of these mutants makes the interpretation of their hydrotropic response complex, and the results do not provide strong evidence for or against a role of auxin and auxin transport in the hydrotropic response. 6.2.4 ABA and the Hydrotropic Response Abscisic acid (ABA) functions mostly in plant responses to dehydrating stresses (Finkelstein et al. 2002), and hence a change in ABA homeostasis could take place under hydrotropic stimulation. Interestingly, mild water stress stimulates primary root elongation (Sharp 2002). This implies that ABA might enhance root growth in search of water under these conditions. Root growth of nhr1 mutants is insensitive to ABA in NM (Eapen et al. 2003), but is highly stimulated by this hormone when seedlings are growing in the screening medium with a water potential gradient (Ponce and Cassab, unpublished observations). This suggests that orthogravitropic growth of mutant roots is stimulated under severe water stress conditions in the presence of exogenous ABA. A putative role of ABA in root gravitropism has been discussed (Feldman et al. 1985), but has not been thoroughly analyzed further (LaMotte and Pickard 2004). Previously, we suggested that ABA might antagonize the early transduction of gravi-induction of hydrotropic-responsive roots (Eapen et al. 2005). In fact, there are some ABA mutants affected either in their

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gravity or waviness response (Lu and Federoff 2000; Kang et al. 2002). On the other hand, it has been postulated that ABA maintains a higher growth rate on the side with lower water potential in hydrotropically responsive roots. This suggestion arose from the observation that the hydrotropic response of roots in an agar KCl system of two Arabidopsis ABA mutants, aba1-1 and abi2-1, was slightly reduced compared with those of wild type (Takahashi et al. 2002). In contrast, both aba1-1 and abi2-1 mutant roots responded positively to hydrotropism in the screening system with a water potential gradient (Eapen et al. 2003). However, there are some differences between these two hydrotropic systems, namely, light conditions, seedling age, and the substrate of the water potential gradient, which may account for this discrepancy. Light has been shown to influence the gravity response in roots, shoots, and other organs (Hangarter 1997). So far, there are no studies that have examined an interaction of ABA with the development of an asymmetric auxin gradient for differential growth, which may be an important factor during hydrofacilitation. Yet, it was recently reported that cells in the columella and quiescent center of Arabidopsis showed low levels of ABA, which were not increased by water stress, suggesting a non-stress-related role for ABA in these cell types (Christmann et al. 2005). Further, there are some reports of ABAactivated gene expression in the root cap (Hong et al. 1988; Nylander et al. 2001), and so the root cap might be required for proper regulation of the ABA-dependent processes of cell division (Dewitte and Murray 2003). Therefore, an important role of ABA in developmental programs such as tropisms is anticipated. A detailed analysis of more ABA mutants and the cloning of the NHR1 gene might thus provide evidence as to whether ABA functions on gravitropic and/or hydrotropic response in roots. 6.2.5 Future Experiments Our understanding of hydrotropism has lagged many years behind that of gravitropism and phototropism. However, the future characterization of the genes in the hydrotropic mutants isolated so far might help to unravel the players in this particular growth response. Many questions remain open, particularly those related with the sensing system for moisture gradient and the mechanism that merges and assesses the diverse stimuli impacting on the root in order to generate the proper tropic response. Thus far, the NHR1 gene seems to block root gravitropic growth and allow roots to direct their growth toward water, since in nhr1 mutant roots gravity response is enhanced (Fig. 6.3). Further, it remains to be determined whether signals such as Ca2+, calmodulin, pH increases, reactive oxygen species, inositol 1,4,5-trisphosphate, auxin, ethylene, flavonoids, cytokinins, and brassinosteroids participate in hydrotropism as they seem to do in gravitropism (reviewed in Chapter 2). Thus, there is considerable potential for further research to uncover the mystery of how roots are able to amplify a signal, such as water.

6.3 Electrotropism The direction of growth of certain plant cells or organs can be modified by an applied electric field. This phenomenon, known as electrotropism, has been observed in fungi

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Figure 6.3. Root hydrotropism opposes gravitropism in Arabidopsis. A. Perception of gravity (a weak signal) appears to occur in columella cells of the root cap, triggered by amyloplasts that can sediment because of gravitational force. After the sensory system identifies this stimulus, it connects with the hormone system (perhaps by transient Ca2+ fluxes and/or alkalinization of columella cytoplasm) and an asymmetrical signal is originated within the columella cells, leading to the lateral movement of auxin. B. Perception of moisture gradients (a strong signal) might take place anywhere in the root cap, but assessment of both gravity and water signals most probably occurs in columella cells, which in turn elicits amyloplast degradation and ultimately initiates the appropriate bending response. How the sensory system for humidity gradients connects with the hormone system remains to be uncovered. NHR1 might inhibit the root gravitropic response when both humidity gradients and gravity sensitiveness merge. ABA and ethylene may positively regulate hydrotropism and inhibit gravi-facilitation. Shaded bar denotes a moisture gradient; solid bar symbolizes lack of a moisture gradient; gray arrows indicate the direction of the auxin transport, and their width correlates with levels of transported auxin; white arrow represents an increase in Ca2+ fluxes and pH in a columella cell.

(McGillavray and Gow 1986) and algae (Brower and Giddings 1980), as well as in pollen tubes (Marsh and Beams 1945), roots (Fondren and Moore 1987; Schrank 1959), and shoots (Schrank 1959; Lee et al. 1983) of higher plants. Upon gravistimulation, the current flow along the upper flank of the distal elongation zone (DEZ) reversed to efflux from the root (Iwabuchi et al. 1989; Collings et al. 1992), and changes in intracellular potentials in this zone occurred within a minute (Ishikawa and Evans 1990a). These changes arose before the development of the gravitropic curvature. Gravistimulation also modified the pattern of electric current surrounding the root tip (Behrens et al. 1982; Björkman and Leopold 1987; Iwabuchi et al. 1989), and within the root cap triggered rapid depolarization of statocytes (Behrens et al. 1985), suggesting that electrical/ionic signals may be an important component of the gravity sensing/response system. Electrotropism was enhanced by treatments that interfere with gravitropism, like decapping the roots or pretreating them with Ca2+ chelator. Likewise, roots of ageotropum were more responsive to electrotropic stimulation than were roots of normal peas (Ishikawa and Evans 1990b), suggesting that the early steps of gravitropism and electrotropism occur by independent mechanisms. Nonetheless, the motor mechanisms of the two responses may have features in common since auxin and auxin transport inhibitors reduced both gravitropism and electrotropism (Moore et al. 1987). The kinetics of electrotropic curvature in Vigna mungo L. roots revealed that curvature

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occurred in the same root toward both the anode and cathode, but the two responses took place in two different regions of the root—the central elongation zone (CEZ) and the DEZ, respectively (Wolverton et al. 2000). Furthermore, both responses are related to the electric field rather than one being a secondary response to induced gravitropic stimulation, since these oppositely directed responses could be reproduced individually by a localized electric field application to the region of response. The electrotropic responses of plant organs described thus far are quite clear, but very little is known about the actual mechanisms of stimulus reception and directional guidance.

6.4 Chemotropism In green plants, the ultimate energy source for growth is light rather than a chemical input. The directional cues from light, supplemented by the orientation afforded by gravitropic growth, help place the aerial part of the plant in the optimal position to intercept sunlight. Gravity and light are thus two of the main directional signals engaged in the regulation of vegetative growth. However, chemicals still act as significant vehicles of communication between individuals in their reproductive behavior or, in the case of insectivorous plants, as signals for obtaining nutrients. Chemotropism is a directional growth response that is driven by a chemical stimulus; the term chemotaxis is used to depict the locomotory responses of motile organisms or gametes. Both forms of response can be positive (toward a beneficial or attractive substance) or negative (away from a harmful or unattractive substance) (Hart 1990). Responses to chemical substances have been well documented in lower plants, particularly in unicellular organisms and gametes. In multicellular organisms there is usually a greater homeostasis of the cellular environment, and chemoresponses to external chemicals seem limited to rather specialized situations. A very wide range of chemicals is implicated in these types of responses, indicative of the variety of adaptive advantages that can be provided in these situations. In higher plants, considerable chemical interaction does indeed occur. Many bacteria and fungi release hormones and hormone-like substances into the soil, and these can have significant effects on root growth (Bilderback 1985). In some older physiological texts, it was suggested that roots can develop chemotropic responses to soil nutrients. However, these suggestions were based upon studies in which chemicals were unilaterally applied to individual roots of several species (Newcombe and Rhodes 1904), and these methods do not constitute a robust chemotropic directional assay (i.e., re-directional growth in response to the repositioning of the stimulus). However, there is a recent report in which the root cap seems to sense extracellular glutamate to trigger a reduction in the rate of cell production and/or cell expansion (Filleur et al. 2005), suggesting a specific response which is likely to involve the action of a specific receptor, but these authors did not report on a directional response to an actual gradient. Certain plant organs, however, seem to develop directional responses to chemical stimuli, such as the trapping organs of insectivorous plants, and pollen tubes during their growth down the style. Mechanical stimuli usually elicit the initial release of insect traps (Chapter 5), but in many cases chemical stimuli seem to enable the subsequent tighten-

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ing of the trap (Ziegler 1962). However, nothing is known about any of the chemoreceptors involved. Since the early work by Molisch and DuBary in the 1890s (Hart 1990), the directional growth of the pollen tube down the style and into the ovary has usually been regarded as the classic example of chemotropic growth in higher plants. Two molecules that have neurotransmitter properties in animals were recently found to be involved in pollen tube growth, GABA (␥-amino butyric acid) and nitric oxide (NO). Through genetic analysis in Arabidopsis, a gradient of GABA was shown to be involved in the final stages of pollen tube guidance to the ovule (Palanivelu et al. 2003). GABA is a four-carbon ␻-amino acid, best known for its role in animal neuronal synapses, but was actually first discovered in plants (Steward et al. 1949). In the pistil, GABA concentrations increase along the pollen tube path, reaching maximal concentrations in the inner integument cells directly surrounding the micropyle, the target of the pollen tube for delivery of sperm cells (Palanivelu et al. 2003). This gradient is interrupted in pollen-pistil interaction 2 (pop2) mutants, resulting in aberrant pollen tube growth and guidance. POP2 encodes a transaminase, which converts GABA to succinate. pop2 pistils have more than 100-fold greater GABA levels than do wild-type tissues, thus reducing the magnitude of the gradient. This elevation apparently results in improper targeting (Palanivelu et al. 2003) (Fig. 6.4A and Color Section). Hence, intracellular degradation of GABA in wildtype pollen tubes by POP2 presumably increases the GABA gradient, allowing tubes to distinguish the micropyle from the rest of the ovule. The fact that a GABA gradient exists along the path of pollen tube supports a chemotropic growth of pollen tubes. In animals, GABA binds to G-protein-coupled GABA receptors; homologs of these proteins have been identified in plants. The heterotrimeric G-protein alpha subunit, GPA1, is expressed in the pollen tube, although it is not known whether it participates in pollen tube guidance (Ma 2003). A role of nitric oxide (NO) was recently demonstrated in the regulation of pollen tube growth in Lilium longiflorum, especially in the reorientation response (Prado et al. 2004) (Fig. 6.4B and Color Section). A NO gradient may play a role in finding a suitable path for the pollen tube, suggesting that this may be another directional response. Nonetheless, so far nothing is known about the nature of the NO chemoreceptor. A female gamethophyte protein from maize has also been shown to be required for pollen tube attraction (Márton et al. 2005), indicating that a wide range of different substances can elicit a chemoresponse in pollen tubes. Furthermore, directional guidance of pollen tubes into the style does not appear to be influenced by gravity, indicating that their chemotropic response might only interact with their hydrotropic, electrotropic, and oxytropic ones (see following section).

6.5 Thermotropism and Oxytropism Temperature exerts clear effects on all aspects of plant growth and development. Root thermotropism was first detected by Barthèlèmy (1884), who performed experiments with hyacinth bulbs floating in water-filled containers (reviewed in Aletsee, 1962). Wortmann (1885) documented both positive and negative thermotropism in several

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Figure 6.4 (also see Color Section). Chemotropism in pollen tubes. A concentration gradient of GABA affects the final stages of directional guidance of pollen tubes to the ovule. A. In the wild-type ovule, the inner integument possess a much higher level of GABA than do other tissues of the ovule or the septum, and the pollen tube grows toward the micropyle. In the pop2 mutant ovule, GABA concentrations in the entire ovule, including the funiculus and outer integument, are very high, and that in the inner integument is even higher. The pollen tube of pop2 mutants fails to direct its growth toward the micropyle, and grows randomly in the ovary (after Ma 2003). B. Lily pollen tube showing three consecutive reorientation responses, which were induced by moving the NO source to the locations marked with arrows. The growth axis of the pollen tube always developed right angles after each challenge by the NO source facing the pollen tube-tip. (From Feijó JA, Costa SS, Prado AM, Becker JD, Certal AC. 2004. Signaling by tips. Current Opinion in Plant Biology 7:589-598, with permission.)

species using seedlings planted in moist sawdust in a metal box heated on one side by a gas burner and cooled on the other side by water. Others followed Wortmann’s work, but a consensus on the existence of thermotropism was not reached until recently. Fortin and Poff (1990) showed that primary roots of maize reoriented from their original vertical direction when exposed to a 4.2oC cm–1 thermal gradient applied perpendicular to the gravity vector, indicating that a thermal gradient can be sensed by roots. This positive root thermotropic curvature might represent the integrated sum of thermotropism and gravitropism (Fortin and Poff 1991). However, when roots were placed horizontally under 1 g with a vertical thermal gradient, the thermal stim-

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ulus at 15 oC was stronger than the gravity stimulus and the root curved toward the top of the dish (Fortin and Poff 1991), providing additional evidence that thermal gradient sensing is accountable for the thermotropic response. The thermoreceptors in plants remain to be discovered but, in animals, temperature-sensitive channels can relay thermal information. Thermotropism could be of adaptive value for root optimal development since horizontal temperature gradients are common in agroecosystems (Fortin and Poff 1990). Molisch (1884) was the first to demonstrate an aerotropic response of plant roots, with Pfeffer (1906) reviewing research on this topic and proposing the term oxytropism. As with many of the tropisms discussed in this chapter, research in oxytropism was ignored for 60 years until several reports of altered root system growth resulting from limiting O2 availability appeared (listed in Porterfield and Musgrave, 1998). Recently, oxytropism has been reexamined in a microrhizotron capable of producing and maintaining an O2 gradient of 0.8 mmol mol–1 mm–1 using two cultivars of pea: Weibul’s Apolo and ageotropum (Porterfield and Musgrave 1998). Oxytropic curvature was seen all along the O2 gradient in both cultivars of pea, with growth toward the higher O2 concentrations regardless of the starting position within the O2 gradient. Roots of ageotropum showed a curvature of 90 degrees into the O2 gradient, in contrast to the gravity-sensing cultivar, which only curved 45 degrees. The curvature of the Weibul’s Apolo cultivar indicates that roots normally integrate both the gravity and oxygen signal, resulting in the diageotropic and plagiotropic growth seen in response to soil flooding (Huck 1970). Oxytropism may allow roots to evade O2-deprived soil strata and may also be the basis of an auto-avoidance mechanism, diminishing the competition between roots for water and nutrients as well as oxygen (Porterfield and Musgrave 1998). In addition, it has been shown that pollen tube guidance is influenced not only by chemical or water gradients, but also by oxygen gradients. Pollen tubes of several species showed a clear tropic response to oxygen gradients in an in vitro system (Blasiak et al. 2001). The biological significance of this phenomenon in vivo has not been analyzed yet, but may be critical for orienting the pollen tubes toward the stigma, or in maintaining basipetal growth in the style.

6.6 Traumatropism Wounding is a mechanical process that harms cells in a localized region, but which also typically results in alterations in the activities of cells in other regions (Imaseki 1985). A less well-known effect of wounding is the induction of differential growth in the wounded organ within the first hour or so after damage. Pfeffer introduced the term traumatropism in 1893 to describe such wound-induced, directional growth responses. Along with his experiments on thigmotropism, Darwin (1881) also analyzed the responses of roots to wounding. The root response to injury was found to be similar to touch, that is, if the injury was close to the root tip, the root curved away from the side that was wounded, occasionally even to the extent of forming a 90-degree curvature; but if the wound was just beyond the tip, the root bent toward the wounded side. Aerial organs, overall, seem to be much less sensitive than roots to injury stimuli. Most of this work has been performed on coleoptiles, in which slicing or abrasion can induce

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positive curvatures of 30 to 40 degrees (Bünning 1959). The negative and positive traumatropic response of both roots and shoots indicates the participation of message transmission from the wound site to the cells showing differential growth. Such a wound message may be transmitted chemically (through the action of hormones or proteins), electrically (through the generation and propagation of action potential), and/or mechanically (through activated mechanosensory Ca2+ channels). It remains to be determined whether only one or the combined action of these three wounding messages regulates traumatropism. The significance of traumatropism to the plant may be to offer initial protection to injury, reinforced later on by the systemic long-distance signaling induced in the wound response (Schilmiller and Howe 2005). 6.7 Overview Sensory systems in plants may consist more of a network of interconnected response chains rather than a series of separate stimulus reception-transduction pathways. Dissection of this complex network will require the development of innovative methodologies in conjunction with present technologies to resolve the mechanism of perception and assessment that controls organ-bending responses to these diverse stimuli.

6.8 Acknowledgments This work is dedicated to Professor Barbara G. Pickard, whose comprehensive knowledge of the field has been the source of many invaluable suggestions from which my studies on hydrotropism have profited greatly. We warmly thank Yoloxóchitl Sánchez for drawing the figures and Manuel Saucedo for his enormous contribution in the isolation of the super-hydrotropic mutants. We are grateful to all past and present members of the laboratory for their contributions and discussions on hydrotropism. We also gratefully acknowledge financial support by the Mexican Council for Science and Technology (CONACYT grant No. 462022Q), the Universidad Nacional Autónoma de México (Dirección General de Asuntos del Personal Académico Grant No. IN224103), and the University of California Institute for Mexico and the United States (UC Mexus).

6.9 Literature Cited Aletsee L. 1962. “Thermotropismus.” In Physiology of movements, Encyclopedia of plant physiology, Vol. 17, Part 2, edited by W. Ruhland, pp. 1–14. Berlin: Springer. Barthèlèmy A. 1884. De l’action de la chaleur sur les phénomènes de végétation. Compte Rendus de l’Academie du Sciences (Paris) 98:1006–1007. Behrens H.M., Gradmann D., Sievers A. 1985. Membrane potential responses following gravistimulation in roots of Lepidium sativum L. Planta 163:463–472. Behrens H.M., Weisenseel M.H., Sievers A. 1982. Rapid change in the pattern of electric current around the root tip of Lepidium sativum L. following gravistimulation. Plant Physiology 70:1079–1083.

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Bilderback D.E. 1985. “Regulators of plant reproduction, growth and differentiation in the environment.” In Hormonal regulation of development, III. Role of environmental factors, Encyclopedia of plant physiology, New Series, Vol. 11, edited by R.P. Pharis and D.M. Reid, pp. 652–706. Berlin: Springer. Björkman T., Leopold A.C. 1987. An electric current associated with gravity sensing in maize roots. Plant Physiology 84:841–846. Blancaflor E.B., Masson P.H. 2003. Plant gravitropism. Unraveling the ups and downs of a complex process. Plant Physiology 133:1677–1690. Blasiak J., Mulcahy D.L., Musgrave M.E. 2001. Oxytropism: a new twist in pollen tube orientation. Planta 213:318–322. Brower D.L., Giddings T.H. 1980. The effects of applied electric fields on Micrasterias. II. The distributions of cytoplasmic and plasma membrane components. Journal of Cell Science 42:279–290 Bünning E. 1959. “Die wirkung von wundreizen.” In Physiology of movements, Encyclopedia of plant physiology, Vol. 17, Part 1, edited by W. Ruhland, pp. 199–134. Berlin: Springer. Christmann A., Hoffmann T., Teplova I., Grill E., Müller A. 2005. Generation of active pools of abscisic acid revealed by in vivo imaging of water-stressed Arabidopsis. Plant Physiology 137:209–219. Collings D.A., White R.G., Overall R.L. 1992. Ionic current changes associated with the gravityinduced bending response in roots of Zea mays L. Plant Physiology 100:1417–1426. Darwin, C. 1881. The power of movement of plants. London: John Murray. Dewitte W., Murray J.A. 2003. The plant cell cycle. Annual Review of Plant Biology 54:235–264. Eapen D., Barroso M.L., Campos, M.E., Ponce G., Corkidi G., Dubrovsky J.G., Cassab G.I. 2003. A no hydrotropic response root mutant that responds positively to gravitropism in Arabidopsis. Plant Physiology 131:536–546. Eapen D., Barroso M.L., Ponce G., Campos M.E., Cassab G.I. 2005. Hydrotropism: root growth responses to water. Trends in Plant Science 10(1):44–50. Feldman L.J., Arroyave N.J., Sun P.S. 1985. Absicic acid, xanthoxin and violaxanthin in the caps of gravistimulated maize roots. Planta 166:483–498. Filleur S., Walch-Liu P., Forde B.G. 2005. Nitrate and glutamate sensing by plant roots. Biochemical Society Transactions 33:283–286. Finkelstein R.R., Gampala S.S.L., Rock C.D. 2002. Abscisic acid signaling in seeds and seedlings. Plant Cell 14:S15–S45. Fondren W.M., Moore R. 1987. Collection of gravitropic effectors from mucilage of electrotropicallystimulated roots of Zea mays L. Annals of Botany 59:657–659. Fortin, M.-C., Poff K.L. 1990. Temperature sensing by primary roots of maize. Plant Physiology 94:367–369. Fortin, M.-C., Poff K.L. 1991. Characterization of thermotropism in primary roots of maize: dependence on temperature and temperature gradient and interaction with gravitropism. Planta 184:410–414. Friml J., Vieten A., Saluer M., Weijers D., Schwarz H., Hamann T, Offringa R., Jurgens G. 2002. Lateral relocation of auxin efflux regulator PIN3 mediates tropism in Arabidopsis. Nature 415:806–809. Hangarter, R. P. 1997. Gravity, light and plant form. Plant Cell and Environment 20:796–800. Hart, J. W. 1990. Plant tropism and growth movement, pp. 172–201. London: Unwin Hyman. Hejnowicz Z., Sondag C., Alt W., Sievers A. 1998. Temporal course of graviperception in intermittently stimulated cress roots. Plant Cell and Environment 21:1293–1300. Hong B., Uknes S., Ho T.-H.D. 1988. Cloning and characterization of a cDNA encoding a mRNA rapidly-induced by ABA in barley aleurone layers. Plant Molecular Biology 11:495–506.

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7

Single-Cell Gravitropism and Gravitaxis Markus Braun* and Ruth Hemmersbach

7.1 Introduction Over the last few decades, single-cell systems have increasingly attracted attention as model organisms for research on gravity-related biological processes. Gravity is the only constant environmental stimulus and provides the most reliable cue that free-swimming and sessile organisms learned to use for orientation. This capacity is found in organisms ranging from single cells up to multicellular animals and plants. In all these systems gravitropism and gravitaxis involve the pure physical step of susception followed by physiological steps comprising gravity perception, signal transduction, and signal transmission, eventually resulting in a gravity vector-related response in the form of a reorientation of growth or movement direction. In this review, free-swimming protozoa like flagellates (e.g., Euglena) and ciliates (e.g., Paramecium and Loxodes) are referred to as single-cell systems, as are tip-growing cell types such as protonemata of mosses and ferns and the rhizoids and protonemata of algae. Although such cells are part of a multicellular organism, they extend away from it. They have to cope directly with the environment and they have to adapt to it in a most beneficial way in order to survive. Communication of single-cell systems with other cells of the organ is limited and they are not integrated into complex signal transduction networks and response pathways required for multicellular response. Thus, for single-celled systems, the stimulus response is dependent only on the cell’s own orientation. All singlecell systems share a number of advantageous features for study. The unobstructed access to the cell body permits a great variety of experimental approaches and allows easy isolation for biochemical and molecular analyses. The signaling pathways are relatively short and all phases occur in a single cell. This chapter accentuates the substantial contribution single-cell systems have added to our understanding of the intracellular mechanisms underlying gravity sensing and the molecular basis of the gravity-dependent signaling pathways.

7.2 Definitions of Responses to Environmental Stimuli that Optimize the Ecological Fitness of Single-Cell Organisms Microorganisms (bacteria and protists), animals, and plants respond to environmental stimuli in a multitude of ways. The capacity of free-living organisms to orient the direction of their movement with respect to the source of an external stimulus is called taxis: positive taxis if the direction of movement is toward the source of the stimulus, negative taxis if it is away from it, and diataxis or transverse taxis if it is at an angle to the stimulus direction. *Corresponding author 141

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Correspondingly, sessile organisms show growth responses called tropisms defining a steady-state bending of an organ with respect to the direction of the stimulus source. The kind of environmental stimulus to which the organisms respond is indicated as a prefix to the appropriate term: for example, phototaxis or phototropism (directional response to light; see also Chapter 4); chemotaxis and chemotropism (directional response to a chemical, e.g., nutrients; Chapter 6); thermotaxis and thermotropism (response to a thermal gradient; Chapter 6). In this chapter we concentrate on directional responses with respect to the gravitational field, which are called gravitaxis and gravitropism. In addition, a special gravitydependent kinetic response has been described for some protists (ciliates). These systems have the capacity to regulate their swimming velocity depending on their swimming direction. They manage to speed up during upward swimming and to decelerate during downward swimming. Consequently, they compensate for at least part of their passive sinking (sedimentation) rate (Machemer et al. 1991; Ooya et al. 1992; HemmersbachKrause et al. 1993a). Gravikinesis is calculated by comparing the upward and downward swimming velocities of a cell (population) with its sedimentation velocity (Machemer and Bräucker 1992): Gravikinesis =

upward swimming rate − downward swimming rate − sedimentation rate 2

7.3 Occurrence and Significance of Gravitaxis in Single-Cell Systems Gravitactic behavior has been reported for several protozoan species. These unicellular organisms are heavier than water and most of the species studied so far show negative gravitaxis, which guides them to the surface. Well-studied examples are the negative gravitaxis of the heterotrophic ciliates Paramecium and Tetrahymena (for review, see Bean 1984; Häder et al. 2005; Hemmersbach and Häder 1999; Hemmersbach et al. 1999), the oxygen-dependent gravitaxis of the microaerophilic ciliate Loxodes (Fenchel and Finlay 1986), and the light-dependent gravitaxis of the autotrophic green algae Euglena and Chlamydomonas (Bean 1984; Häder 1987). These examples clearly show that the direction of gravitaxis increases the ecological fitness of the organisms. Besides ciliates and flagellates (Figure 7.1), gravity effects have also been studied in other unicellular organisms: amoeba, acellular slime molds, swimming reproductive stages such as zoospores and sperm cells, and bacteria. These systems have been exposed to different acceleration levels in order to analyze the impact of gravitational forces on different physiological processes (behaviour, proliferation, etc.). Due to the fact that in these cases a clear hypothesis as to the mechanism of graviperception is still missing, we will only briefly mention these in this chapter (for more details, see Häder et al. 2005) before describing the more thoroughly understood single-cell systems such as Euglena, Loxodes, and Chara. The migrating plasmodium of the single-celled slime mold Physarum polycephalum (Myxomycetes, acellular slime mold) shows gravitaxis and has been used as model system to study the impact of gravity on actomyosin-driven movements (Block et al. 1986).

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Figure 7.1. Unicellular systems used to study gravity sensing. A. Paramecium biaurelia (170 µm), seen in phase contrast. B. Loxodes striatus (150 µm), bright field; arrows indicate Müller organelles (MO). C. Euglena gracilis (50 µm), seen with differential interference contrast.

Though distinct behavioral responses to altered gravitational stimulation have been reported, the gravireceptor candidates in Physarum remain speculative: nuclei and mitochondria have been proposed, both occurring in high numbers in this giant single cell. The accumulation of zoospores near the surface of the medium raises the stillunsolved question of whether this behaviour is guided by gravity or by oxygen (Cameron and Carlile 1977). The gravity-induced, altered physiology of sperm cells is of high interest due to possible impacts on reproduction and thus, for example, food supply during long-term space flights (Engelmann et al. 1992; Tash and Bracho 1999). In the case of bacteria, effects of microgravity on, e.g., growth rate, sporulation and phage productivity have been reported (for reviews, see Mennigmann and Lange 1986; Cogoli and Gmünder 1991); however, reasons and mechanisms remain speculative. According to the model derived by Klaus and coworkers (1997), it seems likely that a “cumulative effect of gravity may have a significant impact on suspended cells via their fluid environment, where an immediate, direct influence of gravity may otherwise be deemed negligible.”

7.4 Significance of Gravitropism in Single-Cell Systems Among the few single-cell systems that respond gravitropically are the rhizoids of characean green algae, which share similar functions with roots of higher plants. They anchor the organism in the substrate by penetrating mud and soil, thereby enabling stabilized upward growth of the shoots. Other gravitropically responding cell types, including moss caulonemata and protonemata and characean protonemata, seek to grow upward in darkness in order to find optimal ecological conditions where the plants harvest light, re-

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generate a multicellular body, and proliferate. These cells elongate by polarized growth or tip growth enabling them to extend rapidly and so penetrate easily all kinds of substrates. When accidentally covered with sediment or sand, protonemata of Chara arise from cells at the nodal complexes of the green thallus by asymmetric cell division. These cells use negative gravitropic tip growth to grow back to the light. Light induces termination of tip growth and a depolarization of the cells. The cells undergo a complex pattern of divisions that reconstitute the green thallus. This process ensures survival of the plant in a difficult environment characterized by unpredictably changing conditions.

7.5 What Makes a Cell a Biological Gravity Sensor? Gravity is a weak but ubiquitous force that acts on masses. Therefore, work has to be done in the form of moving a mass in the gravitational field in order to create sufficient energy to activate a biological sensor. Theoretically, many cells possess organelles with sufficient masses whose gravitationally induced movements could create sufficient energy. However, most cell types of the various tissues usually actively prevent organelles from sedimenting by keeping them in place via cytoskeletal anchorage. This genetically defined high degree of cytoplasmic organization is quite stable, even against moderately increased acceleration forces. Thus, organelles do not move in the gravitational field or their movements are obviously not transduced into a physiological response. Therefore, in addition to the presence of organelles with sufficient mass and size, a gravity-sensing apparatus must exist in a gravisensitive cell type that facilitates sedimentation of specific masses and mediates the activation of gravity-specific receptors (i.e., the transduction of the physical stimulus into a physiological gravity perception signal). As discussed in Chapters 1 and 2, little is known about the mechanism of gravity sensing in higher plant statocytes, and identification of components of a gravity-sensor apparatus is limited to starch-filled amyloplasts. These amyloplasts act as statoliths whose gravity-directed sedimentation precedes graviperception and gravitropic curvature. Several studies suggest the involvement of actin microfilaments that might act as transducers of tensional forces generated by the gravity-induced sedimentation of statoliths (Sievers et al. 1991a; Yoder et al. 2001; Blancaflor 2002; Perbal and Driss-Ecole 2003) to mechano-sensitive receptors in cortical ER membranes or in the plasma membrane of higher plant statocytes (Ding and Pickard 1993; Kiss 2000). However, experimental evidence for the role of the actin cytoskeleton in susception and perception of gravity remains controversial. For example, treating statocytes with actin-disrupting drugs increased the sedimentation rate of statoliths (Sievers et al. 1989) and actually enhanced the gravitropic response (Hou et al. 2003, 2004; Yamamoto and Kiss 2002). In light of this dilemma as to the identity and role(s) of the gravity perception machinery in higher plants, the ease of manipulation afforded by single-cell systems opens up new vistas in order to understand gravity-sensing and gravity-oriented growth. In the following, we summarise the data on single-cell systems that have led to an increasing understanding of the nature of the gravisusception apparatus and the complexity of components (i.e., cytoskeletal elements, membrane compartments, ion channels, and receptor proteins that need to interact to elicit a beneficial orientation response).

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7.6 Gravity Susception—The Initial Physical Step of Gravity Sensing A gravisensitive cell must provide the molecular and cellular conditions that facilitate the activation of a receptor by gravity. The pure physical action that is induced by a change of the direction or amount of acceleration resulting in the activation of a gravireceptor has been termed susception. Sedimentation of masses is a prerequisite for any interaction of gravity with cellular components. Two hypotheses have been put forward to explain how gravity is sensed in a cell (Salisbury 1993; Sack 1997; Kiss 2000). Candidates for sedimentable masses are either intracellular statoliths or the entire protoplast. In higher plants, the statolith-based hypothesis (the starch-statolith theory) is favored since the discovery of starch-filled amyloplasts (statoliths) in specialized columella and endodermal cells of higher plants (reviewed in Sack 1997; see also Chapter 1). However, the absence of obvious statoliths in several graviresponsive cells, including the internodal cells of Chara, and the observation that starchless mutants of Arabidopsis can sense gravity, were decisive for the formulation of the alternative, protoplast-based hypothesis (the protoplast-pressure theory, also discussed in Chapter 1; Pickard and Thimann 1966; Wayne et al. 1990; Wayne and Staves 1996). In the protoplast-pressure theory, the hydrostatic pressure of the entire protoplast is suggested to trigger conformational changes of gravireceptor molecules at the plasma membrane. The cell then perceives the direction of gravity by sensing the differential tension and compression between the plasma membrane and the extracellular matrix at the top and at the bottom of the cell, respectively. In the following, we will provide evidence that both models of gravity sensing are realized in single-cell systems.

7.7 Susception in the Statolith-based Systems of Chara Characean rhizoids and protonemata are among the best-studied gravisensory cell types in which the cytoskeleton-based susception apparatus is well-understood. In downwardgrowing Chara rhizoids, BaSO4-crystal-filled vesicles rather than starch-filled amyloplasts serve as statoliths. The high density of BaSO4 and the vesicle size of 1 to 2 µm make these statoliths ideal for indicating the direction of gravity to the cell. Removal of statoliths from the tip abolishes gravitropic responsiveness (Sievers et al. 1991b), which clearly indicates that gravity signaling is triggered by these intracellular sedimentable particles. Thus, the sensory system for gravitropism is not obscured by alternative and redundant mechanisms as appears to be the case for higher plants. Statoliths in Chara rhizoids do not passively fall into the tip. In fact, they are actively kept in an area 10 to 35 µm above this region (Figure 7.2) (Hejnowicz and Sievers 1981; Braun 2002). Myosin-like proteins, which were found attached to the surface of statoliths (Braun 1996a), interact with predominately axially arranged actin microfilaments to prevent statoliths from settling into the tip by exerting net-basipetal forces (Braun and Wasteneys 1998). In tip-upward-growing protonemata of Chara, actomyosin forces prevent statoliths from sedimenting toward the cell base by acting net-acropetally (Hodick et al. 1998; Braun et al. 2002). Polar organization and gravity-oriented tip growth were shown to be dependent on the

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Figure 7.2. Apical part of a Chara rhizoid. Statoliths sediment onto the subapical plasma membrane upon gravistimulation. The growth direction is reoriented by a reduction of elongation growth of the lower cell flank. Diameter of the cell is 30 µm.

highly dynamic actin cytoskeleton that is regulated by the concerted action of numerous associated proteins (Braun et al. 2004). Inhibitor studies have shown that disrupting the actin cytoskeleton in rhizoids and in protonemata terminated the actomyosin-driven polarized growth and caused statoliths to drop into the tip or toward the nucleus, respectively (Hejnowicz and Sievers 1981; Bartnik and Sievers 1988; Sievers et al. 1996). After removing the drug, statoliths were quickly repositioned and gravity-oriented tip growth was restored as soon as the actin cytoskeleton was rearranged and fully functional (Braun 2001). Experiments conducted under microgravity conditions provided by parabolic flights of sounding rockets (TEXUS, MAXUS) and during Space Shuttle missions (IML-2, S/MM05), as well as experiments in simulated weightlessness provided by twodimensional (fast-rotating) and three-dimensional clinostats, have unravelled the specific contributions of gravity and actomyosin to the interplay of forces that underlie the statoliths-based gravity-sensing (susception) apparatus of Chara rhizoids and protonemata (Buchen et al. 1993, 1997; Cai et al. 1997; Hoson et al. 1997; Braun et al. 2002). When the influence of gravity was abolished during the microgravity phases of sounding rocket flights (Volkmann et al. 1991; Buchen et al. 1993) and randomized during rotation on clinostats (Hoson et al. 1997; Braun et al. 2002), actomyosin forces generated a displacement of statoliths against the former direction of gravity. Thus, in vertically-oriented rhizoids and protonemata at 1 ⫻ g, the statoliths are kept in a dynamically stable equilibrium position by actomyosin forces which exactly compensate the effect of gravity on the statoliths (Figure 7.3). Detailed analysis of the movements of statoliths in microgravity and of statoliths

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Figure 7.3. Gravitropic phases in characean rhizoids and protonemata. In rhizoids, the position of the statoliths (St) is balanced by net-basipetally acting actomyosin forces (Factin) and gravity (Fgravity). Upon gravistimulation, actomyosin forces guide the sedimenting statoliths toward the gravireceptors located in a beltlike area of the plasma membrane 10 to 35 µm above the tip. The Spitzenkörper (SpK) remains arrested at the tip and the calcium gradient (indicated by darker and lighter grey dotted areas) is always highest at the tip. When statoliths contact the gravireceptors (GR), graviperception takes place and is followed by a local reduction of cytosolic Ca2+ that results in differential extension of the opposite cell flanks (double-headed arrows). In upward-growing protonemata (lower row), the effect of gravity on statoliths is compensated by netacropetally acting actomyosin forces. Upon horizontal positioning, sedimenting statoliths are guided to the gravireceptors which are located near the growth center at the tip. This causes a drastic shift of the calcium gradient and the Spitzenkörper (SpK) upward where the new outgrowth occurs. The white arrows point to the area of maximal calcium influx. MT, microtubule; MY, myosin; SpKc, center of the Spitzenkörper.

which were displaced to different cell regions by optical laser tweezers or by centrifugation revealed the surprising complexity of forces by which actomyosin controls statolith positioning (Braun et al. 2002). Individual acropetal and basipetal movements of statoliths can be observed in both cell types, indicating that statoliths interact with the mainly axially oriented actin microfilaments with opposite polarities. When statoliths were centrifuged into the subapical region, statolith transport back to the original position was observed which is not notably influenced by gravity (Sievers et al. 1991b; Braun and Sievers 1993). Active transport is generated along actin microfilaments and statoliths do not sediment onto the lower cell flank until they have reached the statolith region near the tip, where sedimentation is not constrained by microtubules (Braun and Sievers 1994). Microtubules are excluded from the apical region of rhizoids and protonemata. The actomyosin component of movement is always the strongest pointing toward the statolith region. This ensures that statoliths are always kept in, or are retransported to, their original position. In the statolith region itself, however, gravity plays the decisive

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role as an additional passive transport component that contributes to the resting positioning of statoliths as long as the rhizoid tip points downward. Any changes in the orientation of the cells with respect to the direction or the amount of the acceleration results in a disturbance of this balance and a displacement, namely, the lateral sedimentation of statoliths (Figure 7.3). In lateral directions, statolith position is only weakly controlled by the actomyosin system in both cell types (Leitz et al. 1995; Buchen et al. 1997). Recently, the forces required to move statoliths in the lateral direction were determined during the 13-minute microgravity phases of two MAXUS rocket flights. In rhizoids growing vertically downward, lateral acceleration forces in a range of 0.1 ⫻ g were sufficient to displace statoliths toward the membrane-bound gravireceptors. Based on the known size of statoliths and the density differences between statoliths and the surrounding cytoplasm, it was calculated that molecular forces in a range of 2 ⫻ 10–14 N must be exerted on a single statolith in the lateral direction to overcome the cytoskeletal bonds and induce sedimentation, thus eliciting graviperception (Limbach et al. 2005). The need for the complex actomyosin-based control of statoliths position for gravity perception became fully comprehensible only after it was discovered that only the subapical plasma membrane area of the statolith region, 10 to 35µm from the cell tip, is able to trigger the gravitropic response upon statolith sedimentation (Braun 2002). Forcing statoliths to sediment outside these areas by using optical laser tweezers or centrifugation did not result in a gravitropic response (Braun 2002). After reorienting rhizoids by 90 degrees, the statoliths sediment mainly along the gravity vector and settle onto the lower cell flank of the statolith region where graviperception takes place and the graviresponse is initiated. However, when cells were stimulated at angles different from 90 degrees, statoliths did not simply follow the gravity vector (Figure 7.3) (Hodick et al. 1998). Instead, even in almost fully inverted cells, statoliths were actively redirected against gravity and were guided to the small gravisensitive area of plasma membrane in this “statolith region” of the tip. Gravistimulation of tip-upward-growing protonemata causes an actin-mediated acropetal displacement of statoliths into the apical dome, where they sediment very close to the tip (Figure 7.3) (Hodick et al. 1998). In contrast to rhizoids, the gravisensitive plasma membrane area in protonemata is limited to an area 5 to 10 µm basal to the tip (Braun 2002). During the upward bending of protonemata, the statoliths periodically sediment along the gravity vector and leave the gravisensitive site, which deactivates the gravireceptor. The periods when the statoliths exit the gravisensitive region are reflected by phases of straight growth. Actomyosin-mediated transport of statoliths back to the gravisensitive membrane area reinitiates the gravitropic response from time to time until the vertical orientation is resumed (Figure 7.3). On one hand, these actin-mediated susception mechanisms guarantee a highly efficient readjustment of the growth direction and, on the other hand, avoid inexpedient responses to transient stimuli. Gravity sensing and negative gravitropic (upward) response mechanisms in other single-celled systems like the protonemata of mosses, especially Ceratodon purpureus, are less well-understood. It is thought that these cells use starch-filled amyloplasts as statoliths. Should they be using the “protoplast pressure” mechanism for graviperception discussed above (i.e., the density-difference between the cell´s protoplast and the sur-

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Figure 7.4. Models of graviperception in different protists. Ca2+- and K+-mechanoreceptor channels are located in specific areas of the plasma membrane (ant = anterior cell pole). Receptor channels are activated by the mechanical gravity-induced load of the protoplast (direction of gravity indicated by arrows). In Loxodes, specialized gravireceptor organelles (the Müller organelles) complement gravity sensing by using a statolithbased perception mechanism. Activation of the receptor channels affects the membrane potential and regulates the activity of motion organelles: cilia in Paramecium and Loxodes (not shown) and a single flagellum in Euglena. Modified from Hemmersbach and Bräucker (2002).

rounding medium triggers graviperception), graviorientation should be abolished in neutral-buoyancy experiments, whereas an intracellular gravisensor, such as a starch statolith-based system, should still be functional under these conditions. Recent experiments have shown that gravitropism in this moss takes place in media that are denser than the cytoplasm of the apical cell (Sack et al. 2001). This result provides evidence that gravity sensing in Ceratodon strictly relies on sedimentation of intracellular masses rather than on the mass of the entire protoplast. Like in Chara rhizoids and protonemata, the statolith-based sensory system is not complemented by alternative mechanism.

7.8 Susception in the Statolith-based System of Loxodes The ciliate Loxodes (Figure 7.1) maintained its positive gravitaxis when the density of the external medium was in the same range as or even higher than the density of its cytoplasm (1.03 g/cm3). As gravikinesis of Loxodes was slightly reduced under isodensity conditions, Neugebauer et al. (1998) proposed that in Loxodes an intracellular gravisensing mechanism is complemented by a protoplast pressure-based mechanism (Figure 7.4). By analogy to the situation in Chara, Loxodes uses BaSO4 as the statolith material. Loxodes possesses 5 to 25 so-called Müller organelles (Müller vesicles), which are 7- to 10-µm-wide vacuoles containing a body of BaSO4 (3–3.5 µm in diameter) fixed to a mod-

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ified ciliary stick. It has been suggested that gravity is perceived by bending the ciliary complex, which induces changes in membrane potential, regulating the activity of the body cilia (Fenchel and Finlay 1984, 1986).

7.9 Susception in the Protoplast-based Systems of Euglena and Paramecium Starting in the nineteenth century, different hypotheses have been proposed to explain the mechanism of gravitaxis in unicellular organisms either involving a pure physical mechanism, a physiological process, or a combination of both (for reviews, see Bean 1984; Machemer and Bräucker 1992; Häder et al. 2005). Although the physical mechanisms assume a passive alignment of the cell in the water column caused by, for example, the cell being tail-heavy, physiological mechanisms predict the existence of an active gravireceptor. To summarize the results from decades of experimentation—some of which will be mentioned below—a physiological gravity signal transduction pathway exists in unicellular systems and, thus, the existence of a gravity-sensing mechanism can be predicted in free-swimming organisms. For example, after immobilizing Paramecium cells (Figure 7.1), Kuznicki (1968) observed not only sedimentation of the cells but also their variable orientation, in contradiction to a purely physically determined mechanism (buoyancy principle) of gravitaxis (Fukui and Asai 1985). In contrast to the results obtained from Ceratodon and Loxodes, increasing the density of the medium impaired graviorientation of Euglena (Figure 7.1) (density 1.046 g/ml_1) and Paramecium (density 1.054 g/ml_1). The capacity for orientation was completely disturbed under isodensity conditions. In addition to indications for a physiologically guided mechanism of graviperception, these results support the hypothesis of protoplast-based graviperception in Paramecium and Euglena. It was speculated that membrane-located gravisensors are early inventions of evolution, whereas the Müller bodies of Loxodes are later acquisitions due to adaptation to special living conditions (e.g., living in the sediment of lakes).

7.10 Graviperception in the Statolith-based Systems of Chara For graviperception to occur in characean rhizoids, statoliths must be fully settled on the gravisensitive subapical plasma membrane area (Figures 7.2 and 7.4; see also Braun 2002). Recent experiments that have been performed during parabolic flights of the A300 Zero-G aircraft elucidated the mode of gravireceptor activation in characean rhizoids (Limbach et al. 2005). The final curvature angles of flight samples, which experienced several short phases of microgravity, were compared to controls in a 1 ⫻ g reference centrifuge which had experienced the same conditions of the flight, except for the excursions of microgravity. The data revealed that statoliths, which were weightless but still in contact with the plasma membrane during the microgravity phases, still activated the membrane-bound gravireceptor. It therefore could be ruled out that the pressure exerted by the weight of statoliths is required for gravireceptor activation. This finding was supported by control experiments on the ground which demonstrated that increasing the

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weight of sedimented statoliths by lateral centrifugation did not enhance the gravitropic response. However, graviperception was terminated within seconds when the contact of statoliths with the plasma membrane was interrupted by inverting gravistimulated cells. This result provides evidence that graviperception in characean rhizoids relies on direct contact allowing as-yet unknown components on the statoliths’ surface to interact with membrane-bound receptors rather than on pressure or tension which is caused by the weight of the statoliths (Limbach et al. 2005). A mechanoreceptor was postulated to represent the receptor because the gravitropic responses of many plant organs seem to obey the sine law of gravitropism (Galland 2002). The pressure that statoliths exert on the receptor at different gravistimulation angles would explain the sinusoidal dependency, but the observation that the number of statoliths which settle on the receptor area in characean rhizoids decreases with the steepness of the angle can equally well account for this dependency.

7.11 Graviperception in the Statolith-based System of Loxodes In order to show that Müller vesicles function as gravisensory organelles in Loxodes, these structures were destroyed by laser ablation within individual cells. Observation of the manipulated cells revealed that they were no longer able to orient to the gravity vector, though their vitality and swimming velocity were unaffected (Hemmersbach et al. 1998). As discussed earlier in this chapter, it has been proposed that Loxodes perceives gravity through bending of the ciliary complex (Figure 7.4), thus inducing changes in the membrane potential. Such changes modify the activity of the body cilia, affecting the swimming behaviour (Fenchel and Finlay 1984, 1986). It is still unknown whether the gravity-induced “pull” of the BaSO4-granulum is directly transduced via ion channels in the plasma membrane, or whether it requires second messengers. Besides a cellular gravisensor, it is postulated from isodensity experiments that Loxodes also has a protoplast-based perception mechanism (Figure 7.4). In order to determine the minimum acceleration that is necessary to induce a graviresponse, Loxodes, Paramecium, and Euglena were exposed to increasing acceleration steps from microgravity to 1.5 ⫻ g, or vice versa, on a centrifuge microscope in space. With this experimental approach, threshold values for gravitaxis have been determined: Loxodes ⱕ 0.15 ⫻ g, Paramecium: 0.3 ⫻ g (Hemmersbach et al. 1996a), Euglena: ⱕ 0.16 ⫻ g and 0.12 ⫻ g (Häder et al. 1996, 1997). The existence of such thresholds implies that gravitaxis is the result of a physiological signal transduction chain.

7.12 Graviperception in the Protoplast-based Systems of Paramecium and Euglena Due to the lack of distinct statoliths in most protists, it was proposed that the mass of the whole cytoplasm causes a mechanical load on the lower cell membrane, thus stimulating mechano-(gravi)-sensitive ion channels. This sensory mechanism was termed the statocyst hypothesis, which corresponds to the protoplast-pressure theory (for review, see Machemer and Bräucker 1992). Thus, in this model, mechano-sensitivity is a prerequi-

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site for graviperception. In Paramecium and other ciliates, mechano-sensitive ion channels are distributed in a characteristic, bipolar manner in the plasma membrane: mechano-sensitive K+-channels mainly posteriorly and mechano-sensitive Ca2+-channels mainly anteriorly (Figure 7.4; for a full discussion of mechano-sensitive channels, see Chapter 5). In upward-swimming cells, the mechanical load of the protoplast is supposed to induce an outward-directed force onto the lower part of the cell membrane, thus opening mechano-sensitive K+-channels, which hyperpolarizes the cell. In turn, membrane hyperpolarization promotes an increase of swimming velocity, which can be measured in upward-swimming cells. In contrast, in a downward-swimming cell, the swimming velocity is reduced due to depolarization of the membrane potential by mechanical stimulation of mechano-sensitive Ca2+ channels. Such a distinct activation of two kinds of receptor populations delivers a reasonable explanation for gravitaxis and gravikinesis (Baba et al. 1991; Machemer and Bräucker 1992). Correspondingly, in Euglena, stretch-activated gravireceptor channels are proposed to be located asymmetrically in the anterior cell membrane (Figure 7.4). In an upwardswimming (negative gravitactic) Euglena, these channels are closed. If the longitudinal axis of the cell deviates from the vertical or if the cell swims downward, these channels open (Lebert and Häder 1996; Lebert et al. 1997; H¨äder et al. 2005), thereby inducing a depolarization of the plasma membrane, followed by activation of an intracellular signal transduction cascade that results in a reorientation of the cell. If graviperception in these systems occurs via gravisensory channels, it should be possible to measure a “gravireceptor potential.” This was done by intracellular electrophysiological recording in Paramecium and in the ciliate Stylonychia. Depending on the cell’s orientation with respect to the gravity vector, a hyperpolarization or a depolarization was registered after turning a cell upside-down (Gebauer et al. 1999; Krause 2003). In the case of Euglena, a direct measurement of electrical potentials has not been successful. However, an involvement of the membrane potential in graviperception in these cells was supported by experiments in which the lipophilic cation TPMP+ (triphenyl-methylphosphonium) and the ionophore calcimycin were used to alter the cellular membrane potential. Both compounds resulted in a loss of gravitaxis. In another experimental approach, Richter et al. (2001a) used voltage-sensitive dyes such as Oxonol to observe changes in membrane potential depending on the Euglena cell’s orientation within the gravity field. These experiments led to the discovery of a close relationship between precision of orientation and membrane potential. Interestingly, the channel blocker gadolinium blocked gravitaxis in Euglena, suggesting that stretch-activated channels function as gravisensory ion channels in this species (Lebert et al. 1997). Corresponding experiments with Paramecium showed no specific effect on gravitaxis, indicating that gadolinium is not specific for mechano-sensitive channels in this cell (Nagel and Machemer 2000). Regardless of the nature of the mass and the gravireceptor, graviperception can only occur when sufficient energy is provided to the system to overcome thermal noise in the receptor. Graviresponses are found in protists of different size and cell volume, ranging from Euglena gracilis (2.6 ⫻ 103 µm3) to Paramecium caudatum (327 ⫻ 103 µm3) or even the giant ciliate Bursaria truncatella (30 ⫻ 106 µm3). Energetic considerations show that in larger species, gating energy for gravisensory channels is clearly above the ther-

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mal noise level (2 ⫻ 10–21 J). However, for smaller species such as Euglena or even smaller systems such as human cells (lymphocytes), the signal-to-noise ratio may be limiting. This ratio is even more critical if one considers threshold values in the range of 0.1 ⫻ g to 0.3 ⫻ g for graviperception. Therefore, in these systems the involvement of supporting amplifying structures, such as the microfilament system of Loxodes, may facilitate graviperception (Häder et al. 2005).

7.13 Signal Transduction Pathways and Graviresponse Mechanisms in the Statolith-based Systems of Chara Although several studies clearly indicate that gravireceptor molecules are located in the subapical plasma membrane area of the statolith region (Braun 2002), the nature of the membrane-bound gravireceptor and the immediate downstream physiological steps involved in graviperception in rhizoids and protonemata remain to be clarified. Results obtained from the local application of ions and channel blockers by means of microcapillaries suggest that membrane-potential changes might be among the earliest steps following gravireceptor activation (Braun, unpublished results). However, more data have been published in the last decade that illuminate the cellular and molecular processes that underlie the gravitropic response. The smooth, downward curvature response of a rhizoid is best described as “bending by bowing,” whereas the response of a protonema was described as “bending by bulging” (Braun 1996b), referring to the bulge that initially appears on the upper cell flank and indicates the drastic upward shift of the growing tip. The Spitzenkörper (a tip-growth organizing complex consisting of endoplasmic reticulum, actin filaments, and a dense network of vesicles) (Braun 1997) and, consequently, also the center of maximal growth are displaced upon gravistimulation of protonemata by intruding statoliths (Figure 7.3). Although rhizoids can be forced to respond to some extent like protonemata, this can only be done by pushing statoliths aymmetrically into the apical dome with optical tweezers or by centrifugal forces greater than 50 ⫻ g (Braun 2002). There is evidence from centrifugation experiments (Braun 1996b; Hodick and Sievers 1998) and from attaching particles to the surface of gravitropically responding rhizoids (Sievers et al. 1979) that the position of the growth center at the cell tip is relatively stable and that the Spitzenkörper is more tightly anchored by cytoskeletal forces in rhizoids than in protonemata. The specific properties of the actin cytoskeleton, which have been shown to be responsible for Spitzenkörper anchorage, are controlled by calcium. Interestingly, calcium imaging demonstrated a drastic shift of the steep, tip-high calcium gradient toward the upper flank during initiation of the graviresponse in protonemata (Figure 7.3), but not in rhizoids (Braun and Richter 1999). In accordance with this observation, dihydropyridine fluorescence (indicating the tip-focused distribution of putative calcium channels) was also displaced toward the upper flank in graviresponding protonemata, but not in rhizoids (Braun and Richter 1999). Calcium imaging studies in rhizoids conducted by Simon Gilroy and coworkers (unpublished results) indicated that the impact of statolith sedimentation in rhizoids is limited to a local decrease in the concentration of cytosolic calcium in the area where sta-

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toliths sedimented onto the lower subapical cell flank. It is speculated that this reduction is due to a local inhibition of calcium channels that is initiated by statolith-induced gravireceptor activation. The subsequent reduction of the rate of exocytosis of secretory vesicles causes differential growth of the opposite cell flanks and results in the positively gravitropic curvature growth (Figure 7.3). The data obtained from characean protonemata suggest that the early asymmetric distribution of the calcium gradient (which precedes the negative graviresponse in protonemata) results either from statolith-induced repositioning of calcium channels or, more likely, by differential activation and/or inhibition of apical calcium channels (Braun and Richter 1999). Such processes would lead to an asymmetric influx of calcium, thus altering the pattern of exocytosis and causing an asymmetric incorporation of calcium channels. The asymmetric influx of calcium could mediate a repositioning of the Spitzenkörper and the growth center by differentially regulating actin anchorage or the activity of actin-associated proteins along the shifting calcium gradient (Braun and Richter 1999). The resulting polarity change would lead to the new growth direction (Figure 7.3). Additional support for the proposed gravitropic response mechanism in protonemata comes from immunofluorescence labeling of spectrin-like proteins in the actin-rich area that contains the ER aggregate in the center of the Spitzenkörper. The signal, which localizes to the median cell axis during vertical growth, is drastically displaced toward the upper flank—the site of future outgrowth—during initiation of the graviresponse in protonemata, clearly before curvature is recognizable (Braun 2001). In contrast, the same labeling in rhizoids gives a signal that remains symmetrically positioned in the apical dome throughout the graviresponse. These findings confirm that a repositioning of the Spitzenkörper is involved in the negative graviresponse of protonemata, but probably does not play a role in the positive graviresponse of rhizoids (Figure 7.3; Braun 2001). The tendency of protonemata to reorient toward the former growth axis after only short gravistimulation phases indicates that the new growth axis induced by the upward shift of the Ca2+ gradient is rather labile and may require actin anchorage to stabilize the new growth direction (Braun and Richter 1999; Braun 2001).

7.14 Signal Transduction Pathways and Graviresponse Mechanisms in Euglena and Paramecium The important role of gravity for cellular orientation and confirmation of the correctness of the term “gravi” for dedicated tactic and kinetic responses of motile microorganisms were shown by experiments in microgravity and in simulated weightlessness. Exposing a culture of ciliates with a preferred upward orientation to the conditions of microgravity resulted in a random distribution after 80 seconds in a rocket experiment and after 120 seconds in a clinostat experiment (Hemmersbach-Krause et al. 1993b). Similar behavioural responses were observed for Euglena (Vogel et al. 1993). Long-term cultivation of Euglena (Häder et al. 1996), Paramecium, and Loxodes (Hemmersbach et al. 1996a) for up to 12 days in space did not affect the reactivity of the cells to changing accelerations, indicating no persisting adaptation phenomena to microgravity. Hypergravity proved to be an ideal tool to enhance weak responses to gravity as gravitaxis and gravikinesis be-

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come more pronounced (up to 9 µg in the case of Paramecium) (Hemmersbach et al. 1996b; Bräucker et al. 2001). When considering likely elements of the gravitactic signal-transduction pathway, it was assumed that second messengers such as Ca2+, cAMP, cGMP, and calmodulin might play a role in this process, as they are clearly involved in the regulation of ciliary and flagellary activity; in the case of cAMP, a coupling to hyperpolarizing cellular events has been shown (Bonini et al. 1986; Schultz and Schönborn 1994). Application of phosphodiesterase inhibitors (IBMX, caffeine, 8-bromo-cyclo-AMP) to Euglena cultures, with the aim of increasing intracellular cAMP levels, resulted in an increase in the precision of negative gravitaxis, whereas decreasing cAMP levels inhibited the capacity of the cells for gravitactic orientation. Determination of the cAMP levels of Paramecium and Euglena under different acceleration conditions revealed significant accelerationdependent changes (Tahedl et al. 1998; Häder et al. 2005). Indeed, experiments in hypergravity and microgravity showed changes in cAMP in Paramecium according to the predication of the “statocyst” (protoplast-pressure) hypothesis: a decrease in cAMP was found in microgravity where no stimulation of the lower membrane should occur and an increase was found under hypergravity conditions due to an increased mechanical load on the lower membrane compared to 1 ⫻ g conditions (Häder et al. 2005). Direct visualization of second messenger changes in living cells are promising approaches to clarify the exact underlying mechanism and the time course of events. Using the chlorophyll-free flagellate Astasia longa loaded with calcium Crimson dextran and excited by laser light, researchers have investigated a possible correlation between the cytosolic calcium concentration and the orientation of the cell. In this context, a 180-degree turn of a negative-gravitactic Astasia culture induced an increase in the calcium signal, with a maximum after 30 seconds correlated with the reorientation of the cells. When performed in space, this experiment revealed that microgravity-adapted Astasia cells showed an increase in calcium-dependent fluorescence signal when accelerated above their threshold for gravitaxis. An image analysis system established a correlation between the calcium-dependent fluorescence signal and the swimming direction: cells moving in parallel to the acceleration vector showed a low signal and cells moving perpendicular to the vector displayed a high signal (Richter et al. 2001b). Acceleration-dependent changes in intracellular calcium levels in Euglena were also observed in a recent parabolic flight experiment onboard the Airbus A300 Zero-G (Richter et al. 2002). Together, these data support the existence of a calcium influx during reorientation of misaligned cells (Richter et al. 2001b, 2002).

7.15 Conclusions The multitude of significant findings that have increased our knowledge of gravitysensing and gravity-response mechanisms strongly emphasizes the usefulness of unicellular model systems for this field of research. Thus, the details of the molecular and cellular processes of gravity sensing are not obscured by a multiplicity of systems or by redundant processes, as appears to be the case for gravitropism in higher plants (Barlow 1995; Kiss 2000). Although the gravity-dependent responses of single-celled systems are

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obviously very different, the presented experimental data reveal that especially the early phases of gravity sensing share common features. The gravisensory processes can be reduced to two principles: perception via intracellular statoliths and via the whole protoplast. Gravisensory ion channels and cascades of ubiquitous second messengers are predicted to operate in most gravity-dependent signaling pathways, and have been identified in some cases. Finally, the cytoskeleton has been shown to play a master role in the complex process of gravity sensing and graviorientation. One fascinating question, further discussed in Chapters 8 and 9, will hopefully be answered in the near future: What will be the impact of long-term exposure to microgravity conditions in multigeneration experiments on the physiology of specialized gravisensory cells?

7.16 Acknowledgments The authors thank the several space shuttle crews, the teams of EADS ST, Kayser-Threde, Deutsches Zentrum für Luft- und Raumfahrt (DLR), NASA, Swedisch Space Corporation (SSC), Novespace, and the European Space Agency (ESA) for their dedicated work and for stimulating discussions. This work was financially supported by Deutsches Zentrum für Luft- und Raumfahrt (DLR) on behalf of the Bundesministerium für Bildung und Forschung (50WB9998 and 50WB0515).

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Lebert M and Häder D-P. 1996. How Euglena tells up from down. Nature 379:590. Lebert M, Richter P and Häder D-P. 1997. Signal perception and transduction of gravitaxis in the flagellate Euglena gracilis. Journal of Plant Physiology 150:685–90. Leitz G, Schnepf E and Greulich KO. 1995. Micromanipulation of statoliths in gravity-sensing Chara rhizoids by optical tweezers. Planta 197:278–288. Limbach C, Hauslage J, Schaefer C and Braun M. 2005. How to activate a plant gravireceptor— early mechanisms of gravity sensing studied in characean rhizoids during parabolic flights. Plant Physiology 139:1–11. Machemer H and Bräucker R. 1992. Gravireception and graviresponses in ciliates. Acta Protozoologica 31:185–214. Machemer H, Machemer-Röhnisch S, Bräucker R and Takahashi K. 1991. Gravikinesis in Paramecium: theory and isolation of a physiological response to the natural gravity vector. Journal of Comparative Physiology A 168:1–12. Mennigmann HD and Lange M. 1986. Growth and differentiation of Bacillus subtilis under microgravity. Naturwissenschaften 73:415–7. Nagel U and Machemer H. 2000. Effects of gadolinium on electrical membrane properties and behavior in Paramecium tetraurelia. European Journal of Protistology 36:161–8. Neugebauer DC, Machemer-Röhnisch S, Nagel U, Bräucker R and Machemer H. 1998. Evidence of central and peripheral gravireception in the ciliate Loxodes striatus. Journal Comparative Physiology A 183: 303–311. Ooya M, Mogami Y, Izumi-Kurotani A and Baba SA. 1992. Gravity-induced changes in propulsion of Paramecium caudatum: a possible role of gravireception in protozoan behaviour. Journal Experimental Biology 163: 153–167. Perbal G and Driss-Ecole D. 2003. Mechanotransduction in gravisensing cells. Trends in Plant Science 8:498–504. Pickard BG and Thimann KV. 1966. Geotropic response of wheat coleoptiles in absence of amyloplast starch. Journal of General Physiology 49:1065–1086. Richter P, Lebert M, Korn R and Häder D-P. 2001a. Possible involvement of the membrane potential in the gravitactic orientation of Euglena gracilis. Journal of Plant Physiology 158: 35–9. Richter P, Lebert M, Tahedl H and Häder D-P. 2001b. Calcium is involved in the gravitactic orientation in colorless flagellates. Journal of Plant Physiology 158:689–97. Richter PR, Schuster M, Wagner H, Lebert M and Häder D-P. 2002. Physiological parameters of gravitaxis in the flagellate Euglena gracilis obtained during a parabolic flight campaign. Journal Plant Physiology 159: 181–190. Sack FD. 1997 Plastids and gravitropic sensing. Planta 203:S63–S68. Sack FD, Schwuchow JM, Wagner T and Kern V. 2001. Gravity sensing in moss protonemata. Advances in Space Research 27:871–876. Salisbury FB. 1993. Gravitropism: changing ideas. Horticultural Review 15:233–278. Schultz JE and Schönborn C. 1994. Cyclic AMP formation in Tetrahymena pyriformis is controlled by K+-conductances. FEBS Letters 356:322–6. Sievers A, Heinemann B and Rodriguez-Garcia MI. 1979. Nachweis des subapikalen differentiellen Flankenwachstums im Chara-Rhizoid während der Graviresponse. Zeitschrift für Pflanzenphysiologie 91:435–442. Sievers A, Kruse S, Kuo-Huang L-L and Wendt M. 1989. Statoliths and microfilaments in plant cells. Planta 179:275–278. Sievers A, Buchen B, Volkmann D and Hejnowicz Z. 1991a. Role of the cytoskeleton in gravity perception. In The Cytoskeletal Basis for Plant Growth and Form, edited by C.V. Lloyd, pp. 169–182. London: Academic Press.

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Sievers A, Kramer-Fischer M, Braun M and Buchen B. 1991b. The polar organization of the growing Chara rhizoid and the transport of statoliths are actin-dependent. Botanica Acta 104:103–109. Sievers A, Buchen B and Hodick D. 1996. Gravity sensing in tip-growing cells. Trends in Plant Science 1:273–279. Tahedl H, Richter P, Lebert M and Häder D-H. 1998. cAMP is involved in gravitaxis signal transduction of Euglena gracilis. Microgravity Science Technology 11:173–8. Tash JS and Bracho GE. 1999. Microgravity alters protein phosphorylation changes during initiation of sea urchin sperm motility. FASEB Journal 13:S43–S54. Vogel K, Hemmersbach-Krause R, Kühnel C and Häder D-P. 1993. Swimming behavior of the unicellular flagellate, Euglena gracilis, in simulated and real microgravity. Microgravity Science Technology 4: 232–237. Volkmann D, Buchen B, Hejnowicz Z, Tewinkel M and Sievers A. 1991. Oriented movement of statoliths studied in a reduced gravitational field during parabolic flights of rockets. Planta 185:153–161. Yamamoto K and Kiss JZ. 2002. Disruption of the actin cytoskeleton results in the promotion of gravitropism in inflorescence stems and hypocotyls of Arabidopsis. Plant Physiology 128: 669–681. Yoder TL, Zheng H-Q, Todd P and Staehelin L-A. 2001. Amyloplast sedimentation dynamics in maize columella cells support a new model for the gravity-sensing apparatus of roots. Plant Physiology 125:1045–1060. Wayne R, Staves MP and Leopold AC. 1990. Gravity-dependent polarity of cytoplasmic streaming in Nitellopsis. Protoplasma 155:43–57. Wayne R and Staves MP. 1996. A down to earth model of gravisensing or Newton’s Law of gravitation from the apple’s perspective. Physiologia Plantarum 98:917–921.

8

Space-Based Research on Plant Tropisms Melanie J. Correll and John Z. Kiss*

8.1 Introduction—The Variety of Plant Movements Although plants are generally considered stationary organisms, they do move in response to a variety of environmental stimuli. Plants can move their organs and direct their growth to avoid harmful situations or to find important resources for survival. The movements of plants can be classified into three main categories: nastic responses, circumnutations, and tropisms. Nastic movements, sometimes called turgor movements, include motion of plants to external environments that do not depend on the direction of the stimulus source. Examples include motion due to changes in temperature (thermonasty), light (photonasty), touch (haptonasty), humidity (hydronasty), chemicals (chemonasty), as well as other stimuli. Nastic movements may allow the plant to protect itself from harmful elements or improve growth, development, and reproductive opportunities. These movements can be seen in the opening and closing of some flowers due to the light/dark cycles during the day, or the movement of leaves of the “sensitive plant” (Mimosa) when touched (see Chapter 5). The mechanisms and level of interactions of nastic movements with other plant movements are unclear. In contrast, circumnutations are the endogenous oscillatory movements of plants around a central axis. Circumnutation may be a simple consequence of growth and may serve as a mechanism for plant movements to adjust plant growth from movements that overshoot the plumb line of gravity (Barlow et al. 1994; Antonsen et al. 1995; Kiss 2006). The purpose of circumnutation is unknown and may simply be a consequence of general growth processes. In Arabidopsis thaliana, circumnutation of organs has been shown to be modulated by the circadian clock, with greatest movement occurring during dawn (Niinuma et al. 2005). The role of gravity in circumnutation is still unclear, although recent evidence suggests that gravity and circumnutation are inherently linked (Kitazawa et al. 2005; Kiss 2006; Yoshihara and Iino 2006). Tropisms are the directed growth of a plant organ in response to external stimuli. Unlike nastic movements, tropisms depend on the direction of the stimulus. Curvature of a plant organ toward the stimulus is termed positive tropism, and curvature away is termed negative tropism. The best-characterized tropisms are directed growth in response to gravity (gravitropism; see also Chapters 1 and 2), light (phototropism; Chapter 4), water (hydrotropism; Chapter 6), touch (thigmotropism; Chapter 5) and oxygen (oxytropism; Chapter 6), although many other tropisms have been reported. Since several types of plant movements can occur simultaneously, the ability to study one movement without the interactions from other movements is extremely difficult. For example, on Earth, gravity is ubiquitous. Thus, movements induced by gravity inevitably *Corresponding author 161

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interact with movements induced by other stimuli (Barlow 1995). The relatively strong gravitropic responses of plants can mask comparatively weaker tropisms, such as hydrotropism or oxytropism. Therefore, in order to better characterize tropisms, scientists have used several methods to reduce the effects of gravity on plant motion (Figure 8.1). These methods include the use of the following: • Plant mutants that have reduced responses to gravity; • Experimental systems that prevent plants from receiving unidirectional gravity acceleration (i.e., clinostats or random positioning machines); • Microgravity/free-fall environments. The extent to which these techniques reduce the effects of gravity on plant movement has been a topic of much debate, with each method having unique advantages and disadvantages (Sievers and Hejnowicz 1992; Kordyum 1997; Correll and Kiss 2002). In this chapter, experimental results from microgravity experiments are compared with other ground-based studies on plant movements for experiments performed from ~1990 to the present. Although we will concentrate on discussing ground-based and microgravity research on tropisms, other movements such as circumnutation will also be briefly considered.

8.2 The Microgravity Environment The gravitational acceleration on Earth is approximately 9.8 m/s2, defined as 1g. The term microgravity has been used to refer to levels that are less than 1g, typically 10–3g to 10–6g. In this chapter, we consider microgravity to range from approximately 1% of Earth’s gravitational acceleration (0.01g) to approximately 1 millionth of the Earth’s gravitational acceleration (10–6g). Other terms that have been used to describe different levels of gravitational accelerations include hypogravity (accelerations less than 1g but greater than 10–3g), weightlessness (net sum of forces acting on a body equaling zero), zero-g (an object that does not experience any gravitational pull), and hypergravity (accelerations greater than 1g) (Schaefer et al. 1993; Klaus 2001). Since gravity can influence other nongravity-related movements of plants, scientists have attempted to reduce gravitational effects below biologically detectable levels to study certain types of plant movements. One way to mitigate the effects of gravity is to travel away from the major source of gravitational interference, the Earth. Unfortunately, to reduce the effects of gravity on plants to 1 millionth of Earth’s gravity, you would have to travel approximately 6.4 million km away from Earth (Rogers et al. 1997). To put this in context, the moon orbits the Earth at approximately 380,000 km. Thus, studies on plant movement at this distance are unlikely in the near future, so other options are necessary to mitigate the effects of gravity on plants. The most effective way to reduce gravity effects on objects is through the use of free fall. An object is in free fall when it is falling under the sole influence of gravity, thus neglecting air friction. Free fall can be described as a condition where no external forces acting on the body of interest produce stress and that any forces cannot be detected by the

Figure 8.1 (also see Color Section). Potential methods to simulate the microgravity environment of low orbit spaceflight. A. A classic two-dimension clinostat that rotates at 1 to 3 rpm in the direction indicated by the white arrow. Petri dishes on the clinostat contain Arabidopsis seedlings. B. A three-dimensional clinostat, also known as a random positioning machine. The sample is placed on the inner frame, and the randomization is controlled by computer software. C. The free-fall phase (which lasts approximately 25 sec) of a parabolic flight on a NASA airplane. (Note: the flight also contains a hypergravity phase after the free fall.) The methods in A and B attempt to simulate microgravity but can be considered “omnilateral” gravity stimulation, whereas the parabolic flights (C) achieve the actual free-fall state of microgravity as noted by the orientation of the scientist.

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object of interest (i.e., the plant). All of the studies with plants in microgravity discussed in this chapter are actually with plants in free fall. Some mechanisms to create free fall or a microgravity environment include the use of drop towers, parabolic flights of airplanes, rockets, and orbiting satellites such as the Space Shuttle and space stations (Figure 8.1 and Color Section). The quality and duration of microgravity created in these environments are variable. For example, the longest drop tower (located in a mine shaft in Japan) is 460 m in length and only provides approximately 10 sec of free fall (Tokyo 1993). Parabolic flights are when a plane or rocket climbs rapidly at a ~45-degree angle then descends at a ~45-degree angle (Figure 8.1C). The parabolic flight between these two maneuvers can reduce gravitational effects to ~10–2g for about 20 sec for airplanes (Pletser 1995) and ~10–5g for up to several minutes for rockets (European Space Agency 2006). However, since many tropistic responses of plants can take up to several minutes before detection, the short duration of microgravity provided by drop towers and parabolic flights limits the types of studies that can be performed with these methods. Studies with drop towers and parabolic flights have been successfully used with protists to study the gravitaxis (orientation with respect to the gravity vector) (Häder and Hemmersbach 1997; see Chapter 7) or to study elements in the gravity-induced-signal cascade, such as movement of statoliths in plant roots (Volkmann et al. 1991). Parabolic flights are useful, relatively inexpensive laboratories to study the very early phases of gravity responses in plants. However, the best methods for studying tropisms and other plant responses in microgravity are through the use of biosatellites and orbiting spacecraft. Biosatellites (unmanned missions into space) can provide long-term microgravity, but typically missions are of about two weeks’ duration. Orbiting space stations such as Skylab, Mir, or the International Space Station (ISS) have offered the potential to study plant tropisms in a microgravity environment indefinitely. All of the methods used to create a reduced-gravity environment have limitations in their effectiveness as a microgravity laboratory. For example, the quality of acceleration on orbiting craft depends on the orbital motion of the spacecraft, the position of the item on the spacecraft, and the aerodynamic drag on the craft. Luckily, these accelerations are only about 10–6g in magnitude, which is relatively small and possibly below the sensory threshold for plant responses to gravity (Shen-Miller et al. 1968; Sobick and Sievers 1979; Merkys and Laurinavicius 1990; and see below). However, other accelerations can be created by vibrations in orbiting craft from fans, pumps, centrifuges, and crew activity. For the Space Shuttle, these accelerations were found to be about 10–4g, which may affect some experimental results since the minimum threshold values for root and shoot curvature may range in the values of 10–4g and 10–3g, respectively (Shen-Miller et al. 1968; Sobick and Sievers 1979; Merkys et al. 1986; Merkys and Laurinavicius 1990). Therefore, scientists using orbiting spacecraft for microgravity studies should consider these additional accelerations in their analyses. Other issues that influence the quality of experiments on plant tropisms performed in space, aside from gravitational effects, include artifacts from poor gas composition and exchange, lack of sufficient lighting, radiation effects, unscheduled/scheduled power out-

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ages, and lack of biological replicates. Several experiments have noted poor gas composition or exchange rates which resulted in unusual growth responses of plants in microgravity (Porterfield et al. 1997; Kiss et al. 1998, 1999; Perbal et al. 1996). In addition, many experiments performed in space have not been replicated, which limits the scientific interpretation of results (Cogoli 1996). The major impedances to studying plant tropisms in space are the limited opportunities and the associated costs. Recent delays in Space Shuttle flights to the ISS have further limited the amount of science that can be performed in the future. The future outlook for opportunities to study plant movements in space is also discussed in Chapter 9 of this book.

8.3 Ground-based Studies: Mitigating the Effects of Gravity The problems associated with studying biological responses to gravity on space missions have encouraged scientists to develop alternative ground-based methods to simulate microgravity. Devices such as clinostats and random positioning machines (also known as three-dimensional clinostats) have been used to reduce gravity effects on plants for tropism studies (Figure 8.1). A clinostat generally rotates the specimen around one axis whereas a random positioning machine offers three-dimensional rotation (Salisbury 1993; Hoson et al. 1997). The extent to which clinostats simulate microgravity has been debated, since results from clinostats do not always correspond to those from true microgravity experiments (Brown et al. 1976; Sievers and Hejnowicz 1992; Heathcote et al. 1995a; Brown et al. 1996; Kordyum 1997; Klaus 2001). A comparison of experimental results of plant and other organisms in microgravity with results from clinostats has been reviewed (Brown et al. 1976; Albrecht-Buehler 1992; Kordyum 1997) and is discussed in more detail in the subsequent sections of this chapter. Another ground-based attempt to reduce gravity effects on plant movements is through the use of mutants in gravity perception. For example, mutants of Arabidopsis that have reduced activity in the phosphoglucomutase gene (PGM) have reduced responses to gravity (Kiss et al. 1989). The pgm plants have reduced amounts of starch-filled amyloplasts in the cells involved in the perception of gravity. An otherwise cryptic positive, red lightinduced phototropism in roots was discovered with studies using these plants (Ruppel et al. 2001). Interestingly, hypergravity experiments performed with pgm mutants have shown that gravitropic orientation of organs of the pgm plants could be restored with accelerations of 5g for roots and 10g for hypocotyls (Fitzelle and Kiss 2001). Other starchdeficient mutants (e.g., agravitropic pea, Pisum sativum) have been used to study other tropisms such as hydrotropism or oxytropism that may be masked by gravitropic response (Jaffe et al. 1985; Porterfield and Musgrave 1998). Another Arabidopsis mutant in the gravity-perception phase lacks its endodermal layer (scr, SCARECROW mutants), the location of statoliths in shoots (Fukaki et al. 1996; Fukaki et al. 1998). These mutants have been used to study gravitropism and other plant movements such as circumnutation (Kitazawa et al. 2005). Currently, there are only a few mutants that have been identified as being specific to gravity perception in plants, so studies like these are limited.

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8.4 Gravitropism On Earth, the direction of the gravitational acceleration vector can indicate the location of water/nutrients (i.e., underground) and sunlight (i.e., aboveground). Not surprisingly, gravity is a relatively strong stimulus in terms of directing plant growth. Typically, primary roots grow toward the direction of gravitational acceleration (positive gravitropism), and shoots grow away from the direction of gravitational acceleration (negative gravitropism). Other organs in plants, such as inflorescence stems, leaves, and lateral roots and branches also display gravitropic responses, but their orientation may be an intermediate angle relative to the gravity vector (Hangarter 1997; Kiss et al. 2002; Mano et al. 2006). After reorientation, a plant organ will curve to reach the appropriate orientation, called the gravitational set point angle (Mullen and Hangarter 2003). The gravitational set point angle in plants can be observed in the patterns of branching in trees and orientation of lateral and primary roots. The temporal steps of gravitropism can be separated into three main stages: perception, signal transduction, and response. Details of these stages are found in Chapters 1, 2, and 3 and are described in Kiss (2000). Microgravity and simulated-microgravity environments have been used to study each stage of gravitropism as well as gravity-induced morphogenesis of plants and the straightening of a plant organ after a gravitropic response (autotropism) (Stankovic et al. 1998). In the following sections, experiments that use microgravity as a tool to study the elements of gravity perception, signal transduction, and response in gravitropism are described. Gravimorphogenesis of plants has been reviewed in a paper by Takahashi (1997). 8.4.1 Gravitropism: Gravity Perception Several models for gravity perception in plants have been proposed, although two models currently dominate (Perrin et al. 2005; Chapter 1). One model, the starch-statolith hypothesis, suggests that perception of gravity is mediated by the settling of dense organelles (statoliths) after a plant is reoriented (Figure 8.2) (Kiss 2000). In flowering plants, the amyloplasts in specialized cells function as statoliths; in roots, the amyloplasts are in the columella cells; and in shoots they are located in the endodermal cells (reviewed in Masson et al. 2002). The other model for gravity perception in plants suggests that the entire mass of the protoplast is involved in perceiving gravity (Staves et al. 1997). To date, neither of these models has been excluded and both may be acting simultaneously. Alternatively, another as-yet undiscovered mechanism of gravity detection may be involved in gravity perception in plants (reviewed in Wolverton et al. 2002; Chapter 1). Microgravity has been a useful tool to explore the mechanisms of gravity perception in plants. For several plant species, the statoliths (i.e., amyloplasts in plants) in root columella cells of plants grown in microgravity were located in different regions of the cell compared to the location of statoliths in cells of roots from 1g controls (Driss-Ecole et al. 2003). For example, most of the amyloplasts in Arabidopsis roots were located in the proximal two-thirds of the cell in plants grown in microgravity compared to amyloplasts from 1g-grown plants, where most of the amyloplasts were located in the distal one-third of the cell (Briarty et al. 1995).

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Figure 8.2. Electron micrographs of root cap columella cells from Arabidopsis seedlings grown in a 1g control (A) and in microgravity (B) during spaceflight. The amyloplasts (arrows) in the 1g control are in the distal (“bottom”) part of the cell, and the gravity vector is toward the bottom of the figure. In contrast, the amyloplasts (arrows) in the microgravity sample are dispersed throughout the cytoplasm. N = nucleus.

For other plants, such as white clover (Trifolium repens) and lentil (Lens culinaris) roots, amyloplasts were grouped near the center of the cell in microgravity whereas amyloplasts remained close to the distal half in 1g controls (Perbal and Driss-Ecole 1989; Lorenzi and Perbal 1990; Smith et al. 1997). The grouping of amyloplasts was also found in plants grown on clinostats, but to a lesser extent (Smith et al. 1997). Nonrandom grouping of amyloplasts in apical cells from moss (Ceratodon purpureus) was also found in clinostat- and microgravity-treated plants (Kern et al. 2001). Although the mechanisms of the grouping of amyloplasts are unknown, it appears that the statocytes do not move as a group in microgravity (Driss-Ecole et al. 2000). It also appears that amyloplasts relocate within 6 minutes of microgravity treatments (Volkmann et al. 1991; Driss-Ecole et al. 2000). Future experiments in microgravity may help reveal the kinetics and mechanisms of amyloplasts’ movement in response to changes in acceleration and provide a better understanding of the grouping effect of amyloplasts in microgravity. Other studies on gravity perception mechanisms have been performed using plants that have reduced starch content and therefore smaller amyloplasts. Plants grown in microgravity that were then treated to 1g accelerations for 60 minutes showed greater curvature in roots from wild-type plants compared to roots from starch-deficient mutants (Kiss et al. 1998). These results further suggest that the amyloplasts are involved in the mechanisms of gravity perception. However, restoration of the gravitropic response in these mutants could be accomplished by treating plants with hypergravity, indicating that the starchfilled amyloplasts are not required for a gravitropic response (Fitzelle and Kiss 2001). The detailed mechanisms of gravity perception in plants are still unclear (Chapter 1). However, studies in microgravity have shown that amyloplast location in root columella cells moves in response to free fall in a potentially nonrandom orientation. It appears that

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plants which develop in microgravity are still responsive to gravitational acceleration, suggesting that components of gravity perception do not need gravity to be expressed. Future studies in microgravity may identify new mechanisms of gravity perception in plants and identify downstream elements in the gravity signal transduction cascade. 8.4.2 Gravitropism: Signal Transduction Once the gravity signal is perceived in plants, downstream elements in the signaling cascade begin to react while the response (curvature) phase begins. Elements downstream of sensing include changes in calcium, pH, flavanoids, ethylene, relocalization of auxin carriers (PIN proteins), and auxin redistribution which results in differential elongation of a plant organ (Masson et al. 2002; Chapter 2). Since all tropisms result in the differential growth of an organ, many of the downstream elements involved in gravitropism overlap with other plant tropisms. Few studies have been performed in microgravity on the downstream elements in plant tropisms because these elements are difficult to measure on Earth, even without the additional constraints imposed in the space environment. Some of the studies that have been performed using microgravity to identify downstream elements in signal transduction are briefly described here. An experiment with lentil roots was performed to identify the role of actin cytoskeleton in amyloplast movement in microgravity (Driss-Ecole et al. 2000). Amyloplasts from roots treated with a drug that blocks actin polymerization (cytochalasin D) did move in response to microgravity, but the rate of movement was slowed compared to nontreated roots (Driss-Ecole et al. 2000). These authors proposed that the microfilament bundles were not completely depolymerized with the cytocalasin D treatment, which allowed for limited movement of amyloplasts. The limited rate of movement of the amyloplasts in the cytochalasin D samples suggests that the cytoskeleton is involved, at some level, in gravity-induced responses. Other ground-based studies with drugs that disrupt the actin-cytoskeleton have also demonstrated that the cytoskeleton is involved in gravity responses, although results from these studies are often conflicting and depend on the organ, plant species, drug dosage, and experimental conditions (Friedman et al. 2003; Palmieri and Kiss 2005). Auxin transport is another downstream element in tropisms that has been studied using microgravity. Polar auxin transport was inhibited in the internode segments of pea but was promoted in maize (Zea mays) coleoptiles (Ueda et al. 1999). The mechanisms of auxin transport during tropism in microgravity have not been studied but, since seedlings can curve in response to stimuli in microgravity, it appears that auxin transport during tropisms does not need gravity to function. Other studies with maize noted similar amounts of auxin in plants grown in microgravity and 1g controls (Schulze et al. 1992). Calcium redistribution is believed to be involved in signal transduction during tropisms. Free calcium was redistributed in sweet clover (Melilotus alba) root columella cells in microgravity compared to 1g controls and clinorotated seedlings (Hilaire et al. 1995). Fewer calcium precipitates were found with the nucleus and amyloplasts in cells exposed to microgravity compared to 1g and clinorotated controls. The causes of the redistribution of the calcium-associated precipitates in microgravity are unclear. Additional studies on calcium redistribution in microgravity may help further characterize its role in gravity signaling.

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Figure 8.3. Images of roots of lentil seedlings grown in a 1g control (A) and in microgravity (B) during spaceflight. Roots of the 1g control are straight and oriented relative to gravity (toward the bottom of the figure). In contrast, roots of the microgravity-grown seedlings appear to have a more random orientation. Photographs are courtesy of Dr. Dominique Driss-Ecole.

Studies with downstream elements in the signal-transduction cascade of tropisms in microgravity may provide significant contributions to our understanding of gravity effects on plant growth and movement. Currently, plants have been developed with green fluorescent proteins that are expressed along with proteins in the downstream components of gravity signaling (e.g., PIN2-GFP plants) (Abas et al. 2006). Studies monitoring the expression and localization of these proteins in microgravity may prove useful to identify how downstream elements in gravity-induced signaling respond to different gravitational accelerations. 8.4.3 Gravitropism: The Curving Response As the gravity signal-transduction process has reached a significant level of propagation downstream, the response or curvature of the organ can be detected. Without gravity as a cue to direct plant growth, plants appear to grow randomly. For example, roots of cress (Lepidium sativum) plants and hypocotyls of Arabidopsis show a vertical orientation in 1g but appear to grow randomly in microgravity (Figure 8.3) (Johnsson et al. 1996a; Johnsson et al. 1996b; Kiss et al. 1998; Kiss et al. 1999; Kiss 2000). Roots and hypocotyls also grew randomly when grown in simulated microgravity, such as on clinostats or random positioning machines (Kraft et al. 2000). After microgravity-grown plants are exposed to acceleration, their organs will curve in response to the direction of the acceleration vector (Kiss et al. 1998; Kiss et al. 1999). The following questions have been proposed to explore the sensitivity of plants to gravity: 1. What is the minimum time required for a plant to sense and respond to gravity (threshold duration or presentation time)?

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2. What is the minimum gravitational acceleration that can be detected in plants to induce a response (g-threshold)? 3. How do the presentation time and acceleration dose interact to induce a response (threshold dose; gravitational acceleration ⫻ time)? Various doses of acceleration have been provided to plants using clinostats, centrifuges, and microgravity to study the sensitivity of plants to gravitational acceleration (Brown et al. 1995; Volkmann and Tewinkel 1996; Hejnowicz et al. 1998; Perbal et al. 2004). Originally, it was hypothesized that the reciprocity rule for tropism governed gravity-induced curvature in plants. The reciprocity rule suggests that if g (gravitation acceleration) and t (time of stimulation) are varied reciprocally to maintain a constant product, then the curvature or response should be the same (Johnsson et al. 1995). For oat (Avena sativa) coleoptiles, reciprocity appeared to hold true for lower accelerations (0.2 to 1g) and stimulation times of 2 to 65 minutes for plants grown in microgravity and on clinostats (Johnsson 1965; Shen-Miller 1970; Johnsson et al. 1995). However, the sensitivity of oat to gravity also depended on the stage of the plant (i.e, height; Shen-Miller 1970; Johnsson 1965; Johnsson et al. 1995). For example, the threshold dose for taller plants showing detectable curvature was about 55 g·s, whereas for shorter plants it was approximately 120 g·s. Therefore, the developmental age or size of the plant may have an influence on the threshold dose that can be sensed. For other plant species, the reciprocity rule did not represent the response. For example, the curvature of cress roots in microgravity after a dose of 60 g·s for treatments of 0.1g for 600 seconds and 1g for 60 seconds should result in similar magnitudes if reciprocity were valid. However, results indicated that the 0.1g ⫻ 600 seconds treatment had about half (14 degrees) of the curvature compared to roots from the 1g ⫻ 60 seconds treatment (32 degrees; Volkmann and Tewinkel 1996, 1998). Other factors may affect the sensitivity of a plant to gravity, such as the growth history of the plant. For example, roots from cress grown in microgravity were more sensitive to the stimulation time than were roots from plants grown in a 1g centrifuge, with threshold doses of 30 g·s and 60 g·s, respectively (Volkmann and Tewinkel 1996). Enhanced curvature was also found for lentil roots previously grown in microgravity compared to plants that were grown on a 1g centrifuge (Perbal et al. 2004). Surprisingly, roots from transgenic and wild-type rapeseed (Brassica napus) during microgravity had no detectable curvature after 1 hour of centrifugation at 1g, whereas roots from ground controls curved significantly after reorientation (Iversen et al. 1996). These results were not attributed to altered growth rates between microgravity-grown plants and ground controls or the additional accelerations imposed by uncontrolled vibrations in the space craft, but may have been due to other technical difficulties imposed by the spaceflight environment. The threshold duration (i.e., the shortest duration of time capable of eliciting a detectable gravitropic response) in microgravity also appears to be variable. For oat, the response was less than 1 minute at 1g (Brown et al. 1995). These results are consistent with Arabidopsis roots, where less than 1 minute at 1g was also found to cause curvature (Kiss et al. 1989). However, some researchers propose, based on mathematical extrapolation, that the threshold value is 1 g·s, indicating that only 1 second at 1g (Earth) is required to

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elicit a curvature response (Perbal and Driss-Ecole 2002). Future studies in microgravity can clarify the nature of the duration of stimulus needed to elicit a response in plants. The lowest acceleration that a plant can detect has been estimated to be approximately 10–4g for roots and 10–3g for shoots (Shen-Miller et al. 1968; Merkys and Laurinavicius 1990). These results were obtained from studies with oat and lettuce (Lactuca sativa) plants grown on clinostats or in microgravity. Since experimental limitations could not provide the lower levels of gravitational acceleration, these authors used mathematical methods to predict the minimum acceleration that could be detected. Microgravity provided by spaceflight environments may not be low enough to study the minimum accelerations that plants can detect, mainly due to other acceleration on the craft from crew activity and centrifuges. Therefore, directly identifying the lowest magnitude of acceleration that plants can detect may be a challenge with today’s technology. Results from ground-based studies to determine the sensitivity of plants to gravity using clinostats have been conflicting. For example, the curvature of white clover roots due to gravity stimulation after previous growth in microgravity, 1g, or on a 2-D clinostat showed that roots only curved ~35 degrees from clinostat-grown plants, ~75 degrees from 1g-grown plants, and ~60 degrees from microgravity-grown plants after 10 hours (Smith et al. 1999). The authors suggest that the reduced curving response of roots grown on the clinostat treatment compared to both plants grown in microgravity and on 1g controls was due to root cap deterioration. Other researchers have found that roots from microgravity-grown plants are more sensitive to gravity compared to clinostat-grown plants (Lorenzi and Perbal 1990; Perbal and Driss-Ecole 1994; Volkmann and Tewinkel 1996; Perbal et al. 1997). Therefore, the application of clinostats to study gravity responses in plants is limited.

8.5 Phototropism Light is another important stimulus that is involved in determining the direction of plant growth. Therefore, plants have evolved a variety of photoreceptors to sense the quality, quantity, direction, and intensity of light. In flowering plants, the photoreceptors can be grouped into the blue/UV-A photoreceptors (cryptochromes and phototropins) and the red/far-red photoreceptors (phytochromes). Phototropic responses are largely controlled by the phototropins with the other photoreceptors modulating the response (Briggs and Christie 2002). Hypocotyls and stems typically curve toward blue or white light (positive phototropism), whereas roots typically curve away from the light source (negative phototropism). Since all phototropic responses of plants on Earth have to contend with gravity, it is not surprising that phototropism and gravitropism are constantly interacting to influence plant form. Recently, a gene downstream of phytochromes was identified in Arabidopsis that links gravity and light-regulated processes, GIL1 (Gravitropic in the Light 1) (Allen et al. 2006). The hypocotyls of gil1 mutants do not show the random gravitropic orientation found in wild-type plants grown in red or far-red light. Identification of this novel gene is a significant step toward understanding how light and gravity interact to determine plant form. The many interacting effects of phototropism and gravitropism are described in Chapter 4 and reviewed in Correll and Kiss (2002).

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The sensitivity of plants to light is difficult to study on Earth due to the interacting effects from gravity. Therefore, microgravity has been explored to determine the sensitivity of plants to different wavelengths and intensity of light. The first intensive study on plant phototropic responses in microgravity was performed on wheat (Triticum aestivum) seedlings (Heathcote et al. 1995a). Wheat coleoptiles showed a significantly enhanced curvature response to 6 or 9 seconds of photostimulation in microgravity compared to ground controls, although no significant differences in curvature were found after 3, 501, or 1,998 seconds of photostimulation (Heathcote et al. 1995a). It appears that in microgravity the phototropic curvature may be enhanced relative to plants grown in 1g, but this enhancement may also depend on the total fluence of the illumination provided. An enhanced curvature response to illumination was found from clinostat-grown plants relative to both space-grown and ground controls regardless of duration of stimulation (Heathcote et al. 1995a). Since enhanced curving responses were not found for all durations of light stimulation in microgravity, it appears that in this case, again, the clinostat does not effectively represent the microgravity environment provided in space. Other studies with maize also showed an enhanced curvature in response to directional illumination from coleoptiles grown on clinostats relative to 1g controls (Nick and Schäfer 1988). The phototropism of moss also was studied in microgravity during spaceflight (Kern and Sack 2001). Apical cells (protonemata) of moss can display both positive and negative phototropism to red light. The kinetics of alignment in the red light path (~1.5 µmol m–2s–1) was similar between protonemata from moss grown in 1g and in microgravity. However, for dark-grown cultures that were exposed to low irradiance of red light (~50 nmol m–2s–1), more of the protonemata (70%) had aligned in the light path (±45 degrees of the path) in microgravity compared to 1g controls. These results suggest that, on Earth, gravitropism and phototropism both interact at low fluence of red light to orient moss protonemata. Interestingly, dark-grown protonemata grew in spirals in microgravity and an experiment on Space Shuttle mission STS-107 was performed to study this response. However, due to the Space Shuttle Columbia accident, there were relatively few results from this study (Kern et al. 2005). Other experiments have been performed to study phototropism in plants in the microgravity environment, although results have been limited. A future experiment is planned for the ISS using hardware developed by NASA and the European Space Agency (ESA). This experiment, called TROPI for tropisms, will monitor phototropic responses of roots and hypocotyls of Arabidopsis to various gravitational accelerations (Figure 8.4 and Color Section) in both red and blue light (Correll et al. 2005). In addition, gene profile analyses on seedlings will be performed to study the interacting effects of light and gravity on gene expression patterns.

8.6 Hydrotropism, Autotropism, and Oxytropism All other tropisms directly interact with gravitropism, although only a few of these have been studied in space. Roots of some species of plants have shown a positive curvature toward water (i.e., hydrotropism; see Chapter 6) on Earth, but this can be masked by the gravitropic response. For example, hydrotropism in pea seedlings was evident in plants that

Figure 8.4 (also see Color Section). Photographs of the TROPI (for the study of tropisms) hardware and the European Modular Cultivation System (EMCS) for use on the International Space Station. A. The EMCS has standard Experimental Containers (ECs) that hold hardware unique to each specific experiment. In this case, the EC has several cassettes which are for the growth of Arabidopsis seedlings. Prior to launch, seeds are mounted on a sandwich of membranes and filter papers in the cassettes, and the experiment is activated in space by hydration of the seeds. B. An EC being inserted into the EMCS through the centrifuge door. The EMCS provides an incubator, lighting system, and high-resolution video which all are on a centrifuge palette. C. The EMCS with the unit door open to show the two centrifuge palettes (C1, C2). Each centrifuge has slots (ECS = EC slot) for four ECs, and there are two video cameras (Cm) which image the ECs via a polished mirror (M). The EMCS has a controlled gaseous atmosphere (with ethylene scrubbing) and a water delivery system. The centrifuge is off for microgravity experiments, but is capable of accelerations up to 2g for use as a control and for study of the effects of fractional g.

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were grown on a clinostat or in plants that were mutated in their response to gravity (Jaffe et al. 1985; Takahashi and Suge 1991; Takahashi et al. 1996). Hydrotropism was also evident in lateral roots from cucumber (Cucurbita ovifera) seedlings grown on a clinostat or in microgravity (Takahashi et al. 1999). The lateral roots from clinorotated and microgravity seedlings grew at approximately 40- to 50-degree angles from the primary root axis, and this curvature was toward the water source, whereas roots from ground controls grew ~90 degrees from the primary root axis. Therefore, although roots do respond to moisture gradients, it appears that this response is largely masked by gravity-induced responses. The seemingly random orientation of plant roots out of rooting matrices for plants grown in microgravity may in fact be a result of oxytropism (Porterfield and Musgrave 1998). On two space shuttle experiments (STS-54 and STS-68) (Porterfield and Musgrave 1998), roots of Arabidopsis plants grown in microgravity grew out of the rooting matrices in the direction of the oxygen gradient. The enzymatic activity, localization, and expression of mRNA of a protein associated with oxygen stress, alcohol dehydrogenase (ADH), were enhanced in roots grown in microgravity compared to ground controls (Porterfield et al. 1997). This observation suggested that roots may either be oxygen-deprived in microgravity or expressing ADH in response to another stress induced by spaceflight. To test whether oxygen availability was limited to plants during microgravity, a novel sensor was designed that measured oxygen availability (not concentration) in microgravity (Liao et al. 2004). Results using this sensor during parabolic flights showed that oxygen availability changes during periods of microgravity and, therefore, roots from plants grown in microgravity are likely experiencing oxygen deprivation. Experiments on the ground showed that indeed roots from both normal and agravitropic mutants of pea curved in response to oxygen gradients (Porterfield and Musgrave 1998). For example, roots from normal plants curved about half the amount of roots from agravitropic mutants after 48 hours of treatment. These results suggest that roots respond to gradients of oxygen, but this is largely masked by gravitropic responses. Autotropism is the straightening of an organ after the g vector is randomized on a clinostat or is reduced, as is found in microgravity (Stankovic et al. 1998). Cress roots that curved in response to various gravitational accelerations all underwent straightening once the acceleration was removed (Stankovic et al. 2001). Roots from clinorotated plants also showed autotropism, suggesting that the straightening process is a process that does not depend on the prestimulus orientation. Microgravity offers a unique ability to identify and study tropic responses of plants. Studies in microgravity can reduce the interacting effects of gravitropism with other tropisms, allowing the characterization of the tropism of interest without the ever-present gravity responses. It also seems likely that plants curve in response to a variety of as-yet unidentified stimuli which may only be found when grown in low g conditions, or with studies that use plants that lack the ability to perceive gravity.

8.7 Studies of Other Plant Movements in Microgravity The microgravity environment is a tool that can also be used to study a range of other plant movements and growth responses. These other movements of plants have been

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studied (sometimes simultaneously) with tropisms in spaceflight. For example, both gravitropism and circumnutations of lentil roots were measured in an experiment on the space shuttle (Antonsen et al. 1995; Perbal and Driss-Ecole 1994). These authors found that roots in microgravity displayed oscillations in growth along with gravitropic responses that varied as the function of the stimulation time. Ground-based studies with Arabidopsis have shown that roots grown on a random position machine make large loops in the right-handed direction, whereas in 1g, roots display a wavy, right-handed slant (Piconese et al. 2003). Similarly, nastic movements in coleoptiles of wheat seedlings were observed in plants in microgravity (Heathcote et al. 1995b). The extent of the interactions between oscillatory movements and gravitropism is unclear. Some evidence with plants mutated in genes involved in gravity responses has suggested that the oscillatory movements are coupled to gravitropism with auxin distribution being an overlapping element (Piconese et al. 2003; Kitazawa et al. 2005; reviewed in Kiss 2006). In contrast, evidence from spaceflight experiments has indicated that a variety of plants do have circumnutations in microgravity, suggesting that gravity is not necessary for circumnutation (Brown et al. 1990; Heathcote et al. 1995b; Antonsen et al. 1995). Future studies using microgravity may help to define the roles of gravity in these types of plant movements.

8.8 Space Flight Hardware Used to Study Tropisms The types of hardware that have been used to study plant tropisms in microgravity are variable since the spaceflight conditions can influence the hardware design. For example, experiments that did not have crew available, such as experiments performed on sounding rockets, biosatellites, or some space shuttle/station experiments, needed to have fully automated hardware. Other experiments had crew interaction, which increased the complexity associated with these types of studies. Some examples of the different types of plant facilities used in microgravity to study plant tropisms are described in the next section. In addition, hardware for plant growth studies in general has been reviewed (Lork 1988; Porterfield et al. 2003). The use of sounding rockets or biosatellites is a relatively cost-effective way to perform simple experiments in microgravity. Cress plants grown in sounding rockets (TEXUS program) were housed in containers that had temperature recording devices and an automatic fixation system (Tewinkel et al. 1991). The fixation system automatically flooded the containers holding the plants with chemical fixatives. Such in-flight fixation allowed further analysis on the ground to identify plastid location during the excursion into microgravity. Other hardware that has been used to study gravity effects on plant tropisms and growth is the Biological Research in Canister (BRIC) system. The canisters are basically light-tight cylinders that are placed aboard the Space Shuttle for experiments that have little or no crew involvement. One such experiment with the BRIC system studied phototropism and gravitropism interactions in moss. This hardware incorporated LEDs (light emitting diodes) (STS-87) for unilateral illumination and a chemical fixation system for moss plants grown in Petri dishes (Kern et al. 1999).

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Crew involvement in experiments allows for more complex experimental design and analyses. For example, two tropism experiments, one with wheat, called Phototropic Transients (FORTRAN), and another with oat, called Gravitational Threshold (GTHRES), were performed on the space shuttle implementing the International Microgravity Laboratory payload (IML-1; Heathcote et al. 1995a; Brown et al. 1995). These experiments were performed on the Spacelab module of the space shuttle mission STS-42 in 1992. Plants in these experiments were transported to the shuttle in specially designed containers called Plant Carry-On Containers (PCOCs). Once onboard, the containers were stowed in either the Mid-deck Ambient Stowage Insert (MASI) or housed in the Gravitational Plant Physiology Facility (GPPF). In the GPPF, plants were placed in containers called Plant Cubes, which had a window that allowed infrared radiation but no visible light to penetrate, and a second window that allowed blue light to penetrate during photostimulation periods. Photostimulation occurred when plants were placed on the Recording and Stimulus Chamber (REST). A culture rotor, which consisted of two centrifuges, provided a force of 1g, and a test rotor, which provided 0 to 1g, were both available for video recording of curvature responses to different gravitational accelerations. Ground controls for these experiments included a clinostat in a GPPF facility. These procedures required crew time to transport containers and perform the experiments, but also allowed for controls to be grown in space. Biorack is another facility that has been used to study tropisms in the space environment. It was developed by the ESA as a multi-user facility to study a variety of biological materials (Manieri et al. 1996). The Biorack facility had a cooler/freezer, a glovebox, and two incubators. Two types of containers have been available for studying biological specimens in the Biorack facility. Type I containers hold a volume of 65 ml and Type II containers hold a volume of 385 ml. Both Arabidopsis and lentil seedlings have been germinated and grown in Type I containers with Type II containers used as fixation devices (Perbal et al. 1987; Kiss et al. 1999). This hardware allowed for video monitoring of plant growth and curvature responses and, with the aid of the crew, the plants could be chemically fixed to study the movement of amyloplasts in gravity-perceiving cells once the samples had been returned to Earth. Ideally, in these spaceborne experiments, all environmental parameters except the one being studied should be controlled and monitored, including temperature, humidity, gas phase composition, and light intensity. A new facility called the European Modular Cultivation System (EMCS) has been developed by the ESA to study plant growth on the ISS. Applications of the EMCS to other biological systems are also being explored. The EMCS has also addressed the requirement to monitor many of the environmental parameters outlined above (Figure 8.4) (Brinckmann and Brillouet 2000; Brinckmann 2005). The EMCS has an incubator with two centrifuges, atmospheric and humidity control, ethylene scrubbers, video monitoring, and Experiment Containers (ECs) that interface with the Experiment Unique Equipment (EUE). The EUE can be custom-designed for each experiment’s requirements. A project is in development to study tropisms in Arabidopsis seedlings using the EMCS and video images, and frozen tissues will be returned to Earth for analyses (Correll et al. 2005).

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8.9 Future Outlook and Prospects With the advancement of molecular techniques and cellular imaging technologies, many of the key players involved in tropisms have been identified and localized within cells. However, how all these elements integrate and orchestrate tropisms during plant development is far from understood. Microgravity experiments have only begun to clarify the sensitivity of plants to gravity and light. It is likely that different species may be more sensitive to particular environmental stimuli, resulting in an enhanced tropistic response. Future long-term experiments that will use space stations as microgravity laboratories to study tropisms are likely to reveal new information about how plants have evolved in a 1g environment on Earth and to identify new tropistic responses that are masked by gravity. In addition to increasing basic knowledge, this information, in the long term, will aid in the development of plants for use in bioregenerative life support systems during space missions.

8.10 Literature Cited Abas, L., Benjamins, R., Malenica, N., Paciorek, T., Winiewska, J., Moulinier-Anzola, J.C., Sieberer, T., Friml, J., and C. Luschnig. 2006. Intracellular trafficking and proteolysis of the Arabidopsis auxin-efflux facilitator PIN2 are involved in root gravitropism. Nature Cell Biol. 8: 249–256. Albrecht-Buehler, G. 1992. The simulation of microgravity conditions on the ground. Grav. Space Biol. Bull. 5: 3–10. Allen, T., Ingles, P.J., Praekelt, U., Smith, H., and G.C. Whitlam. 2006. Phytochrome-mediated agravitropism in Arabidopsis hypocotyls requires GIL1 and confers a fitness advantage. Plant J. 46: 641–648. Antonsen F., Johnsson, A., Perbal, G., and D. Driss-Ecole. 1995. Oscillatory growth movements of roots in weightlessness. Physiol. Plant. 95: 596–603. Barlow, P.W. 1995. Gravity perception in plants: A multiplicity of systems derived by evolution? Plant Cell Environ. 18: 951–962. Barlow, P.W., Parker, J.S., and P. Brain. 1994. Oscillations of axial plant organs. Adv. Space Res. 14: 49–58. Briarty, L.G. Maher, E.P., and T.-H. Iversen. 1995. “Growth differentiation and development of Arabidopsis thaliana under microgravity conditions.” In C. Mattok (ed.), Biorack on Spacelab IML-1, European Space Agency, Noordwijk, The Netherlands, pp. 141–154. Briggs, W.R., and J.M. Christie. 2002. Phototropins 1 and 2: Versatile plant blue-light receptors. Trends Plant Sci. 7: 204–210. Brinckmann, E. 2005. ESA hardware for plant research on the International Space Station. Adv. Space Res. 36: 1162–1166. Brinckmann, E., and C. Brillouet. 2000. Space plant research on the ISS with the European Modular Cultivation System and with BIOLAB. SAE Tech. Papers Series Paper No. 2000– 01–2472. Brown, A.H., Dahl, A.O., and D.K. Chapman. 1976. Limitation on the use of the horizontal clinostat as a gravity compensator. Plant Physiol. 58: 127–130. Brown, A.H., Chapman, D.K., Lewis, R.F., and A.L. Venditti. 1990. Circumnutations of sunflower hypocotyls in satellite orbit. Plant Physiol. 94: 233–238.

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Brown, A.H., Chapman, D.K., Johnsson, A., and D. Heathcote. 1995. Gravitropic responses of the Avena coleoptile in space and on clinostats. I. Gravitropic response threshold. Physiol. Plant. 95: 27–33. Brown, A.H., Johnsson, A., Chapman, D.K., and D. Heathcote. 1996. Gravitropic responses of the Avena coleoptile in space and on clinostats. IV. The clinostat as a substitute for space experiments. Physiol. Plant. 98: 210–214. Cogoli, A. 1996. Biology under microgravity conditions in Spacelab International Microgravity Laboratory 2 (IML-2). J. Biotechnol. 47: 67–70. Correll, M.J., and J.Z. Kiss. 2002. Interactions between gravitropism and phototropism in plants. J. Plant Growth Regul. 21: 89–101. Correll, M.J., Edelmann, R.E., Hangarter, R.P., Mullen, J.P., and J.Z. Kiss. 2005. Ground-based studies of tropisms in hardware developed for the European Modular Cultivation System (EMCS). Adv. Space Res. 36: 1203–1210. Driss-Ecole, D., Jeune, B., Prouteau, M., Julianus, P., and G. Perbal. 2000. Lentil root statoliths reach a stable state in microgravity. Planta 211: 396–405. Driss-Ecole, D., Lefranc, A., and G. Perbal. 2003. A polarized cell: the root statocyte. Physiol. Plant. 118: 305–312. European Space Agency. 2006. Human Spaceflight: Sounding Rockets. Web page , accessed April 18, 2006. Fitzelle, K.J., and J.Z. Kiss. 2001. Restoration of gravitropic sensitivity in starch-deficient mutants of Arabidopsis by hypergravity. J. Exp. Bot. 52: 265–275. Friedman, H., Vos, J.W., Hepler, P.K., Meir, S., Halevy, A.H., and S. Philosoph-Hadas. 2003. The role of actin filaments in the gravitropic response of snapdragon flowering shoots. Planta 216: 1034–1042. Fukaki, H., Fujisawa, H., and M. Tasaka. 1996. SGR1, SGR2, SGR3: Novel genetic loci involved in shoot gravitropism in Arabidopsis thaliana. Plant Physiol. 110: 945–955. Fukaki, H., Wysocka-Diller, J., Kato, T., Fujisawa, H., Benfey, P.N., and M. Tasaka. 1998. Genetic evidence that the endodermis is essential for shoot gravitropism in Arabidopsis thaliana. Plant J. 14: 425–430. Häder, D.-P., and R. Hemmersbach. 1997. Graviperception and graviorientation in flagellates. Planta 203: S7–S10 Hangarter, R.P. 1997. Gravity, light and plant form. Plant Cell Environ. 20: 796–800. Heathcote, D.G., Brown, A.H., and D.K. Chapman. 1995a. The phototropic response of Triticum aestivum coleoptiles under conditions of low gravity. Plant Cell Environ. 18: 53–60. Heathcote, D.G., Chapman, D.K., and A.H. Brown. 1995b. Nastic curvatures of wheat coleoptiles that develop in true microgravity. Plant Cell Environ. 18: 818–822. Hejnowicz, Z., Sondag, C., Alt, W., and A. Sievers. 1998. Temporal course of graviperception in intermittently stimulated cress roots. Plant Cell Environ. 21: 1293–1300. Hilaire, E., Paulsen, A.Q., Brown, C.S., and J.A. Guikema. 1995. Microgravity and clinorotation cause redistribution of free calcium in sweet clover columella cells. Plant Cell Physiol. 36: 831–837. Hoson, T. Kamisaka, S., Masuda, Y., Yamashita, M., and B. Buchen. 1997. Evaluation of the threedimensional clinostat as a simulator of weightlessness. Planta 203: S187–S197. Iversen, T.-H., Ødegaard, E., Beisvåg, T., Johnsson, A., and O. Rasmussen. 1996. The behaviour of normal and agravitropic transgenic roots of rapeseed (Brassica napus L.) under microgravity conditions. J. Biotechnol. 47: 137–154. Jaffe, M.J., Takahashi, H., and R.L. Biro. 1985. A pea mutant for the study of hydrotropism in roots. Science 230: 445–447.

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Johnsson, A. 1965. Investigations of the reciprocity rule by means of geoptropic and geoelectric measurements. Physiol. Plant. 18: 945–967. Johnsson, A., Brown, A.H., Chapman, D.K., Heathcote, D., and C. Karlsson. 1995. Gravitropic responses of the Avena coleoptiles in space and on clinostats. II. Is reciprocity valid? Physiol. Plant. 95: 34–38. Johnsson, A., Karlsson, C., Chapman, D.K., Braseth, J.D., and T.H. Iversen. 1996a. Dynamics of root growth in microgravity. J. Biotechnol. 47: 155–165. Johnsson, A., Karlsson, C., Iversen, T.H., and D.K. Chapman. 1996b. Random movements in weightlessness. Physiol. Plant. 96: 169–178. Kern, V.D., Sack, F.D., White, N.J., Anderson, K., Wells, W., and C. Martin. 1999. Spaceflight hardware allowing unilateral irradiation and chemical fixation in situ in Petri dishes. Adv. Space Res. 24: 775-778. Kern, V.D., and F.D. Sack. 2001. Effects of spaceflight (STS-87) on tropisms and plastid positioning in protonemata of the moss Ceratodon purpureus. Adv. Space Res. 25: 941-949. Kern, V.D., Smith, J.D., Schwuchow, J.M., and F.D. Sack. 2001. Amyloplasts that sediment in protonemata of the moss Ceratodon purpureus are nonrandomly distributed in microgravity. Plant Physiol. 125: 2085-2094. Kern, V.D., Schwuchow, J.M. Reed, D.W., Nadeau, J.A., Lucas, J., Skripnikov, A., and F.D. Sack. 2005. Gravitropic moss cells default to spiral growth on the clinostat and in microgravity during spaceflight. Planta 221: 149-157 Kiss, J.Z. 2000. Mechanisms of the early phases of plant gravitropism. Crit. Rev. Plant Sci. 19: 551-573. Kiss, J.Z. 2006. Up, down, and all around: How plants sense and respond to environmental stimuli. Proc. Natl. Acad. Sci. USA 103: 829-830. Kiss, J.Z., Hertel R., and F.D. Sack. 1989. Amyloplasts are necessary for full gravitropic sensitivity in roots of Arabidopsis thaliana. Planta 177: 198-206. Kiss J.Z., Katembe, W.J., and R.E Edelmann. 1998. Gravitropism and development of wild-type and starch-deficient mutants of Arabidopsis during spaceflight. Physiol. Plant. 102: 493-502. Kiss, J.Z., Edelmann, R.E., and P.C. Wood. 1999. Gravitropism of hypocotyls of wild-type and starch-deficient Arabidopsis seedlings in spaceflight studies. Planta 209: 96-103. Kiss, J.Z., Miller, K.M., Ogden, L.A., and K.K. Roth. 2002. Phototropism and gravitropism in lateral roots of Arabidopsis. Plant Cell Physiol. 43: 35-43. Kitazawa, D., Hatakeda, Y., Kamada, M., Fujii, N.M., Miyazawa, Y., Hoshino, A., Iida, S., Fukaki, H., Morita, M.T., Tasaka, M., Suge, H., and H. Takahashi. 2005. Shoot circumnutation and winding movements require gravisensing cells. Proc. Natl. Acad. Sci. USA 102: 18742-18747. Klaus, D.M. 2001. Clinostats and bioreactors. Grav. Space Biol. Bull. 14: 55-64. Kordyum, E.L. 1997. Biology of plant cells in microgravity and under clinostating. Intl. Rev. Cytol. 171: 1-78. Kraft, T.F., Van Loon, J.J., and J.Z. Kiss. 2000. Plastid position in Arabidopsis columella cells is similar in microgravity and on a random-positioning machine. Planta 211: 415-422. Liao, J., Liu, G., Monje, O., Stutte, G.W., and D.M. Porterfield. 2004. Induction of hypoxic root metabolism results from physical limitation in O2 bioavailability in microgravity. Adv. Space Res. 34: 1579-1584. Lorenzi, G., and G. Perbal. 1990. Root growth and statocyte polarity in lentil seedling roots grown in microgravity or on a slowly rotating clinostat. Physiol. Plant. 78: 532-537. Lork, W. 1988. Experiments and appropriate facilities for plant physiology research in space. Acta Atronaut. 17: 271-275. Manieri, P., Brinckmann, E., and C. Brillouet. 1996. The Biorack facility and its performance during the IML-2 Spacelab mission. J. Biotechnol. 47: 71-82.

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Mano, E., Horiguchi G., and H. Tsukaya. 2006. Gravitropism in leaves of Arabidopsis thaliana (L.) Heynh. Plant Cell Physiol. 47: 217-223. Masson, P.H., Tasaka, M., Morita, M.T., Guan, C., Chen, R. and K. Boonsirichai. 2002. Arabidopsis thaliana: A model for the study of root and shoot gravitropism. In C.R. Somerville and E.M. Meyerowitz (eds.), The Arabidopsis Book. American Society of Plant Biologists, Rockville, MD, doi: 10.1199/tab.0009, www.aspb.org/publications/arabidopsis/, accessed April 26, 2007. Merkys, A.J., and R.S. Laurinavicius. 1990. Plant Growth in Space. In M. Asashima and G.M. Malacinski (eds.), Fundamentals of Space Biology, pp. 69-83. Japanese Science Society Press, Tokyo; Springer-Verlag, Berlin. Merkys, A., Laurinavicius, R., Bendoraityté, Svegzdiené, D., and O. Rupainiené. 1986. Interaction of growth determining systems with gravity. Adv. Space Res. 6: 71-80. Mullen, J.L. and R.P. Hangarter. 2003. Genetic analysis of the gravitropic set-point angle in lateral roots of Arabidopsis. Adv. Space Res. 31: 2229-2236 Nick, P., and E. Schäfer. 1988. Interaction of gravi- and phototropic stimulation in the response of maize (Zea mays L.) coleoptiles. Planta 173: 213-220. Niinuma, K., Someya, N., Kinura, M., Yamaguchi, I., and H. Hamamoto. 2005. Circadian rhythm of circumnutation in inflorescence stems of Arabidopsis. Plant Cell Physiol. 46: 1423-1427. Palmieri, M., and J.Z. Kiss. 2005. Disruption of the F-actin cytoskeleton limits statolith movement in Arabidopsis hypocotyls. J. Exp. Bot. 56: 2539-2550. Perbal, G., and D. Driss-Ecole. 1989. Polarity of statocytes in lentil seedling roots grown in space (Spacelab D1 mission). Physiol. Plant. 75: 518-524. Perbal, G., and D. Driss-Ecole. 1994. Sensitivity to gravistimulus of lentil seedling roots grown in space during the IML 1 mission of Spacelab. Physiol. Plant. 90: 313-318. Perbal, G., and D. Driss-Ecole. 2002. Contributions of space experiments to the study of gravitropism. J. Plant Growth Regul. 21: 156-165. Perbal, G., Driss-Ecole, D., Rutin, J., and G. Sallé. 1987. Graviperception of lentil seedling roots grown in space (Spacelab D1 Mission). Physiol. Plant. 90: 313-318. Perbal, G., Legué, V., and D. Driss-Ecole. 1996. Growth and gravisensitivity of lentil seedling roots grown in space. In Plant in Space Biology EDSSS, pp. 29-40, Tohoku University, Sendai, Japan. Perbal, G., Driss-Ecole, D., Tewinkel, M., and D. Volkmann. 1997. Statocyte polarity and gravisensing in seedling roots grown in microgravity. Planta 203: S57-S62. Perbal, G., Lefranc, A., Jeune, B., and D. Driss-Ecole. 2004. Mechanotransduction in root gravity sensing cells. Physiol. Plant. 120: 303-311. Perrin, R.M., Young, L.S., Murthy, U.M.N., Harrison B.R, Wang, Y., Will J.L., and P.H. Masson. 2005. Gravity signal transduction in primary roots. Ann. Bot. 96: 737-743. Piconese, S., Tronelli, G., Pippia, P., and F. Migliaccio. 2003. Chiral and non-chiral nutations in Arabidopsis roots grown on the random positioning machine. J. Exp. Bot. 54: 1909-1918. Pletser, V. 1995. Microgravity research during aircraft parabolic flights: The 20 ESA campaigns. ESA Bull. Available on web page , accessed April 24, 2007. Porterfield, D.M., Matthews, S.W., Daugherty, C.J., and M.E. Musgrave. 1997. Spaceflight exposure effects on transcription, activity, and localization of alcohol dehydrogenase in the roots of Arabidopsis thaliana. Plant Physiol. 113: 685-693. Porterfield, D.M., and M.E. Musgrave. 1998. The tropic response of plant roots to oxygen: Oxytropism in Pisum sativum L. Planta 206: 1-6. Porterfield, D.M., Neichitailo, G.S., Mashinski, A.L., and M.E. Musgrave. 2003. Spaceflight hardware for conducting plant growth experiments in space: The early years 1960-2000. Adv. Space Res. 31: 183-193.

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Rogers, M.J., Vogt, G.L., and M.J. Wargo. 1997. Microgravity: A teacher’s guide with activities in science, mathematics, and technology. NASA publication EG-1997-08-1100-HQ. Web page , accessed April 24, 2007. Ruppel, N.J., Hangarter, R.P., and J.Z. Kiss. 2001. Red-light-induced positive phototropism in Arabidopsis roots. Planta 212: 424-430. Salisbury, F.B. 1993. Gravitropism: Changing ideas. Hort. Rev. 15: 233-278 Schaefer, R.L., Jahns, G.C., and D. Reiss-Bubenheir. 1993. Plant response to the microgravity environment. In P.M. Gresshoff (ed.), Plant Responses to the Environment, pp. 59-70, CRC Press, Boca Raton, FL. Schulze, A., Jensen, P.J., Desrosiers, M., Buta, J.G., and R.S. Bandurski. 1992. Studies on the growth and indole-3-acetic acid and abscisic acid content of Zea mays seedlings grown in microgravity. Plant Physiol. 100: 692-698. Shen-Miller, J. 1970. Reciprocity in the activation of geotropism in oat coleoptiles grown on clinostats. Planta 92: 152-163. Shen-Miller, J., Hinchman R.R., and S.A. Gordon. 1968. Thresholds for georesponse to acceleration in gravity-compensated Avena seedlings. Plant Physiol. 43: 338-344. Sievers, A., and Z. Hejnowicz. 1992. How well does the clinostat mimic the effect of microgravity on plant cells and organs? ASGSB Bull. 5: 69-75. Smith, J.D., Todd, P., and L.A. Staehelin. 1997. Modulation of statolith mass and grouping in white clover (Trifolium repens) grown in 1-g, microgravity and on the clinostat. Plant J. 12: 13611373. Smith, J.D., Staehelin, L.A., and P. Todd. 1999. Early root cap development and graviresponse in white clover (Trifolium repens) grown in space and on a two-axis clinostat. J. Plant Physiol. 155: 543-550 Sobick, V., and A. Sievers. 1979. Responses of roots to simulated weightlessness on the fast-rotating clinostat. In R. Holmquist (ed.), COSPAR Life Sci. Space Res., Vol 17, pp. 285-290, Pergamon Press, Oxford. Stankovic, B., Volkmann, D., and F.D. Sack. 1998. Autotropism, automorphogenesis, and gravity. Physiol. Plant. 102: 328-335. Stankovic, B., Antonsen, F., Johnsson, A., Volkmann, D., and F.D. Sack. 2001. Autonomic straightening of gravitropically curved cress roots in microgravity. Adv. Space Res. 27: 915-919. Staves, M.P., Wayne, R., and A.C. Leopold. 1997. The effect of the external medium on gravitropic curvature of rice (Oryza sativa, Poaceae) roots. Am. J. Bot. 84: 1522-1529. Takahashi, H. 1997. Gravimorphogenesis: Gravity-regulated formation of the peg in cucumber seedlings. Planta 203: S164-S169. Takahashi, H., and H. Suge. 1991. Root hydrotropism of an agravitropic pea mutant, ageotropum. Physiol. Plant. 82: 24-31. Takahashi, H., Takano, M., Fujii, N., Yamashita, M., and H. Suge. 1996. Induction of hydrotropism in clinorotated seedling roots of Alaska pea, Pisum sativum L. J. Plant Res. 109: 335-337. Takahashi, H., Mizuno, H., Kamada, M., Fujii, N., Higashitani, A., Kamigaichi, S., Aizawa, S., Mukai, C., Shimazu, T., Fukui, K., and M. Yamashita. 1999. A spaceflight experiment for the study of gravimorphogenesis and hydrotropism in cucumber seedlings. J. Plant Res. 112:497-505. Tewinkel, M., Burfeindt, J., Rank, P., and D. Volkmann. 1991. Automatic fixation facility for plant seedlings in the TEXUS sounding rocket programme. Microgravity Sci. Tec. 4: 216-220. Tokyo, P.H. 1993. Goodness, gracious, great balls of fire! New Sci. 1857:9. Ueda, J., Miyamoto, K., Yuda, T., Hoshino, T., Fuiji, S., Mudai, C., Kamigaichi, S., Aizawa, S., Yoshizaki, I., Shimazu, T., and F. Fukui. 1999. Growth and development, and auxin polar trans-

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9

Plan(t)s for Space Exploration Christopher S. Brown*, Heike Winter Sederoff, Eric Davies, Robert J. Ferl, and Bratislav Stankovic

9.1 Introduction Mankind will explore the solar system. Some of that exploration will be done robotically, allowing vicarious human experience and study of extraterrestrial locations. However, the U.S. space program’s plans are replete with strategies to enable the first-hand human exploration of space. With the human system such an essential part of the long-term plan, technologies to keep humans alive and performing at full capacity in extraterrestrial environments must be developed. To accomplish the long-term goal of a stable human presence on other planetary bodies in the solar system, the development and integration of a dependable life support system is critical. Continued shipment and re-supply of the essentials for human survival— breathable air, clean water, and food—would be risky as well as prohibitively expensive. They are risky in that the shipments would depend on an absolutely fail-proof launch, transit, and landing system. They are expensive in that, with current technologies, the cost to get 1 kg into low Earth orbit is about $10,000. That’s a $3,000 hamburger! Therefore, success of the human expansion into the cosmos must coincide with the development of a life support system that is capable of regenerating all the essentials for survival. Such systems already exist—they are called plants. Using the primary processes of photosynthesis (air revitalization and biomass production) and transpiration (water purification), plants have provided human beings with all the essentials for survival as well as many of the nonessentials that make life interesting. To date, developing the engineering and infrastructure to get us into space has resulted in important advances in transport and propulsion systems. But this is not all that is needed for the successful human exploration of space. As stated by astrophysicist Freeman Dyson: “The chief problem for a manned mission [to space] is not getting there but learning how to survive after arrival. Surviving and making a home away from Earth are problems of biology rather than engineering. Any affordable program of manned exploration must be centered in biology, and its time frame tied to the time frame of biotechnology; a hundred years[...] is probably reasonable. To make human space travel cheap, we will need advanced biotechnology in addition to advanced propulsion systems.” (Dyson 1999)

The development of a plant-based, biologically regenerative life support system is therefore critical to providing the fundamental needs of a human crew. However, to be fully supportive in a potentially changing environment, such a system must also be capable of responding to the changing needs of the personnel onboard. Plants must be capable of quickly reprogramming their metabolism, physiology, and growth to keep the crew *Corresponding author 183

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functioning at full capacity. Developing a bioregenerative life support system based on such “programmable plants” is not only key to human expansion into the solar system, it is now also scientifically tenable by bringing together the emergent disciplines of genomics, nanoscience, biosensor technology, robotics, and intelligent machines. Combining these disciplines with our ever-expanding understanding of how plants sense and respond to their environment, as described in the previous chapters of this book, will allow manipulation of plant growth and development to sustain a human presence in space.

9.2 Human Missions to Space There are multiple reasons for sending humans into space, including scientific research, economic benefit, and psychological satisfaction derived from fulfilling the quest to explore and extend the human frontier. It is possible that in the distant future, mankind will have to colonize space as Earth becomes overcrowded or uninhabitable. It is inevitable that the exploratory missions will become outposts, outposts will become colonies, and colonies will become self-sustaining, independent, and permanent. For this to occur, all the elements of life support and permanent presence must be available and/or produced in situ. Short of this, human settlements in space will remain forever outposts, utterly dependent on Earth. One such space destination is Mars. Overall, the conditions on Mars are not so radically different from Earth (Carr 1996). It has been proposed that biological activity could be supported with some protective structure (Wheeler 1999). Although it is unlikely that there is life on the surface of Mars at present (McKay 1997), there is evidence that it may have existed there in the past (McKay et al. 1996). This compelling, yet controversial, speculation constitutes just one among many of the reasons for going to Mars, i.e., to search for clues of extinct or extant life. Other near-term human settlements in space include a permanent low-Earth-orbit station (currently the International Space Station) and the Moon. The success of these settlements will require a long-term human presence in space and on planetary surfaces other than Earth’s, which, of course, will require a robust, adaptable, and controllable life support system.

9.3 Life Support The four basic functions of any human life support system are to provide: • • • •

Suitable atmosphere; Clean water; Food; Waste processing.

Essentially, the options for life support are either re-supply or regeneration. Re-supply is simple to explain—essential survival supplies are shipped from Earth. As described

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earlier, this is both costly and risky. Regeneration involves physical/chemical methods for the synthesis of O2 from CO2, the production of clean water from wastewater, and the processing of waste (Eckart 1996). However, there are presently no physical/chemical technologies available that allow the synthesis of food from inorganic material. Bioregenerative systems, usually utilizing photosynthetic organisms and supporting engineering components, can provide all of these (Smernoff and MacElroy 1989). Current life support technologies for space exploration are entirely physico-chemical and are part of the re-supply philosophy that has served the U.S. space program well for the last several decades. However, as mission time frames and distances from Earth increase, generative, biologically based life support systems are increasingly favored over other types (Olsen 1982; Schwartzkopf 1992; Fogg 1995). In addition to air, water, food, and waste processing, other needs will come to the fore in a mission of extended length or on a self-sustaining colony. These include usable energy, materials (especially for construction and manufacturing), pharmaceuticals, and specialty chemicals. Although mining and chemical extraction of in situ resources can provide some of these needs (Clark 1989; Meyer and McKay 1989; Zubrin and Wagner 1996), biological systems with plants as an integral component eventually could supply most of them. Plants have other advantages. Their built-in systems for DNA replication, protein synthesis, and biomass production can be exploited to maximize mission success. Plants are solar-powered and naturally self-perpetuating, obviating the need for expensive and potentially unreliable re-supply missions. An added and very important feature of plants is that their seeds are small, easy to transport, tolerant of extreme environments, longlasting, and contain all the information needed for the subsequent plants to go through their entire life cycles. With the continued advances in the understanding of the genetic control underlying plant development and adaptation that can be reliably anticipated in the next 10 to 40 years, a life support system centered on plants could provide most of the resources necessary for mission success and independent sustainability.

9.4 Genomics and Space Exploration The field of plant gravitational and space biology has started to utilize genetic and genomic technology to understand and manipulate the basic mechanisms of plant response and adaptation to space conditions (Lomax et al. 2003). Transgenic Arabidopsis thaliana seedlings flown for a short period on the Space Shuttle were used to study unique space stress conditions (Paul et al. 2001), which in many ways were found to be similar to hypoxic stress. In a later report, genome-wide patterns of expression in space-flown Arabidopsis showed similarities to heat shock (Paul et al. 2005). However, no discernable differences in gene expression were detectable in 23-day-old wheat seedlings grown on the International Space Station when compared with the ground control plants (Stutte et al. 2006), suggesting that many of the stress responses linked to space flight may be more related to specific growth conditions rather than being an inherent component of the space environment. In ground-based studies, whole genome microarrays have been used to monitor global

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gene expression in gravitational and mechanically stimulated Arabidopsis roots in order to identify specificity and analyze temporal resolution for the very early responses to these stimuli at the molecular level (Kimbrough et al. 2004; Chapter 2). Transgenic plants modified in their activation of a very early signaling molecule [inositol-1,4,5triphosphate (InsP3)] were generated and showed a decreased and delayed response to gravity as well as an increased tolerance to other abiotic stresses (Salinas-Mondragon et al. 2005; Perera et al. 2006). Although Arabidopsis serves as an excellent model system, its behavior does not always predict the behavior of crop plants. The genetic differences between plant species that have evolved over millions of years and crop plants obtained by targeted breeding do not always allow for a direct translation of phenotypic effects of biotechnological manipulation. For example, tomato plants transformed with the same gene used to reduce InsP3-mediated signaling in Arabidopsis (discussed above) exhibit extreme drought-tolerance and have a very high stress-tolerant phenotype overall (Perera et al., in preparation; Khodakovskaya et al., in preparation)—phenotypes not readily predicted from the known effects of the transgene in Arabidopsis. Conceptual designs for greenhouses on Mars, on the Moon, and in Earth orbit incorporate artificial light, altered gravity, and low atmospheric pressure and composition, all of which are known to affect plant physiology and development (McKay et al. 1991; Corey et al. 2002). If plants can be developed that can thrive at less than Earth atmospheric pressures, then this will save on the materials and energy costs to build and maintain a “Martian” greenhouse. Using a low-pressure growth chamber, Paul et al. (2004) compared gene expression, as changes in transcript abundance, between wildtype Arabidopsis plants grown under Earthlike atmospheric pressure (101 kPa), low atmospheric pressure (10 kPa), and low oxygen concentration (2% O2 at 101 kPa). They identified genes regulated specifically by hypobaria or hypoxia, and genes regulated by both stresses. Under hypobaria, the most dramatic changes were found in desiccationassociated and abscisic acid-regulated pathways, indicating water stress (Paul et al. 2004). These kinds of assays suggest that plants which evolved on Earth do possess the genetic resources to become adapted to a Martian greenhouse environment. Genetic engineering is now routine for some plant species. Insertion of a single gene or a cluster of genes results in transgenic plants expressing new phenotypic traits. Examples include resistance to insects (Bt corn), resistance to high salt, improved tolerance to chilling, ozone resistance, altered starch concentration and composition, altered lipid composition, enhanced shelf life (Stitt and Sonnewald 1995; Giddings et al. 2000; Daniell et al. 2001; Qi et al. 2004; Khodakovskaya et al. 2006), enhanced vitamin and antioxidant content (Shintani and DellaPenna 1998; Ye et al. 2000; Davuluri et al. 2005), production of edible vaccines and pharmaceuticals (Walmsley and Arntzen 2000; Ma et al. 2005), and production of thermoplastics and industrial oils (Jaworski and Cahoon 2003). Recently the successful expression of an active archaebacterial superoxide reductase from the thermophile Pyrococcus furiosus in plant cells opened up an entire new array of possibilities to generate stress-tolerant plants (Im et al. 2005). The use of genes from extremophiles that have evolved to survive the most extreme physical environments can now be used to greatly extend the adaptive range of successful crop plants for space environments (Grunden and Boss 2004).

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The examples above represent the insertion and expression of a single or very few genes. Because most phenotypic traits are polygenic, i.e., involving more than one gene, successful engineering will require manipulation of a network of complex metabolic and regulatory pathways involving multiple genes and/or gene families (Chen et al. 1998). One can envision a combination of breeding and transgenic technology to allow largescale manipulation of the genome to modulate such complex traits. As our understanding of biological processes in cells and organisms improves, and as plant transformation techniques evolve, we will be able to more efficiently and rationally create new plants for space exploration.

9.5 Nanotechnology Nanotechnology encompasses the development of tools and devices for the manipulation and probing of matter of nanometer-scale dimensions. Recently, the potential of nanoscience and its tools have become clear. A great deal of effort is being expended to develop nanosensors and nanotools. It is entirely possible that such tiny machines will be engineered and inserted into living cells. It is clear that within the foreseeable future we will develop new tools that will be important for space exploration. A critical barrier that must be overcome is the development of nanoscale devices that can be implanted into living cells or (preferably) produced in vivo by co-opting the cellular machinery itself. Recently, a gene from the hyperthermophile Sulfobolus shibatae was altered to produce a protein that adheres to gold or to semiconductor material (McMillan et al. 2002). These two-dimensional, selfassembled lattice structures could act as nano-templates upon which sensors or other devices could be built. Actuators to enable gene expression could be created, and sensors to monitor gene expression at all levels could be developed. Although linking these nanosensors and nanoactuators to external (electronic) devices will be a significant challenge, it is possible.

9.6 Sensors, Biosensors, and Intelligent Machines Precise monitoring and control of all aspects of the human/plant/environment biosphere must be a basic system component of a bioregenerative life support system for human life support in space. The integration of nanobiosensors that are linked telemetrically to Earth as part of a distributed intelligent system will be key to success. Because any program of solar system exploration will involve significant periods with humans not present or unavailable to spend time on plant culture, the integration of remotely controlled and smart nanotechnology to monitor and control plant function and metabolism of plants will be critical. The distributed sensor system must have versatility and adaptability, and be able to monitor both the physiological status of the plants as well as the environment in which they are growing, including the aerial environment and the rhizosphere. Microfabrication

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technologies have allowed the integration of sensor systems into miniature multifunctional configurations for the detection of factors important for plants, such as oxygen, pH (Guice et al. 2005; McNaughton et al. 2005), potassium (Cosofret et al. 1995), iron (Mlika et al. 1997), phosphate (Engblom 1998), and nitrate (Knoll et al. 1994; Barker et al. 1998). Although these devices are small and can even accommodate multiple sensor types, they are far from nanoscale devices and little is known regarding their long-term operation and functionality.

9.7 Plan(t)s for Space Exploration Envisage a scenario where selected plant species would be genetically engineered and remotely controlled to provide clean air, potable water, and food while at the same time acting as a pharmaceutical factory and source of raw materials. Specifically engineered and controlled species would be integrated into a cohesive system of sensor-laden, intelligently controlled structures (greenhouses) to provide for immediate life support needs (air, water, food) while also supplying the necessities and enhancers for long-term presence (clothing, shelter, pharmaceuticals, nutraceuticals, materials, fiber, flavoring, perfumes, soaps). In addition to providing life support requirements, plants are fundamental for the maintenance of the psychological well-being of a crew, particularly on long-term missions or permanent settlements. The concept is graphically shown in Figure 9.1 (see also Color Section). Remote instructions, capable of traveling small or great distances (such as radio signals), are generated and sent to the primary receiver. The primary receiver is a physical device (such as an antenna) located in close proximity to the target plants. The primary receiver processes the incoming instructions and generates a secondary signal to which the plants have been engineered and programmed to respond (such as chemical signals, low-fluence infrared radiation, or other novel electromagnetic wavelengths). The secondary signal is perceived by the plant via existing natural components (such as organelles like plastids or pigments such as phytochrome) or via a yet-to-be-designed component we call a “perceptosome.” We envision perceptosomes as rationally designed, bionanotechnologically derived units within the plants. We anticipate they would be based on peroxisomes—vesicles with highly specialized functions that are designed for post-translational import of signal-tagged proteins synthesized in the cytoplasm. Different perceptosomes would have different receptors for import sequences, and genes would be modified so that the required protein (pigment) had the requisite import molecule attached. Thus, proteins normally located in the cytoplasm or other organelles could be modified to make perceptosomes unique for particular purposes. These perceptosomes would be stimulated by the secondary signal to activate a cascade of intercellular or systemic signaling pathways. The systemic signals would take chemical, physical, or electrical form and would in turn stimulate gene activation in appropriate target tissues. This would result in the initiation of programmed functions such as increased photosynthetic rate, specific secondary product biosynthesis, or induction of flowering. All the steps in this process would be integrated into a sensor-rich, robotically enhanced, and intelligently controlled monitoring and feedback control system. Ultimately,

Figure 9.1 (also see Color Section).

Programmable plants: a concept for human life support in space.

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this type of system will depend on advances in genomics, proteomics, bioinformatics, and nanoscience at a minimum. Understanding and harnessing the biochemical machinery of plant cells to produce said devices in situ will be critical, as will the identification of technology areas that currently limit such processes. The foundational work upon which this scenario can be built has been done. The physiology and metabolism in a variety of space-exposed plants have been studied (Tripathy et al. 1996; Croxdale et al. 1997; Musgrave et al. 1998; Klymchuk et al. 2001; Kuznetsov et al. 2001; Nedukha et al. 2001; Stutte et al. 2006). Extremely early molecular responses to gravitational and mechanical stress have been identified (Kimbrough et al. 2004). Research on molecular aspects of plant stress gene regulation in space-grown plants (Paul et al. 2001; Stankovic 2001) as well as developing chimeric genes for use as biomonitors (Paul and Ferl 2002) has been published. Localized stimuli resulting in systemic regulation of gene expression in plants has been shown, using heat pulses (Stankovic and Davies 1996; Stankovic et al. 2000), electrical stimulation (Stankovic and Davies 1996; Davies and Stankovic 2006), and low-level microwave radiation (Vian et al. 2006). Recently, the groups of Boss and Grunden expressed archaebacterial genes from Pyrococcus furiosus in Arabidopsis cells, resulting in increased survival rate under high temperature (Im et al. 2005). Identification and expression of genes from extremophiles have the potential of generating plants that could survive conditions beyond their adaptive range here on Earth. Finally, Wheeler and his colleagues have published extensively on advanced life support concepts and on the development of a Martian greenhouse (Wheeler 1999). A sensible progression and melding of this foundational work and inclusion of other critical enabling technologies is laid out in the architecture in Figure 9.2, and demonstrates a pathway toward “programmable plants” for human life support in space. As of this writing, technology limitations exist in the scale of plant transformation and in the ability to communicate specific programming instructions to plants. Current genetic engineering technology allows relatively simple transformations of a few genes at a time. Although this can be powerful when artfully employed to modify metabolic pathways, in order to have ever-increasing influence we must move from making one or two modifications at a time to making sweeping changes in plant structure and function. The time may be nearing when we can engineer and introduce minichromosomes into plants, which are capable of carrying stably, from generation to generation, batteries of genes encoding new biochemical pathways. Current human–plant communication technologies allow relatively simple signal transmissions to plants, usually through modification of the environment. Modifying light cycles can, for example, induce flowering, and modification of nutrition can evoke changes in plant form. However, true plant programmability necessitates the development of novel signaling methods that will allow plants to respond to various and specific signals that might be transmitted across interplanetary space. Such a programmable plant needs to respond to remotely applied stimuli in a reversible manner. To provide specificity, the signals applied need to be distinct from the environmental stimuli that plants normally experience. Transduction of the physical stimuli into biological responses will be accomplished through introduction of stimulus-responsive specific molecular switches. In the long term, success of human exploration of space will depend on the ability of

Programmable plants whose development, metabolism, and function are remotely controlled in conjunction with an autonomous life support biosphere on Mars

Ultimate Objective: 2050 – Heritable, in planta devices for monitoring and controlling plant function. Can be controlled externally, or internally (biofeedback loops)

Intelligent, remotely controlled, large scale plant selection/breeding program on Mars

Permanent phytotron complex on Mars, with fully-functional, intelligent control systems. Control locally or from Earth

Long-Term: 2025 – 2050 Manipulation and optimization of plant performance in space

Incorporation of in planta, biologically-based, or man-made nanodevices to monitor and control plant function. Can be controlled externally/ remotely

Plant selection programs on Mars

Phytotron (greenhouse) deployed on Martian surface with machine-based plant biologist monitoring and control systems

Mid-Term: 2015 – 2025 Miniaturized DNA microarray technology to monitor total gene expression remotely, i.e. space

Understand the processes that control plant protein and secondary metabolite production

In planta monitoring of gene expression/ physiological status

Plants with specific, controllable elements

Selective, localized control of specific gene expression

Plant selection for Mars greenhouse conditions

Near-Term: 2007 – 2015 Metabolism and physiology in spaceexposed plants

Gene expression using DNA microarray from space-exposed plants

Genes as biomonitors of plant stress in space

Promoters to induce specific genes

Localized stimulus/ systemic response mechanism in plants

Plant selection/ engineering with specific designed traits

Mars greenhouse on Earth with all Martian conditions using distributed sensor and control methods

Mars “greenhouse” on Earth with selected Mars environmental factors using conventional control methods

Foundation: 2007

Figure 9.2. An architecture for programmable plants for human life support on Mars.

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the inhabitants to live (and thrive) “off the land” and their ability to adapt to the new conditions in which they find themselves. To enable this, technologies must be developed that make solar system exploration a great deal more dependable, economical, and sustainable. The potential for achieving all three—using biology to survive, living (and thriving) off the land, and making the economics more favorable—can be attained by utilizing recent advances in, and combining the potentials of, the fields of genomics, proteomics, nanoscience, biosensor technology, robotics, and intelligent machines.

9.8 Imagine . . . The year is 2057. A ship is hurtling through space 6 weeks from Mars, and its crew looking forward with some anxiety to their nearly 2-year-long stay on the Red Planet. By this time, travel between the third and fourth planet from the Sun takes place, but it is neither routine nor does it meet any kind of normal schedule. The spacefarers’ concern is for the things Earth-dwellers take for granted—fresh food, vitamins, antibiotics, pain killers, paper, construction materials, even flavorings and soaps. What do they do? They send out signals in the form of a sophisticated code toward their destination. Why? Because within specialized chambers on Mars are specialized plants adapted to the conditions found on the Red Planet. One unique adaptation is that they are programmed to receive and respond to these remotely generated signals. Perhaps the instruction will be “Make vitamins” (by this time, the pathways for B12 and D have been engineered into plants, making all 13 essential vitamins available botanically); or “Make acetyl salicylic acid” (aspirin and other naturally occurring secondary products are plant-derived); or “Make penicillin” (the biochemical pathway for this antibiotic and others has been engineered into plants by now); or “Make plastic” (the first plants were engineered more than 60 years earlier to produce a biodegradable thermoplastic); or perhaps “Enter fruit development stage” (the crew is just plain hungry for something fresh). Regardless, the crew can rest assured that their needs will be met and can breathe a sigh of relief, saying “Thank goodness for programmable plants!”

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McKay CP. 1997. The search for life on Mars. Origins of Life and Evolution of the Biosphere 27(1-3): 263-289. McKay DS, Gibson EK, ThomasKeprta KL, Vali H, Romanek CS, Clemett SJ, Chillier SDF, et al. 1996. Search for past life on Mars: possible relic biogenic activity in Martian meteorite ALH84001. Science 273(5277): 924-930. McMillan RA, Paavola CD, Howard J, Chan SL, Zaluzec NJ and Trent JD. 2002. Ordered nanoparticle arrays formed on engineered chaperonin protein templates. Nature Materials 1(4): 247-252. McNaughton BH, Anker JN and Kopelman R. 2005. Magnetic microdrill as a modulated fluorescent pH sensor. Journal of Magnetism and Magnetic Materials 293(1): 696-701. Meyer T and McKay C. 1989. The resources of Mars for human settlement. J. Brit. Interplanetary Soc 42, 147-160. Mlika R, Ben Ouada H, Kalfat R, Gamoudi G, Mhenni F and Jaffrezic-Renault N. 1997. Thin sensitive organic membranes on selective iron-ion sensors. Synthetic Metals 90(3): 239-243. Musgrave ME, Kuang AX, Brown CS and Matthews SW. 1998. Changes in Arabidopsis leaf ultrastructure, chlorophyll and carbohydrate content during spaceflight depend on ventilation. Annals of Botany 81(4): 503-512. Nedukha OM, Kordyum EL, Brown CS and Chapman D. 2001. The interaction of microgravity and ethylene on the ultrastructure cell and Ca2+ localization in soybean hook hypocotyl. J Gravit Physiol 8(1): P49-50. Olsen R. 1982. “Regenerative Life Support Research/Controlled Ecological Life Support System (RSLR/CELSS).” Final Report for NASA Contract NAS2-11148 by Boeing Aerospace, NASA Ames Research Center, Moffett Field, CA. Paul AL, Popp M, Gurley W, Guy C, Norwodd K and Ferl R. 2005. Arabidopsis gene expression patterns are altered during spaceflight. Adv. Space Research 36: 1175-1181. Paul AL, Daugherty CJ, Bihn EA, Chapman DK, Norwood KL and Ferl RJ. 2001. Transgene expression patterns indicate that spaceflight affects stress signal perception and transduction in Arabidopsis. Plant Physiol 126(2): 613-621. Paul AL and Ferl RJ. 2002. Molecular aspects of stress-gene regulation during spaceflight. J Plant Growth Regul 21(2): 166-176. Paul AL, Schuerger AC, Popp MP, Richards JT, Manak MS and Ferl RJ. 2004. Hypobaric biology: Arabidopsis gene expression at low atmospheric pressure. Plant Physiol 134(1): 215-23. Perera IY, Hung CY, Brady S, Muday GK and Boss WF. 2006. A universal role for inositol 1,4,5trisphosphate-mediated signaling in plant gravitropism. Plant Physiol 140(2): 746-760. Qi B, Fraser T, Mugford S, Dobson G, Sayanova O, Butler J, Napier JA et al. 2004. Production of very long chain polyunsaturated omega-3 and omega-6 fatty acids in plants. Nat Biotechnol 22(6): 739-745. Salinas-Mondragon R, Brogan A, Ward N, Perera, IY, Boss WF, Brown CS and Sederoff HW. 2005. Gravity and light: integrating transcriptional regulation in roots. Gravit Space Biol Bull 18(2): 121-122. Schwartzkopf S. 1992. Design of a controlled ecological life support system. BioScience 42: 526-535. Shintani D and DellaPenna D. 1998. Elevating the vitamin E content of plants through metabolic engineering. Science 282(5396): 2098-2100. Smernoff D and MacElroy RD. 1989. Use of Martian resources in a Controlled Ecological Life Support System (CELSS). J. Brit. Interplanetary Soc 42: 179-184. Stankovic B and Davies E. 1996. Both action potentials and variation potentials induce proteinase inhibitor gene expression in tomato. FEBS Lett 390(3): 275-279. Stankovic B, Vian A, Henry-Vian C and Davies E. 2000. Molecular cloning and characterization of a tomato cDNA encoding a systemically wound-inducible bZIP DNA-binding protein. Planta 212(1): 60-66.

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Stankovic B. 2001. 2001: A plant space odyssey. Trends in Plant Science 6(12): 591-593. Stitt M and Sonnewald U. 1995. Regulation of metabolism in transgenic plants. Ann Rev Plant Phys and Plant Mol Biol 46: 341-368. Stutte GW, Monje O, Hatfield RD, Paul AL, Ferl RJ and Simone CG. 2006. Microgravity effects on leaf morphology, cell structure, carbon metabolism and mRNA expression of dwarf wheat. Planta 224(5):1038-1049. Tripathy BC, Brown CS, Levine HG and Krikorian AD. 1996. Growth and photosynthetic responses of wheat plants grown in space. Plant Physiol 110(3): 801-806. Vian A, Roux D, Girard S, Bonnet P, Paladian F, Davies E and Ledoigt G. 2006. Microwave irradiation affects gene expression in plants. Plant Signaling and Behavior 1: 67-69. Walmsley AM and Arntzen CJ. 2000. Plants for delivery of edible vaccines. Curr Opin Biotechnol 11(2): 126-129. Wheeler R. 1999. Proceedings of the First Inflatable Greenhouses for Mars Symposium, Kennedy Space Center. NASA TM 2000-208577, NASA Kennedy Space Center, FL. Ye X, Al-Babili S, Kloti A, Zhang J, Lucca P, Beyer P and Potrykus I. 2000. Engineering the provitamin A (beta-carotene) biosynthetic pathway into (carotenoid-free) rice endosperm. Science 287(5451): 303-305. Zubrin, R. and Wagner, R. 1996. The Case for Mars: The Plan to Settle the Red Planet and Why We Must. New York,The Free Press.

Index

Page numbers followed by an f denote a figure. ABA (abscisic acid), 128-29 Abiotic stress, 186 Ablation. See Laser Abrasion, 134 Abscisic acid. See ABA Acceleration, 170 Actin. See also F-actin networks amyloplast connection with, 112 cytoskeleton, 146, 153 drugs disrupting, 144 filaments, 5f Action potential, 93, 135, 189f Adaptability, 187 ADH (alcohol dehydrogenase), 174 Aequorin-derived signal, 25, 63, 103 Aerial organs, 134 tissue, 108 Agravitropism mutants displaying, 124 roots displaying, 6, 55 Alcohol dehydrogenase. See ADH Algae, 79 Alkalinization, 27 Alleles, 55 Amoeba, 142 Amyloplasts. See also Spheroplasts actin connection with, 112 degradation, 128 distal, 167f impaired, 9 mature, 11 rogue, 13 sedimentation of, 4, 25 weight of, 29 Analysis gene, 37f, 172 genetic, 28-29 global, 32

hydrotropism, 125-26 microgravity, 146 proteomic, 110 Animal cells, 56, 105 Anions, 102 Antibiotics, 191 Aperture, stomatal, 104 Arabidopsis, 84 Atmosphere, 184 Autoinhibitory domain, 96 Autoradiography, 34 Autotropic straightening, 84 Autotropism, 172-74 Auxin accumulation, 36 carriers, 168 efflux, 113 existence of, xiii flavonoids regulating, 61 flow, 57 gradient, 22 influx, 49 mechanisms, 65-67 native, 47 plants resistant to, 66 polar transport of, 48f transport, 59-61 transportation of lateral, 30 transporters within statocytes, 24 Bacteria, 80 Basipetal transmission, 23f Behavior growth, 39 organ, xv Bending, 112. See also Gravitropic bending; Hydrotropic bending BFA body formation, 58 Bifurcation, 36

197

198 Binding sites, 96 Biochemical experiments, 51 fractionation, 29 studies, 8-9 Bioinformatics, 190 Biological Research in Canister. See BRIC Biology plant, 185 tools, 86 Biomass production, 183, 185 Bioregeneration, 177 Biosatellites, 164, 175. See also Communication Biosensor technology, 191 Branching roots, 54 Brassinosteroids, 129 Breeding, 187 BRIC (Biological Research in Canister), 175 Buffeting, 91 Bulk detection, 25 Ca2+, 105 gravisignaling including, 106 L-type, 101 Mechanostimulation response from, 103 wave, 25 Calcium, 63 channel, 154 free, 168 signatures, 104f Calmodulin, 25 Cameleon, 103 cAMP, 155 Carnivorous plants, 92 Carriers auxin, 168 influx, 53-55 Cations, 22 Causation, 7 Cell(s). See also Unicellular systems animal, 56, 105 deformation, 95 flank, 154 gravitaxis in, 142-43 gravitropism in, 143-44 imaging, 177 internodal, 12 maturation, 8-9

INDEX

motor, 93 orientation, 155 parenchyma, 80 research, 85 root cap, 104f subapical region of, 26 suspension, 53 upward-swimming, 152 Central elongation zone. See CEZ Centrifugation, 147, 170 CEZ (central elongation zone), 110f, 131 cGMP, 155 Channel calcium, 154 ion, 97 protein, 96f Chara, 12, 150-51 Chemical messenger, 93 Chemiosmotic model, 53 Chemotaxis, 131 Chemotropism, 131-32 Chromatography, 51 Chromosaponin, 55 Ciliates, 141 Circadian clock, 161 rhythms, 81 Circumnutation, 7, 161 Clinostats, 146. See also Random positioning machines Cloning, 54 Cold stimulus, 31, 108 Coleptiles, 28, 123, 175 Communication, xiv Compounds, 47 Computer software, 163f CREB transcription, 106 Crop response, xv Cryptochromes, 81-82, 171 Cultivation, 92f Curvature hydrotropic, 128 response, 21-22, 35, 84, 169-71, 176 Cytokinin, 38-39 Cytoplasma cations, 22 mass, 151 pH, 24, 112 Cytoskeleton, 15

INDEX

actin, 146, 153 dynamic, 6 forces conveyed via, 98 graviorientation, 156 in gravity perception, 12-14 interactions with, 97f vacuole, 15 Data kinetic, 64 microarray, 108 pharmacological, 105 thigmomorphogenic, 94 Deformation, 95 Dehydrating stress, 128-29 Depolarization membrane, 101 statocyte, 130 Detection, 25 Development endodermis, 7 plant, 170 planta, 54 DEZ (distal elongation zone), 110f, 131 Diacylglycerol, 105 Differential kinetics, 39 Directional cues, 29 Directional guidance, 131 Distal elongation zone. See DEZ DNA binding domain, 32 replication, 185 Doses, 60 Drop towers, 164 Drugs, 144 Dye fluorescent, 57 indo-1, 103 voltage sensitive, 152 EC (Experimental Container), 173f Efflux auxin, 113 IAA, 51-53, 55-56 ion, 93 synthesis, 58-59 Election micrographs, 167f Electrotropism, 129-31 Elongation, 94. See also EZ

199

Elongation zone. See EZ EMCS (European Modular Cultivation System), 173f Endocytosis, 57 Endodermis, 7. See also Hypodermis Endogenous gradients, 57 molecules, 3, 55 Endoplasmic reticulum, 15, 153 Endosomal markers, 56, 58 Engineering genetic, 186, 188 plants, xiv-xv Environment conditions in, 61 microgravity, 162, 174-75 resources of, 11 Enzymes, 97f Ethylene, 109, 168 gravitropism regulated by, 64-65 regulation, 65 Euglena, 150, 151-52, 154 European Modular Cultivation System. See EMCS Evolution conservation, 80 phototropin, 82 Exocytosis, vesicle, 154 Experimental Container. See EC Experiments. See also EC; EMCS biochemical, 51 laser ablation, 10, 111 long-term, 177 microarray, 66 microgravity, 154 patch clamp, 99 proteomic, 34 space, xiv-xv Extents, 184 Extracellular matrix, 100f, 145 EZ (elongation zone), 22, 23f, 38 F-actin networks, 13 Ferns, 79 Filaments, 5f Flagellates, 141 Flavonoid(s), 55, 168 auxin regulated by, 61 synthesis, 32

200 Floral meristem initiation, 56 Flowering, 189f Fluence rates, 81 Fluorescence, 169 Force spectroscopy, 98 Free fall, 162, 167 Funculus, 133f Fungi, 80, 129 Gadolinium, 152 Gametes, 131 Gas composition, 164 Gating ion, 96f membrane, 98-99 Gene activation, 189f analysis, 37f, 172 expression, 33 identification, 66 regulation, 190 superfamily, 52 Genetic analysis, 28-29. See also Transgene Genetic engineering, 186, 188 Genetics, 33, 37f Genetic strategies, 30 Genomics, 32, 184-85, 190-91 Gentle touch, 99-101 Germination, 10 GFP. See Green Fluorescent protein Global analysis, 32 Gradient auxin, 22 light, 79 moisture, 123, 126-27 oxygen, 134 thermal, 133 Grass shoots, 8-9 Gravikinesis, 142 Gravimorphogenesis, 166 Graviperception in chara, 150 in loxodes, 151 mechanisms, 148 models of, 149f Gravireception, 127 Graviresponse mechanisms, 153-55 Gravisensitivity, 156

INDEX

Gravisignaling Ca2+ role in, 106 mechanosignaling modulating, 111 touch interaction with, 110-13 Gravistimulation, 33. See also Photostimulation protein change related to, 109 Gravitational pressure model, 11-12 Gravitaxis in cells, 142-43 positive, 149 Gravitropic bending hydrotropic bending v., 127 inhibited, 49 maximal, 60 restored, 52 Gravitropic stimulation, 62f Gravitropism, xv. See also Circumnutation; Gravisensitivity; Plagiogravitropism; Thigmotropism altering, 37, 105 in cells, 143-44 enhancing, 13 ethylene regulating, 64-65 events of, 3 gynophore, 10 hydrotropism interacting with, 124 kinetics of, 30 masking, 172 mutation affecting, 29 negative, 6, 85 phases of, 35 phototropism interacting with, 83-84 pulvini mediating, 8-9 regulation of, 11 research, 9 root, 27, 32-37, 40, 58 root v. shoot, 7 signal transduction in, 21-22 stem, 81f studies, 15 temporal steps of, 166 Gravity, 171. See also Graviresponse; Gravisignaling; Gravitational pressure model; Gravity perception; Hypergravity; Signal transduction acceleration of, 170 cue, 91

INDEX

field, 9 micro, 86 morphogenesis induced by, 166 omnilateral, 163f phosphorylation induced by, 34 plumb line of, 161 receptors, 24-26 regulation, 33 reorientation, 66 response, 5, 28, 59 root sensitivity to, 14, 65 roots stimulated by, 62f sensing, 111 sensor, 144-45 signal, xiv susception, 145 vector, 4, 10, 63, 67, 148 Gravity perception, 48, 166-68 characterization of, 4-9 cytoskeleton in, 12-14 mechanisms, 3 signal transduction in, 103-7 Gravity set point angle (GSA), 40 Green Fluorescent protein (GFP), 49 Greenhouses, 188 Ground based studies, 165-66 above v. below, 91f Growth asymmetric, 68 behavior, 39 differential, 126-27 habit, 113 history, 170 during phototropism, 67 plant, 61 response, 3, 174-75 root, 28 signal transduction, 82-83 temperature affecting, 132 GSA. See Gravity set point angle Gynophore, 10, 15 Hardware, 175 Heat load, 85 shock, 185 Helices, 100f

Heterologous systems, 52 Higher plants, 11, 130 Humidity, 176 Hydrophobic core, 95 Hydrophobicity, 53 Hydrostatic pressure, 95 Hydrotropic bending, 127 Hydrotropic curvature, 128 Hydrotropism, 129, 165, 172-74 analysis, 125-26 gravitropism interacting with, 124 root, 126f studies, 123 Hypergravity, 154, 162 Hyperpolarization, 152 Hypobaria, 186 Hypocotyls, 6-8, 60, 67, 169 Hypodermis, 100 Hypogravity, 162 Hypo-osmotic stress, 98-99 Hypoxia, 186 Hypoxic stress, 185 IAA (Indole-3-acetic acid), 47, 109 efflux, 51-53, 55-56 influx carriers, 53-55 mechanisms, 50f redistribution, 57 IBA. See Indole-3-butyric acid Imaging, 177 Indo-1, 103 Indole-3-butyric acid (IBA), 47 Influx auxin, 49 carriers, 53-55 Infrared radiation, 176 Insectivorous plants, 131 InsP3 (1,4,5-trisphosphate), 26-27 Intercellular communication, xiv statoliths, 156 International Space Station. See ISS Internodal cells, 12 Invertebrate organisms, 96-97 Ion channels, 97 efflux, 93 gating, 96f

201

202 Ion (continued) signaling, 103 Iron, 188 Isodensity condition, 149 ISS (International Space Station), 164, 184 Kinetic(s), 59 data, 64 differential, 39 of gravitropism, 30 reduction in, 27 Laboratory microgravity, 177 settings, 113 Laser ablation, 10, 111 light, 155 tweezers, 147 Latrunculin B, 14 Life support, 183 Light, 14, 108, 171 gradient, 79 interception, 85 laser, 155 nondirectional, 84 perception, 80-82 red, 82, 86 signaling intermediate mutants, 83 visible, 176 Lignin, 109 Live imaging, 56 Loxodes, 149-51 MAP kinase, 106 Markers, 37f, 56, 58 Mass cytoplasm, 151 root, 113 Matrix, 100f, 145 Maturation, 8-9 Mature zone (MZ), 23f Mechanical force, 97f Mechanical rotation, 106 Mechanisms adaptation, 109 auto-avoidance, 134 auxin, 65-67 graviperception, 148

INDEX

graviresponse, 153-55 gravity perception, 3 IAA, 50f mechano-sensitive ion channels, 26 molecular, xiv, 21, 22, 40 polarity, 58 redundant, 15 root, 6-8 synthesis, 68 transport, 47 Mechanoreceptor, 99-101 Mechanoresponses, 91-95, 110 Mechano-sensitive ion channels as gravity receptors, 24-26 mechanisms, 26 Mechanosensor, 101-2, 113 Mechanosignaling, 111 Mechanostimulation, 103 Media, 126f Membrane depolarization, 101 gating, 98-99 plasma, 146f potential, 102, 150 tension, 96f, 99 trafficking, 7, 95 Metabolite, 55 Microarray data, 108 experiments, 66 Microbeams, 5 Microfabrication technology, 187-88 Micrographs, 167f Microgravity, 86, 169f analysis, 146 environment, 162, 174-75 experiments, 154 laboratories, 177 statocytes movement in, 167 studies, 171 Microorganisms, 141, 154 Microtubules. See MT Mir, 164 Mitochondria, 143 Models, 53 Moisture, 123, 126-27 Molds. See Slime molds Molecular cloning, 54

mechanisms, xiv, 21, 22, 40 weight, 35 Molecules endogenous, 3, 55 signaling, 186 Monocotyledons, 14 Morphogenesis, 91, 166. See also Gravimorphogenesis; Thigmomorphogenesis Morphology. See also Photomorphogenesis altered, 5 root cap, 36 Mosses, 79 Motility, 112, 131, 154 MT (microtubule), 13, 147f Mud, 143 Multimodular proteins, 98 Mutagenesis, 101 Mutant(s), 6, 162. See also Regenerative life support agravitropic, 124 arabidopsis, 84 cytokinin deficient, 39 double, 53 double v. single, 30 gravitropism affected by, 29 isolation, 125 light-signaling intermediate, 83 lines, 52 phenotypes, 61 starch deficient, 165 MY (myosin), 147f Myosin. See MY MZ. See mature zone Nanoscience, 184, 190, 191 Nanotechnology, 187 Naphthylphthalamic acid (NPA), 51, 60 Nastic responses, 161, 175 Negativity gravitropism, 6, 85 phototropism, 79 Nitric oxide. See NO Nitrite, 188 NM (normal nutrient medium), 125 NO (nitric oxide), 132 Noise, 14, 152 Normal nutrient medium. See NM NPA. See Naphthylphthalamic acid

Nr, 64 Nutrients, 14. See also NM 1,4,5-trisphosphate. See InsP3 Organelles, 3. See also Statoliths Organisms. See also Microorganisms invertebrate/vertebrate, 96-97 sessile, xiii, 141 Organs aboveground, 22 aerial, 134 behavior of, xv bending of, 112 signal transduction in, 39-40 vertically oriented, 38 Orientation. See also Reorientation cell, 155 nonrandom, 167 root, 4 Oscillation, 175 Osmotic pressure, 26 Ovule, 133f Oxonol, 152 Oxygen, 188 gradient, 134 stress, 174 Oxytropism, 132-34, 165, 172-74 Parabolic flights, 146 Paramecium, 150, 151-52 Parenchyma cells, 80 tissue, 60 Pathogens, 108 PCOCs (Plant Carry-on Containers), 175 Peg formation, 10 Perceptosome, 188 Peroxisomes, 188 pH, 168, 188 changes in, 63 cytoplasmic, 24, 112 lipidity, 102 low, 53 signal transduction contribution from changes in, 27-28 Pharmacology data, 105 studies, 24 Phenotypes, xiii

203

204 Phenotypes (continued) mutant, 61 polygenic, 187 Phosphate, 188 Phosphorylation gravity induced, 34 protein, 59-61 Photolysis, 47 Photomorphogenesis, 81 Photoreceptor pigments, 80 Photostimulation, 79 Photosynthesis, 21, 84, 183 Phototropin, 82 Phototropism, 129, 171-72 altering, 29 gravitropism interacting with, 83-84 growth during, 67 negative, 79 phytochromes role in, 86 positive, 85 root, 81f Phytochromes, 82, 86, 188 pI, 35 PIN1 cycling, 57f Plagiogravitropism, 40 Plant(s) adaptability, 187 auxin resistant, 66 biology, 185 carnivorous, 92 development, 170 engineering, xiv-xv form, 84-85, 171 growth, 61 higher, 11, 130 insectivorous, 131 mechanoresponses, 91-95 mechanosensor, 101-2, 113 programmable, 184, 189f, 191f seed, 79 sensory system in, 135 species, 48 stress, 190 tissue, 47, 52, 65 transgenetic, 186 tropisms, 1 wild-type, 127 Planta development, 54

INDEX

Plant Carry-on Containers. See PCOCs Polarity, 102 auxin, 57 mechanisms, 58 Polarization. See Hyperpolarization Pollen tubes, 123, 130 Positivity gravitaxis, 149 phototropism, 85 Potassium, 188 Potential. See also WP action, 93, 135, 189f membrane, 102, 150 Presentation time, 169 Pressure, 26 Pressure theory, 145 Programming plants, 184, 189f, 191f Protein(s), 22, 51. See also Green Fluorescent protein channel, 96f control, 191 dynamic, 13 encoded, 30 gravistimulation related to change to, 109 identification, 33 J-domain, 29 localization, 56 multimodular, 98 phosphorylation, 59-61 PIN, 168 proteolytic degradation of, 68 receptor, 144 signal transduction implying involvement of, 28-32 structure, 83 synthesis, 185 transmembrane, 95 wall-modifying, 108 zinc-finger, 31 Proteolytic degradation, 68 Proteomic(s), xiii, 32, 190, 191 analysis, 110 experiments, 34 Proteosome, 66 Proton(s) caged, 28 pumping, 113

INDEX

Protonemata, 141, 153 Protoplast hydrostatic pressure from, 95 pressure theory, 145, 148 weight, 12 Pulvini, 79 gravitropism mediated by, 8-9 oat, 34 Radial swelling, 94 Radiation, 164, 176 Random positioning machines, 165, 169 Reciprocity rule, 170 Red light, 82, 86 Redundancy, 32 Regenerative life support, 183 Reorientation gravity, 66 root, 57 Replicability DNA, 185 research, 165 Reproductive functions, 21 Research cell, 85 gravitropism, 9 orbit-based, xv replicability, 165 signal transducer, 97 space, 161 touch, 109 Resistance, 54 Resources, 11 Response. See also Curvature; Mechanoresponses crop, xv curvature, 21-22, 35, 84, 169-71, 176 gravity, 5, 28, 59 growth, 3, 174-75 nastic, 161, 175 physiological, 15 touch, 92-94, 107 Reverse genetics, 33, 37f Rhizoids, 141, 148 protonemata responding like, 153 Rhizosphere, 187 Robots, 183, 191 Room temperature, 31

Root(s). See also Root cap agravitropic, 6, 55 branching, 54 decapping, 130 gravitropism, 7, 27, 32-37, 40, 58 gravity sensitivity of, 14, 65 gravity stimulated, 62f growth, 28 horizontal, 50f hydrotropism, 126f mass, 113 mechanisms, 6-8 orientation, 4 phototropism, 81f reorientation, 57 shoots v., 48 signal transduction in, 22-24 stems v., 23f system, 92f thermotropism, 132-34 Root cap cells, 104f longitudinal section of, 5f morphology, 36 ROS sensitivity, 107 Rotation, 106 Sachs, Von, 124f Screening approach, 31 Sedimentation amyloplast, 4, 25 rate, 144 Seed germination, 10 plants, 79 Seedling, 169f homozygous, 50f size, 67 Selective resistance, 54 Septum, 133f Sequence similarity, 54 Sessile organisms, xiii, 141 Shock, 185 Shoot(s) grass, 8-9 gravitropism, 7 roots v., 48 statocytes, 38

205

206 Signal. See also Transduction pathway aequorin-derived, 25, 63 molecules, 186 pathways, 61-64, 153-55 research, 97 transducer, 168-69 Signal transduction, 3 bifurcation, 36 in gravitropism, 21-22 in gravity perception, 103-7 growth, 82-83 insP3 role in, 26-27 in organs, 39-40 pH changes contributing to, 27-28 proteins implicated in, 28-32 in roots, 22-24 vesicular trafficking connected to, 38 Silver-staining, 35 Skylab, 164 Slime molds, 142 Soap, 191 Software. See also Hardware; Programming plants computer, 163f Soil, 91, 143 Solar-tracking, 85 Somatal openings, 80 Sounding rockets, 175 Space experiments, xiv-xv research, 161 Spectroscopy, 98 Sperm, 142 Spheroplasts, 99 Spitzenkörper. See SpK SpK (Spitzenkörper), 147f, 153 Starch mutants deficient in, 165 sheath, 60 Starch-statolith hypothesis, 4, 9-11 Statocytes auxin transporters within, 24 depolarization, 130 microgravity movement by, 167 shoot, 38 Statoliths, 3, 111, 149-50. See also Starchstatolith hypothesis intercellular, 156 susception, 145-49

INDEX

weight, 150 Status, 187 Stem(s) gravitropism, 81f roots v., 23f Stimuli, xiii Stomatal aperture, 104 Strategies, 30 Streaming velocity, 12 Stress(es) abiotic, 186 dehydrating, 128-29 hypo-osmotic, 98-99 hypoxic, 185 oxygen, 174 plant, 190 transfer of, 97 water, 85, 125 Structure protein, 83 tissue, 21 Studies biochemical, 8-9 gravitropism, 15 ground-based, 165-66 hydrotropism, 123 microgravity, 171 pharmacological, 24 transgenetic, 63 unicellular systems, 143f Substrate, 97f Susception gravity, 145 statoliths, 145-49 Suspension cells, 53 Symbiont elicitors, 104 Synthesis efflux, 58-59 enhanced, 64 flavonoid, 32 mechanisms, 68 protein, 185 Synthetic compounds, 47 Systems. See also EMCS heterologous, 52 root, 92f sensory, 135 unicellular, 143f

INDEX

TCH. See Touch Technology, 177, 185 biosensor, 191 microfabrication, 187-88 Temperature, 176 growth effects due to, 132 room, 31 Tendrils, 94 Thermal gradient, 133 Thermal noise, 152 Thermotropism, 132-34 Thigmomorphogenesis, 94 Thigmotropism, 94, 113 Threshold duration, 169. See also Presentation time Timing, 49 Tissue aerial, 108 overexpression, 59 parenchyma, 60 plant, 47, 52, 65 structure, 21 Tools, 174 biology, 86 nanotechnology, 187 Topology, 30 Touch. See also Gentle touch gravisignaling interaction with, 110-13 perception, 95-98 research, 109 response, 92-94, 107 Transcriptional profiling, 107-10 Transduction pathway, 107 Transgene plants, 186 studies, 63 Transpiration, 183 Transport machinery, 53, 59-61 Traumatropism, 134-35 TROPI, 173f Tropism(s) concepts of, xiii plant, 1 Turgor loss, 93 2D-G3. See Two-dimensional gel electrophoresis

Two-dimensional gel electrophoresis (2D-G3), 34 Unicellular systems, 143f Up-regulation, 32 Vacuole, cytoskeleton, 15 transport, 8 Vasculature, 93 Vector gravity, 4, 10, 63, 67, 148 information, 40 Vectorial stimuli, xiii Velocity, 12 Vertebrates, 96-97 Vesicle exocytosis of, 154 inhibitors, 56 movements, 58 trafficking, 38 Vibration, 170 Video, 176 Vitamins, 191 Voltage dependence, 102 dye sensitive to, 152 sensitivity, 101 Water, 14. See also WP; WSM stress, 85, 125 Water potential. See WP Water stress medium. See WSM Weight amyloplast, 29 molecular, 35 protoplast, 12 statoliths, 150 Weightlessness, 162 Wild-type levels, 31 plants, 127 WP (water potential), 126f WSM (water stress medium), 125 Zero-g, 162 Zoospores, 142

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